Tmem2 Regulates Embryonic Vegf Signaling by Controlling Hyaluronic Acid Turnover

Article

Tmem2 Regulates Embryonic Vegf Signaling by Controlling Hyaluronic Acid Turnover Graphical Abstract

Authors Jessica E. De Angelis, Anne K. Lagendijk, Huijun Chen, ..., Carol Wicking, Benjamin M. Hogan, Kelly A. Smith

Correspondence [email protected] (B.M.H.), [email protected] (K.A.S.)

In Brief Vegf signaling is required for primary and secondary angiogenesis. De Angelis, Lagendijk et al. show that HA degradation is essential for proper Vegf signaling and that Tmem2 regulates HA turnover.

Highlights d

Tmem2 regulates Vegf signaling to control angiogenesis

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Tmem2 controls degradation of extracellular HA

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HA turnover by hyaluronidase or digested HA rescues angiogenesis and Vegf signaling

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Overexpression of Vegfc can bypass the need for Tmem2 turnover of extracellular HA

De Angelis et al., 2017, Developmental Cell 40, 123–136 January 23, 2017 Crown Copyright ª 2017 Published by Elsevier Inc. http://dx.doi.org/10.1016/j.devcel.2016.12.017

Developmental Cell

Article Tmem2 Regulates Embryonic Vegf Signaling by Controlling Hyaluronic Acid Turnover Jessica E. De Angelis,1,5 Anne K. Lagendijk,1,5 Huijun Chen,1 Alisha Tromp,1 Neil I. Bower,1 Kathryn A. Tunny,2 Andrew J. Brooks,2 Jeroen Bakkers,3,4 Mathias Francois,1 Alpha S. Yap,1 Cas Simons,1 Carol Wicking,1 Benjamin M. Hogan,1,* and Kelly A. Smith1,6,* 1Institute

for Molecular Bioscience Institute, Translational Research Institute The University of Queensland, Brisbane, QLD 4072, Australia 3Department of Cardiac Development and Genetics, Hubrecht Institute, University Medical Centre Utrecht, Utrecht 3584 CT, the Netherlands 4Department of Medical Physiology, University Medical Centre Utrecht, Utrecht 3584 EA, the Netherlands 5Co-first author 6Lead Contact *Correspondence: [email protected] (B.M.H.), [email protected] (K.A.S.) http://dx.doi.org/10.1016/j.devcel.2016.12.017 2Diamantina

SUMMARY

Angiogenesis is responsible for tissue vascularization during development, as well as in pathological contexts, including cancer and ischemia. Vascular endothelial growth factors (VEGFs) regulate angiogenesis by acting through VEGF receptors to induce endothelial cell signaling. VEGF is processed in the extracellular matrix (ECM), but the complexity of ECM control of VEGF signaling and angiogenesis remains far from understood. In a forward genetic screen, we identified angiogenesis defects in tmem2 zebrafish mutants that lack both arterial and venous Vegf/Vegfr/Erk signaling. Strikingly, tmem2 mutants display increased hyaluronic acid (HA) surrounding developing vessels. Angiogenesis in tmem2 mutants was rescued, or restored after failed sprouting, by degrading this increased HA. Furthermore, oligomerized HA or overexpression of Vegfc rescued angiogenesis in tmem2 mutants. Based on these data, and the known structure of Tmem2, we find that Tmem2 regulates HA turnover to promote normal Vegf signaling during developmental angiogenesis.

INTRODUCTION Angiogenesis is the formation of new vessels from pre-existing vasculature. It is essential in tissues greater than 1 mm3 in size for growth and occurs via growth-factor-induced sprouting of endothelial cells (ECs). A non-cell autonomous source of vascular endothelial growth factor (VEGF) activates ECs via VEGFR2/KDR promoting pERK1/2, PI3K, AKT, and other intracellular signaling in different contexts, inducing proliferation and migration of ECs (Herbert and Stainier, 2011). In zebrafish, primary angiogenesis (initial sprouting from the dorsal aorta) is regulated by Vegfa and secondary angiogenesis (sprouting from the cardinal vein) by Vegfc. In arterial angiogenesis, Vegfa

acts to promote intracellular Plcg signaling, which is essential for all arterial angiogenesis (Lawson et al., 2003). Despite the divarication of EC signaling in response to Vegf in many systems, signaling in zebrafish vasculature during both primary and secondary sprouting angiogenesis occurs almost exclusively via pErk1/2 (Shin et al., 2016a, 2016b). In response to active signaling in ECs, angiogenesis initiates at approximately 22 hr post-fertilization (hpf) in the zebrafish trunk, progresses in an anterior to posterior manner during the extension of the developing tail, and has formed a lumenized and functional blood vasculature by the second day of development. Tmem2 was identified from a forward genetic screen for zebrafish mutants with cardiac looping defects (Smith et al., 2011; Totong et al., 2011), and recently it has described to play a role in skeletal muscle morphogenesis (Ryckebusch et al., 2016). The Tmem2 protein is a single-pass transmembrane domain protein and shows enriched expression during heart development in the endocardium and trabeculae in zebrafish and mice (Smith et al., 2011). The nearest homolog to Tmem2, Kiaa1199, was recently suggested to selectively degrade hyaluronic acid (HA) molecules from high molecular weight chains to lower molecular weight forms (Yoshida et al., 2013). The molecular function of Tmem2, any relationship with HA degradation, and the mechanisms by which it controls vertebrate developmental processes remain to be characterized. HA is an extracellular matrix (ECM) component required for cardiovascular development (Camenisch et al., 2000; Lagendijk et al., 2013). HA is a glycosaminoglycan formed in linear strands of alternating glucuronic acid and N-acetylglucosamine repeats and is known as a ‘‘space-filling’’ molecule for its high affinity for water. HA has been applied clinically for diverse uses, including treatment of osteoarthritis, in various eye surgical procedures (Kleinberg et al., 2011), and as an injectable filler in cosmetic surgeries (Kim and Sykes, 2011). HA is synthesized in a two-step process: firstly, involving UDP-glucose 6-dehydrogenase and, secondly, by the HA synthases, Has1, Has2, and Has3. The Has enzymes are membrane-bound and synthesize HA into the ECM as long continuous strands of various sizes up to 10 MDa. HA is subject to high turnover with specific degrading enzymes, called hyaluronidases (HA’se), known to regulate HA depolymerization. Interestingly, this enzymatic activity generates

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small oligosaccharide fragments, collectively known as o-HA, which are shown to possess bioactive properties (Rodgers et al., 2006; West et al., 1985). Has2 deficient mice die during early embryogenesis from vascular and cardiac defects (Camenisch et al., 2000) and knockdown of hyal1 (a HA’se required for degradation of HA) decreases the angiogenic potential of cells in culture as well as microvessel growth in tumor xenograft models (Tan et al., 2011). Nevertheless, the developmental and mechanistic roles of HA, HA-degrading enzymes, and HA-synthesizing enzymes are not completely understood. We report here the characterization of a specific angiogenesis defect in tmem2 mutants. We find that loss of Tmem2 function in zebrafish results in loss of endothelial pErk1/2 signaling and subsequent loss of angiogenic sprouts. We observe dynamic and restricted expression patterns for functional Tmem2 protein in vivo, including in the vascular EC membrane. In tmem2 mutants, using a biosensor, histochemical staining, and enzymatic manipulation of HA turnover, we find that increased perivascular HA is responsible for reduced EC Erk signaling and stalled angiogenesis. We show that degrading mutant HA by exogenously introducing HA’se or increasing Vegfc levels can rescue tmem2 mutant vascular phenotypes and restore angiogenesis. Our findings introduce an in vivo regulator of angiogenesis in Tmem2 and identify a mechanism whereby ECM turnover of HA controls Vegf signaling and developmental angiogenesis. RESULTS tmem2 Is Required for Arterial and Venous Sprouting Angiogenesis in the Zebrafish In a forward genetic screen for mutants that display defective secondary angiogenesis, we identified two independent alleles of a mutant with no venous sprouting. Using a whole-genome sequence-based mapping and mutation detection approach (Koltowska et al., 2015b), both alleles mapped to chromosome 5 and identified putative mutations in the gene tmem2. The tmem2uq1ks allele encoded a predicted premature stop codon (Q616X) and the tmem2uq2ks allele a predicted splice acceptor site mutation at the boundary of intron 6/exon 7 (Figures S1A– S1D). We had previously isolated a tmem2hu5935 mutant from an earlier forward genetic screen for cardiac morphogenesis defects (Smith et al., 2011), which possesses a premature stop codon at position 985, truncating the protein (K328X). To verify this potential role in angiogenesis, we focused on our previously characterized allele, which is utilized for all of the following analyses. We crossed the tmem2hu5935 mutant with the Tg(kdrl:EGFP) transgenic line and examined vascular development and found dramatic defects in vascular development in concert with the previously described cardiac defects. A reduction in the number of intersegmental vessels (ISVs) was observed in tmem2hu5935 mutant embryos at both primary (arterial; 28 hpf) and secondary (venous; 50 hpf) sprouting stages (Figures 1A– 1C), with variable expressivity of the phenotype. At 48 hpf, tmem2hu5935 mutants presented with shunting of the circulation between the dorsal aorta (DA) and the posterior cardinal vein (PCV), accompanied by a failure of the caudal vein plexus to remodel (data not shown). Given that both arterial and venous sprouting are affected, we examined arterial-venous specification, but in situ hybridization (ISH) for arterial markers, hey2, 124 Developmental Cell 40, 123–136, January 23, 2017

efnb2a (data not shown), and the venous marker dab2 showed no differences between sibling and mutant embryos at 24 hpf (Figure S1E). Live time-lapse imaging of mutant and sibling embryos during primary sprouting (Movie S1) showed that, in mutant embryos, angiogenesis appeared to be correctly segmented, and sprouting could initiate from the DA; however, angiogenesis arrests during sprouting. Tmem2 Is Required for Normal Initiation of Developmental Vegf Signaling Given the well-established role for Vegf signaling in promoting both primary and secondary angiogenesis, we examined whether tmem2 loss of function resulted in reduced Vegf signaling. We took advantage of the variable expressivity of the tmem2hu5935 mutant phenotype to investigate whether a mild reduction in Vegf signaling could enhance the tmem2 ISV defect. We carefully titrated the Vegf receptor inhibitor, SU5416 (Fong et al., 1999), to suboptimal concentrations that had no effect on angiogenic sprouting in sibling embryos. At these concentrations, the tmem2 mutant phenotype was significantly and synergistically enhanced, suggesting that Tmem2 may act in the Vegf pathway (Figures S1F and S1G). To investigate signaling more directly, we examined the intracellular effector for Vegfr-driven primary and secondary angiogenic signaling, phosphorylated Erk1/2 (pErk) (Le Guen et al., 2014; Shin et al., 2016a, 2016b). We performed whole-mount immunofluorescence (IF) analysis at the onset of arterial (22 hpf) and venous (32 hpf) sprouting stages (Figures 1D and 1E). Quantification showed significantly fewer pErk-positive cells in tmem2hu5935 mutant embryos compared with siblings at 22 hpf during primary sprouting and 32 hpf during secondary sprouting (Figures 1F and 1H). Furthermore, fluorescence intensity of the cells that were positive for pErk (quantified relative to pErk-positive neural tube axon body intensity in the same embryos; neurons were not significantly different between sibling and mutant embryos [data not shown]) was significantly reduced in tmem2hu5935 mutant embryos in both the DA (at 22 hpf) and cardinal vein (at 32 hpf) compared with siblings (Figures 1G and 1I). To investigate whether the reduction in Vegf signaling was a consequence of altered gene expression within the Vegf pathway, we examined tmem2 mutants for Vegf pathway components by ISH (Figures S1H and S1I) and microarray (data not shown) and found no evidence for tmem2/Vegf pathway crossregulation at the level of gene expression. Overall, these data demonstrate that Vegf signaling is reduced in the tmem2hu5935 mutant context and that this is occurring independent of transcriptional expression for key genes in the Vegf pathway. Given previous studies of tmem2 phenotypes (Smith et al., 2011; Totong et al., 2011), this may suggest Tmem2 functions at the level of growth factor signaling and plausibly might equate to a general defect in growth factor signaling. Beyond VEGF signaling, BMP signaling plays a role in angiogenesis in contexts other than ISV sprouting (Wiley et al., 2011). Using the Tg(hsp70:nog3)fr14 line to inhibit Bmp signaling, and by examination of phosphorylated Smad1/5/8 (pSmad1/5/8; the downstream effectors of Bmp signaling), we found no evidence for a role for Tmem2 in modulating the BMP pathway (Figure S2). These findings are in agreement with Bmp signaling being necessary for ventral angiogenesis from the caudal vein plexus

Figure 1. Tmem2 Is Required for Angiogenesis and Vegf Signaling (A) Lateral composite views of sibling and tmem2hu5935 mutant embryos crossed with the Tg(kdrl:mCherry) line (white) show a failure to complete angiogenesis at both arterial (28 hpf) and venous (50 hpf) sprouting stages. Yellow and white asterisks indicate the absence of sprouts in the mutants. Scale bar, 100 mm for whole embryo and 25 mm for zoom. (B) Column scatterplots depicting the number of segments with T-branching arterial ISVs per embryo at 28 hpf (n = 17 siblings, n = 29 mutants). (C) Scatterplots depicting the number of segments with venous ISVs per embryo at 50 hpf (n = 10 siblings and n = 11 mutants). (D and E) Maximum projection for whole-mount pErk immunofluorescence staining (white and heatmap) in sibling and tmem2hu5935 mutants on the Tg(kdrl:EGFP) background (green) at (D) 22 hpf and (E) 32 hpf. Average projections of fluorescence intensity for pErk are represented by heatmaps, where blue=low and red=high. Scale bar, 10 mm. pErk-positive cells indicated with white arrowheads; open arrowheads depict staining in corpuscle of Stannius. (F and G) Quantification of pErk-positive endothelial cells across four somites showed (F) significantly fewer pErk-positive endothelial cells in tmem2hu5935 mutant embryos compared with siblings at 22 hpf during primary sprouting (n = 8 sibling, n = 8 mutants) and (G) significantly lower fluorescence intensity (normalized to neuronal expression) of pERK-positive endothelial cells in mutants (n = 37) compared with siblings (n = 57) at 22 hpf. (H and I) Similarly, (H) there were significantly fewer pERK-positive endothelial cells in the PCV across eight somites of mutants (n = 8) at 32 hpf compared with siblings (n = 8; p < 0.01) and (I) significantly lower fluorescence intensity of pERK-positive cells in mutants (n = 20) compared with siblings (n = 52) at 32 hpf. DA, dorsal aorta; PCV, posterior cardinal vein. **p < 0.01, ***p < 0.001, ****p < 0.0001. Error bars represent ±SD.

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Figure 2. Tmem2 Expression Is Enriched in the Vasculature, the C Terminus Is Extracellular, and a Secreted C-Terminal Fragment Can Rescue the Angiogenesis Defect (A) ISH of the zebrafish trunk showing tmem2 expression is ubiquitous at 24 hpf and becomes enriched in venous endothelium at 32 hpf. (A0 ) tmem2 expression is also observed in angiogenic sprouts (black arrows). (legend continued on next page)

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but not dorsal sprouting angiogenesis from the PCV (Wakayama et al., 2015; Wiley et al., 2011). Tmem2 Is Expressed Ubiquitously during Primary Sprouting and Enriched in the Vasculature during Secondary Sprouting To examine where tmem2 is expressed at the onset of these phenotypes, in situ hybridization was performed at 24 and 32 hpf in wild-type embryos. Consistent with previous findings (Smith et al., 2011; Totong et al., 2011), tmem2 was expressed ubiquitously up to and during arterial ISV sprouting stages (24 hpf; Figure 2A). After this time, tmem2 becomes enriched in venous ECs of the cardinal vein, venous sprouts, and the caudal vein plexus (Figures 2A and 2A0 ). Together, these data demonstrate that tmem2 is expressed at the time of primary and secondary sprouting. To further determine protein localization and protein dynamics for Tmem2 in relation to angiogenesis, we generated a bacterial artificial chromosome (BAC) transgenic line by replacing the tmem2 stop codon with an mCherry cassette, creating a Tmem2-mCherry fusion protein (Tg(tmem2BAC: tmem2-mCherry)uq3ks). To determine whether this expression represents functional protein, we crossed it to the mutant and looked for phenotypic rescue. Embryos sorted for Tmem2mCherry expression had a wild-type phenotype, whereas 22% of mCherry-negative embryos from the same clutch developed the tmem2hu5935 mutant phenotype (Figure S3). Hence, this tool permits examination of functional Tmem2-mCherry fusion protein localization during development. To examine in more detail the dynamic changes in Tmem2 expression between 24 and 48 hpf, we performed live confocal imaging of Tg(tmem2BAC:tmem2-mCherry)/Tg(fli1a: EGFP)y1 transgenic embryos. Tmem2-mCherry protein localizes ubiquitously at 24 hpf but was progressively cleared from the trunk as angiogenesis proceeded (Figure 2B and Movie S2). By 48 hpf, Tmem2-mCherry became enriched in the cardinal vein, caudal vein, and venous sprouts as well as other tissues such as the lateral line primordium (Figures 2B and 2C, Movie S2). To determine the subcellular localization of Tmem2, we imaged Tg(tmem2BAC:tmem2-mCherry);Tg(fli1a:EGFP) transgenic embryos at high magnification in developing venous sprouts. We observed mCherry fluorescence adjacent to cytoplasmic GFP from the Tg(fli1a:EGFP) reporter, consistent with Tmem2 residing at the plasma membrane (Figure 2C). Overall, these

observations show that Tmem2 protein localization changes dynamically as development proceeds and is intimately associated with vascular endothelium, particularly the venous ECs, during sprouting. Tmem2 C-Terminus Extracellular Function Is Sufficient for Angiogenesis, and Tmem2 Can Act Cell Autonomously from Endothelial Cells Tmem2 contains a single-pass transmembrane domain, with a large C terminus (92% of the 1,390 amino acid protein) bioinformatically predicted to reside outside the cell (Hogan et al., 2003). To confirm the orientation of membrane-localized Tmem2, we transfected mammalian cells with a C-terminally tagged Tmem2-GFP fusion construct and subjected cells to a permeabilization assay. Unpermeabilized cells transfected with GFP only showed minimal fluorescence using a red fluorescent probe against GFP. Upon permeabilization, GFP detection was markedly increased. In contrast, Tmem2-GFP transfectants showed distinct red fluorescence at the plasma membrane both with and without permeabilization, demonstrating that the C terminus of Tmem2 projects extracellular to the plasma membrane (Figure S4A). A recognized phenomenon of membrane-tethered proteins is ectodomain shedding (Hayashida et al., 2010). The orientation of the majority of Tmem2 into the extracellular space, combined with the dynamic localization of protein around migrating ECs (Movie S2 and Figures 2B and 2C), led us to investigate whether Tmem2 may be a candidate for cleavage of the C-terminal domain. To test this, western blot analysis was performed on Tg(tmem2BAC:tmem2-mCherry) embryo lysates during relevant angiogenesis stages (19, 22, 28, and 32 hpf). Blotting for the C-terminal mCherry-tag showed bands at approximately 180 kDa in size (estimated full-length size of Tmem2-mCherry fusion protein) as well as smaller-sized bands that changed in proportion over developmental time. These were not seen in mCherry-only control lysates (Figure 2D) and are thus consistent with Tmem2 being cleaved and modified outside the cell. We next tested which regions of Tmem2 were necessary for its activity during angiogenesis. Given the localization and fragmentation of the C terminus, we tested a C-terminal fragment for activity. We generated an N-terminally truncated C-terminal-only mutant form of Tmem2 (termed c-tmem2) and the same truncation mutant with a synthetic secretion sequence (sc-tmem2) (Anisimov et al., 2009). mRNA injections into tmem2hu5935 mutants

(B) Lateral trunk views of Tg(tmem2BAC:tmem2-mCherry) embryos showing comparable expression with tmem2 RNA. Right-side panels showing overlay with Tg(fli1a:EGFP). Tmem2-mCherry protein was observed in angiogenic sprouts (white asterisks; white arrowheads depict iridophore autofluorescence). (C) Single z scans of two venous sprouts showing Tmem2-mCherry enrichment. Closer analysis (right panels) indicate mCherry tag is extracellular (white arrowheads). Scale bar, 10 mm and 5 mm for zoom. (D) Western blot analysis for mCherry showing bands consistent with full-length Tmem2-mCherry protein (red asterisk) as well as variable-sized fragments that form over developmental time. Staged Tg(kdrl:mCherry) lysates are controls. (E) Column scatter graph representing the number of ISVs at 48 hpf per embryo in uninjected siblings (n = 13) and tmem2hu5935 mutants (n = 89) as well as tmem2hu5935 mutants injected with full-length tmem2 mRNA (yellow squares; n = 49), c-tmem2 mRNA (gray triangles; n = 51), and sc-tmem2 mRNA (red diamonds; n = 66). (F) Schematic of Tmem2 protein and blow-up of the region where three newly identified tmem2 mutant alleles derived from forward genetic screens are located (tmem2hu4800, tmem2uq1ks, and tmem2uq2ks) as well as mutations generated for analysis in (G). (G) Scatterplots representing the number of ISVs at 48 hpf per embryo in uninjected siblings (n = 25), tmem2hu5935 mutants (n = 68), and tmem2hu5935 mutants injected with full-length tmem2 mRNA (yellow squares; n = 38) or mRNA of the PbH1 domain mutants tmem2W429A (gray triangles; n = 43), tmem2R439A (red triangles; n = 17), and tmem2F608A (blue diamond; n = 14), where none of the PbH1 mutants showed capability to rescue. ****p < 0.0001; ns, not significant. Error bars represent ±SD.

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showed that full-length tmem2 was capable of rescuing angiogenesis (Figure 2E). c-tmem2 mRNA did not rescue angiogenesis in tmem2hu5935 mutants; however, sc-Tmem2 was sufficient to rescue the tmem2hu5935 angiogenesis defect to the same extent as full-length tmem2 (Figure 2E) indicating sufficiency for the C-terminal extracellular protein during angiogenesis. Next, we characterized another previously reported tmem2 mutant allele, tmem2hu4800 (Smith et al., 2011), which also shows angiogenesis defects. The molecular lesion has not been previously reported, and we found that this mutation results in an amino acid substitution V426E in one of the C-terminal PbH1 domains (Figure 2F), a conserved domain predicted to be involved in extracellular cleavage of polysaccharide substrates (Mitchell et al., 2015). To test further whether these PbH1 domains were essential for Tmem2 function, we performed site-directed mutagenesis on important residues within the PbH1 domains. Aromatic and charged residues are conserved across homologous proteins possessing PbH1 domains and are needed for catalytic activity (Mayans et al., 1997). We generated three different mutant constructs with single amino acid substitutions to alanine residues (W429A, R434A, and F608A; Figure 2F) to test if they are required for Tmem2 activity. mRNA injection into tmem2hu5935 mutant embryos showed that, while the full-length tmem2 mRNA was capable of rescuing, no significant difference in the number of ISVs was observed between uninjected mutant embryos compared with tmem2W429A, tmem2R434A, or tmem2F608A mRNA-injected embryos (Figure 2G). These data demonstrate that the C terminus of Tmem2 is sufficient for angiogenesis when secreted into the extracellular space and that single missense substitutions in PbH1 domains abolish function. We hypothesized that if extracellular activity of Tmem2 regulates angiogenesis, then Tmem2 function would need to occur in close proximity to sprouting vessels, if not cell autonomously. To determine whether endothelial expression of tmem2 in the tmem2hu5935, mutant background was sufficient to permit angiogenesis, Tg(fli1ep:GalFF); Tg(uas:tmem2-gfp) lines were generated in the tmem2hu5935; Tg(kdrl:mCherry) background. EC autonomous expression of Tmem2-GFP was sufficient to generate significantly more ISVs in mutant embryos compared with embryos negative for Tmem2-GFP (Figure S4B). Hence, a source of Tmem2 local to sprouting vessels is sufficient for angiogenesis. HA Is Specifically Increased in tmem2hu5935 Mutants Given C-terminal localization and function of Tmem2, combined with the role of the PbH1 domains in degradation of extracellular proteins, we hypothesized that Tmem2 may play a role in ECM turnover. The closest homolog of Tmem2, Kiaa1199, is proposed to degrade HA (Yoshida et al., 2013), hence, to investigate whether HA was perturbed in tmem2hu5935 mutants, we generated a fluorescent reporter for HA localization. Similar to a previously described approach in cultured cells (Zhang et al., 2004), we fused the HA-binding domain of mouse Neurocan to GFP (termed here ssNcan-GFP) to examine HA localization (Figure 3A). In mRNA-injected embryos, we observed ssNcan-GFP accumulation in regions consistent with extracellular ECM (Figure 3B), at the dorsal side of the DA and around the PCV (Figure 3B and Movie S3). To confirm this was bone fide HA localization, we performed careful controls. Firstly, we generated GFP mRNA with a synthetic secretion sequence (ss-GFP) and found that while both ss-GFP 128 Developmental Cell 40, 123–136, January 23, 2017

and ssNcan-GFP localized to the somite boundaries, only ssNcan-GFP concentrated at perivascular locations (Figure S5A). Secondly, we injected the HA-degrading enzyme, HA’se, into embryos expressing either ss-GFP or ssNcan-GFP. We saw a significant decrease in ssNcan-GFP expression in embryos 6 hr after HA’se injection that was not observed in the ss-GFP expressing embryos (Figures S5B and S5C). Upon injection into tmem2hu5935 mutants, dramatically increased ssNcan-GFP was observed around the major vessels compared with sibling controls (Figure 3C and Movies S3 and S4). To quantify this, fluorescence intensity of ssNcan-GFP around the major vessels was measured in sibling and tmem2hu5935 mutant embryos. Although no difference in ssNcan-GFP levels was detectable at 22 hpf (prior to arterial sprouting; data not shown), by 28 hpf, significantly higher fluorescence was observed in mutant embryos compared with siblings (Figure 3D). Importantly, independent direct histochemical staining for HA confirmed this increase in HA around the major vessels (Figure 3E). To investigate whether this increase was a general effect on the ECM or specific to HA, we analyzed fibronectin (Fn). Unlike HA, we saw no difference in Fn fluorescence levels during angiogenesis (Figures 3F and 3G). This was confirmed by western blot analysis whereby no difference in Fn levels was observed at 26 or 50 hpf between sibling and mutant embryos (data not shown). Degradation of HA in tmem2hu5935 Mutants Rescues the Angiogenesis Phenotype Given that tmem2hu5935 mutant embryos have increased HA, we hypothesized that, if this was the cause of the angiogenesis defect, treatment with HA’se to depolymerize endogenous HA should rescue the tmem2hu5935 mutant phenotype. To test this, tmem2hu5935 embryos were injected between 16 and 17 hpf with HA’se at sites around the location of the developing ISVs. Embryos were imaged between 38 and 42 hpf for ISV quantification and subsequently genotyped (Figure 4A). Remarkably, the angiogenesis defect in tmem2hu5935 mutant embryos was significantly rescued following HA’se injection (Figures 4B–4D). Notably, the cardiac defect was not rescued in tmem2hu5935 mutants upon HA’se injection and, instead, all embryos injected with HA’se developed marked cardiac edema and cardiac morphogenesis defects (Figure S5D and Movies S5 and S6). Furthermore, no rescue of the cardiac defect was observed even when careful dose response experiments were performed, including concentrations where the angiogenesis defects were no longer rescued (Figure S6). The phenotype of the tmem2hu5935 mutant was clearly distinct from HA’se-injected embryos, indicating that a fine balance and spatially controlled levels of HA is necessary for normal cardiac development (Movies S5 and S6). Importantly, HA’se injection did not increase angiogenesis or induce ectopic sprouting, indicating that HA degradation is not instructive but permissive for angiogenesis. o-HA Promotes Vegf Signaling during Angiogenesis HA’se rescue experiments demonstrate that excess HA is causal of the tmem2hu5935 mutant angiogenesis phenotype. Whether this is due to a physical impediment from excess space-filling polymeric HA or whether degraded HA molecules, known as o-HA, are permissive for angiogenesis was unclear (Pardue et al.,

Figure 3. Hyaluronic Acid Is Increased in Tmem2 Mutants (A) Schematic representing the strategy for the HA biosensor. mRNA encoding the HA-binding domain of mouse Neurocan fused to GFP (ssNcan-GFP) was injected into 1- to 2-cell stage embryos, grown, and imaged at the relevant time points. (B) Lateral views and zoom-in of 30 hpf Tg(kdrl:mCherry) embryos injected with ssNcan-GFP. White arrowheads indicate expression surrounding the axial vessels in the trunk and black arrowhead expression in the heart. Scale bar, 100 mm for whole embryo and 25 mm for the zoom. (C) ssNcan-GFP is increased and expanded in tmem2hu5935 mutants compared with siblings around the major axial vessels during sprouting angiogenesis. Maximum intensity projections. Scale bar, 15 mm. (D) Quantification showing significantly higher fluorescence intensity of ssNcan-GFP in mutant (n = 15) compared with sibling (n = 15) embryos. (E) Cross-sections of embryos histochemically stained for HA, counterstained for neural red, show increased HA accumulation around the major vessels of 48 hpf tmem2hu5935 mutant. (F) Maximum intensity projections of immunofluorescence staining for fibronectin in sibling and tmem2hu5935 mutants, where blue=low and red=high. Scale bar, 15 mm. (G) Quantification of fibronectin fluorescence intensity showing no significant difference between mutant (n = 8) and sibling (n = 7) embryos. DA, dorsal aorta; PCV, posterior cardinal vein; NC, notochord. ****p < 0.0001. Error bars represent ±SEM.

2008). To distinguish between these possibilities, o-HA was generated by digesting commercially purchased macromolecular HA (10 kDa in size) using HA’se. Digestion was followed by heat inactivation to ensure no enzymatic contribution of HA’se to phenotypic rescue. Subsequently, o-HA was injected into 16- to

18-somite-staged embryos from a tmem2+/hu5935 incross and embryos were imaged at 38–42 hpf, followed by genotyping (Figure 4A). o-HA significantly rescued the tmem2hu5935 mutant angiogenesis phenotype, while undigested 10 kDa HA (data not shown) or heat-inactivated HA’se (heat inact) did not (Figures Developmental Cell 40, 123–136, January 23, 2017 129

Figure 4. Depolymerized HA Rescues the tmem2 Mutant Phenotype (A) Experimental design for hyaluronidase (HA’se) or o-HA injection. (B) Lateral views of entire trunks of sibling and tmem2hu5935 mutant embryos post-injection and imaged at 38–42 hpf. Yellow asterisks indicate the absence of sprouts in uninjected mutants. Representative images demonstrate phenotypic rescue of tmem2hu5935 mutants following injection with HA’se or o-HA. Scale bar, 150 mm. (C) Scatterplots summarizing unaffected siblings in the uninjected (n = 23), HA’se injected (n = 23), o-HA injected (n = 13) or hyaluronidase heat-inactivated (HA’se inact) (n = 26) embryos. (D) Uninjected tmem2hu5935 mutants (n = 19) have significantly fewer ISVs that uninjected siblings. Mutants injected with HA’se (n = 17) or o-HA (n = 14) have significantly rescued phenotypes compared with uninjected mutants. Heat-inactivated HA’se injected mutants were not significantly different from uninjected mutants. ****p < 0.0001; ns, not significant. Error bars represent ±SEM.

4B–4D). This demonstrates that the defect in ISV sprouting was not caused by an accumulation of HA but, rather, an absence of o-HA, which is necessary to permit developmental angiogenesis. To confirm that the reduction in Vegf signaling in tmem2hu5935 mutants was caused by a deficiency of HA turnover, wholemount IF for pErk signaling was performed on sibling and mutant embryos following dextran control or HA’se injection. Embryos were injected at 16–18 somites, fixed 3 hr post-injection, stained, and imaged for pErk induction. pErk levels were reduced in dextran-injected mutant embryos but were rescued to wildtype levels in mutants following HA’se injection (Figures 5A– 5D). To next investigate whether o-HA acts via the Vegf pathway or independently, we performed a series of epistasis experiments. Firstly, we injected HA’se and o-HA into vegfaa morpholino (MO) or kdr/kdrl double MO-injected embryos that have a total block in Vegf signaling. Neither HA’se nor o-HA could rescue either of the morphant phenotypes (Figures 5E and 5F), indicating that HA acts upstream of, or requires intact Vegf 130 Developmental Cell 40, 123–136, January 23, 2017

receptor signaling. Next, we investigated whether o-HA was capable of overcoming dampened Vegf signaling by using suboptimal concentrations of the Vegf receptor inhibitor SU5416 (Figures 5G and 5H). Interestingly, concentrations of SU5416 titrated to cause a mild angiogenesis defect were rescued by injection of o-HA (Figure 5H). However, a severe block in Vegf signaling (by using higher concentrations of SU5416) was not rescued by o-HA (Figure 5I), as was observed in the kdr/kdrl morphant scenario. Together, these data indicate that o-HA can enhance angiogenesis by augmenting Vegfa/Kdr/Kdrl output. HA’se Treatment in tmem2hu5935 Mutants Lacking Intersegmental Vessels Induces Angiogenesis to Restore Trunk Vasculature To further examine the capability of HA’se and HA turnover to control angiogenesis and Vegf signaling, we tested a model of Vegf-signaling-mediated restoration of failed angiogenesis. To investigate whether a repression of Vegf signaling, followed by

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derepression, could control the temporal progression of angiogenesis in the zebrafish trunk, we incubated embryos in SU5416 from the 16-somite stage until 30 hpf. This led to a complete block in primary angiogenesis, which occurs during this period of development. The inhibitor was then replaced or washed to DMSO for a further 18 hr before embryos were imaged and quantified at 48 hpf. We observed that embryos treated with a continuous dose of SU5416 formed no ISVs (neither arterial nor venous) but ISV sprouting was restored in embryos when SU5416 had been removed at 30 hpf, 8 hr after sprouting angiogenesis should have occurred under normal wild-type conditions (Figures 6A–6C). Given this capacity for Vegf signaling to re-initiate failed angiogenesis, we asked if HA’se and HA modulation could do the same. tmem2hu5935 mutant embryos were selected for displaying a strong loss of trunk vasculature at 28 hpf and injected with HA’se. As with removal of SU5416, we observed a strong rescue of arterial sprouting by 48 hpf in this assay, suggesting restoration of Vegfr signaling upon degradation of HA (Figures 6D–6F). This same phenomenon was also observed when injecting o-HA (Figures 6D–6F). To confirm that restoration of angiogenesis from HA’se treatment in tmem2hu5935 mutant embryos is indeed contingent on Vegf signaling, we repeated these experiments and added either DMSO or the inhibitor SU5416 at the time of HA’se treatment (28 hpf; Figure 7A). In accordance with Tmem2/HA acting via Vegf signaling, the restoration of angiogenesis did not occur in the presence of SU5416 (Figure 7B). Overexpression of Vegfc Can Rescue Angiogenesis in tmem2 Mutants To determine whether an increase in Vegf signaling was sufficient to rescue the tmem2 mutant phenotype, we took advantage of a previously published Vegfc overexpression line (Koltowska et al., 2015a). Vegfc has been shown to signal through tip-cell-expressed Vegfr3 (Flt4) as well as Vegfr2 and to modulate primary angiogenesis phenotypes in a number of studies (Gore et al., 2011; Hogan et al., 2009; Koltowska et al., 2015a; Villefranc et al., 2013). We crossed tmem2+/hu5935 carriers

to Tg(hsp70:gal4) and Tg(10xuas:vegfc)uq2bh transgenic lines. The combined presence of these two transgenes permits temporally inducible Vegfc expression following heat shock. Progeny from crosses between Tg(hsp70:gal4)/tmem2+/hu5935 and Tg(10xuas:vegfc)/tmem2+/hu5935 were collected, heat shocked at 28 hpf, and then imaged at 3 days post-fertilization. Sibling embryos overexpressing Vegfc presented with the characteristic Vegfc overexpression phenotype (ectopic sprouting of venous ISVs but no impact on the local arteries (Koltowska et al., 2015a; Le Guen et al., 2014)). tmem2hu5935 mutants did not shown any increased venous sprouting but showed a clear and significant rescue of the arterial ISV phenotype upon Vegfc overexpression (Figures 7C and 7D). These data further confirm that loss of angiogenesis in tmem2hu5935 mutants occurs due to reduced Vegf signaling in arterial ECs. DISCUSSION Here, we uncover a role for Tmem2 as a regulator of angiogenesis. Our studies indicate that Tmem2 localizes to EC membranes and that the extracellular domain is sufficient for function in angiogenesis when secreted from the cell. We find that Tmem2 is necessary to reduce perivascular HA but not to generally degrade the ECM. Importantly, enzymatic degradation of this excess HA in tmem2 mutants rescues angiogenesis. While the increase in HA could cause a physical block, in fact we found that o-HA, when exogenously introduced into mutant embryos, also rescues angiogenesis. Our findings suggest that normal depolymerization of HA enhances Vegf signaling to control angiogenesis in vivo during embryonic development. Tmem2 is a type II transmembrane domain protein and the functional C terminus resides extracellularly when localized at the plasma membrane. The C terminus is annotated to contain PbH1 domains, a structure imputed with catalytic activity for polysaccharide substrates (http://www.ebi.ac.uk/ interpro/entry/IPR006626). The closest homolog of Tmem2, Kiaa1199 (also called Tmem2L), contains these PbH1 domains and has been shown to exhibit specific HA binding

Figure 5. Rescue of the tmem2hu5935 Mutant Phenotype by Degrading HA Requires the Vegf Pathway (A) Whole-mount pErk immunofluorescence staining (white) in sibling and tmem2hu5935 mutants on the Tg(kdrl:EGFP) background (green) at 22 hpf, injected just 3 hr prior with dextran. Scale bar, 10 mm. (B) Graph depicting the significant reduction in fluorescence intensity of pErk-positive endothelial cells in mutants (n = 43) versus sibling embryos (n = 56) following dextran injection. (C) Whole-mount pErk immunofluorescence staining in sibling and mutant embryos 3 hr after HA’se injection. Scale bar, 10 mm. (D) Scatterplots showing no significant difference in fluorescence intensity of pErk-positive endothelial cells in mutants (n = 90) versus sibling embryos following HA’se injection (n = 89). (E) Untreated vegfaa morphants (n = 8) had fewer ISVs compared with uninjected controls (n = 10) and this reduction in sprouting was not significantly different following HA’se (n = 8), o-HA (n = 11) or HA’se inact treatment (n = 9). (F) Likewise, untreated kdr/kdrl double morphants (n = 25) had no detectable ISVs compared with normal uninjected controls (n = 11), and this loss of sprouting was not significantly different following HA’se (n = 17), o-HA (n = 20) or HA’se inact treatment (n = 20). (G) Design of rescue experiments for low and high dose SU5416 treatments shown in (H) and (I). (H) Scatterplot for DMSO-treated embryos injected with HA’se (n = 27), o-HA (n = 29), or HA’se inact (n = 23) showing no difference from uninjected (n = 8) embryos. Low-dose (0.8 mM) SU5416 mildly but significantly impaired ISV sprouting in uninjected embryos (n = 21). SU5416-treated embryos injected with HA’se (n = 40) or HA’se inact (n = 46) were not significantly different from uninjected but o-HA injected embryos (n = 53) were significantly rescued (p < 0.0001). (I) At a higher dose (1 mM SU5416), DMSO treatment again had no effect on embryos not injected (n = 8), injected with HA’se (n = 12), o-HA (n = 16), or HA’se inact (n = 11). At the higher dose, all treatments (uninjected [n = 13], HA’se [n = 12], o-HA [n = 18] and HA’se inact [n = 11]) were significantly different from DMSO with no rescue in any of these conditions observed. For all scatterplots uninjected (white circles), HA’se injected (yellow squares), o-HA injected (gray triangles), HA’se inact (red diamonds). **p < 0.01, ****p < 0.0001. Error bars represent ±SD.

132 Developmental Cell 40, 123–136, January 23, 2017

Figure 6. ISV Sprouting Can Be Restored by Derepressing Vegf Signaling or Treating with HA’se/o-HA (A) Experimental design to examine the effect of repressing and derepressing Vegf signaling on ISV sprouting using the inhibitor, SU5416. (B) Lateral view images of Tg(kdrl:mCherry) embryos treated according to scheme in (A). Scale bar, 25 mm. (C) Scatterplots showing SU5416 treatment from 16 somites to 48 hpf causes complete failure of ISV formation (n = 10) compared with normal sprouting after similar DMSO treatment (n = 13). By contrast, removal of SU5416 at 30 hpf, after ISV sprouting has failed, restores ISV sprouting (n = 10). Addition of SU5416 from 30 hpf had no significant effect on ISV number (n = 10). (D and E) Lateral view images showing the phenotype of sibling or tmem2hu5935 mutant embryos at (D) 28 hpf pre-injection and (E) 42 hpf following HA’se or o-HA injection. Scale bar, 100 mm. (F) Scatterplot demonstrating no difference in number of ISVs in sibling embryos injected with dextran (n = 11), HA’se (n = 10), or o-HA (n = 8) at 28 hpf and measured at 42 hpf; however, tmem2hu5935 mutant embryos have significant restoration of ISV number following injection of HA’se (n = 9) or o-HA (n = 10) compared with dextran control (n = 20). ****p < 0.0001. Error bars represent ±SD.

and degrading capability (Yoshida et al., 2013). Based on sequence similarity and the functional evidence showing that reduced HA degradation is responsible for the Tmem2 mutant phenotype, it seems highly likely that Tmem2 is capable of performing the same direct role. Tmem2 was first identified for its role in cardiac morphogenesis, whereby tmem2 mutants have a cardiac looping defect resulting from an expanded atrioventricular canal (Smith et al., 2011; Totong et al., 2011). This was shown to correlate with increased cardiac jelly (ECM between the myocardium and endocardium), while in the maternal zygotic mutant, cardiac progenitors fail to migrate for cardiac disc fusion (Smith et al., 2011; Totong et al., 2011). These phenotypes are both associated with abnormal ECM or HA (Garavito-Aguilar et al., 2010; Trinh and

Stainier, 2004). More recently, Ryckebusch et al. (2016) have described a role for Tmem2 in muscle formation and attributed this to a disruption in the cell-matrix interaction. Although HA was not analyzed in this context, the correlation of the muscle phenotype with disturbed ECM organization is consistent with what we report here. Given our current data, it will be interesting to determine the relative contribution of the ECM as an active source of signaling molecules and growth factor regulation rather than its roles as a passive substrate during cardiovascular development. A chick chorioallantoic membrane assay previously implicated a pro-angiogenic role for o-HA (West et al., 1985). Whether this pro-angiogenic capability controls developmental processes and how it does so mechanistically remains Developmental Cell 40, 123–136, January 23, 2017 133

Figure 7. Restoration of ISV Sprouting by HA’se Requires Vegf Signaling, and Vegf Signaling Is Sufficient to Rescue the tmem2 Mutant Phenotype (A) Experimental strategy for HA’se and Vegf pathway epistasis experiments. (B) Scatterplots demonstrating that, as observed in Figure 6D, the significant reduction in ISV number in tmem2 mutant embryos (n = 14) compared with siblings (n = 11) is rescued when HA’se is injected into mutants (n = 16) compared with siblings in the presence of DMSO (n = 18). When embryos are incubated with SU5416 from the time of injection, tmem2 mutants (n = 21) have significantly fewer ISVs than siblings (n = 10) when injected with dextran but tmem2 mutants (n = 21) also have significantly fewer ISVs than siblings (n = 9) when injected with HA’se, demonstrating that rescue by HA’se requires intact Vegf signaling. (C) Lateral view images of sibling and tmem2hu5935 mutants on the Tg(kdrl:EGFP) background either negative for Tg(hsp70l:gal4)/Tg(10xuas:vegfc) (control) or positive for Tg(hsp70:gal4)/Tg(10xUAS:vegfc) (vegfc O/E). Scale bar, 100 mm. (D) Scatterplot showing number of ISVs in sibling (open circles; n = 10) and tmem2hu5935 mutants (red squares; n = 22) negative for vegfc O/E, showing a significant reduction in ISV number. tmem2hu5935 mutants positive for vegfc O/E (green triangles, n = 10) had significantly more ISVs than tmem2hu5935 mutants negative for vegfc O/E and were not significantly different from siblings negative for vegfc O/E. **p < 0.01, ****p < 0.0001; ns, not significant. Error bars represent ±SD.

unknown. Here, we provide evidence that depolymerized HA is needed to permit normal Vegf signaling through the Kdr/Kdrl receptor to activate intracellular pErk. This requirement is observed for both Vegfa-regulated arterial sprouting as well as Vegfc-regulated venous sprouting stages. Furthermore, we demonstrate that overexpression of Vegfc is sufficient to rescue primary arterial sprouting, which is consistent with previous data showing that Vegfc is capable of phosphorylating KDR/VEGFR2 and enhancing phenotypes in Kdrl mutant zebrafish (Joukov et al., 1997; Villefranc et al., 2013). While dependency on the Flt4 receptor was not formally tested here, Flt4 is dispensable for primary angiogenesis in zebrafish (Hogan et al., 2009; Kok et al., 2015) and its presence was not sufficient to compensate for Kdr/Kdrl loss upon supplementation with o-HA in this study. The additional level of control over Vegf signaling reported here is consistent with strict control of this pathway in the embryo. Heterozygous Vegfa mutant mice are embryonic lethal (Carmeliet et al., 1996), and overexpression of Vegf ligands causes severe hyperbranching or vasculogenesis de134 Developmental Cell 40, 123–136, January 23, 2017

fects resulting in embryonic lethality (Le Guen et al., 2014; Liang et al., 2001). A previous study has shown in vitro evidence that o-HA can induce Vegfa transcriptionally; however, our expression data are not consistent with this (Figure S1) (Rodgers et al., 2006). In fact, our data show that o-HA can restore Vegf signaling without causing overexpression-associated phenotypes, demonstrating a permissive role for Tmem2 and o-HA. Exactly how o-HA achieves this mechanistically is not known. Our data demonstrate the effect is occurring upstream of receptor signaling. It is therefore tempting to speculate that o-HA may be involved in liberating Vegf molecules from the ECM, assisting in ligand dimerization, or facilitating ligand presentation to receptors; however, further studies are now needed. Of note, HA modulation is already used in the clinic during ocular surgery (Kleinberg et al., 2011), subcutaneous infusions (Wasserman, 2014), cosmetic surgery (Kim and Sykes, 2011), and osteoarthritis treatment (Hunter, 2015). A deeper understanding of the molecular activity of HA and how it abrogates Vegf signaling may have repercussions on current therapeutics.

EXPERIMENTAL PROCEDURES Zebrafish Lines and Husbandry Animal work followed guidelines of the University of Queensland animal ethics committee. Transgenic lines used are detailed in Supplemental Experimental Procedures. Embryo Manipulation and Expression Analysis MO-kdr, MO-kdrl, and MO-vegfaa were used as previously described (Duong et al., 2014; Le Guen et al., 2014). For SU5416 treatments, embryos were dechorionated and incubated in E3 embryo medium containing 1% DMSO at the 15-somite stage. SU5416 concentrations used were 0.25–0.35 mM for suboptimal (Figure S1), 0.8 mM for low, and 1.0 mM for high dose concentrations (Figures 5, 6, and 7). For o-HA and HA’se treatments, embryos were embedded in 1% low-melting agarose and injected with 1 nL of reagents at three sites along the length of the embryo at 15- to 18-somite stages. For o-HA and HA’se preparation, see Supplemental Experimental Procedures. Genotyping and Construct Generation Full-length tmem2 cDNA (Totong et al., 2011) was used to generate c-tmem2 and sc-tmem2 cDNA and cloned into the pCS2+ vector. ssNcan-GFP and ss-GFP were cloned using the Gateway system. Human Tmem2 cDNA (from IMAGE clone 9007233) was cloned into the pEGFP-N1 (Promega). tmem2hu5935 genotyping was performed as previously described (Smith et al., 2011). Primers used for cloning are described in Supplemental Experimental Procedures. In Situ Hybridization, Histology, and Western Analysis ISH analysis was carried out as previously described (Smith et al., 2008). HA histochemistry staining was performed as previously described (Lagendijk et al., 2011). Probe details and the immunohistochemistry procedure are described in Supplemental Experimental Procedures. For western analysis, Tg(tmem2BAC:tmem2-mCherry) and Tg(kdrl:mCherry) embryos were collected in radioimmunoprecipitation assay buffer and run on 8%–15% gradient gels (Bio-Rad). Protein was transferred and blotted on polyvinylidene fluoride membranes (Bio-Rad) with antibodies against rabbit anti-dsRed (Clontech), and horseradish peroxidase-conjugated goat anti-rabbit (Invitrogen). gels were visualized using a Bio-Rad Gel Doc system. Imaging, Quantification, and Statistical Analysis Live and fixed embryos were mounted laterally in 1% low-melting agarose and imaged using a Zeiss LSM 710 FCS confocal microscope. Imaging for ISV quantification purposes and fluorescence intensity quantification for the HA biosensor was performed on a Zeiss Axiovert 200 spinning disc. Imaging for quantification of fluorescent intensities was performed on a Zeiss LSM 710 FCS confocal microscope. Images were processed using ImarisX64 7.70 and/or ImageJ 1.47 (NIH) software. For detailed information on quantification, see Supplemental Experimental Procedures. All statistical analyses were performed using t test (for two groups) or one-way ANOVA (for three or more groups) with Tukey multiple comparison analysis using Prism 6 software (GraphPad Software). SUPPLEMENTAL INFORMATION Supplemental Information includes Supplemental Experimental Procedures, six figures, and six movies and can be found with this article online at http:// dx.doi.org/10.1016/j.devcel.2016.12.017.

(FT110100496), B.M.H. in part by an ARC Future Fellowship (FT100100165), and an National Health and Medical Research Council (NHMRC)/National Heart Foundation Career Development Fellowship (1083811), M.F. by an NHMRC Australia Career Development Fellowship (1011242), and A.Y. by an NHMRC Research Fellowship (1044041). This research was supported in part by NHMRC project grants (1046028 and 1106800). Imaging was performed in the Australian Cancer Research Foundation’s Dynamic Imaging Facility at the Institute of Molecular Bioscience, established with the generous support of the Australian Cancer Research Foundation. We thank Debbie Yelon for sharing reagents, Katarzyna Koltowska and Sungmin Baek for assistance with pErk staining, Marah Heijkoop for technical assistance, and Kylie Georgas for graphic design. Received: June 15, 2015 Revised: October 18, 2016 Accepted: December 16, 2016 Published: January 23, 2017 REFERENCES Anisimov, A., Alitalo, A., Korpisalo, P., Soronen, J., Kaijalainen, S., Leppanen, V.M., Jeltsch, M., Yla-Herttuala, S., and Alitalo, K. (2009). Activated forms of VEGF-C and VEGF-D provide improved vascular function in skeletal muscle. Circ. Res. 104, 1302–1312. Camenisch, T.D., Spicer, A.P., Brehm-Gibson, T., Biesterfeldt, J., Augustine, M.L., Calabro, A., Jr., Kubalak, S., Klewer, S.E., and McDonald, J.A. (2000). Disruption of hyaluronan synthase-2 abrogates normal cardiac morphogenesis and hyaluronan-mediated transformation of epithelium to mesenchyme. J. Clin. Invest. 106, 349–360. Carmeliet, P., Ferreira, V., Breier, G., Pollefeyt, S., Kieckens, L., Gertsenstein, M., Fahrig, M., Vandenhoeck, A., Harpal, K., Eberhardt, C., et al. (1996). Abnormal blood vessel development and lethality in embryos lacking a single VEGF allele. Nature 380, 435–439. Duong, T., Koltowska, K., Pichol-Thievend, C., Le Guen, L., Fontaine, F., Smith, K.A., Truong, V., Skoczylas, R., Stacker, S.A., Achen, M.G., et al. (2014). VEGFD regulates blood vascular development by modulating SOX18 activity. Blood 123, 1102–1112. Fong, T.A., Shawver, L.K., Sun, L., Tang, C., App, H., Powell, T.J., Kim, Y.H., Schreck, R., Wang, X., Risau, W., et al. (1999). SU5416 is a potent and selective inhibitor of the vascular endothelial growth factor receptor (Flk-1/KDR) that inhibits tyrosine kinase catalysis, tumor vascularization, and growth of multiple tumor types. Cancer Res. 59, 99–106. Garavito-Aguilar, Z.V., Riley, H.E., and Yelon, D. (2010). Hand2 ensures an appropriate environment for cardiac fusion by limiting Fibronectin function. Development 137, 3215–3220. Gore, A.V., Swift, M.R., Cha, Y.R., Lo, B., McKinney, M.C., Li, W., Castranova, D., Davis, A., Mukouyama, Y.S., and Weinstein, B.M. (2011). Rspo1/Wnt signaling promotes angiogenesis via Vegfc/Vegfr3. Development 138, 4875–4886. Hayashida, K., Bartlett, A.H., Chen, Y., and Park, P.W. (2010). Molecular and cellular mechanisms of ectodomain shedding. Anat. Rec. 293, 925–937. Herbert, S.P., and Stainier, D.Y. (2011). Molecular control of endothelial cell behaviour during blood vessel morphogenesis. Nat. Rev. Mol. Cell Biol. 12, 551–564.

AUTHOR CONTRIBUTIONS

Hogan, M.C., Griffin, M.D., Rossetti, S., Torres, V.E., Ward, C.J., and Harris, P.C. (2003). PKHDL1, a homolog of the autosomal recessive polycystic kidney disease gene, encodes a receptor with inducible T lymphocyte expression. Hum. Mol. Genet. 12, 685–698.

J.D.A., A.K.L., B.M.H., and K.A.S. designed, performed, and analyzed experiments and co-wrote the manuscript. H.C., A.T., N.B., and J.B. designed and performed experiments. C.W., M.F., C.S., and A.Y. designed and analyzed experiments.

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ACKNOWLEDGMENTS

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A.K.L. was supported by a University of Queensland Postdoctoral Fellowship, K.A.S. by an Australian Research Council (ARC) Future Fellowship

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