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GIPC proteins negatively modulate Plexind1 signaling during vascular development

  1. Jorge Carretero-Ortega  Is a corresponding author
  2. Zinal Chhangawala
  3. Shane Hunt
  4. Carlos Narvaez
  5. Javier Menéndez-González
  6. Carl M Gay
  7. Tomasz Zygmunt
  8. Xiaochun Li
  9. Jesús Torres-Vázquez  Is a corresponding author
  1. New York University Langone Medical Center, United States
  2. New York University School of Medicine, United States
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Cite this article as: eLife 2019;8:e30454 doi: 10.7554/eLife.30454

Abstract

Semaphorins (SEMAs) and their Plexin (PLXN) receptors are central regulators of metazoan cellular communication. SEMA-PLXND1 signaling plays important roles in cardiovascular, nervous, and immune system development, and cancer biology. However, little is known about the molecular mechanisms that modulate SEMA-PLXND1 signaling. As PLXND1 associates with GIPC family endocytic adaptors, we evaluated the requirement for the molecular determinants of their association and PLXND1’s vascular role. Zebrafish that endogenously express a Plxnd1 receptor with a predicted impairment in GIPC binding exhibit low penetrance angiogenesis deficits and antiangiogenic drug hypersensitivity. Moreover, gipc mutant fish show angiogenic impairments that are ameliorated by reducing Plxnd1 signaling. Finally, GIPC depletion potentiates SEMA-PLXND1 signaling in cultured endothelial cells. These findings expand the vascular roles of GIPCs beyond those of the Vascular Endothelial Growth Factor (VEGF)-dependent, proangiogenic GIPC1-Neuropilin 1 complex, recasting GIPCs as negative modulators of antiangiogenic PLXND1 signaling and suggest that PLXND1 trafficking shapes vascular development.

https://doi.org/10.7554/eLife.30454.001

Introduction

Angiogenic sprouting, the formation of new vessels via branching of pre-existing ones, drives most of the life-sustaining expansion of the vascular tree. Sprouting angiogenesis is also pivotal for recovery from injury and organ regeneration and is often dysregulated in disease (Carmeliet, 2003; Ramasamy et al., 2015; Cao, 2013; Fischer et al., 2006). Among the pathways that modulate this process, SEMA-PLXN signaling plays a prominent role. In particular, the vertebrate-specific PLXND1 receptor transmits paracrine SEMA signals from the somites to the endothelium to shape the stereotypical and evolutionarily conserved anatomy of sprouts in the trunk’s arterial tree (Torres-Vázquez et al., 2004; Zygmunt et al., 2011; Gitler et al., 2004; Zhang et al., 2009; Gu et al., 2005; Childs et al., 2002; Epstein et al., 2015). For instance, we have shown that antiangiogenic Plxnd1 signaling antagonizes the proangiogenic activity of the VEGF pathway to regulate fundamental features of vascular development (Torres-Vázquez et al., 2004; Zygmunt et al., 2011; see also Moriya et al., 2010; Fukushima et al., 2011; Kim et al., 2011). Specifically, Plxnd1 signaling acts cell autonomously in the endothelium to spatially restrict the aorta’s angiogenic capacity to form sprouts, thereby determining both the positioning and abundance of the aortic Segmental (Se) vessels. Plxnd1 signaling also guides Se pathfinding, thus shaping these vascular sprouts.

Upon ligand binding, PLXNs act as guanosine triphosphatase-activating proteins (GAPs) and modulate integrin-mediated cell adhesion, cytoskeletal dynamics, and both ERK and MAPK signaling. However, there is limited molecular understanding of how PLXND1 exerts its vascular effects. Although a handful of putative intracellular effectors and modulators have been identified for this receptor (e.g., endocytic adaptors of the GIPC family, small GTPases belonging to the Ras and Rho families, GTPase regulators, and cytoskeletal proteins), clear functional links between these candidates and in vivo PLXND1-dependent angiogenic patterning remain to be established (Zygmunt et al., 2011; Moriya et al., 2010; Fukushima et al., 2011; Choi et al., 2014; Horowitz, 2007; Uesugi et al., 2009; Gay et al., 2011; Sakurai et al., 2010; Sakurai et al., 2011; Wang et al., 2012; Deloulme et al., 2015; Worzfeld et al., 2014; Tata et al., 2014; Burk et al., 2017; Aghajanian et al., 2014; Wang et al., 2013; Pascoe et al., 2015).

To address this gap, we focused on elucidating the relationship between GIPCs and antiangiogenic PLXND1 signaling. Based on our identification of GIPC1/Synectin as a PLXND1-binding protein, we previously showed by X-ray crystallography and co-immunoprecipitation (CoIP) assays that all three GIPC proteins can physically interact with the intracellular tail of the PLXND1 receptor and elucidated the structural organization of the GIPC-PLXND1 complex (Gay et al., 2011; Shang et al., 2017).

Here, we have further evaluated the molecular determinants of the PLXND1-GIPC interaction in the native environment of mammalian cells, firmly establishing a critical role for the last few amino acids of PLXND1 for the formation of the PLXND1-GIPC complex. Moreover, we defined the vascular role of PLXND1-GIPC association using zebrafish and cultured human umbilical vein endothelial cells (HUVECs) as model systems. Our results suggest that GIPCs act redundantly to limit the antiangiogenic signaling of the PLXND1 receptor.

In contrast, prior vascular studies of GIPCs have primarily centered on the role of GIPC1 as a positive modulator of VEGF (Vascular Endothelial Growth Factor) activity. The dominant model is that GIPC1 promotes arterial branching morphogenesis and arteriogenesis by associating with the cytosolic tail of the VEGF co-receptor NEUROPILIN-1 (NRP1), thereby facilitating the trafficking, and thus the signaling, of the VEGF receptor VEGFR2 (FLK1/KDR) (Waters, 2009; Habeck et al., 2002; Wang et al., 2006; Ren et al., 2010; Covassin et al., 2009; Lanahan et al., 2013; Lawson et al., 2003; Hermans et al., 2010; Chittenden et al., 2006; Dedkov et al., 2007; Lanahan et al., 2010, see also Burk et al., 2017).

Hence, our identification of GIPCs as negative intracellular regulators of PLXND1 signaling outlines a novel molecular mechanism by which these endocytic adaptors promote angiogenic development of the vertebrate arterial tree (Chittenden et al., 2006; Lanahan et al., 2010; Fantin et al., 2011; Lampropoulou and Ruhrberg, 2014; Herzog et al., 2011; Plein et al., 2014), highlighting the potential importance of PLXND1 trafficking for proper vascular development.

Results

Isolation of GIPC1/Synectin as a PLXND1-binding protein and dissection of the molecular determinants required for PLXND1-GIPC complex formation

To uncover intracellular modulators and effectors of PLXND1, we isolated proteins that bind to the cytosolic tail of wild-type (WT) mouse PLXND1 (C-mPLXND1WT) using a yeast two-hybrid screen (Fields and Song, 1989). One of the recovered PLXND1-interacting preys (Gay et al., 2011) harbored the entirety of GIPC1 (GAIP interacting protein C-terminus or Synectin, among many other names), the founder of the three-membered family of GIPC endocytic adaptors (Katoh, 2013; De Vries et al., 1998a; Gao et al., 2000; Naccache et al., 2006; Reed et al., 2005). Remarkably, GIPC1, like PLXND1, modulates vascular development and acts within endothelial cells (Zygmunt et al., 2011; Zhang et al., 2009; Moraes et al., 2013; Moraes et al., 2014).

GIPCs are dimeric proteins with a central PDZ (PSD-95/Dlg/ZO-1) domain that binds to the C-terminal PDZ-Binding Motif (PBM) of transmembrane proteins harboring the type I PDZ-binding consensus sequence (S/T)-X-A/V/L/I. GIPCs also contain GH (GIPC Homology) domains at their amino (GH1), and carboxyl (GH2) ends. The GH1 domain promotes self-dimerization, while the GH2 domain binds the globular domain of MYOSIN6 (MYO6), the retrograde actin-based motor driving endocytic vesicle internalization (Shang et al., 2017; Katoh, 2013; Naccache et al., 2006; Salikhova et al., 2008Lou et al., 2001; Cai and Reed, 1999; Gao et al., 2000; Aschenbrenner et al., 2003). The divergent C-terminal region of Plxns (T-segment; 40–60 aa long) likely mediates subfamily-specific interactions between Plxns and other proteins. Indeed, the PLXND1-specific carboxyl sequence SEA-COOH is a consensus GIPC1-binding motif that serves as the PBM in the GIPC1-binding protein NRP1, a VEGF co-receptor (Gay et al., 2011; Lanahan et al., 2013; Lanahan et al., 2010; Wang et al., 2006; Horowitz and Seerapu, 2012; Lanahan et al., 2014; Chittenden et al., 2006).

To understand how PLXND1 and GIPC1 interact physically we performed a crystallographic study with bacterially purified recombinant mouse proteins. This structural analysis revealed that the receptor’s nonameric C-terminal sequence NIYECYSEA-COOH (residues 1917–1925) is the primary GIPC-Binding Motif (GBM) and makes N- and C-terminal contacts with GIPC1’s GH1 and PDZ domains. A pair of receptor’s helices (residues 1893–1908) and the GIPC1’s PDZ domain form a secondary heteromeric interface. Pulldown experiments of GIPC1 by MYO6 in the presence of PLXND1 indicate that MYO6 associates with PLXND1-bound GIPC1 and that deletion of the receptor’s canonical PBM (PLXND1ΔSEA) or I1918A-Y1919A GBM substitutions significantly reduces PLXND1-GIPC1 binding. Conversely, the GIPC1 G114Y mutation disrupting the interaction between PLXND1’s I1918/Y1919 GBM residues and GIPC1’s GH1 hydrophobic pocket also impairs PLXND1-GIPC1 binding (Shang et al., 2017).

Using exogenous expression of murine proteins and a eukaryotic cellular environment, we interrogated the requirement for PLXND1’s canonical PBM and nonameric GBM and also tested the involvement of GIPC1’s PDZ and GH1 domains. To this end, we performed CoIP experiments with COS7 cells, which lack endogenous expression of both PLXND1 (Gu et al., 2005Uesugi et al., 2009; Sakurai et al., 2010; Takahashi et al., 1999) and NRP1 (Tordjman et al., 2002) (Figure 1; see also Supplementary file 1 and Supplementary file 2).

PLXND1’s C-terminal GBM and GIPC's GH1 and PDZ domains are the key molecular determinants of the PLXND1-GIPC interaction.

(A) Diagrams of the wild-type (WT) and truncated V5-tagged (red) forms of the cytosolic tail of murine PLXND1 (V5-C-mPLXND1) used for co-immunoprecipitation experiments. Color-coding is used to highlight the following domains and motifs. GAP1 and GAP2 (Guanosine triphosphatase-Activating Protein domains 1 and 2; black), RBD (Rho GTPase-Binding Domain; green), T-segment (C-terminal segment, includes the GBM; blue) and, GBM (GIPC-Binding Motif; magenta); see (Gay et al., 2011). (B) Diagrams of the wild-type (WT) and truncated FLAG-tagged (purple) forms of murine GIPC1 (FLAG-mGIPC) used for co-immunoprecipitation experiments. Domains indicated as follows: PDZ (PSD-95/Dlg/ZO-1; green) and GH (GIPC Homology domain) 1 (blue) and 2 (orange); see (Katoh, 2013). (C) Western blots (top) and their quantification (bottom, bar graphs). Numbers indicate lane positions. Left-side Western blot (IPFLAG): FLAG immunoprecipitates and V5 co-immunoprecipitates showing interactions between the indicated V5-C-mPLXND1 and FLAG-mGIPC forms. Right-side Western blot (TCL), expression levels of these proteins in total cell lysates as detected with V5 and FLAG antibodies. Quantifications. n = 3 independent experiments for each protein pair. Left-side bar graph (C). Means of percentual V5/FLAG relative binding [(V5CoIP/V5TCL)/(FLAGIP/FLAGTCL)] between the indicated protein pairs from the IPFLAG Western blot (top left) and the relative abundance of the expression levels of these proteins from the TCL Western blot (top right). Error bars, ± SEM. V5/FLAG relative binding was significantly different (p<0.05) between V5-C-mPLXND1 forms and FLAG-mGIPC, F(2, 6)=22.376, p=0.002, as determined by a one-way ANOVA test. A Tukey post hoc analysis was conducted to determine whether the percentual V5/FLAG relative binding between the three tested pairs of proteins was significantly different (p<0.05; asterisks). Right-side bar graph (C). Means of percentual V5TCL/FLAGTCL relative abundance between the indicated protein pairs from the TCL Western blot (top right). Error bars, ± SEM. A Kruskal-Wallis H test was conducted to determine significant differences (p<0.05) in V5TCL/FLAGTCL relative abundance between the indicated protein pairs. Distributions of V5TCL/FLAGTCL relative abundance were not similar for all groups. The medians of V5TCL/FLAGTCL relative abundances were 92.64 (for V5-C-mPLXND1WT/FLAG-mGIPC1), 87.79 (for V5-C-mPLXND1ΔCYSEA/FLAG-mGIPC1), and 96.22 (for V5-C-mPLXND1ΔGBM/FLAG-mGIPC1), but were not statistically significantly different between them (Ramasamy et al., 2015), χ(2)=0.8, p=0.670. (D) Western blots: Top (IPFLAG), FLAG immunoprecipitates and their V5 co-immunoprecipitates showing interactions between the indicated V5-C-mPLXND1 and FLAG-mGIPC forms; bottom, (TCL) detection of the expression levels of these proteins in total cell lysates using antibodies against V5 and FLAG. n = 3 independent experiments for each protein pair. For additional data and statistical comparisons related to this figure, see Supplementary file 1, Supplementary file 2 and Supplementary file 8.

https://doi.org/10.7554/eLife.30454.002

To address the requirement for PLXND1’s canonical PBM and nonameric GBM, we used V5-tagged PLXND1 cytosolic tails (Figure 1A), either with (V5-C-mPLXND1WT) or without (V5-C-mPLXND1ΔCYSEA) the canonical PBM. For GIPC1, we used a FLAG-tagged version of the full protein, FLAG-mGIPC1WT (Figure 1B). We found that deletion of PLXND1’s canonical PBM (V5-C-mPLXND1ΔCYSEA) significantly reduced binding to FLAG-mGIPC1WT (Figure 1C, lanes 2–3 and graph below), consistent with the existence of additional contacts, as suggested by our crystallographic findings (Shang et al., 2017; see also Lee and Zheng, 2010). Accordingly, the PLXND1 form lacking the GBM motif (V5-C-mPLXND1ΔGBM) shows a further reduction in FLAG-mGIPC1WT-binding capacity (Figure 1C, lane 4 and graph below). Importantly, the minimum GIPC1-binding capacity of the V5-C-mPLXND1ΔGBM form observed under these protein over-expression conditions suggests that the secondary GIPC-binding interface formed by the C-terminal pair of helices of PLXND1 (Shang et al., 2017) plays a minor role as a molecular determinant for PLXND1-GIPC1 complex formation.

To test the involvement of the two GIPC1 domains that interact with PLXND1 in the crystal structure (Shang et al., 2017), we used FLAG-tagged GIPC1 fragments harboring the GH1 and PDZ domains individually (FLAG-mGIPC1GH1 and FLAG-mGIPC1PDZ, respectively) and V5-C-mPLXND1WT. Our CoIP data confirmed that each of these domains is sufficient for PLXND1 association (Figure 1D). The ability of GIPC1’s GH1 domain to interact with PLXND1 is consistent with prior observations implicating GIPC1’s N-terminal region in binding to other transmembrane proteins (Naccache et al., 2006; Giese et al., 2012).

Taken together, the data from our yeast two-hybrid (Gay et al., 2011) and crystallographic (Shang et al., 2017) studies demonstrate that PLXND1 and GIPC1 can interact directly and that NRP1 is dispensable for their interaction. These findings are consistent with our CoIP of PLXND1-GIPC1 complexes from COS7 cells, which lack endogenous NRP1 expression (see Tordjman et al., 2002). Together with our dissection of the molecular determinants of PLXND1-GIPC1 complex formation, the high sequence identity of GIPCs, and our observation that GIPC2 and GIPC3 also associate with PLXND1 (Shang et al., 2017), these results establish the critical importance of PLXND1’s GBM and GIPC's GH1 and PDZ domains for PLXND1-GIPC complex formation.

Zebrafish plxnd1skt6 homozygous mutants, which express a Plxnd1 receptor with a predicted impairment in GIPC binding, display angiogenesis deficits with low frequency

To determine the role that GIPC binding exerts on antiangiogenic PLXND1 signaling, we sought to specifically impair PLXND1’s ability to associate with GIPC endocytic adaptors in an in vivo model of vascular development. To do this, we performed CRISPR/Cas9-based genome editing (Auer and Del Bene, 2014; Auer et al., 2014; Chang et al., 2013; Cong et al., 2013; Cong and Zhang, 2015; Gagnon et al., 2014; Hill et al., 2014; Hruscha et al., 2013; Hwang et al., 2013; Irion et al., 2014; Kimura et al., 2014; Mali et al., 2013; Talbot and Amacher, 2014) of the last coding exon of the zebrafish plxnd1 locus to introduce disrupting mutations into the receptor’s GBM (NIYECSSEA-COOH, canonical PBM underlined; Figure 2A). The resulting plxnd1skt6 allele encodes a Plxnd1 receptor missing the PBM because of replacement of the five C-terminal residues by a stretch of 31 amino acids (Figure 2B; see also Supplementary file 1 and Supplementary file 8). Because adding just a single C-terminal residue to the PBM of proteins that interact with PDZ domain-containing partners is sufficient to block their cognate association (Rickhag et al., 2013; Saras et al., 1997; Cao et al., 1999; Garbett and Bretscher, 2012), and deletion of PLXND1’s PBM reduces GIPC binding significantly (Figure 1A–C; see also Shang et al., 2017), the plxnd1skt6 mutant allele is expected to encode a Plxnd1 receptor with reduced or null GIPC-binding ability.

Figure 2 with 2 supplements see all
The plxnd1skt6 allele encodes a functional Plxnd1 receptor putatively impaired in GIPC binding, and its homozygosity induces angiogenesis deficits with low frequency.

(A, B) Diagrams of the cytosolic tails of the zebrafish Plxnd1 proteins encoded by the WT (A) and plxnd1skt6 mutant (B) alleles including their C-terminal amino acid sequences. Color-coding is used to highlight the following domains and motifs. GAP1 and GAP2 (Guanosine triphosphatase-Activating Protein domains 1 (left) and 2 (right); black), RBD (Rho GTPase-Binding Domain; green), T-segment (C-terminal segment, includes the GBM; blue), and GBM (GIPC-Binding Motif; magenta). In the WT protein diagram (A), the canonical PBM (PDZ-Binding Motif) is underlined. In the mutant protein diagram (B), the thin horizontal red bar denotes the amino acid sequence replacing the PBM. (C–H) Confocal lateral images of the trunk vasculature (green) of 32 hpf embryos (region dorsal to the yolk extension). Anterior, left; dorsal, up. Scale bars (white horizontal lines), 100 μm. Genotypes indicated on top of each image in yellow font. Angiogenesis deficits are indicated with asterisks as follows: white (DLAV gaps), magenta (truncated Se). In the WT image (C), the vessels are designated with the white font as follows: DLAV (Dorsal Longitudinal Anastomotic Vessel), Se (Segmental Vessel), DA (Dorsal Aorta), and PCV (Posterior Cardinal Vein). The homozygous WT and homozygous plxnd1skt6 mutant embryos (F–H) are siblings derived from the incross of plxnd1skt6/+heterozygotes. (I) Bar graph. Percentage of Se-DLAV in 32 hpf embryos of the indicated genotypes belonging to each of the following four phenotypic classes. Truncated: maximal (red; includes missing Se), moderate (yellow), and minimal (gray). Non-truncated: Full (black). There was no statistically significant difference in the distribution of the four phenotypic classes between WT and plxnd1skt6 mutants as assessed by a two-sided Fisher Exact test, p=0.05905. Quantifications. To determine whether plxnd1skt6 complements the plxnd1fov01b null, we analyzed vascular patterning (C–E) and scored Se-DLAV angiogenesis (C, E) in embryos of the following three genotypes: WT (124 Se-DLAV, 11 embryos, an average of 11.27 Se-DLAV/embryo), plxnd1fov01b homozygotes (12 embryos), and plxnd1fov01b/plxnd1fov01b transheterozygotes (162 Se-DLAV, 16 embryos, an average of 10.13 Se-DLAV/embryo). All the WT and transheterozygotes displayed proper vascular patterns indistinguishable from each other and lacked Se-DLAV truncations. All the plxnd1fov01b mutants displayed hyperangiogenic vascular mispatterning. To determine how Plxnd1’s inability to interact with GIPCs impacts angiogenic growth, we scored Se-DLAV angiogenesis (F–I) in sibling embryos of the following two genotypes: Homozygous WT (126 Se-DLAV, 12 embryos, an average of 10.5 Se-DLAV/embryo) and plxnd1skt6 homozygous mutants (124 Se-DLAV, 12 embryos, an average of 10.33 Se-DLAV/embryo). For additional data, graphs and statistical comparisons related to this figure, see Figure 2—figure supplement 1 and Supplementary file 3.

https://doi.org/10.7554/eLife.30454.003

In the trunk of WT embryos, Se sprouts arise bilaterally from the Dorsal Aorta (DA) just anterior to each somite boundary (SB) at 21 h post-fertilization (hpf). Se sprouts grow dorsally with a chevron shape connecting with their ipsilateral neighbors above the spinal cord’s roof to form the paired Dorsal Longitudinal Anastomotic Vessels (DLAVs) by 32 hpf (Isogai et al., 2003) (Figure 2C). In contrast, null plxnd1 alleles, such as plxnd1fov01b, show a recessive and hyperangiogenic mutant phenotype characterized by a dramatic mispatterning of the trunk’s arterial tree anatomy. As such, the homozygous mutants form too many, and misplaced, Se sprouts that branch excessively and interconnect ectopically (Figure 2D) as a result of increased proangiogenic VEGF activity (Torres-Vázquez et al., 2004; Zygmunt et al., 2011).

GIPCs limit or promote the activity of their transmembrane consorts by regulating their surface expression, G-protein signaling, and vesicular trafficking (Lanahan et al., 2010; De Vries et al., 1998a; Naccache et al., 2006; Reed et al., 2005; Giese et al., 2012; Lou et al., 2001; Bunn et al., 1999; Wieman et al., 2009; Blobe et al., 2001; Chak and Kolodkin, 2014; Hu et al., 2003; Booth et al., 2002; Jeanneteau et al., 2004a; Arango-Lievano et al., 2016; Varsano et al., 2012). Whether GIPC binding limits, versus enables, PLXND1 signaling gives different predictions regarding the nature of the plxnd1skt6 allele, which can be assessed by its ability to complement the plxnd1fov01b null mutation, the vascular phenotype of plxnd1skt6 homozygotes and their sensitivity to antiangiogenic compounds. If GIPCs limit PLXND1’s antiangiogenic signaling, plxnd1skt6 should be a plxnd1fov01b-complementing hypermorph, and plxnd1skt6 homozygotes have angiogenesis deficits and SU5416 hypersensitivity. If GIPCs directly enable PLXND1’s antiangiogenic signaling (as in Burk et al., 2017), then plxnd1skt6 should be a null incapable of complementing plxnd1fov01b, and plxnd1skt6 mutants show excessive and disorganized angiogenesis and SU5416 hyposensitivity. If PLXND1 indirectly exerts a GIPC-dependent antiangiogenic effect by sequestering GIPCs to antagonize GIPC-dependent proangiogenic VEGFR2/Nrp1-mediated VEGF signaling (see Gay et al., 2011; Lanahan et al., 2013; Lanahan et al., 2010; Wang et al., 2006; Horowitz and Seerapu, 2012; Lanahan et al., 2014; Chittenden et al., 2006), then plxnd1skt6 should be a hypomorph with partial or full plxnd1fov01b complementing capacity, and plxnd1skt6 homozygotes should have hyperangiogenic or normal vascular phenotypes and reduced or normal SU5416 sensitivity. Finally, if GIPCs are irrelevant for PLXND1 signaling, then plxnd1skt6 should fully complement plxnd1fov01b, and plxnd1skt6 homozygotes should show proper vascular patterning, normal angiogenic growth, and WT-like SU5416 sensitivity.

A comparative analysis of WT embryos, plxnd1fov01b homozygous mutants, and plxnd1skt6/plxnd1fov01b transheterozygotes (Figure 2C–E) revealed that, like the WT, the transheterozygotes display a properly patterned arterial tree and normal Se and DLAV angiogenic development (Figure 2C,E; see also Supplementary file 3 and Materials and methods). These observations argue against the model that GIPCs directly enable PLXND1’s antiangiogenic signaling (see Burk et al., 2017). We conclude that the novel plxnd1skt6 allele complements the plxnd1fov01b null and encodes a functional receptor capable of antiangiogenic signaling.

Further support for the notion that reducing Plxnd1’s ability to associate with GIPCs does not inactivate the receptor comes from experiments driving mosaic transgenic endothelial expression of N-terminally tagged forms of zebrafish Plxnd1 in plxnd1fov01b null mutants (Figure 2—figure supplement 2). Briefly, we found that the WT form (2xHA-Plxnd1WT) and a receptor lacking the entire GBM because of removal of the nine C-terminal residues (2xHA-Plxnd1ΔGBM) had the same capacity to cell-autonomously rescue the vascular defects of plxnd1fov01b homozygotes (Figure 2—figure supplement 2D–E,G–H,J). In other words, endothelial cells within Se exogenously expressing either receptor form were absent from ectopic Se, had WT shapes concordant with their position within Se and DLAVs, and, when found within the base of sprouts, were positioned correctly just anterior to somite boundaries. These are the same features that WT endothelial cells display when transplanted into plxnd1fov01b hosts; see (Zygmunt et al., 2011).

This experiment does not clarify whether impairing Plxnd1’s GIPC-binding ability either increases or does not affect Plxnd1’s signaling. To address this point, we compared homozygous WT and homozygous plxnd1skt6 mutant sibling embryos derived from incrosses of plxnd1skt6/+heterozygotes. We found that plxnd1skt6 homozygous mutants showed an adequately organized vascular tree and a low frequency of Se and DLAV truncations (Figure 2G–I; see also the Materials and methods section ‘Quantification of angiogenesis deficits in the trunk’s arterial tree of WT and mutant zebrafish embryos’). However, the distributions of complete and truncated vessels and both the penetrance and expressivity of these defects were not statistically significantly different between WT and plxnd1skt6 mutants (Figure 2I, Figure 2—figure supplement 1 and Supplementary file 3). The unique presence of angiogenic deficits in the mutants, such as maximal and moderate Se and DLAV truncations (Figure 2H–I), suggests that the plxnd1skt6 allele is hypermorphic.

Homozygous plxnd1skt6 mutants are hypersensitive to the antiangiogenic drug SU5416

The results above show that the receptor encoded by the plxnd1skt6 allele is active, but do not fully clarify the nature of the plxnd1skt6 allele and the precise role of GIPCs in antiangiogenic PLXND1 signaling. Hence, we compared the sensitivity of homozygous WT and homozygous plxnd1skt6 mutants to the antiangiogenic compound SU5416, a tyrosine kinase inhibitor that preferentially targets VEGR2 (Herbert et al., 2012; Covassin et al., 2006; Fong et al., 1999.

If GIPCs limit PLXND1 antiangiogenic signaling, then plxnd1skt6 homozygotes should be hypersensitive to SU5416. In contrast, plxnd1skt6 mutants should be hyposensitive to SU5416 if GIPCs bind PLXND1 to directly enable the receptor’s antiangiogenic signaling (as in Burk et al., 2017) or if PLXND1 indirectly exerts a GIPC-dependent antiangiogenic effect by sequestering rate-limiting GIPCs away from proangiogenic VEGFR2/Nrp1 signaling (see Gay et al., 2011; Lanahan et al., 2013; Lanahan et al., 2010; Wang et al., 2006; Horowitz and Seerapu, 2012; Lanahan et al., 2014; Chittenden et al., 2006). Finally, if GIPCs play no role in PLXND1 signaling, then WT and plxnd1skt6 mutants should have similar SU5416 sensitivities.

As a first step, we evaluated the impact of 18–32 hpf SU5416 treatments (0.1–1 μM range) on WT angiogenesis (not shown), which defined a suboptimal SU5416 dose (0.2 μM; in 0.1% DMSO vehicle) capable of inducing minimal antiangiogenic effects on Se and DLAV development by 32 hpf (as in Covassin et al., 2006; Stahlhut et al., 2012; see also Torres-Vázquez et al., 2004; Zygmunt et al., 2011; Childs et al., 2002; Isogai et al., 2003; Zygmunt et al., 2012; Yokota et al., 2015). Accordingly, we treated homozygous WT and homozygous plxnd1skt6 mutants with 0.2 μM SU5416 (experimental group) or 0.1% DMSO (control group); see Figure 3, Figure 3—figure supplement 1 and Supplementary file 4.

Figure 3 with 1 supplement see all
plxnd1skt6 mutants are hypersensitive to the antiangiogenic drug SU5416.

(A–D) Confocal lateral images of the trunk vasculature (green) of 32 hpf embryos (region dorsal to the yolk extension). Anterior, left; dorsal, up. Scale bars (white horizontal lines), 100 μm. Treatments (DMSO or SU5416) indicated on top, genotypes (WT or plxnd1skt6) indicated on the left. Angiogenesis deficits are indicated with asterisks as follows: white (DLAV gaps), magenta (truncated Se). In the image of the DMSO-treated WT (C), the vessels are designated with the white font as follows: DLAV (Dorsal Longitudinal Anastomotic Vessel), Se (Segmental Vessel), DA (Dorsal Aorta), and PCV (Posterior Cardinal Vein). (E) Bar graph. Percentage of Se-DLAV in 32 hpf embryos of the indicated genotype and treatment combinations belonging to each of the following four phenotypic classes. Truncated: maximal (red; includes missing Se), moderate (yellow), and minimal (gray). Non-truncated: Full (black). The distributions of these four phenotypic classes were statistically significantly different between all the possible pairwise comparisons of the four combinations of treatments and genotypes. Significance values were calculated using a two-sided Fisher Exact test and significant differences (p<0.0083) assigned using a Bonferroni-type adjustment for six pairwise genotype comparisons (0.05/6 = 0.0083). Quantifications. We scored Se-DLAV angiogenesis (A–E) in embryos of the following four combinations of treatments and genotypes: DMSO-treated WT (312 Se-DLAV, 28 embryos, an average of 11.14 Se-DLAV/embryo), DMSO-treated plxnd1skt6 (284 Se-DLAV, 26 embryos, an average of 10.92 Se-DLAV/embryo), SU5416-treated WT (322 Se-DLAV, 29 embryos, an average of 11.10 Se-DLAV/embryo), and SU5416-treated plxnd1skt6 (320 Se-DLAV, 28 embryos, an average of 11.43 Se-DLAV/embryo). For additional data, graphs and statistical comparisons related to this figure, see Figure 3—figure supplement 1 and Supplementary file 4.

https://doi.org/10.7554/eLife.30454.006

DMSO and SU5416 had no apparent effects on embryonic shape, size, somite number, cardiac contractility, and circulation (not shown). DMSO slightly impaired Se and DLAV angiogenesis in both genotypes, consistent with its effects at higher doses (Hallare et al., 2006; Chen et al., 2011; Maes et al., 2012). The weak angiogenic impairment induced by DMSO was greater in plxnd1skt6 mutants, albeit without a significant penetrance difference (Figure 3A–B,E, Figure 3—figure supplement 1A and Supplementary file 4).

As expected, SU5416 increased the frequency and severity of angiogenesis deficits in both genotypes (Figure 3C–E, Figure 3—figure supplement 1 and Supplementary file 4). WT embryos and plxnd1skt6 mutants treated with DMSO and SU5416 exhibited statistically significantly different distributions of Se and DLAV truncations (Figure 3E and Supplementary file 4). Notably, DMSO-treated WT embryos and plxnd1skt6 mutants showed angiogenic deficits at statistically significantly different proportions, but only when comparing the ‘minimal’ category or the collective grouping of the three defective categories. In contrast, SU5416-treated WT embryos and plxnd1skt6 mutants displayed angiogenic deficits at statistically significantly different proportions when comparing the three defective categories, either individually or as a group (Figure 3E and Supplementary file 4). For example, while SU5416 induced a non-significant penetrance difference in Se-DLAV truncations between WT embryos (89.7%) and plxnd1skt6 mutants (100%) (Figure 3—figure supplement 1A), it significantly and disproportionally increased the frequency and severity of Se-DLAV angiogenesis deficits in plxnd1skt6 mutants (Figure 3E, Figure 3—figure supplement 1B and Supplementary file 4). In particular, SU5416 disproportionally increased the frequency of the most severe class of angiogenesis deficit in plxnd1skt6 mutants (maximal Se-DLAV truncation), a defect unique to SU5416-treated embryos of both genotypes (Figure 3E).

These effects indicate that plxnd1skt6 mutants are hypersensitive to SU5416. This conclusion fits the prediction of a single hypothesis, namely that GIPC binding limits PLXND1 signaling, and implies that the plxnd1skt6 allele hypermorphically increases antiangiogenic PLXND1 signaling.

Zebrafish gipc mutants display angiogenesis deficits in the trunk’s arterial tree

Mammalian studies implicate GIPC1 in promoting the NRP1-mediated proangiogenic activity of VEGF in the arterial tree (Waters, 2009; Habeck et al., 2002; Wang et al., 2006; Ren et al., 2010; Covassin et al., 2009; Lanahan et al., 2013; Lawson et al., 2003; Hermans et al., 2010; Chittenden et al., 2006; Dedkov et al., 2007; Lanahan et al., 2010). Vertebrates have three GIPCs, which are highly identical (Katoh, 2013), suggesting their potential functional redundancy. Indeed, in assays with recombinant mouse proteins purified from bacteria, all GIPCs can bind to PLXND1 (Shang et al., 2017). Impairing GIPC-PLXND1 complex formation at the receptor level, as in plxnd1skt6 mutants, yields angiogenesis deficits (Figure 2G–I) and SU5416 hypersensitivity (Figure 3). These defects fit the hypothesis that GIPCs promote arterial angiogenic development, at least in part, by limiting antiangiogenic PlxnD1 signaling. Given these findings, we anticipated that genetic inactivation of gipcs would similarly impair angiogenic development in the zebrafish arterial tree.

To define the vascular roles of zebrafish GIPCs, we used CRISPR/Cas9-based genome editing (Auer and Del Bene, 2014; Auer et al., 2014; Chang et al., 2013; Cong et al., 2013; Cong and Zhang, 2015; Gagnon et al., 2014; Hill et al., 2014; Hruscha et al., 2013; Hwang et al., 2013; Irion et al., 2014; Kimura et al., 2014; Mali et al., 2013; Talbot and Amacher, 2014) to make the first gipc1/synectin (gipc1skt1 and gipc1skt2), gipc2 (gipc2skt3 and gipc2skt4), and gipc3 (gipc3skt5) zebrafish mutants. Our novel gipc alleles are putative nulls predicted to encode short, truncated proteins lacking all domains (Figure 4—figure supplement 1; see also Supplementary file 8), and are therefore unable to interact with themselves, each other, Plxnd1, and Myo6; see (Shang et al., 2017).

Given the reported coexpression of GIPC1 and GIPC2 in the endothelium (Wang et al., 2006; Chittenden et al., 2006; Jiang et al., 2013; Butler et al., 2016), we focused on the vascular phenotype of the corresponding single and double zebrafish mutants in both zygotic and maternal-zygotic (MZ) combinations at 32 hpf. We scored 112 gipc mutants (69 embryos without maternal and zygotic gipc1 activity, including 19 animals devoid of zygotic gipc2 activity; plus 24 embryos lacking zygotic gipc1 activity, including 13 without gipc2 activity). This analysis revealed that gipc mutants display recessive Se angiogenic deficits of partial penetrance (Figure 4, Figure 4—figure supplement 2 and Figure 4—figure supplement 3; see also Figure 6C, Supplementary file 5, Supplementary file 6 and Materials and methods). Specifically, the Se vessels were sometimes missing or truncated, leading to corresponding DLAV gaps. At times, they were abnormally thin and straight-shaped. Importantly, every gipc mutant lacked supernumerary, ectopic, or misguided Se vessels, the hallmark defects of zebrafish and mice Plxnd1 nulls (Torres-Vázquez et al., 2004; Zygmunt et al., 2011; Gitler et al., 2004; Zhang et al., 2009; Gu et al., 2005; Worzfeld et al., 2014) and, reportedly, of murine Gipc1 mutants (Burk et al., 2017). We also found that as maternal or zygotic gipc dosage decreases, the penetrance and expressivity of Se truncations tended to increase (Figure 4—figure supplement 3). Together, these findings indicate that gipc1 and gipc2 act redundantly and that, at the zygotic level, loss of gipc1 causes a greater angiogenesis deficit than the removal of gipc2 (Figure 4, Figure 4—figure supplement 2 and Figure 4—figure supplement 3).

Figure 4 with 3 supplements see all
gipc mutants display angiogenesis deficits.

(A) Confocal lateral images of the trunk vasculature (green) of 32 hpf embryos (region dorsal to the yolk extension). Anterior, left; dorsal, up. Scale bars (white horizontal lines), 100 μm. Genotypes indicated on top of each image in yellow font. Angiogenesis deficits are indicated as follows: white asterisks (DLAV gaps), magenta asterisks (truncated Se), white greater-than sign (thin Se). Maternal-zygotic (MZ) removal of gipc activity is denoted by the designation ‘MZ’ in superscript. In the WT image (top left), the vessels are designated with the white font as follows: DLAV (Dorsal Longitudinal Anastomotic Vessel), Se (Segmental Vessel), DA (Dorsal Aorta), and PCV (Posterior Cardinal Vein). (B) Bar graph. Percentage of Se in 32 hpf embryos of the indicated genotypes belonging to each of the following four phenotypic classes. Truncated: severe (includes missing Se), medium (yellow), and weak (gray). Non-truncated: complete (black). Significance values were calculated using a two-sided Fisher Exact test and significant differences (p<0.0033) assigned using a Bonferroni-type adjustment for 15 pairwise genotype comparisons (0.05/15 = 0.0033). Brackets and asterisks indicate pairs of genotypes with significantly different distributions of these four phenotypic classes. Quantifications. We scored Se angiogenesis in embryos of the following six genotypes: WT (138 Se, 12 embryos; an average of 11.5 Se/embryo), gipc1skt1 (130 Se, 11 embryos; an average of 11.8 Se/embryo), gipc1skt1(MZ) (380 Se, 33 embryos; an average of 11.5 Se/embryo), gipc2skt3/skt4(130 Se, 11 embryos; an average of 11.8 Se/embryo), gipc1skt1; gipc2skt3/skt4 (152 Se, 13 embryos; an average of 11.6 Se/embryo), and gipc1skt1(MZ); gipc2skt3/skt4 (220 Se, 19 embryos; an average of 11.5 Se/embryo). For additional data, graphs, and statistical comparisons related to this figure, see Figure 4—figure supplement 2, Figure 4—figure supplement 3 and Supplementary file 5. Please note that given the use of different scales for scoring angiogenesis deficits, it is unfeasible to compare the quantifications in Figure 4 and Figure 5 directly.

https://doi.org/10.7554/eLife.30454.008

Overall, the angiogenic deficits of the single (gipc1 and gipc2) and double (gipc1; gipc2) zebrafish mutants resemble those found in gipc1 morphants (but without the delayed vasculogenesis, small aortic lumen, and abnormal body shape of the latter) (Ren et al., 2010; Hermans et al., 2010; Chittenden et al., 2006) and postnatal Gipc1 mutant mice (impaired arterial branching and arteriogenesis) (Ren et al., 2010; Chittenden et al., 2006; Dedkov et al., 2007; Lanahan et al., 2010; Moraes et al., 2013; Lanahan et al., 2014; Paye et al., 2009). They are also similar to the vascular defects found in kdrl mutant fish (in which inactivation of a VEGFR2 ohnolog leads to reduced VEGF signaling) (Habeck et al., 2002; Covassin et al., 2009; Bussmann et al., 2008) and plxnd1skt6 homozygotes.

These observations are consistent with the involvement of GIPCs in VEGF-induced proangiogenic VEGFR2/Nrp1-mediated signaling (see Gay et al., 2011; Lanahan et al., 2013; Lanahan et al., 2010; Wang et al., 2006; Horowitz and Seerapu, 2012; Lanahan et al., 2014; Chittenden et al., 2006) and support the hypothesis that GIPCs limit PLXND1 antiangiogenic signaling. Finally, these observations directly argue against the possibility that GIPC-PLXND1 binding directly promotes PLXND1’s antiangiogenic signaling (as in Burk et al., 2017) or that GIPCs are not involved in PLXND1 signaling.

Reducing Plxnd1 signaling ameliorates the angiogenic deficits of maternal-zygotic (MZ) gipc1 mutants

The hypothesis that the angiogenesis deficits of gipc mutants result, at least in part, from increased antiangiogenic PLXND1 signaling predicts that reducing the latter via heterozygosity for the null plxnd1fov01b allele might have a restorative effect on the angiogenic growth of zebrafish gipc mutants.

To test this, we compared the Se and DLAV angiogenesis phenotypes of gipc1skt1(MZ) and gipc1skt1(MZ); plxnd1fov01b/+ mutant siblings (Figure 5 and Figure 5—figure supplement 1; see also Materials and methods). Consistent with the proposed hypothesis, we found that plxnd1fov01b heterozygosity led to a dramatic (Figure 5A–B) and statistically significant difference in distribution of the four phenotypic classes between both genotypes (Figure 5C), with a statistically significant reduction in both the penetrance and expressivity of Se and DLAV truncations in gipc1skt1(MZ) mutants (Figure 5—figure supplement 1).

Figure 5 with 1 supplement see all
plxnd1 heterozygosity suppresses the angiogenesis deficits of gipc1skt1(MZ) mutants.

(A, B) Confocal lateral images of the trunk vasculature (green) of 32 hpf embryos (region dorsal to the yolk extension). Anterior, left; dorsal, up. Scale bars (white horizontal lines), 100 μm. Genotypes indicated on top of each image in yellow font. Angiogenesis deficits are indicated with asterisks as follows: white (DLAV gaps), magenta (truncated Se). (C) Bar graph. Percentage of Se-DLAV in 32 hpf embryos of the indicated genotypes belonging to each of the following four phenotypic classes. Truncated: maximal (red; includes missing Se), moderate (yellow), and minimal (gray). Non-truncated: full (black). There was a statistically significant difference (bracket with an asterisk) in distribution of the four phenotypic classes between gipc1skt1(MZ) and gipc1skt1(MZ); plxnd1fov01b/+ embryos, as assessed by a two-sided Fisher Exact test (p<0.05). Quantifications. We scored Se-DLAV angiogenesis in embryos of the following two genotypes: gipc1skt1(MZ) (390 Se-DLAV, 36 embryos, an average of 10.83 Se-DLAV/embryo) and gipc1skt1(MZ); plxnd1fov01b/+ (410 Se-DLAV, 38 embryos, an average of 10.79 Se-DLAV/embryo). For additional data, graphs, and statistical comparisons related to this figure, see Figure 5—figure supplement 1 and Supplementary file 6. Please note that given the use of different scales for scoring angiogenesis deficits, it is unfeasible to compare the quantifications in Figure 4 and Figure 5 directly.

https://doi.org/10.7554/eLife.30454.012

Double maternal-zygotic (MZ) gipc mutants display excessive angiogenesis in the absence of Plxnd1 signaling

To further explore the relationship between GIPCs and PLXND1 signaling, we used a validated splice-blocking morpholino to determine the vascular consequences of removing plxnd1 activity (Torres-Vázquez et al., 2004; Morcos, 2007; Stainier et al., 2017) from WT embryos and gipc1skt1(MZ); gipc2skt(Fischer et al., 2006)(MZ) double MZ mutants (Figure 6).

Removal of plxnd1 activity from gipc1skt1(MZ); gipc2skt4(MZ) maternal-zygotic (MZ) double mutants yields a phenotype similar to that of plxnd1 nulls.

(A–D) Confocal lateral images of the trunk vasculature (green) of 32 hpf embryos (region dorsal to the yolk extension). Anterior, left; dorsal, up. Scale bars (white horizontal lines), 100 μm. Morpholino injection (un-injected or injected with plxnd1 morpholino) indicated on top, genotypes (WT or gipc1skt1(MZ); gipc2skt4(MZ)) indicated on the left. The un-injected WT picture (A) shows the names of the major vessels in white font: DLAV (Dorsal Longitudinal Anastomotic Vessel), Se (Segmental Vessel), DA (Dorsal Aorta), and PCV (Posterior Cardinal Vein). Vascular defects highlighted as follows: truncated or missing Se (magenta asterisk), thin Se (white greater/less-than signs), DLAV gaps (white asterisk). Quantifications. The following number of embryos were analyzed: WT (four embryos), WT injected with plxnd1 morpholino (four embryos; 4/4 showed a vascular phenotype similar to that of plxnd1fov01b nulls), gipc1skt1(MZ); gipc2skt4(MZ) (12 embryos; 7/12 showed angiogenesis deficits), and gipc1skt1(MZ); gipc2skt4(MZ) injected with plxnd1 morpholino (11 embryos; 11/11 showed a vascular phenotype similar to that of plxnd1fov01b nulls).

https://doi.org/10.7554/eLife.30454.014

In contrast to the normal vasculature of WT embryos and the properly organized but hypoangiogenic vascular tree observed in gipc1skt1(MZ); gipc2skt(MZ) double MZ mutants (Fischer et al., 2006) (Figure 6A,C), we found that all the plxnd1 morphants exhibited the hyperangiogenic phenotype of plxnd1 nulls, namely excessive and disorganized Se angiogenesis (Figure 6B,D). The vascular phenotypes of plxnd1 morphants from these two backgrounds appeared qualitatively similar. This observation fits the notion that the angiogenesis deficits of gipc mutants are, at least partially, a result of increased PLXND1 signaling, consistent with the ameliorating effect of plxnd1fov01b heterozygosity in the angiogenic deficits of MZ gipc1 mutants (Figure 5 and Figure 5—figure supplement 1).

GIPC depletion potentiates SEMA3E-induced, PLXND1-dependent responses in HUVECs

To interrogate the role of GIPCs in endothelial PLXND1 signaling, we exploited the HUVEC model system in which PLXND1 signaling can be tuned by adjusting the exogenous supply of its canonical ligand, SEMA3E (Sakurai et al., 2010; Tata et al., 2014). This model offers well-defined molecular and cellular readouts of SEMA3E-induced PLXND1 signaling, such as a decreased level of phospho-active ERK1/2 (pERK1/2; Extracellular signal-related kinase 1 and 2) and cell collapse, caused by cytoskeletal disassembly and lowered adhesion to the substrate (Moriya et al., 2010; Sakurai et al., 2010; Wang et al., 2012; Tata et al., 2014; Aghajanian et al., 2014; Moriya et al., 2010; Aghajanian et al., 2014). Finally, these cells can be cultured without exogenous VEGF, which may have confounding effects (see Lanahan et al., 2013; Lanahan et al., 2010; Chittenden et al., 2006; Stahlhut et al., 2012; Lawson et al., 2002; Rossi et al., 2016; Jin et al., 2017; Lin et al., 2013; Carmeliet et al., 1996; Shalaby et al., 1997; Koch and Claesson-Welsh, 2012; Ruhrberg et al., 2002; Prahst et al., 2008; Soker et al., 1998; Yoshida et al., 2015; Lee et al., 2007; Domigan et al., 2015).

If GIPC negatively modulates PLXND1 signaling, then GIPC depletion should potentiate SEMA3E-induced responses in a PLXND1-dependent manner. Alternatively, if GIPCs directly promote PLXND1 signaling, then GIPC depletion should decrease SEMA3E-induced responses in a PLXND1-dependent fashion. Finally, if GIPCs are indirect players in PLXND1 biology or GIPCs only have PLXND1-independent roles, then GIPC depletion should not affect SEMA3E-induced, PLXND1-dependent responses. To distinguish between these possibilities, we used an immortalized HUVEC line (HUVEC/TERT2), shRNA-mediated GIPC knockdowns, and genome editing to knockout (KO) PLXND1 (see Zygmunt et al., 2011; Wang et al., 2014; Shalem et al., 2014; Sanjana et al., 2014; Ulrich et al., 2016); see Figure 7, Figure 7—figure supplement 1 and Figure 7—figure supplement 2. This approach minimized noise from genetic variability, enabled long-term antibiotic selection for maximal GIPC knockdown, and enabled the isolation, sequencing, and protein-level validation of PLXND1 KO clones (see Supplementary file 1, Supplementary file 8 and Figure 7—figure supplement 2). We accounted for potential off-target effects from constitutive expression of gene-specific shRNAs and guide RNAs (gRNAs) using validated, non-targeting shRNAs (Zygmunt et al., 2011; Wang et al., 2014; Ulrich et al., 2016) as controls.

Figure 7 with 4 supplements see all
GIPC depletion potentiates SEMA3E-induced, PLXND1-dependent ERK inactivation in HUVEC/TERT2 cells.

(A–D) Representative Western blot of active ERK1/2 (pERK) and total ERK1/2 (ERKTotal) from total cell lysates of HUVEC/TERT2 cells under the four conditions (shRNA and gRNA combinations) and the three ligand treatments indicated. Conditions. Control (A; bold black font), PLXND1 loss (B; bold red font), GIPC loss (C; bold green font), and GIPC-PLXND1 double loss (D; bold blue font). Treatments. Vehicle (-) and 2 nM SEMA3E for the indicated times. Cells stably carried a vector coexpressing the Cas9 nuclease and the indicated gRNAs. The alleles of the PLXND1 gRNAs (KOs) are stable and defined (see Supplementary file 1 and Supplementary file 8). (E) Bar graph. Means of percentual relative ERK activity (pERK/ERKTotal) under the described conditions (color coded as above) and treatments. Error bars, ± SEM. Relative ERK activity. Statistically significant differences between pairwise combinations of conditions and treatments are indicated (brackets and asterisks). Quantifications. n = 4 independent experiments per PLXND1 gRNA KO (for a pooled total of 8 experiments); n = 4 independent experiments per non-targeting gRNA (for a pooled total of 8 experiments). One-way ANOVA tests were conducted to determine whether relative ERK activity was significantly different between cells in the control, PLXND1 loss, GIPC loss, and GIPC-PLXND1 double loss conditions across each treatment. There were no outliers in the data, as assessed by inspection of a boxplot. Relative ERK activity data were normally distributed, for each treatment, as determined by Shapiro-Wilk’s test (p>0.05) except for the SEMA3E 15 min treatment; p=0.031. There was homogeneity of variances, as assessed by Levene's test (p>0.05) for equality of variances in all conditions. One-way ANOVA tests summary. Relative ERK activity was not statistically significantly different between conditions under vehicle treatment (F(3, 28)=0.004, p=1). Relative ERK activity was statistically significantly different between conditions under SEMA3E 15 min treatment (F(3, 28)=10.291, p<0.0005), effect size was ω2 = 0.46. Relative ERK activity was statistically significantly different between conditions under SEMA3E 45 min treatment (F(3, 28)=28.738, p<0.0005), effect size was ω2 = 0.72. Summary of the four statistically significant differences revealed by Tukey post hoc analysis (between conditions under SEMA3E 15 min treatment). Control versus PLXND1 loss was statistically significantly different (p<0.05): (95% CI (17.2450 to 67.2680); p=0.001). Control versus GIPC- PLXND1 double loss was statistically significantly different (p<0.05): (95% CI (−80.9622 to −12.0628); p=0.005). GIPC loss versus PLXND1 loss was statistically significantly different (p<0.05): (95% CI (17.1753 to 86.0747); p=0.002). GIPC loss versus GIPC-PLXND1 double loss was statistically significantly different (p<0.05): (95% CI (−90.0872 to −21.1878); p=0.001). Summary of the five statistically significant differences revealed by Tukey post hoc analysis (between conditions under SEMA3E 45 min treatment). Control versus PLXND1 loss was statistically significantly different (p<0.05): (95% CI (−59.2432 to −16.5068); p<0.0005). Control versus GIPC-PLXND1 double loss was statistically significantly different (p<0.05): (95% CI (−57.3307 to −14.5943); p<0.0005). GIPC loss versus PLXND1 loss was statistically significantly different (p<0.05): (95% CI 39.9193 to 82.6557); p<0.0005). GIPC loss versus GIPC-PLXND1 loss was statistically significantly different (p<0.05): (95% CI-80.7432 to −38.0068); p<0.0005). Control versus GIPC loss was statistically significantly different (p<0.05): (95% CI (2.0443 to 44.7807); p=0.028). For additional data, graphs, and statistical comparisons related to this figure, see Supplementary file 7.

https://doi.org/10.7554/eLife.30454.015

Using this setup, we measured the effects of SEMA3E treatment on relative pERK abundance over time in control cells and cells with GIPC loss, PLXND1 loss, and GIPC-PLXND1 double loss (Figure 7; see also Figure 7—figure supplement 1 and Figure 7—figure supplement 2). As expected, SEMA3E stimulation reduced relative pERK abundance in control cells (Figure 7A, black bars in Figure 7E). Importantly, we found that this effect is PLXND1-dependent because PLXND1 loss abrogated it (Figure 7B, red bars in Figure 7E). GIPC loss significantly potentiated the SEMA3E-induced decrease in relative pERK abundance at 45 min (Figure 7C, green bars in Figure 7E). Moreover, this potentiating effect is also PLXND1-dependent, because pERK levels failed to decrease under SEMA3E stimulation in GIPC-PLXND1 double loss cells (Figure 7D, blue bars in Figure 7E). Importantly, cells with GIPC-PLXND1 double loss did not show an intermediate relative level of pERK abundance between that of cells with PLXND1 loss (Figure 7B, red bars in Figure 7E) and cells with GIPC loss (Figure 7C, green bars in Figure 7E). Instead, the relative pERK abundance of SEMA3E stimulated cells with PLXND1 loss (Figure 7B, red bars in Figure 7E) and with GIPC-PLXND1 double loss (Figure 7D, blue bars in Figure 7E) were not statistically significantly different. These quantitative findings align with the qualitatively similar vascular phenotypes of plxnd1 morphants in the WT and gipc1skt1(MZ); gipc2skt(MZ) backgrounds (Fischer et al., 2006) (Figure 6B,D).

The results of additional cell culture experiments with primary HUVEC agree with these findings (Figure 7—figure supplement 3; see also Figure 7—figure supplement 4). In these cells, addition of exogenous SEMA3E induces a PLXND1-dependent collapsing response (Tata et al., 2014). As expected, SEMA3E induced the collapse of HUVEC treated with the non-targeting, shRNA control (Figure 7—figure supplement 3A,D) and this morphological change did not occur in PLXND1-depleted cells (Figure 7—figure supplement 3C,F). GIPC depletion potentiated the SEMA3E-induced morphological response, resulting in cell hypercollapse (Figure 7—figure supplement 3B,E).

In summary, the results of HUVEC assays (Figure 7 and Figure 7—figure supplement 3) support the hypothesis that GIPC depletion potentiates antiangiogenic PLXND1 signaling, consistent with the angiogenesis deficits and SU5416 hypersensitivity of plxnd1skt6 homozygous mutants (Figures 23). Given the demonstrated capacity of GIPCs and PLXND1 to bind directly to each other independently of NRP1 (Gay et al., 2011Shang et al., 2017) (Figure 1), the alignment of the plxnd1skt6 and HUVEC data argues that GIPCs bind to PLXND1 to directly limit its antiangiogenic signaling. Furthermore, the role of GIPCs as negative regulators of antiangiogenic PLXND1 signaling is also consistent with the angiogenesis deficits of gipc mutants (Figure 4) and the ameliorating effect that the partial removal of Plxnd1 signaling exerts on the angiogenic deficits of MZ gipc1 mutants (Figure 5).

Discussion

Our past crystallographic observations (Shang et al., 2017), together with the results of our CoIP, zebrafish, and cell culture experiments, and the consensus regarding the vascular and molecular roles of GIPC1 and PLXND1 in zebrafish (Torres-Vázquez et al., 2004; Zygmunt et al., 2011; Childs et al., 2002; Ren et al., 2010; Hermans et al., 2010; Chittenden et al., 2006; Yokota et al., 2015; Alvarez et al., 2007; Goi and Childs, 2016; Minchin et al., 2015) and mammals (Gitler et al., 2004; Zhang et al., 2009; Gu et al., 2005; Moriya et al., 2010; Fukushima et al., 2011; Kim et al., 2011; Worzfeld et al., 2014; Aghajanian et al., 2014; Ren et al., 2010; Chittenden et al., 2006; Lanahan et al., 2010; Gao et al., 2000; Moraes et al., 2013; Paye et al., 2009; Kanda et al., 2007; Degenhardt et al., 2010; Liu et al., 2016; Hegan et al., 2015) lead us to conclude that GIPCs association with PLXND1 negatively modulates the receptor’s antiangiogenic activity. This conclusion also fits with data from additional studies of the mammalian PLXND1 and PLXNB1 receptors. Briefly, BAC-based transgenic expression of full-length PLXND1 tagged at its C-terminus (a modification predicted to disable PBM-dependent binding events (Rickhag et al., 2013; Saras et al., 1997; Cao et al., 1999; Garbett and Bretscher, 2012) fully rescues the lethality, cardiac, vascular, and skeletal defects of murine Plxnd1 knockouts. In contrast, mutations that eliminate PLXND1’s GAP function inactivate the receptor (Worzfeld et al., 2014). Similarly, PLXNB1 harbors a C-terminal PBM with specificity for guanine nucleotide exchange factors, and removal of PLXNB1’s PBM does not inactivate the receptor (Oinuma et al., 2004).

Beyond its involvement in developmental angiogenesis, the GIPC-based modulation of endothelial PLXND1 signaling might promote the stabilization, repair, homeostasis, and arteriogenic remodeling of vessels, particularly in contexts with minimal proangiogenic stimulation. These conditions are found in the quiescent vascular beds of adults and are a hallmark of several diseases (see Carmeliet, 2003; Carmeliet, 2005).

We highlight that our results and conclusion challenge the model, recently proposed by Burk et al., that GIPC1 enables PLXND1 activity (Burk et al., 2017). These authors primarily address how Gipc1 and Plxnd1 modulate axonal circuit development using mouse embryos and neuronal explants. They propose that GIPC1’s PDZ domain and PLXND1’s canonical PBM drive GIPC1-PLXND1 heteromerization and, based on ex vivo forced expression experiments, conclude that the PBM is essential for PLXND1 activity. Finally, using a Plxnd1 null allele and floxed Gipc1 allele that might yield a truncated protein retaining PLXND1 binding (see Shang et al., 2017; Moraes et al., 2013), they peripherally explore the vascular role of the GIPC1-PLXND1 interaction using a qualitative phenotypic analysis that omits Plxnd1 mutant homozygotes.

Yet prior reports describe the viability, cardiovascular defects, and other abnormalities of Plxnd1 knockouts (Gitler et al., 2004; Zhang et al., 2009; Gu et al., 2005; Fukushima et al., 2011; Kim et al., 2011; Worzfeld et al., 2014; Aghajanian et al., 2014; Kanda et al., 2007; Degenhardt et al., 2010; Liu et al., 2016) and Gipc1 mutants (Ren et al., 2010; Chittenden et al., 2006; Dedkov et al., 2007; Lanahan et al., 2010; Moraes et al., 2013; Hegan et al., 2015) as contrastingly different, and the molecular roles for PLXND1 (Moriya et al., 2010; Aghajanian et al., 2014) and GIPC1 (Ren et al., 2010Paye et al., 2009; Chittenden et al., 2006; Herzog et al., 2011; Moraes et al., 2014) as distinct. Given that the molecular regulation of vascular development is highly conserved (Hogan and Schulte-Merker, 2017), we favor the view that GIPCs limit, in both zebrafish and mammals, the antiangiogenic signaling of the PLXND1 receptor.

In all likelihood, the proangiogenic function of GIPCs is multifaceted and involves additional mechanisms besides the negative modulation of PLXND1 signaling. First, GIPCs physically interact with many other PDZ-binding transmembrane proteins and with the retrograde motor MYO6. Some of these GIPC-binding proteins, for example, NRP1, are expressed in endothelial cells and have vascular functions (Shang et al., 2017; Katoh, 2013; Gao et al., 2000; Naccache et al., 2006; Salikhova et al., 2008; Lou et al., 2001; Cai and Reed, 1999). Second, the angiogenesis deficits of gipc1; gipc2 double mutants are more penetrant and severe than those of plxnd1skt6 homozygotes, in agreement with the notion that the vascular defects of gipc mutants are partially a result of enhanced Plxnd1 signaling.

We highlight that the identification of GIPCs as the first negative intracellular regulators of PLXND1 signaling expands the vascular roles of GIPCs beyond the prevailing notion that GIPC1 promotes arterial branching by facilitating, in an NRP1-dependent manner, proangiogenic VEGF signaling (Chittenden et al., 2006; Lanahan et al., 2010; Fantin et al., 2011; Lampropoulou and Ruhrberg, 2014; Herzog et al., 2011; Plein et al., 2014). Importantly, our findings do not argue against the possibility that GIPCs might also, either directly or indirectly, modulate PLXND1 signaling in connection with NRP1. For example, proangiogenic VEGF signaling via GIPC-NRP1-VEGFR2 complexes might trigger posttranslational modifications in GIPC and PLXND1 that counteract the antiangiogenic activity of the latter by promoting GIPC-PLXND1 interactions or the degradation of the PLXND1 receptor. Alternatively, GIPCs might mediate the formation of novel complexes containing receptors from both pathways (for instance, PLXND1-GIPC-NRP1 and PLXND1-GIPC-NRP1-VEGFR2) that perform still uncharacterized functions (see Chauvet et al., 2007; Bellon et al., 2010).

Our findings suggest the potential involvement of endocytosis and recycling, intracellular processes that regulate other pathways (Lanahan et al., 2010; Naccache et al., 2006; Reed et al., 2005; Salikhova et al., 2008; Hu et al., 2003; Jeanneteau et al., 2004a; Barbieri et al., 2016; Villaseñor et al., 2016; Koch et al., 2014), in modulation of PLXND1 signaling (see also Burk et al., 2017; Salikhova et al., 2008; Steinberg et al., 2013). We favor the model that GIPCs and MYO6 work together to facilitate PLXND1’s endocytic trafficking based on the following observations. GIPCs are MYO6-dependent internalization adaptors (Katoh, 2013; Wollscheid et al., 2016). Crystallographic findings indicate that the binding of PLXND1 conformationally releases GIPC1's domain-swapped autoinhibited dimer, promoting its interaction with MYO6 (Shang et al., 2017). Also, PLXND1, GIPC1, and MYO6 colocalize within intracellular puncta in SEMA3E-stimulated cells (Shang et al., 2017; see also Burk et al., 2017). Finally, the arterial tree deficits of Gipc1 and Myo6 mutants are similar (see Chittenden et al., 2006; Lanahan et al., 2010).

GIPCs (and MYO6) might limit PLXND1 signaling by restricting the amount of time that the activated receptor spends at the cell surface. This mechanism could serve a dual role of regulating access of the receptor to its catalytic targets such as the Rap1 GTPase, found at the inner cell membrane (Wang et al., 2012; Worzfeld et al., 2014; Wang et al., 2013), as well as shaping the inactivation kinetics of ligand-bound PLXND1. We note that recent structural studies indicate that mammalian MYO6 also functions as a ubiquitin receptor, thereby suggesting a potential link between GIPCs and the proteasome-mediated degradation of their cargo (He et al., 2016). Alternatively, GIPCs might function in a non-endocytic manner to limit PLXND1 signaling independently of MYO6. For example, GIPCs might recruit a third protein, which in turn, directly antagonizes PLXND1 signaling (see Lampropoulou and Ruhrberg, 2014; De Vries et al., 1998a; Lou et al., 2001; Wieman et al., 2009; Cai and Reed, 1999; Kofler and Simons, 2016; Guo and Vander Kooi, 2015; De Vries et al., 1998b; Fischer et al., 1999; Fischer et al., 2003; Jeanneteau et al., 2004b; Jean-Alphonse et al., 2014; Lin et al., 2006). Finally, the interaction between PLXND1 and GIPCs might directly influence cell morphology via modulation of cytoskeletal dynamics. Support for this notion comes from genetic experiments in Drosophila and mammalian proteomic studies that implicate GIPCs and MYO6 in actin network stabilization (Djiane and Mlodzik, 2010; O'Loughlin et al., 2018; Isaji et al., 2011; Noguchi et al., 2006).

The mechanisms that regulate GIPC-PLXND1 interaction are unclear. SEMA3E stimulates colocalization of GIPC1 and PLXND1 (Burk et al., 2017; Shang et al., 2017), but the kinetics of their association are not yet defined. Another open question is whether non-canonical PLXND1 ligands (SEMA3A, SEMA3C, SEMA3D, SEMA3G, and SEMA4A) (Epstein et al., 2015; Liu et al., 2016; Hamm et al., 2016) also promote GIPC-PLXND1 complex formation. How SEMA-induced changes in the conformation and oligomerization of PLXND1 (see Pascoe et al., 2015) impact the receptor’s ability to interact with GIPCs, or the specificity of their interaction is also unexplored. Finally, reversible posttranslational modifications might modulate the GIPC-PLXND1 interaction. Because phosphorylation of PDZ domains modulates the recruitment of their partners (Lee and Zheng, 2010; Liu et al., 2013), this is a plausible mechanism for GIPC-based control of GIPC-PLXND1 interactions. At the PLXND1 level, the phosphorylation and S-palmitoylation of the receptor’s cytosolic tail could also play a modulatory role. Plxns are phosphorylated (Cagnoni and Tamagnone, 2014; Franco and Tamagnone, 2008), and the GBM of PLXND1 harbors a conserved tyrosine located within a consensus Src family kinase phosphorylation site. This residue plugs into a hydrophobic GIPC pocket between the GH1 and PDZ domains (Gay et al., 2011; Shang et al., 2017). On the other hand, some type I transmembrane proteins, including Plxns, are S-palmitoylated (Holland and Thomas, 2017; Blaskovic et al., 2013), and S-palmitoylation of the carboxy tail of the GIPC1-binding dopamine Drd3 receptor buries the PBM within the cell membrane to prevent the Drd3-GIPC1 interaction (Arango-Lievano et al., 2016).

In conclusion, we have identified a novel role for the GIPCs as pioneer negative regulators of SEMA-PLXND1 signaling. Given the prominent role of this pathway in shaping organogenesis of cardiovascular, nervous, and other systems and its central importance in cancer biology (Moriya et al., 2010; Gay et al., 2011; Valdembri et al., 2016; Oh and Gu, 2013; Gaur et al., 2009; Gu and Giraudo, 2013; Neufeld et al., 2016; Bielenberg and Klagsbrun, 2007), our findings suggest a broad human health relevance for therapeutic targeting of the SEMA-PLXND1 pathway at the GIPC level.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional information
Genetic reagent
(Danio rerio)
Tg(kdrl:HsHRAS-mCherry)s896DOI:10.1101/gad.1629408ZFIN ID: ZDB-
ALT-081212–4
Transgenic insertion
Genetic reagent
(Danio rerio)
Tg(fli1a:EGFP)y1PMID:12167406ZFIN ID: ZDB-
ALT-011017–8
Transgenic insertion
Genetic reagent
(Danio rerio)
Tg(fli1a:GAL4FF)ubs4DOI:10.1016/j.devcel.
2011.06.033
ZFIN ID: ZDB-
ALT-110921–1
Transgenic insertion
Genetic reagent
(Danio rerio)
Tg(flt1:nlsmCherry)skt7This paperTransgenic insertion.
Made using Torres-Vázquez
lab plasmid #1208
Genetic reagent
(Danio rerio)
gipc1skt1This paperPutative null mutant allele
Genetic reagent
(Danio rerio)
gipc1skt2This paperPutative null mutant allele
Genetic reagent
(Danio rerio)
gipc2skt3This paperPutative null mutant allele
Genetic reagent
(Danio rerio)
gipc2skt4This paperPutative null mutant allele
Genetic reagent
(Danio rerio)
gipc3skt5This paperPutative null mutant allele
Genetic reagent
(Danio rerio)
plxnd1fov01bPMID: 11861480
DOI:10.1016/j.devcel.
2004.06.008
ZFIN ID: ZDB-ALT
-010621–6
Null mutant allele
(point mutation)
Genetic reagent
(Danio rerio)
plxnd1skt6This paperHypermorphic mutant
allele
Cell line
(Cercopithecus aethiops)
COS-7 (Monkey
Kidney Fibroblasts)
American Type
Culture Collection
Cat. #CRL-1651.
RRID:CVCL_0224
https://www.atcc.org/products/All/CRL-1651.aspx
Cell line
(Homo sapiens)
HUVEC/TERT2
(Immortalized Human
Umbilical Vein Endothelial
Cells)
American Type
Culture Collection
Cat. #CRL-4053.
RRID:CVCL_9Q53
https://www.atcc.org/Products/All/CRL-4053.aspx
Cell line
(Homo sapiens)
HUVEC (Normal
Primary Human Umbilical
Vein Endothelial Cells)
Lifeline Cell TechnologyCat. #FC-0003https://www.lifelinecelltech.com/shop/cells/human-endothelial-cells/umbilical-vein-endothelial-cells/huvec-fc-0003/
Cell line
(Homo sapiens)
Non-targeting gRNA1.
Pool of HUVEC/TERT2
cells.
This paperDerived from HUVEC/TERT
2 cell line (ATCC CRL4053).
Cells were grown under blasticidin
(4 μg/ml) selection and used
between 7th and 10th passages.
Cells stably coexpress
Cas9 nuclease and non-targeting
gRNA1 (from Torres-Vázquez
lab plasmid #1859)
Cell line
(Homo sapiens)
Non-targeting gRNA2.
Pool of HUVEC/TERT2
cells.
This paperDerived from HUVEC/TERT
2 cell line (ATCC CRL4053).
Cells were grown under
blasticidin (4 μg/ml) selection
and used between 7th-10th
passages.
Cells are stably coexpressing
Cas9 nuclease and non-
targeting gRNA2 (from
Torres-Vázquez lab plasmid #1860)
Cell line
(Homo sapiens)
PLXND1 gRNA KO1.
Monoclonal PLXND1 KO
HUVEC/TERT2 cell line.
This paperBiallelic (transheterozygous)
PLXND1 knockout line. Derived
from HUVEC/TERT 2 cell line
(ATCC CRL4053). Cells were
grown under blasticidin (4 μg/ml)
selection and used between
7th-10th passages.
Cells are stably coexpressing
Cas9 nuclease and PLXND1
gRNA KO1 (from Torres-Vázquez
lab plasmid #1846)
Cell line
(Homo sapiens)
PLXND1 gRNA KO2.
Monoclonal PLXND1 KO
HUVEC/TERT2 cell line.
This paperBiallelic (transheterozygous)
PLXND1 knockout line. Derived
from HUVEC/TERT 2 cell line
(ATCC CRL4053). Cells were
grown under blasticidin (4 μg/ml)
selection and used between
7th-10th passages.
Cells are stably coexpressing
Cas9 nuclease and PLXND1
gRNA KO2 (from Torres-Vázquez
lab plasmid # 1847)
Cell line
(Homo sapiens)
HEK293T
(embryonic kidney cells)
Matthias
Stadtfeld lab, NYU
Recombinant
DNA reagent
V5-C-mPLXND1WTThis paperTorres-Vázquez lab
plasmid #862. Vector
backbone: pcDNA3.1/
nV5-DEST-V5
Recombinant
DNA reagent
V5-C-mPLXND1ΔCYSEAThis paperTorres-Vázquez lab
plasmid #863. Vector
backbone: pcDNA3.1/
nV5-DEST-V5
Recombinant
DNA reagent
V5-C-mPLXND1ΔGBMThis paperTorres-Vázquez lab
plasmid #1774. Vector
backbone: pcDNA3.1/
nV5-DEST-V5
Recombinant
DNA reagent
FLAG-mGIPC1WTDOI:10.1091/mbc.12.3.615Torres-Vázquez lab
plasmid #864. Vector
backbone: pFLAG-CMV1
Recombinant
DNA reagent
FLAG-mGIPC1GH1DOI:10.1091/mbc.12.3.615Torres-Vázquez lab
plasmid #868. Vector
backbone: pFLAG-CMV2
Recombinant
DNA reagent
FLAG-mGIPC1PDZDOI:10.1091/mbc.12.3.615Torres-Vázquez lab
plasmid #866. Vector
backbone: pFLAG-CMV3
Recombinant
DNA reagent
2xHA-Plxnd1WTThis paperGAL4-responsive,
Gateway and IRES-based
bicistronic vector for Tol2-
mediated zebrafish transgenesis.
Torres-Vázquez lab plasmid #1414
Recombinant
DNA reagent
2xHA-Plxnd1ΔGBMThis paperGAL4-responsive, Gateway
and IRES-based bicistronic
vector for Tol2-mediated
zebrafish transgenesis.
Torres-Vázquez lab plasmid #1685
Recombinant
DNA reagent
lentiCRISPR v2-BlastAddgeneCat. #83480A gift from Mohan Babu.
https://www.addgene.org/83480/
Recombinant
DNA reagent
Non-targeting gRNA1This paperTorres-Vázquez lab
plasmid #1859. Vector
backbone: lentiCRISPR
v2-Blast
Recombinant
DNA reagent
Non-targeting gRNA2This paperTorres-Vázquez lab
plasmid #1860. Vector
backbone: lentiCRISPR
v2-Blast
Recombinant
DNA reagent
PLXND1-KO1This paperTorres-Vázquez lab
plasmid #1846. Vector
backbone: lentiCRISPR v2-Blast
Recombinant
DNA reagent
PLXND1-KO2This paperTorres-Vázquez lab
plasmid #1847. Vector
backbone: lentiCRISPR v2-Blast
Recombinant
DNA reagent
Control shRNA Lentiviral
Particles-A (Non-targeting
control shRNA)
Santa Cruz
Biotechnology
Cat. #sc-108080Encodes a non-targeting
shRNA sequence, will not
lead to the specific degradation
of any known cellular mRNA
Recombinant
DNA reagent
GIPC shRNA (h)
Lentiviral Particles
Santa Cruz
Biotechnology
Cat. #sc-35475-VshRNA pool (three
target-specific constructs
against human GIPC1 that
encode 19–25 nt
(plus hairpin) shRNAs).
Target sequences
(sense sequences 5’ to 3’):
(1) CUGACGAGUUCGUCUUUGA
(2) CCACCACUUUCCACCAUCA
(3) CUGAAUUUGCUGUCUUGAA
Recombinant
DNA reagent
GIPC2 shRNA (h)
Lentiviral Particles
Santa Cruz
Biotechnology
Cat. # sc-75132-VshRNA pool (three
target-specific constructs
against human GIPC2 that
encode 19–25 nt (plus hairpin)
shRNAs). Target sequences
(sense sequences 5’ to 3’):
(1) CAGACGAAUUUGUCUUUGA
(2) GGACACCUUUACUAACUCU
(3) CCAACUUUCUCUCUUUGUA
Recombinant
DNA reagent
GIPC3 shRNA (h)
Lentiviral Particles
Santa Cruz
Biotechnology
Cat. #sc-62376-VshRNA pool (three
target-specific constructs
against human GIPC3 that
encode 19–25 nt (plus hairpin)
shRNAs). Target sequences
(sense sequences 5’ to 3’):
(1) CCUUCAUCAAGAGAAUCAA
(2) GGAGUUUGCACGCUGUUUA
(3) GACAAGUUCCUCUCUAGAA
Recombinant
DNA reagent
Plexin-D1 shRNA
(h) Lentiviral Particles
Santa Cruz
Biotechnology
Cat. #sc-45585-VshRNA pool (three
target-specific constructs
against human PLXND1 that
encode 19–25 nt (plus hairpin)
shRNAs). Target sequences (sense
sequences 5’ to 3’):
(1) GUCAAGAUAGGCCAAGUAA
(2) CCAUGAGUCUCAUAGACAA
(3) CCACAGACAGUUUCAAGUA
Sequenced-based
reagent
plxnd13207-3462 morpholinoDOI:10.1016/j.devcel.
2004.06.008
Validated splice-blocking
morpholino against zebrafish
plxnd1. Synthesized by
GENE TOOLS, LLC).
Sequence (5’ to 3’):
CACACACACTCACGTTGATGATGAG
AntibodyChicken anti-GFPInvitrogenCat. #A10262IF (1:1,000); zebrafish
AntibodySheep anti-mCherryHolger Knaut lab, NYUIF (1:1,000); zebrafish.
Custom made antibody
AntibodyMouse anti-pFAK Tyr397MilliporeCat. #05–1140IF (1:1,000); zebrafish
AntibodyRabbit anti-GIPC1Proteintech GroupCat. #14822–1-AP.
RRID:AB_2263269
WB (1:3,000). This antibody
detects GIPC1, GIPC2,
and GIPC3 (our data)
AntibodyRabbit anti-GIPC2AbcamCat. #ab175272WB (1:5,000). This antibody
detects GIPC1 and
GIPC2 (our data)
AntibodyRabbit anti-GIPC3AbcamCat. #ab186426WB (1:5000). This antibody
is specific for GIPC3 (our data).
Validated against HeLa TCL
(positive control; a gift from
Mamta Tahiliani’s lab, NYU)
AntibodyMouse anti-PLXND1R and D SystemsCat. #MAB41601
Clone #752815
WB (1:250). Lyophilized
reagent reconstituted in
200 μl of sterile PBS
(GIBCO, Cat. #10010–023)
AntibodyRabbit anti-Phospho-
p44/42 MAPK (Erk1/2)
(Thr202/Tyr204) (D13.14.4E)
XPTM
Cell Signaling TechnologyCat. #4370S.
RRID:AB_2315112
WB (1:20,000)
AntibodyMouse anti-p44/42
MAPK (Erk1/2) (L34F12)
Cell Signaling TechnologyCat. #4696SWB (1:10,000)
AntibodyRabbit anti-GAPDH
(D16H11)
Cell Signaling TechnologyCat. #5174P.
RRID:AB_10622025
WB (1:20,000)
AntibodyMouse anti-FLAG M2SIGMA-ALDRICHCat. #F3165,
clone M2.
RRID:AB_259529
WB (1:20,000)
AntibodyRabbit anti-V5-Tag
(D3H8Q)
Cell Signaling TechnologyCat. #13202S.
RRID:AB_2687461
WB (1:10,000)
Peptide,
recombinant protein
Human Semaphorin 3ER and D SystemsCat. #3239-S3BWorking concentration
of 2 nM (prepared in 1xPBS
with 0.1%BSA (SIGMA_ALDRICH,
Cat.A8022)
Chemical
compound, drug
SU5416SIGMA-ALDRICHCat. #S8442Working concentration
of 0.2 μM in fish water.
From 10.5 mM stock solution
in DMSO (SIGMA-ALDRICH,
Cat. #D8418)
Chemical
compound, drug
Gelatin, from porcine skinSIGMA-ALDRICHCat. # G1890-100GWorking concentration
of 0.1% (prepared in distilled
water and then autoclaved)
Chemical
compound, drug
Blasticidin S HCl, powderThermoFisher ScientificCat. #R21001From a stock solution
of 10 mg/ml. Prepared in
UltraPure Distilled water
(Invitrogen Cat. # 10977–015)
Chemical
compound, drug
Puromycin DihydrochlorideThermoFisher ScientificCat. #A1113803From a stock solution of 10 mg/ml

Biochemistry and cell culture experiments

Yeast two-hybrid screen

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The pGBK-T7 and pACT2 vectors (Clonetech Laboratories, Inc) were used, respectively, for the bait and the preys. The bait consisted of a 96 kDa fusion protein harboring N-terminally the Myc-tagged DNA-binding domain of GAL4 and the cytosolic tail of mPLXND1 (632 aa) at its carboxyl end. A pre-transformed cDNA library from E11 stage mouse embryos undergoing PLXND1-dependent vascular and neuronal development (Gitler et al., 2004; van der Zwaag et al., 2002) (Clonetech Laboratories, Inc) was used to make prey proteins fused to the C-terminus of an HA-tagged GAL4 activation domain. The screening (2.8 × 106 preys) was performed by Dualsystems Biotech AG and yielded 15 bait-dependent clones.

Co-immunoprecipitation of mGIPC-mPLXND1 complexes in COS7 cells

Details for the cell line used

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COS-7 cells (Monkey Kidney Fibroblasts) purchased from the American Type Culture Collection (ATCC); ATCC #CRL-1651 (unrecorded lot number, thus the Certificate of Analysis containing detailed authentication and mycoplasma contamination information is unretrievable from ATCC’s website).

Cell culture and transfection

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COS-7 cells were cultured in 10 cm diameter culture dishes (Falcon #379096) with Dulbecco's modified Eagle's medium (DMEM; Corning cellgro #10–013-CV) and 10% fetal bovine serum (FBS; Gemini Bio-Products #100–106). Cells were grown using a 37°C humidified 5% CO2 atmosphere. Transfections were performed at 70% confluency with Lipofectamine2000 (Thermofisher Scientific #11668027) according to the manufacturer’s specifications using 2.5 ug of each plasmid. Four hours post-transfection the medium was replaced with fresh complete DMEM. Expression vectors and their V5-tagged mPLXND1 and FLAG-tagged mGIPC protein payloads are described in Supplementary file 1 and Supplementary file 8.

Immunoprecipitation and western blotting

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48 post-transfection, whole cell lysates were prepared as follows. Dishes were placed on ice, rinsed once with PBS (phosphate-buffered saline) and lyzed in 1 ml of ice-cold lysis buffer (50 mM Tris pH 7.5, 5 mM EDTA, 150 mM NaCl, 1% Triton X-100) supplemented with the EDTA-free protease inhibitor cocktail c0mplete (Roche #12683400). FLAG-tagged GIPC proteins were immunoprecipitated overnight at 4°C with rocking using an anti-FLAG antibody (SIGMA #F1804) and 30 ul of Protein A/G PLUS-Agarose (Santa Cruz Biotechnology #sc-2003). Next day, the protein immunoprecipitates were washed 3x with washing buffer (50 mM Tris-HCl pH 7.5 and 150 mM NaCl). Samples were loaded onto pre-casted NuPAGE 4–12% Bis-Tris protein gels (Invitrogen #NP0335BOX). The immunoprecipitated FLAG-GIPC1 forms and the interacting co-immunoprecipitated V5-tagged mPLXND1 proteins were detected by Western blotting using anti-FLAG (SIGMA #F1804) and anti-V5 antibodies (CST #13202S), respectively.

Quantification of protein-protein interactions between V5-C-mPLXND1 forms and FLAG-mGIPC1WT from FLAG immunoprecipitates and V5 co-immunoprecipitates

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Data were collected from three independent experiments for each protein interaction and analyzed using a one-way ANOVA test followed by a Tukey post hoc analysis. See Figure 1C (left-side bar graph and legend) and Supplementary file 2. Data analysis was carried out using the SPSS Statistics 23.0 software package and the Laerd Statistics tutorial (Statistics, 2015a).

Quantification of the relative abundance of V5-C-mPLXND1 forms and FLAG-mGIPC1WT from total cell lysates (TCL)

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Data were collected from three independent experiments for each protein interaction and analyzed using a Kruskal-Wallis H test. See Figure 1C (right-side bar graph and legend) and Supplementary file 2. Data analysis was carried out using the SPSS Statistics 23.0 software package and the Laerd Statistics tutorial (Statistics, 2015b).

Experiments with HUVEC/TERT2 cells

Details for the cell line used

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Immortalized HUVEC cells (HUVEC/TERT2) purchased from the American Type Culture Collection (ATCC); ATCC #CRL-4053 (authenticated via STR profiling and mycoplasma negative).

Construction of vectors for Cas9 and gRNA coexpression

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We used restriction cloning (BsmBI) to clone into the lentiviral vector lentiCRISPR v2-Blast (a gift from Mohan Babu; Addgene plasmid # 83480) pairs of aligned oligos to make the following gRNAs: non-targeting gRNA1, non-targeting gRNA2, PLXND1-KO1, and PLXND1-KO2 (Supplementary file 1); as in (Shalem et al., 2014; Sanjana et al., 2014). The four resulting vectors were propagated in Stbl3 bacteria (ThermoFisher Scientific #C737303) at 32°C.

Cell culture, lentivirus production, and infection with lentiCRISPR v2-Blast vectors for Cas9 and gRNA coexpression

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Immortalized HUVEC cells were cultured in low serum medium optimized for human endothelial cells and without human growth factors (VascuLife EnGS Endothelial Medium Complete Kit from Lifeline Cell Technology #LL-0002). Cells were grown in 10 cm diameter culture plates (Falcon #379096) pre-covered with 0.1% gelatin (SIGMA #G1890-100G) and incubated in a humidified 5% CO2 atmosphere at 37°C. Human embryonic kidney 293 T cells (HEK293T; gift of Matthias Stadtfeld; NYU) were cultured in high D-Glucose Dulbecco's modified Eagle's media (Gibco # 10313–021) containing 10% FBS and 2 nM L-glutamine (Gibco #25030–081). Plates (10 cm) of human embryonic kidney 293T (90% confluence) were individually transfected with one of the four lentiCRISPR v2-Blast constructs along with lentivirus packaging and envelope plasmids (gift of Matthias Stadtfeld; NYU) using TransIT-293 reagent (Mirus Bio #Mir2700). Each plate was transfected with 15 μg of the lentiCRISPR v2-Blast construct, 0.75 μg of each lentivirus packaging vector (tat, rev, and gag/pol) and 1.5 μg of pVSV-G. Viral supernatants were harvested and filtered through 0.45 μm at 48 and 72 h. HUVEC cells were infected at ~50% confluency using 100 μl of viral supernatants and 5 μg/ml polybrene (Millipore #TR-1003-G) for 2 consecutive days. Pools of stable HUVEC cells were selected for 5 days with 4 μg/ml blasticidin (ThermoFisher Scientific # R21001). Pilot studies confirmed that these selection conditions killed 100% of the uninfected cells.

Isolation of monoclonal cell populations of PLXND1 knockout cells infected with lentiCRISPR v2-Blast vectors for Cas9 and gRNA coexpression

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Pools of stable HUVEC cells were harvested using trypsin (Gibco #25300–054) and grown at a very low density in 15 cm diameter culture plates (Corning #430599). Twelve individual cells from each of the PLXND1-KO1 and PLXND1-KO2 gRNAs expressing lentiCRISPR v2-Blast vectors were grown to 100% confluence in 96-well plates. These cells were then expanded in larger plates until confluent 10 cm plates were obtained. The resulting monoclonal cell populations were used for characterizing the DNA sequence (GENEWIZ performed sequencing) and protein level via Western blot to identify PLXND1 knockout lines. Two of these (PLXND1 gRNA KO1 and KO2) were used for the experiments presented here.

shRNA infection

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Blasticidin-resistant stable cells infected with lentiCRISPR v2-Blast vectors for Cas9 and gRNA coexpression were infected as follows. Briefly, pools of cells expressing a non-targeting vector (gRNA1 or gRNA2) and monoclonal PLXND1 KO lines (PLXND1 gRNA KO1 or KO2) were infected at ~ 50% confluency using 5 μg/ml polybrene (Millipore #TR-1003-G) for 2 consecutive days. Infections were performed with the puromycin-resistant shRNA lentiviral particles, with the non-targeting control, GIPC1/2 pool (Santa Cruz Biotechnology #sc-108080, sc-35475-V, and sc-75132-V). The medium was changed 24 h after the second infection, and infected cells were selected with 1.5 μg/ml puromycin (ThermoFisher Scientific #A1113803) for 72 hr.

Detection of phospho-active (pERK) and total ERK (ERKTotal) and quantification of relative ERK activity

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Cells were stimulated with either vehicle or 2 nM recombinant human Semaphorin 3E (R and D Systems #3239-S3B) for 15 and 45 mins. pERK and ERKTotal levels were measured using Western blots from total cell lysates (TCLs) prepared as follows. Cell plates were placed on an ice bed, rinsed with PBS and lyzed in 250 μl of lysis buffer (50 mM Tris pH 7.5, 0.5 mM EDTA, 150 mM NaCl, 1 mM EDTA, 1% SDS) supplemented with protease (cOmplete, Roche #12683400) and phosphatase (1 mM sodium fluoride, 10 mM β-glycerophosphate, and 1 mM sodium vanadate) inhibitors. Lysates were loaded onto NuPAGE 4–12% Bis-Tris protein gels (Invitrogen #NP0336BOX) and proteins transferred to Immobilon-P membranes (Millipore; #IPVH00010). pERK and ERKTotal were immunodetected with rabbit and mouse antibodies against ERK1/2 (CST #4370S and #4696S, respectively) and revealed with Western Lightning PLUS-ECL (PerkinElmer #NEL103001EA). Relative ERK activity was calculated ratiometrically as pERK/ERKTotal.

Experiments with primary HUVEC cells

Details for the cell line used

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Normal primary human umbilical vein endothelial cells (HUVEC) were purchased from Lifeline Cell Technology; #FC-0003 (STR profiling data unavailable, mycoplasma negative).

Cell collapse assays

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Cells between passages 2 and 6 were infected with shRNA lentiviral particles and puromycin-selected for 48 h as described above. Cells were divided into two pools, one for validating the knockdown and the other for performing the cellular morphology assay. Cells for the latter experiment were starved for 6 h and then treated with either vehicle or 10 nM recombinant human Semaphorin 3E (R and D Systems #3239-S3B) for 45 min. Cells were washed with ice-cold 1xPBS, fixed at room temperature (RT) for 15 min with 4% PFA in PBS. Cells were then washed 2x with PBS and permeabilized with 0.1% Triton-X100 in PBS for 5 min at RT. Next, cells were incubated for 20 min at RT in 20 μg/ml phalloidin-tetramethylrhodamine B isothiocyanate (SIGMA #P1951) to label the F-actin cytoskeleton. Cells were next washed 3x in PBS, incubated for 5 min at RT in 0.5 μg/ml DAPI (Molecular Probes #D1306) to visualize nuclei, and kept in PBS at 4°C until imaging. Fluorescent images were acquired with an Eclipse Ti-E inverted microscope (Nikon) using the 10X objective (NA 0.3).

Cell collapse data collection and evaluation

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Images were collected from 50 to 100 cells from each of three independent experiments per knockdown and treatment condition. Images were processed with FIJI (https://imagej.net/Fiji). As automated extraction of cellular contours was unfeasible, cells were qualitatively classified as uncollapsed, collapsed, or hyper-collapsed based on their size and morphology (Figure 7—figure supplement 3).

Western blot validation of GIPC knockdowns and PLXND1 knockout in primary and immortalized HUVEC

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Western blots of TCLs were performed with rabbit antibodies for GIPC1 (Proteintech Group; 14822–1-AP) and GIPC2 (Abcam #ab175272). A mouse antibody was used for PLXND1 (R and D Systems #MAB41601). GAPDH (loading control) was detected with a rabbit antibody (CST; #5174P).

Zebrafish experiments

Genome editing for making gipc and plxnd1 mutants

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gRNA design and genomic target site selection were performed using the CHOPCHOP and CRISPRscan web tools (Labun et al., 2016; Montague et al., 2014; Moreno-Mateos et al., 2015). gRNAs were in vitro transcribed from PCR-assembled oligo templates (Integrated DNA Technologies) as in (Gagnon et al., 2014) (see Supplementary file 1). G0 fish were made by delivering 1 nl of a genome editing mix containing S. pyogenes Cas9 nuclease and one or more gene-specific gRNAs (125 pg/per gRNA) into the cytoplasm of one cell stage Tg(fli1a:EGFP)y1 embryos (see Auer and Del Bene, 2014; Auer et al., 2014; Chang et al., 2013; Cong et al., 2013; Cong and Zhang, 2015; Gagnon et al., 2014; Hruscha et al., 2013; Hwang et al., 2013; Irion et al., 2014; Kimura et al., 2014; Mali et al., 2013; Talbot and Amacher, 2014). Cas9 was provided as mRNA (300 pg; in vitro transcribed from the pST1374-NLS-flag-linker-Cas9 vector (Shen et al., 2013), a kind gift from Xingxu Huang; Addgene plasmid #44758) or protein (500 pg; PNA Bio #CP01). Genome editing of G0 embryos was assessed by evaluating PCR products amplified with BioReady Taq DNA Polymerase (Bulldog Bio #BSAX050) from gDNA of 4–12 pooled individuals via agarose gel electrophoresis and single-colony Sanger sequencing. For the latter, PCR products were TOPO-TA cloned into the pCRII-TOPO or pCR2.1-TOPO vector (ThermoFisher Scientific; #450640 or #450641), and 8–20 individual colonies were sequenced. To identify F1 heterozygous mutant carriers, gDNA from individuals was PCR-amplified, and the biallelic sequence trace analyzed using the Poly Peak Parser and TIDE web tools (Hill et al., 2014; Brinkman et al., 2014). The molecular nature of the mutant alleles of F1 parents was confirmed using single-colony Sanger sequencing of PCR products amplified from fin clip-derived gDNA GENEWIZ performed sequencing.

Forced endothelial expression of 2xHA-Plxnd1 forms in plxnd1fov01b mutants

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We used the GAL4/UAS system (Scheer and Campos-Ortega, 1999) and Tol2-based transient transgenesis (Kikuta and Kawakami, 2009) to drive forced mosaic endothelial-specific expression of 2xHA-tagged forms of zebrafish Plxnd1 (2xHA-Plxnd1WT or 2xHA-Plxnd1ΔGBM) and the green fluorescent marker EGFP. Briefly, 1 nl of a 100 pg of Tol2 mRNA and 20 pg of vector DNA solution was injected into the cytoplasm of one-cell plxnd1fov01b mutants carrying both the endothelial Tg(fli1a:GAL4FF)ubs4 GAL4 driver (Zygmunt et al., 2011) and the red nuclear arterial Tg(flt1:nls-mCherry)skt7 reporter (this study). Faithful coexpression of 2xHA-Plxnd1 and EGFP was accomplished using an IRES (internal ribosomal entry site element)-based bicistronic UAS cassette (Kwan et al., 2007). Embryos were fixed at 32 hpf, immunostained and imaged as described below.

Quantification of the vascular patterning activity of exogenous 2xHA-Plxnd1 forms in plxnd1fov01b mutants

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Arterial endothelial clones with exogenous expression of 2xHA-Plxnd1 forms (EGFP+) that occupied the Se, DLAV, or both (Se-DLAV) positions were qualitatively scored as exhibiting a WT-like phenotype if they were properly shaped and positioned according to the following definitions. For Se sprouts: unbranched chevron morphology and, if connected to the DA, a base located just anterior to the somite boundary. For DLAVs: unbranched linear morphology parallel to the roof of the spinal cord (Figure 2—figure supplement 2D–J and Supplementary file 3).

Embryo treatments with SU5416 and DMSO

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SU5416 (SIGMA #S8442) was prepared and used as in Covassin et al. (2006) and Stahlhut et al. (2012). Briefly, a 200 μM SU5416 stock solution in DMSO (SIGMA #D8418; vehicle) was dissolved in fish water to a final concentration of 0.2 μM SU5416 (a suboptimal dose) and 0.1% DMSO. Control, vehicle-only (0.1% DMSO) treatments were also performed. Homozygous WT and homozygous plxnd1skt6 mutant embryos were manually dechorionated before receiving the SU5416 and DMSO treatments using a common solution for both genotypes. Embryos were treated from 18 to 32 hpf to specifically target both Se and DLAV angiogenesis (see Torres-Vázquez et al., 2004; Zygmunt et al., 2011; Childs et al., 2002; Isogai et al., 2003; Zygmunt et al., 2012; Yokota et al., 2015), and then fixed for immunostaining.

Quantification of angiogenesis deficits in the trunk’s arterial tree of WT and mutant zebrafish embryos

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Embryos carrying the Tg(fli1a:EGFP)y1 vascular reporter were fixed at 32 hpf, genotyped using a tail biopsy, immunostained to visualize both the vasculature and the somite boundaries and confocally imaged as described below. Scoring of angiogenesis deficits was performed bilaterally in the ~six somite-long region dorsal to the yolk extension using one of the two following scoring methods and with knowledge of the genotype. Scoring of ‘Se and DLAV truncations’: this scoring method emphasizes truncations found within the dorsal side of the trunk’s arterial tree. This method was used for Figure 2C,E,F–I (see also Figure 2—figure supplement 1 and Supplementary file 3), Figure 3 (see also Figure 3—figure supplement 1 and Supplementary file 4), and Figure 5 (see also Figure 5—figure supplement 1 and Supplementary file 6). The classification of Se and DLAV truncations is based on the relative span of Se and DLAVs along the dorsoventral and anteroposterior axes, respectively, using as reference the following landmarks. The horizontal myoseptum and the actual (or expected) level of the DLAV. Four phenotypic classes are used to define Se and DLAV spans. The three Se-DLAV truncation categories are as follows: maximal (includes both Se that are missing and those that fail to grow dorsally past the horizontal myoseptum); moderate (includes Se that grow dorsally past the horizontal myoseptum but not further than half the distance between the horizontal myoseptum and the level of the DLAV); and minimal (Se that grow dorsally past half the distance between the horizontal myoseptum and the level of the DLAV, but which form an incomplete DLAV). An incomplete DLAV is one that fails to span the distance between the anteriorly and posteriorly flanking ipsilateral somite boundaries. Non-truncated Se-DLAV: full (Se that grow dorsally to the level of the DLAV and that form a complete DLAV). Scoring of ‘Se truncations’: this scoring method emphasizes truncations found within the ventral side of the trunk’s arterial tree. This method was used for Figure 4 (see also Figure 4—figure supplements 23 and Supplementary file 5). The classification of Se truncations is based on the relative span of Se vessels measured along the dorsoventral axis. Briefly, the length of a perpendicular line traced between the actual (or expected) Se sprouting site and the level of the DLAV was assigned a value of 100%. Four phenotypic classes are used to define Se span. Se truncation categories: severe (0–25%, includes missing Se), medium (26–50%), and weak (51–75%). Non-truncated: complete (76–100%). Note that both the Se-DLAV and Se scales used for quantifying angiogenesis deficits are based on the relative span of the vascular structures scored, which eliminates the noise that variations in size between embryos with similar body proportions would otherwise introduce.

Comparisons of angiogenesis deficits between genotypes and treatments

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Significance statistical values were calculated using a two-sided Fisher Exact test with the aid of the SISA web tool (http://www.quantitativeskills.com/sisa/statistics/fiveby2.htm). When performing more than one pairwise comparison (as in Figures 34), significant differences were assigned using a Bonferroni adjustment. This conservative adjustment reduces the likelihood of obtaining false-positive results (type I errors or the rejection of true null hypotheses) when simultaneously applying many statistical tests to a dataset (Statistics, 2016). The Bonferroni adjustment is commonly used to compare drug effects and in angiogenesis studies (see Hamada, 2018; Weichand et al., 2017; Basagiannis et al., 2016).

Morpholino injection and analysis of plxnd1 morphants

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A previously validated splice-blocking plxnd1 morpholino (Gene Tools, LLC) that faithfully phenocopies the vascular defects of the plxnd1fov01b null allele (plxnd13207-3462; see Torres-Vázquez et al., 2004) was used. plxnd13207-3462 morpholino (2.5 ng) was injected into each of the one-cell stage embryos used in the experiment. The morpholino was injected into both WT gipc1skt1(MZ)gipc2skt4(MZ) double mutants carrying the Tg(fli1a:EGFP)y1 reporter. Both un-injected and embryos injected with the plxnd1 morpholino were fixed at 32 hpf to evaluate the trunk vasculature.

Fixing, immunostaining, and mounting of zebrafish embryos

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Embryos were fixed overnight at 4°C in 4% paraformaldehyde (Santa Cruz #sc-281692) dissolved in PBS (1x phosphate buffered saline pH 7.5). Embryos were then washed 6x with PBST (PBS with 0.2% Tween; Sigma #P1379) for 10 min/each and placed in blocking solution (PBST with 1% bovine serum albumin; Sigma #A8022) for 1 h. Embryos were then incubated overnight at 4°C with primary antibodies diluted in blocking solution and then washed with PBST 6x for 10 min/each. Embryos were then incubated overnight at 4°C with fluorescent secondary antibodies diluted in blocking solution for 2 h at RT or overnight at 4°C. Embryos were then washed in PBST 4x for 15 min/each and mounted in an aqueous 0.5% agarose solution before imaging. Primary antibody dilutions used as follows. To label somite boundaries, mouse anti-pFAK Tyr397 (Millipore #05–1140 at 1:1000); to enhance the visualization of EGFP, chicken anti-GFP (Invitrogen #A10262 at 1:1000); to improve mCherry detection, sheep anti-mCherry (custom-made at 1:1000; a gift from Holger Knaut). The following secondary antibody dilutions were used: Alexa Fluor 488 goat anti-chicken (Invitrogen #A11039 at 1:1000), Cy3 donkey anti-sheep (Jackson ImmunoResearch #713-165-147 at 1:1000), and Alexa Fluor 647 donkey anti-mouse (Invitrogen #A31571 at 1:1000).

Confocal imaging of zebrafish embryos

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Confocal images of fixed immunostained embryos were taken with a Leica TCS SP5 microscope using a water dipping 40x lens (0.8 NA). The 488, 561, and 647 nm laser lines were used. Images were processed with FIJI (https://imagej.net/Fiji).

Genotyping of gipc and plxnd1 mutants and Tg(fli1a:GAL4FF)ubs4 transgenics

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Genotyping was done using PCR products amplified from gDNA (see Supplementary file 1 for primers), with plxnd1fov01b genotyping as in Zygmunt et al. (2011). All PCR products were treated with ExoSAP-IT (ThermoFisher Scientific; #78201.1 ML) before sequencing. PCR conditions included BioReady Taq DNA Polymerase (Bulldog Bio #BSAX050), used according to the manufacturer’s recommendations. Abbreviations. Size: length of the PCR amplicon from the WT (wild-type) or M (mutant) allele. Notes: N-PCR (nested PCR). Genotyping methods: AGE (agarose gel electrophoresis), HMA (heteroduplex mobility assay (Ota et al., 2013), and Sanger sequencing (sequencing; performed by GENEWIZ). Detailed protocols are available upon request.

AllelePCR conditions and commentsGenotping methods
Primer pair
(ID#s)
Align
(ToC)
Extend
(secs)
CyclesSize in bp
(WT/M)
NotesAGEHMASequencing
Primer ID#
gipc1skt112891–12889622534372/368N-PCR
(if fixed)
n/aFor hetz12891
13655–13653641940300/296n/aFor hetz13653
gipc1 skt212891–12889573034372/379n/an/aFor hetz12891
gipc2 skt313667–13422601435218/216n/an/an/a13421
gipc2 skt413667–13422601435218/172n/aYesn/a13421
gipc3 skt513510–1351267.72538377/337n/aYesn/a13512
plxnd1skt611844–1184565.93034442/438N-PCRn/an/an/a
12532–1184557.73034262/258n/an/a12532
plxnd1fov01b13498–10167631638519/519N-PCRn/an/a13646
13646–1364757.51138395/395
Tg(fli1a:
GAL4FF)ubs4
11730–1173157.54535402Only Tg presenceYesn/an/a

Zebrafish lines and maintenance

Zebrafish mutant alleles

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The plxnd1 (plxnd1skt6), gipc1 (gipc1skt1 and gipc1skt2), gipc2 (gipc2skt3 and gipc2skt4), and gipc3 (gipc3skt5) alleles were made via genome editing for this study (above) and are described here at the genomic DNA (Supplementary file 1) and protein (Figure 2B, Figure 4—figure supplement 1 and Supplementary file 8) levels. The null plxnd1fov01b allele is described in Torres-Vázquez et al. (2004). All gipc alleles are predicted nulls encoding small truncated proteins lacking all three domains based on the sequence of the targeted genomic regions (the corresponding cDNAs were not sequenced). The potential instability of mutant gipc transcripts and the occurrence of transcriptional adaptation in gipc mutants were left untested (see El-Brolosy and Stainier, 2017).

Zebrafish transgenic lines

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The red fluorescent reporters Tg(kdrl:HsHRAS-mCherry)s896 and Tg(flt1:nls-mCherry)skt-1 were used to visualize, respectively, endothelial cell membranes (Chi et al., 2008) and arterial endothelial cell nuclei (this study; see also Bussmann et al., 2010). The green fluorescent cytosolic reporter Tg(fli1a:EGFP)y1 was used to label the endothelium (Lawson and Weinstein, 2002). The Tg(fli1a:GAL4FF)ubs4 endothelial GAL4 line (Zygmunt et al., 2011) was used to drive expression of UAS constructs and was genotyped via PCR (see Supplementary file 1 and the Materials and methods section ‘Genotyping of gipc and plxnd1 mutants and Tg(fli1a:GAL4FF)ubs4 transgenics’).

Ethics statement 

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Zebrafish embryos and adults were kept and handled using standard laboratory conditions at New York University and under IACUC-app roved animal protocols (#151202–01 and #170103–01).

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Decision letter

  1. Deborah Yelon
    Reviewing Editor; University of California, San Diego, United States
  2. Marianne E Bronner
    Senior Editor; California Institute of Technology, United States

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

Thank you for sending your article entitled "GIPC proteins pattern the vasculature by limiting the antiangiogenic activity of the Semaphorin receptor PlexinD1" for peer review at eLife. Your article has been evaluated by three peer reviewers, and the evaluation is being overseen by a Reviewing Editor and Marianne Bronner as the Senior Editor.

The reviewers have discussed the reviews with one another, and their discussion has raised a number of issues (elaborated below) that would need to be addressed through revisions to your manuscript. Since some of these revisions would likely require you to perform new experiments, we ask that you respond to this letter within the next two to three weeks with a specific action plan and timetable for the completion of the additional work. Please bear in mind that the standard timeline for a revision for eLife is only 2-3 months. The Reviewing Editor and reviewers will then consider your plans, assess the feasibility of an effective response to their concerns, and offer a binding recommendation (such as an official invitation to revise your manuscript for eLife). Alternatively, if you would prefer to submit your manuscript elsewhere, rather than revising it for eLife, please let us know.

Summary:

In this interesting study, Carretero-Ortega and colleagues identify the GIPC endocytic adaptors as the first negative intracellular regulators of PlxnD1 signaling, expanding our knowledge of the network controlling early arterial angiogenesis. The authors first use biochemical assays with truncated forms of PlxnD1 to identify a GIPC-Binding Motif (GBM) in the cytosolic tail of the protein, which they find to be essential for GIPC binding. In addition, they confirm the involvement of the PDZ and GH1 domains of GIPCs in the formation of the PlxnD1-GIPC1 complex. In order to characterize the role of GIPCs on the vascular activity of PlxnD1, the authors compare the effects of focused, endothelium-specific overexpression of PlxnD1 and truncated forms of PlxnD1 that do not bind GIPCs. Finally, they use CRISPR/Cas9-mediated genome editing to generate the first gipc1, gipc2, and gipc3 zebrafish mutants, and they analyze their vascular phenotypes and make a case for their angiogenic defects being caused by increased PlxnD1 signaling. Finally, they show that GIPC depletion increases Sema3E-induced PlxnD1-dependent responses in HUVEC cells. Overall, the studies are straightforward, and the data have the potential to have an important impact on our understanding of angiogenesis signaling networks. However, there are several issues, described below, that need to be addressed more thoroughly in order to solidify support for the authors' conclusions.

Essential revisions:

1) This manuscript nicely uses an in vivo assay (Figure 2) to test whether GIPC-binding deficient mutants of PlxnD1 lose activity for vessel patterning. Surprisingly, both wild type PlxnD1 and PlxnD1ΔGBM are equally effective at restoring vessel patterning in PlxnD1 mutants. The authors interpret this to suggest that GIPCs are not required for PlxnD1 activity. The authors then suggest that either 'PlxnD1ΔGBM will be more potent than PlxnD1WT' or that 'both PlxnD1's activity and potency will be independent of GIPC-binding'. These two hypotheses are poorly worded as it is not clear what 'potency' and 'activity' refer to. By potency, do the authors mean that PlxnD1ΔGBM is insensitive to inhibition and therefore more active? What phenotype would a 'more active' PlxnD1 mutant have? For the second hypothesis, do the authors mean that while GIPC binds PlxnD1, it has no influence on activity? Also, and importantly, when comparing the influence of PlxnD1-WT and PlxnD1ΔGBM on "angiogenic potential", the authors speculate on the meaning of an observed trend that is not statistically significant. Could the authors increase the sample size for these observations? Would they then find evidence for a statistically significant difference? Finally, the conclusion that PlxnD1 signaling acts to limit endothelial proliferation should be confirmed through direct analysis of proliferation (e.g. by EdU labeling).

2) The intersegmental vessel truncation observed in zygotic gipc1 or gipc2 single mutants is not statistically significant. Therefore, rather than illustrating these so prominently in Figure 3, it might be more worthwhile to focus on the significant effects observed in double mutants and MZ mutants, which show a phenotype that has low penetrance. The compression of the graph's X-axis in Figure 3B to emphasize this phenotype is unusual and could be misleading. At least 80% of the embryo SeV are normal, and the way that the data are presented may give the impression that the phenotype is stronger than it actually is. It would also be helpful to provide statistical analysis in Figure 4—figure supplement 3 and to adjust the X-axis in Figure 4—figure supplement 3B. Since these results are key to the authors' conclusions, significance and robustness need to be clearly shown, particularly as the results in this manuscript do not agree with those in a published article on the same topic in mouse (Burk et al., 2017) It should be straightforward (in principle) for the authors to increase their n and procure a more robust result.

3) Even given its low penetrance, the gipc mutant phenotype is interesting as it is very different from the mouse GIPC knockout vascular phenotype that shows a PlxnD1-like phenotype in equivalent vessels (as opposed to fewer vessels in fish gipc mutants). Burk et al., 2017, claim (in their Figure 10D) that, in mouse embryos, loss of Gipc1 leads to ectopic ISV sprouting, the same phenotype observed in PlexinD1 mutants (not shown) and in Gipc1;PlexinD1 double heterozygote embryos. Although this is illustrated only briefly and not very convincingly by Burk et al., this published result seems to be in strong contrast to the gipc mutant phenotypes illustrated in Figure 3 of this manuscript. Why are fish and mouse null Gipc mutants different? An insightful discussion is needed, and the authors do not provide much material in this regard. It would also be helpful if the previous name of Gipc1 (Synectin) were to be mentioned in this manuscript.

4) The claim of Figure 4, that the angiogenesis defects of gipc mutants are partially due to increased PlxnD1 signaling, is based on the analysis of zygotic gipc1;plxnD1 double mutants. However, as gipc1 zygotic mutants had no statistically significant Se defect, this experiment might not be suitable for supporting the authors' claim. Notably, the double mutants are analyzed only qualitatively here, and no quantitation is provided. Importantly, if GIPC acts as a repressor of PlxnD1 activity, one would expect a rescue in the double mutant. Therefore, it is possible to argue that the authors have shown that, even though GIPC binds PlxnD1, its activity is not required to influence PlxnD1 signaling. However, the authors do not reach this conclusion. Perhaps PlxnD1 is epistatic to GIPC (i.e. acts downstream of GIPC). Even though GIPC binds PlxnD1, it could act upstream and/or in a different pathway to control angiogenesis of the SeV (such as in the neuropilin pathway or other pathways where GIPC is known to act). Finally, the fact that GIPC and PlxnD1 mouse double mutants (Burk et al.), have identical and not opposite phenotypes, unlike in the fish, is not discussed in any depth.

5) The data presented in Figure 6 clearly demonstrate that loss of GIPC potentiates loss of Sema3E-induced collapse. The authors conclude that this is PlxnD1-dependent because siRNA against PlxnD1 fails to induce the collapse. However, double knockdown of GIPC and PlxnD1, followed by Sema3E addition, is necessary to make this conclusion, and this experiment was not performed. Without double knockdown data, there is no evidence that PlxnD1, GIPC and Sema3E interact in collapse, as this not been tested in the same cells. The same argument can be made for the ERK data – again, PlxnD1-GIPC double knockdown, with and without Sema3E, in the same cells is a necessary experiment to support the conclusion. Alternatively, could the authors corroborate the ERK activation results (Figure 6) in zebrafish? Since all the zebrafish mutants are available, the use of HUVECs seems less relevant. ERK antibody staining has been extensively described in zebrafish, and it could be used to investigate ERK activation in the arteries of gipc mutants, as well as in gipc;plexind1 double mutants. This type of analysis has the potential to strengthen the authors' claims.

6) The authors seem to assume that GIPCs can mediate PlxnD1 activity only (or primarily) via direct binding through the identified domains. Although this was the way that the authors initially identified GIPCs (via 2-hybrid screening), can they truly rule out other possibilities? Additionally, the authors do not seem to consider other potential roles of GIPCs that are PlxnD1-independent. Although they state in their Results section that "the angiogenesis deficits of gipc mutants are, at least partially, due to increased PlxnD1 signaling", they do not assess or discuss additional possibilities. One relatively easy experiment that the authors could perform in order to address these questions would be to overexpress plexind1 (WT and truncated forms) in endothelial cells of gipc single and double mutants. Alternatively, the authors could consider the possible relevance of the known interactions of GIPC with Neuropilin. While Neuropilin can act as a Plexin coreceptor, it can also act in a Plexin-independent manner. This is of relevance especially in experiments analyzing growth cone collapses or other effects on morphology, where Neuropilin has been shown to act independently of Plexin. Can the authors distinguish the negative regulation of Plexin from a potential positive regulation of Neuropilin? This is of course hampered by the caveat that there are no reliable Neuropilin reagents in zebrafish available as morpholino and mutant data only partially match. Maybe different downstream effectors could be affected pharmacologically? At a minimum, it would be helpful for the authors to provide more discussion of alternate possibilities that could explain the observed ways in which GIPC function affects PlxnD1 activity.

[Editors' note: the authors’ plan for revisions was approved and the authors made a formal revised submission.]

Thank you for submitting your revised manuscript entitled "GIPC proteins pattern the vasculature by limiting the antiangiogenic signaling of the Semaphorin receptor Plexind1" for consideration by eLife. Your revised submission has been reviewed by two out of the three original peer reviewers, and the evaluation has been overseen by a Reviewing Editor and a Senior Editor. The reviewers have opted to remain anonymous.

Our decision has been reached after a process of initial independent review followed by extensive discussion between the reviewers and consultation with the editors. The initial independent reviews are reprinted at the bottom of this email, and the summary of the reviewers' discussion is provided in the next paragraph. Based on the overall consensus emerging from this discussion, we regret to inform you that your work will not be considered further for publication in eLife.

To summarize the reviewers' discussion: Both reviewers appreciate the improvements made during the revision of this manuscript, including the changes to the text and the addition of new data. However, both reviewers ultimately concluded that the revised material did not provide sufficient evidence in support of the primary conclusions of the manuscript. While much of the data in the paper is robust, the in vivo interaction of PlexinD1 and GIPC has not been robustly demonstrated via analysis of the PlexinD1 skt6 mutant. Specifically, reviewers were dissatisfied with the degree of statistical significance present in the data that was meant to demonstrate the effect of the skt6 mutation on angiogenesis, as well as in the data that was meant to demonstrate the rescue of the null allele by skt6. The lack of robust statistical significance here throws the conclusion that GIPCs are direct, negative modulators of PlexinD1 signaling into question. Among other concerns (see reviews below), reviewers were uncertain about the conclusions reached from the SU5416 treatments: they wondered whether the Bonferroni type adjustment is appropriate in this case and whether developmental delay could confound the interpretation of these data. Reviewers agreed that the strongest elements of the manuscript are the GIPC mutant data, both alone and when bred to PlexinD1 mutants. Also, the in vitro experiments looking at pERK after Sema3e treatment in GIPC/ PlexinD1 knockdown/knockouts are convincing. However, these experiments show association and not binding/mechanism. While there is clearly an association of PlexinD1 and GIPC and pERK, it remains unclear whether this interaction requires direct binding.

Reviewer #1:

The authors claim that the plxnd1 skt6 allele encodes a hypomorphic Plxnd1 receptor which induces angiogenesis deficits in homozygous mutant embryos.

However in the Figure 2 legend they clearly state that there is no statistical difference in phenotype occurrence between Wt and mutant embryos.

In the Results section, the authors claim that there is no statistical difference in Se phenotypes in Wt and plxnd1 skt6 siblings (without clarifying whether this denotes heterozygous or homozygous mutant embryos).

If the observed effect is not statistically significant, it cannot be used to claim a angiogenesis defect with low penetrance.

Additionally I disapprove of the quantification, as the authors should have analyzed always the exact same region and number of Se-DLAVs per embryo and not a varying average of 10.5 per embryo.

For me it is not clearly stated whether the less than 5% of affected Se-DLAVs are of all counted Se-DLAVs or are number of embryos with affected Se-DLAVs. I assume the first, as it would be difficult to get to 5% of 12 embryos. However it would be important to know how many of the embryos do show vessel truncations in one or two vessels.

The total number of only 12 embryos analyzed for each genotype is also really low.

The experiments addressing hypersensitivity to Su5416 have theoretically been done under the same conditions as when assessing the phenotype of homozygous plxnd1skt6 mutants.

Why do the authors suddenly see dramatic difference in angiogenesis defects now in 40% of the vessels in 0.1% DMSO? Why does the DMSO also affect WT development? How are changes to more than 80% affected vessels only achieving one star significance? another explanation would be that the embryos have not really reached 32hpf but are a little developmentally delayed. The experimental procedure does not state whether and how the embryos were dechorionated for treatment from 18hpf on.

SU5416 has been previously used in zebrafish at concentrations of 1 µM and was shown to completely block Vegf mediated ISV sprouting. Here the authors use 0.2µM and show an effect on Wt ISV migration, which is slightly enhanced in the mutants, but there is no evidence that this does not just reflect a developmental delay caused by the chemical treatment. The authors do need to control for this alternative hypothesis. Also the embryo in Figure 3B seems to be "larger" or at higher magnification than the others, as the distance from the PCV to the top of the DLAV is larger than in the other three images. also in the supplement it is evident that already 90% of the Wt embryos are affected by the SU5416 treatment. The experimental conditions should have been chosen so that the Wt is not affected at all.

In sum for essential revision1B we requested to show a statistically significant effect on angiogenesis. Unfortunately, the data presented are still not convincing.

Likewise, why is there such a big variance in phenotype distribution when scoring gipc1 skt1(MZ) mutant embryos? Compare Figure 4B with Figure 5C.

It would also be helpful if the authors would indicate e.g. in Figure 2 or 3 and clearly define in the text what constitutes a maximal, a moderate or a minimal Se-truncation.

The authors claim that Gipc binding limits the antiangiogenic signaling of the PlxnD1 receptor. However, there are too many assumptions and too little data for such a statement.

For one: what is the antiangiogenic Plxnd1 signaling: where is that shown just based on a poorly controlled hypersensitivity to VegfR2 receptor inhibition, which was done for a very long time period? or based on the proposed hypomorphic identity of the plxnd1 skt6 allele, which as mentioned above is not clearly a significant phenotype.

The analysis of the gipc mutants is worthwhile, however there is too little evidence supporting the Gipc/Plexin interaction on a phenotypic level, with regards to their collaborating role in angiogenesis.

An alternative explanation for the partial reduction in phenotype would be that while in gipc1 skt1 Mz embryos gipc2 and gipc3 can still bind to Wt Plexin and therefore are less available for interaction with another receptor (like Nrp1) in the plxnd1 fov01b heterozygous less plexin molecules are available to limit the gipc signaling pool. This would also explain the reduction in phenotype, and be based on a Plexin/gipc interaction, but say nothing about the actual signaling cascades required for generating an angiogenesis phenotype (see our comment regarding essential revision 4B).

As the interaction studies between Gipc1/2 and plxnd1 are only analyzed qualitatively, meaning that you find massive ectopic sprouting, how were you expecting to see a double phenotype in case of no interaction? the gipc1skt1(Mz); gipc2 skt3/skt4 double phenotype did affect less than 20% of the embryos, only about 10% were affected medium to severely. that would mean that in case of NO interaction you would have found 10% of the Se's being truncated, but still having massive ectopic filopodia and sprouts. Given that there would be additional ectopic sprouts coming from the neighboring Se's, I doubt that this would have been detectable. I therefore believe that the experiment is not suitable to disprove alternative hypothesis…

In sum there is not enough evidence supporting a model whereby Gipc limits Plexind1 antiangiogenic signaling, as Gipc could still act on angiogenesis in a Plexin independent manner, eg via neuropilin (see essential revisions 6!). I don't think that the authors have sufficiently experimentally addressed this question. However, this affects the core of the authors claims by directly affecting the functionality of the Plexin/Gipc interactions in angiogenesis.

Reviewer #2:

The authors have cleared up misconceptions created by language and elegantly used hypotheses to set up outcomes of their experiments and what this means. This is a large improvement over the previous version of the manuscript.

The new skt6 allele of plexinD1 was used in very clear experiments to show it is functional and hypermorphic. There is also a plexinD1 null HUVEC line generated as well for the cell culture experiments.

Graphs have been improved. Numbers increased and emphasized.

Improved discussion of Burk results in context of the results of this manuscript.

Overall, there is a huge improvement to the manuscript. Many figures are edited or completely new and there is new data to answer my major points.

[Editors’ note: what now follows is the decision letter after the authors submitted for further consideration.]

Thank you for submitting your rebuttal letter regarding the manuscript entitled "GIPC proteins pattern the vasculature by limiting the antiangiogenic signaling of the Semaphorin receptor Plexind1" for consideration by eLife. I have read your rebuttal letter and conferred with the Reviewing Editor. We both agree that it would be appropriate for you to submit a revised version of the manuscript that addresses the comments of the reviewers.

We would like to clarify that both reviewer 1 and 2 were in good agreement during the second round of review. Therefore, we urge you to address their comments to the best of your ability. However, given your concerns we will not return the revised manuscripts to either reviewer, but will instead ask previous reviewer 3 (who did not respond to our previous request to re-review), plus a new reviewer. Of course we cannot guarantee the outcome but will do all that we can to be fair.

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

Thank you for resubmitting your work entitled "GIPC proteins pattern the vasculature by limiting the antiangiogenic signaling of the Semaphorin receptor Plexind1" for further consideration at eLife. Your revised article has been favorably evaluated by Marianne Bronner (Senior Editor), a Reviewing Editor, and two reviewers. One of these reviewers (Reviewer #3) had reviewed the original version of this manuscript, and the other (Reviewer #4) was a newly recruited reviewer.

As you can see from the reviewers' comments (copied below), the reviewers feel that the manuscript has been improved, but both agree that there are some remaining issues that need to be addressed before acceptance. The reviewers have discussed these issues with each other after submitting their reviews, and they agree that it is particularly important for the manuscript to be thoroughly edited, with the goal of creating a more succinct and streamlined experience for the reader. The reviewers also agree that you should make a series of specific adjustments to the text and figures in an effort to clarify or modify the message, in accordance with each of the individual points that have been raised in the reviews below. Finally, regarding reviewer #4's suggestion that you perform an additional co-IP experiment (see comment #1 from reviewer #4 below), the reviewers agree that this is not essential. If you have the data at hand, please do include it, but it will also be acceptable to simply make changes in the text describing the biochemical effects of this mutation. In that case, the idea that this mutant version of the zebrafish protein loses its ability to interact with Gipc proteins should be presented as speculation, not as fact.

Reviewer #3:

After carefully reading the full history of this manuscript, I tend to agree with the previous assessment of reviewer #2.

I believe the revised version provided by the authors successfully addressed the essential concerns raised during the first round of review. Moreover, I understand that the data added in the revised version was approved in advance in the context of the revision plan.

While I agree with reviewer #1, that some issues of statistical significance were not fully solved, the authors did a good job not only in providing extensive explanations for the observed results, but also by listing alternative hypotheses. In my view, the faithful presentation of the results, and the efforts made to explain them, are valuable, despite of the phenotypes themselves being not hugely penetrant.

As agreed by all 3 reviewers, the findings of this manuscript are important and add to our understanding of the mechanisms underlying Plexind1 signalling. Since most of the initial concerns were addressed properly, and the findings are carefully interpreted, I tend to support publication.

I do think however that the title/impact statement/conclusions could be toned down a bit to reflect more accurately the main findings.

Reviewer #4:

Previous work by Dr. Torres-Vazquez demonstrated a requirement for Semaphorin3E/PlexinD1 signaling in constraining segmental vessel sprouting and migration in zebrafish embryos, and two papers (Burk et al., 2017; Shang et al., 2017) more recently demonstrated an interaction between the C-terminus of PlexinD1 and the GIPC family of endocytic adaptor proteins. The goal here is to determine how GIPC proteins modulate PlexinD1 function in endothelial cells (ECs). Strengths of this work include the use of both cell culture and zebrafish models; the generation of multiple new zebrafish mutants; genetic interaction studies in both models; and clear data presentation. The authors conclude that GIPC functions as a negative regulator of PlexinD1, serving to dampen its antiangiogenic activity. In aggregate, the presented data strongly support this conclusion, which contrasts with findings in Burk et al. However, a major problem is that an assumption is made regarding the biochemical consequences of a key plxnd1 mutant without direct presentation of experimental data to support this assumption.

1) The CRISPR/Cas9-generated plxnd1skt6 mutant harbors a deletion that removes the final 5 C-terminal amino acids and extends the protein with 31 amino acids. The authors assume that this mutant will abrogate interaction with Gipc proteins because 1) deletion of these 5 amino acids in human PLXND1 decreases interaction with human GIPC1 (as shown in Figure 1); and 2) addition of a single amino acid to the C-terminus of other PDZ-binding proteins abrogates their interaction with PDZ proteins (cite published work). However, deletion of these 5 amino acids in the human protein decreases but does not abolish PLXND1/GIPC1 interaction (Figure 1), and the authors do not test whether the extended Plxnd1 protein generated in these zebrafish mutants does indeed fail to bind Gipc proteins. As the conclusions are based entirely on this important assumption and this assumption is repeatedly stated as fact (for example, subsection “Homozygous plxnd1skt6 mutants are hypersensitive to the antiangiogenic drug SU5416”; Figure 2 title), this should be tested with at least one recombinant Gipc protein as in Figure 1.

2) In Figure 2—figure supplement 2, the authors show that mosaic EC-specific expression of wild type or C-terminally truncated PlexinD1 (missing the 9 C-terminal amino acids that comprise the entire GIPC binding motif) can similarly rescue segmental vessel development in plxnd1fov01b null mutants. They interpret this to mean that Gipc1 binding is not required for antiangiogenic activity of Plxnd1 (truncated version is fully functional) and use these data to support their claim that the plxnd1skt6 allele is hypermorphic due to inhibited interaction with Gipc1. However, the 9 amino acid deletion is not truly comparable to the 5 amino acid deletion/31 amino acid insertion in the mutant. If the authors can show that the skt6 protein is deficient in Gipc1 binding (Comment #1), this would minimize this concern.

https://doi.org/10.7554/eLife.30454.030

Author response

Essential revisions:

1A) This manuscript nicely uses an in vivo assay (Figure 2) to test whether GIPC-binding deficient mutants of PlxnD1 lose activity for vessel patterning. Surprisingly, both wild type PlxnD1 and PlxnD1ΔGBM are equally effective at restoring vessel patterning in PlxnD1 mutants. The authors interpret this to suggest that GIPCs are not required for PlxnD1 activity. The authors then suggest that either 'PlxnD1ΔGBM will be more potent than PlxnD1WT' or that 'both PlxnD1's activity and potency will be independent of GIPC-binding'. These two hypotheses are poorly worded as it is not clear what 'potency' and 'activity' refer to. By potency, do the authors mean that PlxnD1ΔGBM is insensitive to inhibition and therefore more active? What phenotype would a 'more active' PlxnD1 mutant have? For the second hypothesis, do the authors mean that while GIPC binds PlxnD1, it has no influence on activity?

We revised the text to highlight that PlxnD1 is antiangiogenic, avoiding the use of the terms “activity” and “potency.” As agreed, we thus instead speak of “Plxnd1 signaling”. Given the proposed plan for addressing essential revision 1B (below), we stated how each of the three competing hypotheses makes different forecasts regarding the nature of the plxnd1skt6allele (predicted to be deficient in GIPC-binding and made via genome editing). These hypotheses are assessed by the ability of the plxnd1skt6 allele to complement the null plxnd1fov01b mutation and by the vascular phenotype and sensitivity to the antiangiogenic compound SU5416 of plxnd1skt6 homozygotes.

In the revised manuscript, this information is found in the Results’ section entitled “Zebrafish mutants in which plxnd1 function is provided solely by a receptor deficient in GIPC binding display angiogenesis deficits with low penetrance and are hypersensitive to the antiangiogenic drug SU5416,” and in Figures 2-3, Figure 2—figure supplement 1 and Figure 3—figure supplement 1. Finally, the plxnd1skt6 allele designation follows ZFIN-approved nomenclature.

1B) Also, and importantly, when comparing the influence of PlxnD1-WT and PlxnD1ΔGBM on "angiogenic potential", the authors speculate on the meaning of an observed trend that is not statistically significant. Could the authors increase the sample size for these observations? Would they then find evidence for a statistically significant difference?

Given that increasing the sample size for the transgenic rescue experiment was unlikely to resolve the issue, we did not pursue this direction. Briefly, few cells express the exogenous receptor. Hence, the assay is too noisy which makes the evaluation of differences in angiogenic potential an impractical proposition.

Instead, as proposed before, we characterized the novel zebrafish plxnd1skt6mutant allele that we made via genome editing (formerly referred as plxnd1ΔGBM allele in our prior reply). This allele is predicted to yield a Plxnd1 protein in which a scrambled amino-acid sequence replaces the PBM. Using this allele, we performed the following three experiments.

1) Performed complementation assays between our novel plxnd1skt6 allele and the null mutant plxnd1fov01b to define the vascular phenotypes of the trans-heterozygotes (plxnd1skt6 complements plxnd1fov01b).

2) Determined the phenotype of plxnd1skt6 homozygotes (they exhibit angiogenesis deficits at low frequency).

3) To further characterize both the functionality and nature of the plxnd1skt6 allele we decided, for simplicity, to compare the sensitivity of WT embryos and plxnd1skt6 mutants to the antiangiogenic compound SU5416, rather than performing the optional cell transplantation experiment outlined previously (plxnd1skt6 mutants are hypersensitive to SU5416).

These observations led us to conclude that the receptor deficient in GIPC binding encoded by the plxnd1skt6 allele is both functional and hypermorphic. In the revised manuscript, these findings and conclusions are described in the Results’ section entitled “Zebrafish mutants in which plxnd1 function is provided solely by a receptor deficient in GIPC binding display angiogenesis deficits with low penetrance and are hypersensitive to the antiangiogenic drug SU5416” and in Figures 2-3, Figure 2—figure supplement 1 and Figure 3—figure supplement 1.

1C) Finally, the conclusion that PlxnD1 signaling acts to limit endothelial proliferation should be confirmed through direct analysis of proliferation (e.g. by EdU labeling).

This comment relates to the transgenic rescue experiment now found in Figure 2—figure supplement 2, and directly related to the graph in panel K (now inexistent). We believe that the reviewer’s comment arose from the potential ambiguity of the original text, which could be read as overstating one of our conclusions. Since prior studies already established a connection between Plxnd1 signaling and proliferation, we indicated in our prior reply that the EdU labeling experiment was unnecessary. More importantly, the relevant panel K is no longer part of the revised Figure 2—figure supplement 2, since the plxnd1skt6 data supports better the notion that Plxnd1 proteins deficient in GIPC binding are hypermorphic. Accordingly, we revised the Results’ section entitled “Zebrafish mutants in which plxnd1 function is provided solely by a receptor deficient in GIPC binding display angiogenesis deficits with low penetrance and are hypersensitive to the antiangiogenic drug SU5416” and the legend of Figure 2—figure supplement 2.

2A) The intersegmental vessel truncation observed in zygotic gipc1 or gipc2 single mutants is not statistically significant. Therefore, rather than illustrating these so prominently in Figure 3, it might be more worthwhile to focus on the significant effects observed in double mutants and MZ mutants, which show a phenotype that has low penetrance. The compression of the graph's X-axis in Figure 3B to emphasize this phenotype is unusual and could be misleading. At least 80% of the embryo SeV are normal, and the way that the data are presented may give the impression that the phenotype is stronger than it actually is.

We modified Figure 3A (now Figure 4A) to show only the images of the vascular phenotypes of the four recommended genotypes. Accordingly, we presented the phenotypes of gipc1 and gipc2 single zygotic mutants as Supplementary Data (Figure 4—figure supplement 2).

As recommended for Figure 3B (now Figure 4), we adjusted the X-axis to 100% and showed the “No truncation” phenotypic category as a black bar. We made similar modifications to other graphs related to vascular phenotypes (see Figures 2I, 3E, 4B, 5, Figure 2—figure supplement 1, Figure 3—figure supplement 1, Figure 4—figure supplement 3 and Figure 5—figure supplement 1). We further improved all phenotypic graphs by displaying significant differences using the standard brackets and asterisks (rather than with text; see Figures 3E, 4B, 5C, Figure 3—figure supplement 1, Figure 4—figure supplement 3, and Figure 5—figure supplement 1B).

We note that, for simplicity and readability, the graphs of vascular phenotypes show one kind of significant difference. Supplementary Files 3-6 show the remaining significant differences.

2B) It would also be helpful to provide statistical analysis in Figure S2 and to adjust the X-axis in Figure S2B.

We calculated significant differences and adjusted the X-axis for both panels in now Figure 4—figure supplement 3. We applied similar changes to Figures 3E, 4B, 5C, Figure 3—figure supplement 1, Figure 4—figure supplement 3 and Figure 5—figure supplement 1B. We also now show the “No truncation” phenotypic category as a black bar in Figure 4—figure supplement 3B. We applied similar changes to Figures 2I, 3E, 4B, 5, Figure 2—figure supplement 1, Figure 3—figure supplement 1, Figure 4—figure supplement 3 and Figure 5—figure supplement 1.

2C) Since these results are key to the authors' conclusions, significance and robustness need to be clearly shown, particularly as the results in this manuscript do not agree with those in a published article on the same topic in mouse (Burk et al., 2017) It should be straightforward (in principle) for the authors to increase their n and procure a more robust result.

We agree with the reviewers that it is important to highlight the solid significance and robustness of our results, which are based on the analysis of one hundred twelve embryos (sixty-nine embryos without maternal and zygotic gipc1 activity, including nineteen animals devoid of zygotic gipc2 activity; plus twenty-four embryos lacking zygotic gipc1 activity, including thirteen without gipc2 activity). Hence and as stated before, the analysis of more embryos is unwarranted. We highlight that we found that gipc mutants display recessive arterial angiogenic deficits of partial penetrance. Moreover, none of the gipc mutants analyzed phenocopied, even partially, the hyperangiogenic and mispatterned vasculature of plxnd1fov01b nulls.

Accordingly, this information is mentioned in detail in the revised manuscript as follows. First, in the Results’ sections entitled “Zebrafish gipc mutants display angiogenesis deficits in the trunk’s arterial tree,” “Reducing Plxnd1 signaling ameliorates the angiogenic deficits of maternal-zygotic gipc1 mutants”, and “Double maternal-zygotic (MZ) gipc mutants display excessive angiogenesis in the absence of Plxnd1 signaling.” Second, in the corresponding figures: Figures 4-6, Figure 4—figure supplement 2, Figure 4—figure supplement 3 and Figure 5—figure supplement 1). Finally, in Supplementary Files 5-6. We note that the incorporation of essential revisions 2A-2B (now Figure 4—figure supplement 3; above) further highlights the significant differences in penetrance and expressivity between gipc mutants.

3A) Even given its low penetrance, the gipc mutant phenotype is interesting as it is very different from the mouse GIPC knockout vascular phenotype that shows a PlxnD1-like phenotype in equivalent vessels (as opposed to fewer vessels in fish gipc mutants). Burk et al., 2017, claim (in their Figure 10D) that, in mouse embryos, loss of Gipc1 leads to ectopic ISV sprouting, the same phenotype observed in PlexinD1 mutants (not shown) and in Gipc1;PlexinD1 double heterozygote embryos. Although this is illustrated only briefly and not very convincingly by Burk et al., this published result seems to be in strong contrast to the gipc mutant phenotypes illustrated in Figure 3 of this manuscript. Why are fish and mouse null Gipc mutants different? An insightful discussion is needed, and the authors do not provide much material in this regard.

We agree with the reviewers. Accordingly, we provide further details in two sections of the revision. First, we do this briefly in the last paragraph of the Results’ section entitled “Zebrafish gipc mutants display angiogenesis deficits in the trunk’s arterial tree.” Second, we address in great depth the potential reasons behind the discrepancy in the seventh, eighth and ninth paragraphs of the Discussion section. Briefly, we do not favor the possibility that gipc genes play distinct roles in the zebrafish and mice. Instead, we believe that their functions are conserved and list seven points that invite a reinterpretation of the findings by Burk et al. (see also our answer to essential revision 4C).

3B) It would also be helpful if the previous name of Gipc1 (Synectin) were to be mentioned in this manuscript.

The original text highlighted that GIPC1 and Synectin are synonymous. Now for emphasis, we repeatedly mention this equivalency in the revision. Specifically, Synectin is now mentioned six times throughout the main body of the manuscript.

4A) The claim of Figure 4, that the angiogenesis defects of gipc mutants are partially due to increased PlxnD1 signaling, is based on the analysis of zygotic gipc1;plxnD1 double mutants. However, as gipc1 zygotic mutants had no statistically significant Se defect, this experiment might not be suitable for supporting the authors' claim. Notably, the double mutants are analyzed only qualitatively here, and no quantitation is provided.

In the revised manuscript we addressed this point via the following two experiments. First, and using a quantitative phenotypic analysis, we determined that plxnd1 heterozygosity ameliorates the angiogenesis deficits of gipc1skt1(MZ)maternal-zygotic mutants. We describe these findings in the Result’s section entitled “Reducing Plxnd1 signaling ameliorates the angiogenic deficits of maternal-zygotic gipc1 mutants”, Figure 5 and Figure 5—figure supplement 1 (see also Supplementary File 6).

Second, we compared the vascular phenotype of gipc1skt1(MZ); gipc2skt4(MZ)maternal-zygotic double mutants with and without morpholino-induced plxnd1 knockdown at the 32 hpf stage. Importantly, this morpholino faithfully phenocopies the vascular phenotype of plxnd1fov01b null mutants, namely Se vessel over-sprouting and disorganization (see reference #5 in the revision). Since it is impossible to score the abundance and length of Se vessels at 32 hpf, we evaluated the vascular abnormalities qualitatively. We found that gipc1skt1(MZ); gipc2skt4(MZ)maternal-zygotic double mutants injected with the plxnd1 morpholino phenocopy the excessive and disorganized angiogenesis of plxnd1 nulls. We describe these findings in the Results’ section entitled “Double maternal-zygotic (MZ) gipc mutants display excessive angiogenesis in the absence of Plxnd1 signaling” and in Figure 6.

4B) Importantly, if GIPC acts as a repressor of PlxnD1 activity, one would expect a rescue in the double mutant. Therefore, it is possible to argue that the authors have shown that, even though GIPC binds PlxnD1, its activity is not required to influence PlxnD1 signaling. However, the authors do not reach this conclusion. Perhaps PlxnD1 is epistatic to GIPC (i.e. acts downstream of GIPC). Even though GIPC binds PlxnD1, it could act upstream and/or in a different pathway to control angiogenesis of the SeV (such as in the neuropilin pathway or other pathways where GIPC is known to act).

To clarify, the model that GIPCs simultaneously promotes pro-angiogenic VEGF signaling and limits anti-angiogenic PLXND1 signaling predicts that the double gipc mutant/plxnd1 morphant would have a plxnd1 null phenotype. The results of the revised experiment above agree with this prediction, supporting this model. However, if for example, the double gipc mutant/plxnd1 morphant had a phenotype like that of gipc mutants, the model would be invalid. None of these potential results argues against the possibility that GIPC also acts in a PLXND1-independent manner. Please see also our reply to essential revision 4A.

4C) Finally, the fact that GIPC and PlxnD1 mouse double mutants (Burk et al.), have identical and not opposite phenotypes, unlike in the fish, is not discussed in any depth.

We rectified this omission in the revised manuscript. Please see our answer to essential revision 3A for further details.

5A) The data presented in Figure 6 clearly demonstrate that loss of GIPC potentiates loss of Sema3E-induced collapse. The authors conclude that this is PlxnD1-dependent because siRNA against PlxnD1 fails to induce the collapse. However, double knockdown of GIPC and PlxnD1, followed by Sema3E addition, is necessary to make this conclusion, and this experiment was not performed. Without double knockdown data, there is no evidence that PlxnD1, GIPC and Sema3E interact in collapse, as this not been tested in the same cells. The same argument can be made for the ERK data – again, PlxnD1-GIPC double knockdown, with and without Sema3E, in the same cells is a necessary experiment to support the conclusion.

As suggested and previously agreed to, we performed one combined GIPC/PLXND1 loss of function experiment (using pERK abundance as a readout of PLXND1 activity; see Figure 7 and Figure 7—figure supplements 1-2 in the revision). Importantly, the results of this experiment indicate that loss of GIPC potentiates the SEMA3E-induced/PLXND1-dependent decrease in pERK abundance. The cell collapse data presented in the original Figure 6 is now found in Figure 7—figure supplement 3 (see also Figure 7—figure supplement 4). We describe the pERK and cell collapse experiments in the revised Results’ section entitled “GIPC depletion potentiates SEMA3E-induced, PLXND1-dependent responses in HUVECs”.

5B) Alternatively, could the authors corroborate the ERK activation results (Figure 6) in zebrafish? Since all the zebrafish mutants are available, the use of HUVECs seems less relevant. ERK antibody staining has been extensively described in zebrafish, and it could be used to investigate ERK activation in the arteries of gipc mutants, as well as in gipc;plexind1 double mutants. This type of analysis has the potential to strengthen the authors' claims.

We did not perform the suggested experiment with fish since we believe that the cell culture model is the best option to answer this question. Briefly, prior studies indicate that VEGF signaling via VEGFR2 and Nrp1 up-regulates pERK levels and that, in this context, GIPC1 is required for maximal VEGF-dependent pERK up-regulation. Also, PLXND1 signaling antagonizes VEGF signaling in part by reducing pERK levels. Importantly, in cell culture, we can activate PLXND1 signaling in the absence of exogenous VEGF stimulation, allowing us to use the relative abundance of pERK as a quantifiable PLXND1-specific signaling output. In contrast, in the embryo, VEGF signaling must be active to enable both endothelial ERK phosphorylation and Se angiogenesis. Moreover, endothelial pERK immunofluorescence is variable within and across WT embryos in our hands. Accordingly, we experimented with using HUVECs. We describe the pERK experiment in the revised Results’ section entitled “GIPC depletion potentiates SEMA3E-induced, PLXND1-dependent responses in HUVECs” and in Figure 7 and Figure 7—figure supplements 1-2. Importantly, the results corroborate the proposed model. Please see our answer to essential revision 5A.

6A) The authors seem to assume that GIPCs can mediate PlxnD1 activity only (or primarily) via direct binding through the identified domains. Although this was the way that the authors initially identified GIPCs (via 2-hybrid screening), can they truly rule out other possibilities?

Of course not. Our results do not rule out the possibility that GIPCs might also influence PLXND1 signaling independently of the formation of GIPC-PLXND1 complexes and we now highlight this scenario in the eleventh paragraph of the revised Discussion section.

See also the fourth paragraph in the Results’ section entitled “Isolation of GIPC1/Synectin as a PLXND1-binding protein and dissection of the molecular determinants required for PLXND1-GIPC complex formation” and remaining of the revised Discussion section.

6B) Additionally, the authors do not seem to consider other potential roles of GIPCs that are PlxnD1-independent. Although they state in their Results section that "the angiogenesis deficits of gipc mutants are, at least partially, due to increased PlxnD1 signaling", they do not assess or discuss additional possibilities.

We revised both the Introduction (penultimate paragraph) and the Discussion (tenth and eleventh paragraphs) sections to provide further details about the relationship between GIPC1 and NRP1-mediated VEGF signaling and to highlight that GIPC proteins have multiple partners, including additional molecules involved in the modulation of vascular development. See also our reply to essential revision 6A.

6C) One relatively easy experiment that the authors could perform in order to address these questions would be to overexpress plexind1 (WT and truncated forms) in endothelial cells of gipc single and double mutants. Alternatively, the authors could consider the possible relevance of the known interactions of GIPC with Neuropilin. While Neuropilin can act as a Plexin coreceptor, it can also act in a Plexin-independent manner. This is of relevance especially in experiments analyzing growth cone collapses or other effects on morphology, where Neuropilin has been shown to act independently of Plexin. Can the authors distinguish the negative regulation of Plexin from a potential positive regulation of Neuropilin? This is of course hampered by the caveat that there are no reliable Neuropilin reagents in zebrafish available as morpholino and mutant data only partially match. Maybe different downstream effectors could be affected pharmacologically? At a minimum, it would be helpful for the authors to provide more discussion of alternate possibilities that could explain the observed ways in which GIPC function affects PlxnD1 activity.

It was unfeasible for us to do the experiments mentioned above (we lacked the required stocks/reagents and building them would take a few zebrafish generations). Hence, we instead strengthened the Discussion (see tenth and eleventh paragraphs) to specify that in all likelihood, the proangiogenic function of GIPCs is multifaceted and involves additional mechanisms besides the negative modulation of PlxnD1 signaling. See also our reply to essential revisions 6A and 6B.

However, we highlight that the angiogenesis deficits and SU5416 hypersensitivity of plxnd1skt6 mutants support the model that GIPCs directly influence PLXND1 signaling. This new data is found in the Results’ section of the revision entitled “Zebrafish mutants in which plxnd1 function is provided solely by a receptor deficient in GIPC binding display angiogenesis deficits with low penetrance and are hypersensitive to the antiangiogenic drug SU5416” (see Figures 2-3). Further support for this model comes from the experiments described in the Results’ section entitled “GIPC depletion potentiates SEMA3E-induced, PLXND1-dependent responses in HUVECs” (see Figure 7).

[Editors' note: the authors’ plan for revisions was approved and the authors made a formal revised submission.]

Thank you for submitting your revised manuscript entitled "GIPC proteins pattern the vasculature by limiting the antiangiogenic signaling of the Semaphorin receptor Plexind1" for consideration by eLife. Your revised submission has been reviewed by two out of the three original peer reviewers, and the evaluation has been overseen by a Reviewing Editor and a Senior Editor. The reviewers have opted to remain anonymous.

Our decision has been reached after a process of initial independent review followed by extensive discussion between the reviewers and consultation with the editors. The initial independent reviews are reprinted at the bottom of this email, and the summary of the reviewers' discussion is provided in the next paragraph. Based on the overall consensus emerging from this discussion, we regret to inform you that your work will not be considered further for publication in eLife.

To summarize the reviewers' discussion: Both reviewers appreciate the improvements made during the revision of this manuscript, including the changes to the text and the addition of new data. However, both reviewers ultimately concluded that the revised material did not provide sufficient evidence in support of the primary conclusions of the manuscript. While much of the data in the paper is robust, the in vivo interaction of PlexinD1 and GIPC has not been robustly demonstrated via analysis of the PlexinD1 skt6 mutant. Specifically, reviewers were dissatisfied with the degree of statistical significance present in the data that was meant to demonstrate the effect of the skt6 mutation on angiogenesis, as well as in the data that was meant to demonstrate the rescue of the null allele by skt6. The lack of robust statistical significance here throws the conclusion that GIPCs are direct, negative modulators of PlexinD1 signaling into question. Among other concerns (see reviews below), reviewers were uncertain about the conclusions reached from the SU5416 treatments: they wondered whether the Bonferroni type adjustment is appropriate in this case and whether developmental delay could confound the interpretation of these data. Reviewers agreed that the strongest elements of the manuscript are the GIPC mutant data, both alone and when bred to PlexinD1 mutants. Also, the in vitro experiments looking at pERK after Sema3e treatment in GIPC/ PlexinD1 knockdown/knockouts are convincing. However, these experiments show association and not binding/mechanism. While there is clearly an association of PlexinD1 and GIPC and pERK, it remains unclear whether this interaction requires direct binding.

Reviewer #1:

The authors claim that the plxnd1 skt6 allele encodes a hypomorphic Plxnd1 receptor which induces angiogenesis deficits in homozygous mutant embryos.

However in the Figure 2 legend they clearly state that there is no statistical difference in phenotype occurrence between Wt and mutant embryos.

In the Results section, the authors claim that there is no statistical difference in Se phenotypes in Wt and plxnd1 skt6 siblings (without clarifying whether this denotes heterozygous or homozygous mutant embryos).

If the observed effect is not statistically significant, it cannot be used to claim a angiogenesis defect with low penetrance.

Thank you for the time and effort that you have dedicated to the review of our manuscript.

1.1) As stated in the text, the frequency, penetrance and expressivity (Figure 2I, Figure 2—figure supplement 1) of the defects of plxnd1skt6 mutants are low and not statistically significantly different from WT. We can edit the text to clarify further how these observations fit with the proposed nature of the allele. By scoring the progeny derived from an incross of plxnd1skt6/+heterozygotes, we found that only the plxnd1skt6 homozygous mutants, but not their homozygous WT siblings, had “angiogenesis deficits with low penetrance”. This statement is neither self-contradictory nor misleading since the definition of penetrance (percentage of individuals with a given genotype displaying a phenotype associated with that genotype) is independent of the concept of statistical significance. Finally, we state: “…the unique presence of angiogenic deficits in the mutants…is consistent with the notion that the plxnd1skt6 allele is hypermorphic”. However, to add more clarity and nuance, we propose to revise this text to read instead: “…the unique presence of angiogenic deficits in the mutants, such as maximal and moderate Se and DLAV truncations, suggests that the plxnd1skt6 allele could be hypermorphic”.

1.2) To clarify, we claim that the plxnd1skt6 behaves like a hypermorph (gain of function; but see typo-1). The bases for this assessment is the data on Figures 2-3. For example: “…the hypothesis that GIPCs limit PLXND1’s antiangiogenic signaling foretells that plxnd1skt6 should be a plxnd1fov01b complementing hypermorph and that its homozygous mutants will display angiogenesis deficits and hypersensitivity to antiangiogenic compounds”. Hence, the definitive characterization of the plxnd1skt6 allele as hypermorphic is based on the hypersensitivity of the homozygous mutants to SU5416. Thus, we state: “…the angiogenesis deficits of plxnd1skt6 homozygotes and their hypersensitivity to the antiangiogenic drug SU5416 support the model that GIPC binding limits the antiangiogenic signaling of the PLXND1 receptor. In our interpretation, it is the loss of this inhibiting effect what confers to the plxnd1skt6 allele its demonstrable hypermorphic character.” See also authors’ reply 6.4. For additional comments regarding the SU5416 hypersensitivity of plxnd1skt6 mutants, see authors’ replies 3.1-3.5, 4.1-4.4, 6.2-6.4, 7.2 and 9.2.

Additionally I disapprove of the quantification, as the authors should have analyzed always the exact same region and number of Se-DLAVs per embryo and not a varying average of 10.5 per embryo.

For me it is not clearly stated whether the less than 5% of affected Se-DLAVs are of all counted Se-DLAVs or are number of embryos with affected Se-DLAVs. I assume the first, as it would be difficult to get to 5% of 12 embryos. However it would be important to know how many of the embryos do show vessel truncations in one or two vessels.

The total number of only 12 embryos analyzed for each genotype is also really low.

2.1) To clarify, we always analyzed the same anatomical region. In the Materials and methods section entitled “Quantification of angiogenesis deficits in the trunk’s arterial tree of WT and mutant zebrafish embryos” we say: “…Scoring of angiogenesis deficits was performed bilaterally in the ~six somite-long region dorsal to the yolk extension.” This method is used widely in the field. The small differences in the average number of Se-DLAVs analyzed (see Supplementary File 3, first table) are due to biological variability within and across genotypes. Briefly, embryos of the same stage are of similar, but not identical, size. The yolk extension’s length can vary slightly. Finally, the left and right sides of the embryo are in approximate register. Thus, our images capture 5 or 6 Se vessels per side (see the first table in Supplementary File 3).

2.2) To clarify, the quantification in Figure 2I reflects the percentage of affected Se-DLAVs of all counted Se-DLAVs. This information is provided in the legend of Figure 2.

2.3) To clarify, 3 out of 12 embryos show vessel truncations. This information is in the second table of Supplementary File 3 (as indicated in the legend of Figure 2).

2.4) The reviewer says that the number of embryos analyzed in Figure 2I is low. We analyzed 12 WT homozygotes and 12 plxnd1skt6 homozygous mutants, or 124-126 Se-DLAVs/genotype (Supplementary File 3). It is unclear why the reviewer considers these numbers insufficient because the reviewer offers no justification for this comment. This comment is inconsistent with the reviewer’s praise (reviewer’s comment #7) of Figure 4B (based on the analysis of 11-33 embryos/genotype). The reviewer’s comment is also inconsistent with the editorially approved plan of our revision. Briefly, in essential revision 4A (related to a different experiment), we agreed to compare two sets of 10 embryos without objection. Finally, we are aware that increasing sample size would allow us to detect a smaller effect size. However, we thought it was more relevant to test our hypothesis further using the SU5416 assay because lowering VEGF signaling would be very illuminating, as outlined in the authors’ reply 7.2 (please see also authors’ replies 1.2, 4.2-4.4 and 6.4).

The experiments addressing hypersensitivity to Su5416 have theoretically been done under the same conditions as when assessing the phenotype of homozygous plxnd1skt6 mutants.

Why do the authors suddenly see dramatic difference in angiogenesis defects now in 40% of the vessels in 0.1% DMSO? Why does the DMSO also affect WT development? How are changes to more than 80% affected vessels only achieving one star significance? another explanation would be that the embryos have not really reached 32hpf but are a little developmentally delayed. The experimental procedure does not state whether and how the embryos were dechorionated for treatment from 18hpf on.

3.1) To clarify, the data in Figures 2-3 was scored and analyzed identically (see legends and Materials and methods). We can add to the latter section the dechorionation procedure. In the legends of Figures 2-3 we indicate that the quantifications are for Se-DLAV angiogenesis. We can edit the Material and Methods to add that embryos were manually dechorionated before the treatments.

3.2) The reviewer incorrectly states that 40% of the vessels in DMSO-treated WT embryos show angiogenesis defects. Instead, the graph (Figure 3E; see also the first table of Supplementary File 4) shows that 15.3% of Se-DLAVs have an angiogenesis deficit. Most defects are minimal (13.1%), the rest are moderate (2.2%). The graph’s legend reads: “Percentage of Se-DLAV in 32 hpf embryos of the indicated genotype and treatment combinations belonging to each of the following four phenotypic classes”. The ~40% (35.7%) value that the reviewer cites is for the penetrance (percentage of embryos with Se-DLAV truncations).

3.3) The effects of DMSO are minimal (see above) and controlled for in our experiments. DMSO impacts embryonic development at high concentrations, as shown by zebrafish toxicology studies (see PMIDs 16125774, 21356178 and 23082109). Although zebrafish angiogenesis studies typically use only 0.1% DMSO as a drug vehicle (references #94 and #161), this dose can weakly impact WT vascular development, as our data shows. Thus, comparing the effects of DMSO and SU5416 (dissolved in DMSO) is an excellent experimental design feature of our study (and one that is absent from most other studies).

3.4) There is no evidence that DMSO or SU5416 induce developmental delay at the doses used. We can edit the text to highlight this information. Specifically, these treatments did not induce apparent effects on embryonic morphology and size, somite number, cardiac contractility and circulation (defects induced by high DMSO doses reported in toxicological studies). We also note that we used a suboptimal SU5416 dose. Together, these observations strongly suggest that SU5416 treatment induces specific effects, namely an increase in the frequency and severity of vascular defects (over those induced by DMSO only treatment). Moreover, the experiments with the two genotypes were run side by side using common dilutions. We can edit the Results to highlight these observations.

3.5) In Figure 3E the depiction of significant differences with one star is correct. As the legend states, the graph shows that “the distributions of the four phenotypic classes were statistically significantly different between all the possible pairwise comparisons of the four combinations of treatments and genotypes.” We also state that, the “…Significance values were calculated using two-sided Fisher Exact tests …”. In this statistical test, the level of significance (p-value) does not indicate the strength of the difference between groups. Thus, significant differences must be labeled with one star, regardless of the actual magnitude of the p-value (Laerd Statistics (2016). Fisher's exact test using SPSS Statistics. Statistical tutorials and software guides. Retrieved from https://statistics.laerd.com/). See Supplementary File 4 for the significance values.

SU5416 has been previously used in zebrafish at concentrations of 1 µM and was shown to completely block Vegf mediated ISV sprouting. Here the authors use 0.2µM and show an effect on Wt ISV migration, which is slightly enhanced in the mutants, but there is no evidence that this does not just reflect a developmental delay caused by the chemical treatment. The authors do need to control for this alternative hypothesis. Also the embryo in Figure 3B seems to be "larger" or at higher magnification than the others, as the distance from the PCV to the top of the DLAV is larger than in the other three images. also in the supplement it is evident that already 90% of the Wt embryos are affected by the SU5416 treatment. The experimental conditions should have been chosen so that the Wt is not affected at all.

In sum for essential revision1B we requested to show a statistically significant effect on angiogenesis. Unfortunately, the data presented are still not convincing.

4.1) There is no evidence that DMSO or SU5416 induce developmental delay. Moreover, we controlled for the effects of DMSO and used a suboptimal SU5416 dose. See the authors’ replies 3.3-3.4, 6.3 and 9.2.

4.2) Using an SU5416 treatment that affects the WT is practical and valid. First, 0.1% DMSO is the recommended dose for dissolving SU5416 (which is a water-insoluble compound). Second, and as our data shows, 0.1% DMSO alone has a weak effect on angiogenesis in the WT. Hence, even an ineffectively low dose of SU5416 dose (dissolved in DMSO by necessity) would still affect the WT due to DMSO’s “background” effects. Third, we controlled for the effects of DMSO (see authors’ replies 3.3-3.4). Finally, it is valid to start with a condition affecting the WT. Briefly, our experimental design (Figure 3) follows the logic of enhancer and suppressor screens, where one tests the sensitivity to an insult (a drug or a mutation). One can start with genotypes that display phenotypes that are not significantly different (like WT and plxnd1skt6; see Figure 2I) and challenge with a second insult (like SU5416; see Figure 3). Alternatively, one can start with an abnormal phenotype (like that of SU5416-treated WTs) and ask if a second insult (plxnd1skt6 homozygosity) modifies the phenotype (Figure 3). Either way, the conclusion is the same: SU5416-treated plxnd1skt6 mutants have more severe angiogenesis deficits than SU5416-treated WTs.

4.3) Our data show a statistically significant effect on angiogenesis. SU5416-induced vascular defects are more frequent and severe in plxnd1skt6 mutants than in WT embryos. SU5416 treatment induces defects in 89.7% of WTs and 100% of plxnd1skt6 mutants (a non-significant difference in penetrance; see Figure 3—figure supplement 1A and Supplementary File 4). However, the critical point is that this 10.3% penetrance difference cannot account for the significant increase in the frequency and severity of angiogenesis deficits observed in SU5416-treated plxnd1skt6 mutants (Figure 3E, Figure 3-supplement 1B and Supplementary File 4). To elaborate, the graph in Figure 3E (26-29 embryos analyzed) shows the “Percentage of Se-DLAV in 32 hpf embryos of the indicated genotype and treatment combinations belonging to each of the following four phenotypic classes…The distributions of these four phenotypic classes were statistically significantly different between all the possible pairwise comparisons of the four combinations of treatments and genotypes”. Figure 3—figure supplement 1 shows the penetrance (A) and expressivity (B) of Se-DLAV truncations for treated embryos. Note, for example, that for SU5416 treatment the penetrance of defects is not significantly different between WT and mutants, yet their expressivity is. Supplementary File 4 (second to third pages) shows eight tables of significance values (p) related to various comparisons of distributions, penetrance, and expressivity. Please note that in seven out of eight comparisons of significance values, the SU5416 treatment induces a significant difference between WT and plxnd1skt6mutants (with mutants being more affected). Again, the exception is for the penetrance of Se-DLAV truncations (first table on the third page) and, meaningfully, the expressivities are significantly different (second and third tables on the third page). Finally, we highlight the comment by reviewer #2 stating “the new skt6 allele of plexinD1 was used in very clear experiments to show it is functional and hypermorphic”.

4.4) The hypersensitivity of plxnd1skt6 mutants to SU5416 supports the model that GIPCs limit Plxnd1 antiangiogenic signaling. As explained above, our data and statistical analysis show that DMSO and SU5416 treatments impair angiogenesis, with SU5416 inducing defects with both higher frequency and severity. Both DMSO and SU5416 have stronger effects on plxnd1skt6 homozygotes. These mutants express only a Plxnd1 receptor that is deficient in GIPC binding. The hypothesis that GIPCs limit antiangiogenic signaling predicts that plxnd1skt6 mutants should have potentiated antiangiogenic Plxnd1 signaling and thus be hypersensitive to antiangiogenic insults, precisely what our data shows. Accordingly, we highlight that the GIPC-based modulation of endothelial PLXND1 signaling is likely to be also of particular relevance in contexts with minimal proangiogenic stimulation.

4.5) We agree that the embryo in Figure 3B does appear larger. We could show the image of a different embryo instead. However, we note that all the images were taken using the same magnification. Importantly, our angiogenesis quantification scales (see authors’ reply 5.2) are based on the relative span of the vascular structure(s) scored, precisely to eliminate the noise that variations in embryo size would otherwise introduce.

Likewise, why is there such a big variance in phenotype distribution when scoring gipc1 skt1(MZ) mutant embryos? Compare Figure 4B with Figure 5C.

It would also be helpful if the authors would indicate e.g. in Figure 2 or 3 and clearly define in the text what constitutes a maximal, a moderate or a minimal Se-truncation.

5.1) We clarify that it is invalid to compare the phenotype distributions of the gipc1skt1(MZ)mutants shown in Figures 4B and 5C because the angiogenesis deficits in these figures were scored using different scales, as indicated in their respective figure legends. Specifically, Figure 4B and Supplementary File 5 show classes of Se truncations (for example, in gipc1skt1(MZ) mutants 15.53% of the Se vessels are truncated). In contrast, Figure 5C and Supplementary File 6 show classes of Se-DLAV truncations (for example, in gipc1skt1(MZ)mutants 18.7% of Se-DLAV are truncated).

5.2) To clarify, the definitions for the truncation classes for both scoring scales are in the manuscript. In the Materials and methods, the section entitled “Quantification of angiogenesis deficits in the trunk’s arterial tree of WT and mutant zebrafish embryos” describes the definitions used for the scoring of “Se truncations” and “Se and DLAV truncations”. The Se truncation scale emphasizes ventral defects (Se truncation scale), while the Se-DLAV truncation scale emphasizes dorsal defects. Importantly, we do not compare the data in Figure 4 with that of Figure 5, as this would be inappropriate.

The authors claim that Gipc binding limits the antiangiogenic signaling of the PlxnD1 receptor. However, there are too many assumptions and too little data for such a statement.

For one: what is the antiangiogenic Plxnd1 signaling: where is that shown just based on a poorly controlled hypersensitivity to VegfR2 receptor inhibition, which was done for a very long time period? or based on the proposed hypomorphic identity of the plxnd1 skt6 allele, which as mentioned above is not clearly a significant phenotype.

6.1) We speak of “antiangiogenic Plxnd1 signaling” because multiple published studies in both zebrafish and mice justify the use of this term. Moreover, we agreed to use this term in the editorially approved plan for our revision. First, inactivation of Plxnd1 increases angiogenesis (leads to the formation of more, ectopic, excessively branched and misguided Se sprouts). See the Introduction and Results and references cited therein. Second, ligand stimulation of PlxnD1 exerts a molecular effect opposite to that induced by proangiogenic VEGF signaling (reduces ERK activity, rather than increasing it). See Results and references cited therein. Finally, we agreed to use the term “Plxnd1 signaling”. In the editorially approved plan for our revision, part of our answer to essential revision 1A reads: “We will revise the text to highlight that PlxnD1 is anti-angiogenic, avoiding the use of the terms “activity” and “potency”. We will instead speak of “PlxnD1 signaling”.

6.2) To clarify, our use of the term “antiangiogenic Plxnd1 signaling” is unrelated to the proposed nature of the plxnd1skt6 allele and the hypersensitivity of plxnd1skt6 homozygotes to SU5416 (VegfR2 inhibitor).

6.3) The 14-hr span of our low dose SU5416 treatment (from 18 to 32 hpf) is within the range of similar studies and is designed to target both Se and DLAV angiogenesis specifically. It is unclear why the reviewer considers the treatment overtly long. In reference #94 (Figure 1 and Materials and methods) the authors treat for 12 hours longer (24-hours; from shield stage (6 hpf) to 30 hpf). In reference #161 (Materials and methods) the authors treat for 2 hours less (12-hr; from 18 to 30 hpf). Furthermore, we used a lower SU5416 dose than the studies cited above, which diminishes the likelihood of non-specific effects. For references about the timing of Se and DLAV angiogenesis, please see references #5, 6, 10, 82 and PMIDs 22899709 and 26588168.

6.4) To clarify, we do not propose that the plxnd1skt6 allele is hypomorphic. On the contrary, our data support the notion that this mutant allele is hypermorphic. Please see the authors’ replies 1.1-1.2). Please see also the comment by reviewer #2 saying that: “the new skt6 allele of plexinD1 was used in very clear experiments to show it is functional and hypermorphic”.

The analysis of the gipc mutants is worthwile, however there is too little evidence supporting the Gipc/Plexin interaction on a phenotypic level, with regards to their collaborating role in angiogenesis.

An alternative explanation for the partial reduction in phenotype would be that while in gipc1 skt1 Mz embryos gipc2 and gipc3 can still bind to Wt Plexin and therefore are less available for interaction with another recptor (like Nrp1) in the plxnd1 fov01b heterozygous less plexin molecules are available to limit the gipc signaling pool. This would also explain the reduction in phenotype, and be based on a Plexin/gipc interaction,but say nothing about the actual signaling cascades required for generating an angiogenesis phenotype (see our comment regarding essential revision 4B).

7.1) To clarify, we do not say that the role of GIPCs and Plxnd1 is collaborative. We state that GIPCs limit Plxnd1 signaling; see manuscript’s title, manuscript’s impact statement and at various points throughout the text.

7.2) The reviewer’s alternative model about why plxnd1 heterozygosity suppresses the angiogenesis deficits of gipc1skt1(MZ) mutants (Figure 5) can explain this genetic interaction, but not the data in Figure 3 (the SU5416 hypersensitivity of plxnd1skt6 mutants). In the reviewer’s model, GIPCs are rate limiting and PlxnD1 acts like a sink for GIPCs, thereby reducing the amount of GIPC available for VEGFR2/Nrp1 signaling. Given that the receptor encoded by plxnd1skt6 is deficient in GIPC binding, it should not act as a sink for GIPCs (or do so minimally). Hence, the reviewer’s model predicts that in plxnd1skt6 mutants the pool of GIPC available for VEGFR2/Nrp1 signaling should be greater and thus the mutants should be insensitive or less sensitive to the antiangiogenic VEGFR2 inhibitor. However, Figure 3, Figure 3—figure supplement 1 and Supplementary File 4 show that plxnd1skt6 mutants are instead hypersensitive to SU5416 (the opposite result predicted by the reviewer’s hypothesis). In contrast, the model that GIPC binding to Plxnd1 (see Figure 1) limits the antiangiogenic activity of the latter is consistent with all the zebrafish (Figures 2-6 and corresponding figure supplements and supplementary files) and cell culture data (Figure 7 and corresponding figure supplements and supplementary files) in our manuscript. See also authors’ reply to the summary of the reviewers’ discussion (E), authors’ replies 1.2, 3.5, 4.3-4.4, 7.3, 8.1-8.2, 9.1-9.2 and the comment by reviewer #2 (comment #12).

7.3) We highlight that our results and interpretations do not argue against the notion that GIPCs also have vascular effects that are PlxnD1-independent or Nrp1-dependent, as repeatedly stated in the manuscript.

As the interaction studies between Gipc1/2 and plxnd1 are only analyzed qualitatively, meaning that you find massive ectopic sprouting, how were you expecting to see a double phenotype in case of no interaction? the gipc1skt1(Mz); gipc2 skt3/skt4 double phenotype did affect less than 20% of the embryos, only about 10% were affected medium to severely. that would mean that in case of NO interaction you would have found 10% of the Se's being truncated, but still having massive ectopic filopodia and sprouts. Given that there would be additional ectopic sprouts coming from the neighboring Se's, I doubt that this would have been detectable. I therefore believe that the experiment is not suitable to disprove alternative hypothesis…

8.1) The HUVEC data in Figure 7E addresses the effect of simultaneous inactivation of PLXND1 and GIPC quantitatively. Importantly, PLXND1 inactivation and the double loss of PLXND1 and GIPC induce molecular phenotypes that are not statistically significantly different. At 15 and 45 min of SEMA3E stimulation, the relative ERK activity of GIPC knockdown cells (green bars) is significantly different from that of PLXND1 KO (knockout) cells treated with either non-targeting shRNAs (red bars) or GIPC shRNAs (blue bars). However, the relative ERK activity of PLXND1 KO (knockout) cells treated with non-targeting shRNAs (red bars) or treated with GIPC shRNAs (blue bars) is not significantly different. In other words, PLXND1 KO cells with GIPC knockdown (blue bars) do not show an “intermediate” pERK level between that of PLXND1 KO cells (red bars) and GIPC knockdown cells (green bars). Instead, they show a level of relative ERK activity like that of PLXND1 KO cells (red bars).

8.2) The results of the qualitative phenotypic analysis of MZ gipc1; gipc2 double mutants with and without morpholino induced plxnD1 knockdown are consistent with the model (rather than prove it). Our manuscript says that the fact that double maternal-zygotic (MZ) gipc mutants show excessive angiogenesis in the absence of Plxnd1 signaling is “consistent with the model that the angiogenesis deficits of gipc mutants are, at least partially, due to increased PLXND1 signaling”. Please see also the authors’ reply 7.2. Moreover, the experiment followed the editorially approved plan of our revision. Specifically, in our reply to essential revision 4A, we said that “…it is impossible to score the abundance and length of Se vessels at 32 hpf, we must evaluate the vascular abnormalities qualitatively”.

In sum there is not enough evidence supporting a model whereby Gipc limits Plexind1 antiangiogenic signaling, as Gipc could still act on angiogenesis in a Plexin independent manner, eg via neuropilin (see essential revisions 6!). I don't think that the authors have sufficiently experimentally addressed this question. However, this affects the core of the authors claims by directly affecting the functionality of the Plexin/Gipc interactions in angiogenesis.

9.1) We provide strong experimental support for the model that GIPCs directly limit antiangiogenic Plxnd1 signaling. Our model does not imply that GIPCs lack Neuropilin-dependent roles in angiogenesis and experiments directly addressing the involvement of GIPCs in Neuropilin-mediated VEGFR2 signaling are not part of the editorially approved plan of our revision. As summarized in the authors’ reply 7.2, the results of our fish and cell culture experiments support the notion that GIPCs directly limit antiangiogenic Plxnd1 signaling. Our model and results are compatible with the notion that GIPCs also play Plxnd1-independent vascular roles, including Neuropilin-dependent functions (see authors’ reply 7.3).

Moreover, in essential revision 6C of the editorially-approved plan for our revision, a reviewer wonders whether we could distinguish the negative regulation of Plexin from a potential positive regulation of Neuropilin. This reviewer acknowledges the caveat that “there are no reliable Neuropilin reagents in zebrafish available,” concluding that we should “provide more discussion of alternate possibilities that could explain the observed ways in which GIPC function affects PlxnD1 activity”. Accordingly, the Introduction and Discussion detail the relationship between GIPC1 and NRP1-mediated VEGF signaling and highlight that GIPCs have many partners, including other modulators of vascular development. See also our reply to essential revision 6A in the editorially approved plan for our revision and authors’ reply to the summary of the reviewer’s discussion (E).

9.2) The SU5416 hypersensitivity of plxnd1skt6 mutants and the results of the HUVEC experiments directly argue that GIPCs act in a PLXND1-dependent fashion. Note that the HUVEC experiments were done without exogenous VEGF stimulation to avoid simultaneous activation of Neuropilin1/GIPC1-mediated VEGFR2 signaling. To review the arguments related to the SU5416 hypersensitivity, please see the authors’ reply 7.2 (and also the authors’ replies 1.2, 4.3-4.4). Importantly, the cell culture experiments with HUVECs provide a unique opportunity to interpret PLXND1 signaling without exogenous VEGF. Given that GIPC1 association with Neuropilin1 is thought to facilitate VEGFR2 signaling, the reviewer’s hypothesis predicts that GIPC depletion should not affect SEMA3E-induced, PLXND1-dependent responses in HUVECs. However (and in support of our model), GIPC depletion potentiates SEMA3E-induced, PLXND1-dependent responses, leading to a further decrease in relative pERK abundance and cell hypercollapse (see Figure 7 and Figure 7—figure supplements 1-4).

Reviewer #2:

The authors have cleared up misconceptions created by language and elegantly used hypotheses to set up outcomes of their experiments and what this means. This is a large improvement over the previous version of the manuscript.

The new skt6 allele of plexinD1 was used in very clear experiments to show it is functional and hypermorphic. There is also a plexinD1 null HUVEC line generated as well for the cell culture experiments.

Graphs have been improved. Numbers increased and emphasized.

Improved discussion of Burk results in context of the results of this manuscript.

Overall, there is a huge improvement to the manuscript. Many figures are edited or completely new and there is new data to answer my major points.

We thank you for your help and are delighted to know that you appreciate how all the new reagents, experiments, and data that we collected and how these satisfactorily test the proposed model.

[Editors’ note: the author responses to the re-review follow.]

I have read your rebuttal letter and conferred with the Reviewing Editor. We both agree that it would be appropriate for you to submit a revised version of the manuscript that addresses the comments of the reviewers.

We would like to clarify that both reviewer 1 and 2 were in good agreement during the second round of review. Therefore, we urge you to address their comments to the best of your ability. However, given your concerns we will not return the revised manuscripts to either reviewer, but will instead ask previous reviewer 3 (who did not respond to our previous request to re-review), plus a new reviewer. Of course we cannot guarantee the outcome but will do all that we can to be fair.

The re-revised manuscript addresses the comments that reviewer 1 and reviewer 2 provided after our initial revision. Hence, the present manuscript includes several changes. Please note that these modifications include an increased number of references and changes to their numbering.

We also modified two figures based on the feedback of these two reviewers. Briefly, the image in Figure 3B was changed. Also, the penetrance graph in Figure 5—figure supplement 1A was updated by including an asterisk to indicate that the penetrance of Se-DLAV truncations is significantly different between the two genotypes under.

Moreover, we also updated Supplementary Files 3-6. We inserted comments into each of these re-revised Supplementary Files explaining the rationale behind the changes that we performed.

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

As you can see from the reviewers' comments (copied below), the reviewers feel that the manuscript has been improved, but both agree that there are some remaining issues that need to be addressed before acceptance. The reviewers have discussed these issues with each other after submitting their reviews, and they agree that it is particularly important for the manuscript to be thoroughly edited, with the goal of creating a more succinct and streamlined experience for the reader.

Done. We edited the text in several places as follows.

Results’ second section (entitled “Zebrafish plxnd1skt6 homozygous mutants, which express a…”): We revised the last line of the third paragraph and revised the sixth (last) paragraph.

Results’ third section (entitled “Homozygous plxnd1skt6 mutants are hypersensitive…”): We revised the first line of the third paragraph and the last three lines (fourth and fifth paragraphs).

Results’ seventh (last) section (entitled “GIPC depletion potentiates …”): We performed several minor edits to the third, fourth and fifth paragraphs.

Discussion section: We edited the first line of the first paragraph and the last line of the third paragraph.

The reviewers also agree that you should make a series of specific adjustments to the text and figures in an effort to clarify or modify the message, in accordance with each of the individual points that have been raised in the reviews below.

Done. See our replies regarding the comments of reviewer 3 and reviewer 4.

Finally, regarding reviewer 4's suggestion that you perform an additional co-IP experiment (see comment #1 from reviewer 4 below), the reviewers agree that this is not essential. If you have the data at hand, please do include it, but it will also be acceptable to simply make changes in the text describing the biochemical effects of this mutation. In that case, the idea that this mutant version of the zebrafish protein loses its ability to interact with Gipc proteins should be presented as speculation, not as fact.

We do not have such CoIP data. Thus, we changed the Abstract by editing the line that said “Zebrafish with a truncated Plxnd1 receptor that is deficient in GIPC binding…” to “We found that zebrafish that endogenously express a Plxnd1 receptor putatively impaired in GIPC binding…” (fifth line). We also made other minor revisions to the Abstract to improve its readability.

Second, we also changed the title of the Results second section to “Zebrafish plxnd1skt6 homozygous mutants, which express a Plxnd1 receptor with a predicted impairment in GIPC binding, display angiogenesis deficits with low frequency”. We also changed the end of the first paragraph of this section to: “…the plxnd1skt6 mutant allele is expected to encode a Plxnd1 receptor with reduced or null GIPC binding ability.”

Third, we edited the title of Figure 2, which now reads: “The plxnd1skt6 allele encodes a functional Plxnd1 receptor putatively impaired in GIPC binding, and its homozygosity induces angiogenesis deficits with low frequency.”

Reviewer #3:

After carefully reading the full history of this manuscript, I tend to agree with the previous assessment of reviewer #2.

I believe the revised version provided by the authors successfully addressed the essential concerns raised during the first round of review. Moreover, I understand that the data added in the revised version was approved in advance in the context of the revision plan.

While I agree with reviewer #1, that some issues of statistical significance were not fully solved, the authors did a good job not only in providing extensive explanations for the observed results, but also by listing alternative hypotheses. In my view, the faithful presentation of the results, and the efforts made to explain them, are valuable, despite of the phenotypes themselves being not hugely penetrant.

As agreed by all 3 reviewers, the findings of this manuscript are important and add to our understanding of the mechanisms underlying Plexind1 signalling. Since most of the initial concerns were addressed properly, and the findings are carefully interpreted, I tend to support publication.

I do think however that the title/impact statement/conclusions could be toned down a bit to reflect more accurately the main findings.

Done. The impact statement now reads: “Zebrafish mutants and human endothelial cell experiments reveal that GIPC family endocytic adaptors bind to the Semaphorin receptor PLEXIND1, a critical regulator of vascular development, to negatively modulate its signaling.”

https://doi.org/10.7554/eLife.30454.031

Article and author information

Author details

  1. Jorge Carretero-Ortega

    Department of Cell Biology, Skirball Institute of Biomolecular Medicine, New York University Langone Medical Center, New York, United States
    Contribution
    Conceptualization, Formal analysis, Funding acquisition, Methodology, Project administration, Resources, Supervision, Visualization, Writing—original draft, Writing—review and editing, Investigation
    For correspondence
    carretero_2000@yahoo.com
    Competing interests
    No competing interests declared
  2. Zinal Chhangawala

    Department of Cell Biology, Skirball Institute of Biomolecular Medicine, New York University Langone Medical Center, New York, United States
    Contribution
    Conceptualization, Formal analysis, Investigation, Resources, Methodology, Writing—review and editing, Visualization
    Competing interests
    No competing interests declared
  3. Shane Hunt

    Department of Cell Biology, Skirball Institute of Biomolecular Medicine, New York University Langone Medical Center, New York, United States
    Contribution
    Resources, Investigation, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  4. Carlos Narvaez

    Department of Cell Biology, Skirball Institute of Biomolecular Medicine, New York University Langone Medical Center, New York, United States
    Contribution
    Resources, Investigation, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  5. Javier Menéndez-González

    Department of Cell Biology, Skirball Institute of Biomolecular Medicine, New York University Langone Medical Center, New York, United States
    Contribution
    Resources, Investigation, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  6. Carl M Gay

    Department of Cell Biology, Skirball Institute of Biomolecular Medicine, New York University Langone Medical Center, New York, United States
    Contribution
    Resources, Investigation, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  7. Tomasz Zygmunt

    Department of Cell Biology, Skirball Institute of Biomolecular Medicine, New York University Langone Medical Center, New York, United States
    Contribution
    Resources, Investigation, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  8. Xiaochun Li

    Department of Population Health, New York University School of Medicine, New York, United States
    Contribution
    Formal analysis, Resources, Writing—review and editing
    Competing interests
    No competing interests declared
  9. Jesús Torres-Vázquez

    Department of Cell Biology, Skirball Institute of Biomolecular Medicine, New York University Langone Medical Center, New York, United States
    Contribution
    Conceptualization, Formal analysis, Funding acquisition, Methodology, Project administration, Resources, Supervision, Visualization, Writing—original draft, Writing—review and editing, Investigation
    For correspondence
    jtorresv@med.nyu.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3808-3978

Funding

National Institutes of Health (1R01HL133687-01)

  • Jesús Torres-Vázquez

Consejo Nacional de Ciencia y Tecnología (187031)

  • Jorge Carretero-Ortega

Consejo Nacional de Ciencia y Tecnología (203862)

  • Jorge Carretero-Ortega

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

Acknowledgements

Funding sources: This work was supported by postdoctoral fellowships from Mexico’s Consejo Nacional de Ciencia y Tecnología (CONACyT 187031 and 203862, to JC-O) and by the National Heart Lung and Blood Institute, NIH (1R01HL133687-01, to JT-V). For reagents and protocols: Moses Chao (FLAG-mGIPC1 construct; see Lou et al., 2001), Holger Knaut (mCherry antibody), Gregory Palardy and Ajay Chitnis (zebrafish genome editing protocol with Cas9 protein), and Xingxu Huang (pST1374-NLS-flag-linker-Cas9 vector; Addgene plasmid #44758; see Shen et al., 2013). For useful discussions: Knaut lab members and other members of the Skirball Institute’s Developmental Genetics Program. Microscopy: We thank Yang Deng, Michael Cammer and NYU School of Medicine’s Microscopy Laboratory for data collected with the Leica SP5 confocal system (grant NCRR S10 RR024708). We thank Agnel Sfeir, Mamta V Tahiliani and Matthias Stadfeld for access to the Eclipse Ti-E inverted microscope (Nikon) used for collecting HUVEC collapse data and for reagents.

Senior Editor

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

Reviewing Editor

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

Publication history

  1. Received: August 9, 2017
  2. Accepted: April 15, 2019
  3. Version of Record published: May 3, 2019 (version 1)

Copyright

© 2019, Carretero-Ortega et al.

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

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