The endosomal RIN2/Rab5C machinery prevents VEGFR2 degradation to control gene expression and tip cell identity during angiogenesis

We first depleted Rab5C from human umbilical vein endothelial cells (HUVECs) with shRNA pools (leading to ~ 90% reduction at the mRNA level; Fig S1A), and assessed the cell-surface levels of VEGFR2 by flow cytometry. Intriguingly, VEGFR2 surface levels in steady-state HUVECs (growing in the presence of VEGF) were consistently reduced two-fold in sh_Rab5C cells, compared to HUVECs expressing scrambled sequences as a control (sh_Ctrl) (Fig. 1a). The decrease in VEGFR2 was further confirmed by quantification of the total VEGFR2 protein levels by Western blotting (Fig. 1b). In contrast, VEGFR1 protein levels were not affected, suggesting that VEGFR1 is not regulated by Rab5C (Fig. 1b; Fig S1B). We then investigated if and how Rab5C regulates VEGFR2 endocytic recycling using flow cytometry. To test whether the internal pool of VEGFR2 can be recruited normally to the plasma membrane, we starved HUVECs for 30 min, a time-frame that allows mobilization of the internal VEGFR2 pool to the plasma membrane but is too short to induce strong changes in de novo protein synthesis. Starvation significantly increased the cell-surface levels of VEGFR2 compared to those in steady-state in sh_Ctrl but not sh_Rab5C cells, indicating that the recruitment from intracellular compartments was impaired (Fig. 1c, d). VEGF induced VEGFR2 endocytosis in both sh_Ctrl and sh_Rab5C cells, although in the latter the internalization was slightly reduced (Fig. 1c, e). The endosomal pool of VEGFR2 is protected from degradation in the absence of ligand, while VEGF stimulation results in rapid degradation [16, 22]. Therefore, we hypothesized that the reduced levels of VEGFR2 in Rab5C-depleted cells were due to increased degradation. To assess VEGFR2 degradation, HUVECs were deprived of growth factors overnight and then stimulated with VEGF, which resulted in a rapid decrease of up to 40% of VEGFR2 levels in 30 min (Fig. 1f), consistent with previous reports [16, 22]. Strikingly, VEGFR2 degradation was enhanced twofold in Rab5C-depleted cells, confirming our hypothesis that the reduction in VEGFR2 levels in these cells is caused by increased degradation (Fig. 1f). Moreover, inhibition of lysosomal proteases blocked VEGF-induced degradation, and nullified the differences in VEGFR2 degradation between sh_Ctrl and sh_Rab5C cells (Fig S1C). In HUVECs that were maintained in the absence of VEGF, VEGFR2 levels declined only modestly (20% over 120 min), and this was not significantly altered by depletion of Rab5C (Fig. 1g).

In summary, these results show that Rab5C protects the endosomal VEGFR2 pool from VEGF-induced lysosomal degradation.

We next investigated if and how Rab5C regulates VEGF signaling. For this purpose, sh_Ctrl and sh_Rab5C HUVECs were starved overnight and then stimulated with VEGF, whereafter AKT phosphorylation was assessed as a read-out for PI3-K-mediated VEGF signaling. VEGF triggered robust and persistent AKT phosphorylation within 10 min in sh_Ctrl cells, which was almost completely abolished in Rab5C-depleted cells, indicating that Rab5C is required for VEGF-induced PI3-K signaling (Fig. 2a). We also investigated VEGF-induced ERK-1/2 phosphorylation. In sh_Ctrl cells, robust induction of ERK-1/2 phosphorylation was observed within 5 min after VEGF addition and persisted for up to 30 min, while this was significantly reduced in sh_Rab5C cells (Fig. 2b).

We then assessed if and how the reduced VEGFR2 signaling observed upon depletion of Rab5C impacts on VEGF-regulated gene expression by mRNA sequencing analysis (RNA-seq). Differential expression was assessed by empirical Bayes analysis followed by correction of p values for multiple testing using the Benjamini–Hochberg false discovery rate (FDR), with a cut-off adjusted p value of 0.05. Using these criteria, the expression of 40 genes was significantly induced and 1 gene was repressed in sh_Ctrl cells after 1 h (t = 1) of VEGF stimulation, while after 4 h (t = 4) the expression of these genes was largely back to base-line levels (Table S1 and Fig. 2c). These kinetics are comparable with those found in previous reports [39‐41]. The induced target genes include well-known immediate VEGF targets, 58% of which encoding transcriptional regulators such as Krüppel-like factors, nuclear receptors of the NR4A family, and members of the Fos/Jun and early growth response protein families, also consistent with earlier findings (Table S1 and Fig. 2c) [39‐41]. Strikingly, expression of a large number of VEGF target genes (34 out of 41) was altered in sh_Rab5C cells (Fig. 2c), and geneset enrichment analysis revealed that at t = 1, expression of the ‘VEGF transcriptome’ was significantly different with respect to that in sh_Ctrl cells (p = 3.12*10^–4, FDR = 1.52*10^–2).

We then assessed the role of Rab5C in sprouting angiogenesis, using the fibrin bead assay [42, 43]. Quantification of the extent of sprouting after 48 h revealed that depletion of Rab5C strongly impaired sprouting, by affecting both the numbers of formed sprouts and their length (Fig. 2d–g). In addition, tip cells clearly formed multiple filopodia in the sh_Ctrl population, a process stimulated by VEGF signaling, whereas sh_Rab5C tip cells were on average more ‘blunted’ (Fig. 2d).

Together, these data show that Rab5C promotes endosomal VEGFR2 signaling and full induction of the immediate VEGF-induced transcriptome, and that Rab5C is required for efficient sprouting angiogenesis.

Because VEGF signaling and the expression of certain VEGF targets is important for the induction and maintenance of tip cells during sprouting angiogenesis, we next addressed whether the reduced sprouting observed in the absence of Rab5C is due to differences in VEGF-induced tip cell specification. For this purpose, we first investigated the expression of a panel of genes commonly associated with tip cell identity, including ANGPT2, APLN, DLL4, NID2, NRP2, PDGFB, TIE1, UNC5B, VEGFR2, and VEGFR3 [6, 7]. Intriguingly, depletion of Rab5C significantly suppressed the transcription of ANGPT2, APLN, DLL4, NID2, UNC5B, and VEGFR3, while the expression of NRP2, TIE1, and VEGFR2 was mildly but not significantly decreased (Fig. 3a). Of note, PDGFB expression appeared not to be regulated at all by Rab5C.

We then assessed tip cell formation in mosaic sprouting assays, by differentially labeling sh_Ctrl and sh_Rab5C cells using two distinct CellTracker dyes, mixing them in a 1:1 ratio, and determining the fraction of tip cells formed by each population after 48 h (Fig. 3b). The knockdown of Rab5C significantly reduced the numbers of tip cells in this assay (Fig. 3b). Similar results were obtained using a pool of mixed shRNAs (Fig. 3b), two different individual shRNAs (Fig S2A–C), and when the dyes were swapped between conditions (Fig S2B).

To address the role of Rab5C in an in vivo model system for sprouting angiogenesis, we next analyzed if Rab5C regulates intersegmental vessel (ISV) development in zebrafish. The zebrafish rab5c gene is highly homologous to its human counterpart, suggesting an important function in the vasculature [35, 37, 38]. We first generated a dominant-negative (DN) mutant Rab5C protein, carrying a single substitution (S35N) in the GTP-binding pocket (Fig S3A). DN Rab mutants sequester GEFs but fail to bind GTP, and therefore cannot be activated [44]. The mutant was fused to mCherry and expressed in HUVECs to test its ability to localize on endosomes. Wild-type (WT) Rab5C localized predominantly on endosomes, some of which were positive for EEA-1, while occasional distribution to the Golgi network was also observed, using Trans-Golgi Network (TGN)46 as a marker (Fig S3B). In contrast, the mutant localized predominantly at the Golgi and hardly on endosomes (Fig S3B). We then cloned the mCherry-tagged WT or DN human RAB5C gene into a construct that integrates via Tol2-mediated transgenesis and is driven by the zebrafish fli-1a promoter, to achieve expression specifically in vessels (Fig. 3c) [45]. The construct was injected into single-cell stage Tg(kdrl:GFP)^s843 zebrafish embryos expressing GFP in all vascular endothelial cells, and mCherry-positive endothelial cells were examined in developing ISVs at 30–32 h post-fertilization (hpf) (Fig. 3d, e) [46, 47]. As these injections result in mosaic expression of mCherry-Rab5C in endothelial cells, mCherry-positive endothelial cells were scored for their position within the developing vessels (tip, stalk, or dorsal aorta; Fig S4A). The distribution of WT-Rab5C positive cells was similar to that of mCherry-expressing cells (Fig. 3f and Fig S4B). However, cells expressing DN-Rab5C were less commonly found in the tip cell position (22% versus 56% in the mCherry control), and most DN-Rab5C positive cells localized to the stalk or the dorsal aorta (Fig. 3f and Fig S4B).

Altogether, our findings indicate that Rab5C is required for tip cell formation, with Rab5C expressing cells having a competitive advantage for tip cell positioning over cells with impaired Rab5C function.

We showed by mosaic overexpression of DN-Rab5C that Rab5C activation is required for tip cell formation in vivo. While these experiments allowed us to identify the preference for tip or stalk cell position in a competitive situation, we also analyzed the effects of DN-Rab5C overexpressed simultaneously in all endothelial cells, by generating transgenic Tg(kdrl:GFP)^s843 embryos with vascular-specific expression of human mCherry-WT-Rab5C (Tg(fli1a:mCherry-WT-hRAB5C)^mu227, or mCherry-DN-Rab5C (Tg(fli1a:mCherry-DN-hRAB5C)^mu228. Similar to our findings in HUVECs, we observed localization of WT Rab5C protein in rapidly moving vesicles in endothelial cells in vivo (Fig. S4C), while the DN mutant was mostly absent from these vesicles but was instead retained in a perinuclear compartment, most likely the Golgi (Fig. S4C). Indeed, quantification confirmed that the number of mCherry-positive vesicles in DN-Rab5C-expressing ISV cells was strongly decreased, compared to WT-Rab5C expressing cells (Fig. S4D).

We then scored ISV formation at 30–32 hpf, a time-point at which normal ISV endothelial cells have migrated dorsally and interconnected, to form the dorsal longitudinal anastomotic vessel (DLAV) [46, 47]. The survival of DN-Rab5C-expressing embryos was compromised at later stages but not before 30–32 hpf, thus allowing analysis of the developing ISVs. Without WT cells taking over the tip cell function, we observed an impairment of endothelial cell migration in the ISVs of embryos expressing DN-Rab5C (Fig. 4a). While in WT-Rab5C-expressing embryos ISV development was characterized by 83.3% reaching the DLAV and 16.7% migrating nearly to the top, DN-Rab5C-overexpressing endothelial cells migrated only to the level of the horizontal myoseptum (‘half’) in 31% of the embryos, with 47.3% reaching the ISV length, and only 21.7% reaching the DLAV (Fig. 4b).

To disrupt zebrafish Rab5c function in vivo via additional methods, we also used Morpholino (MO)-mediated block of rab5c translation (rab5c ATG MO) or splicing (rab5c e2i2 MO), as well as CRISPR/Cas9-mediated genome modification to induce mutations in the rab5c gene (Fig. S5A–E). A heterozygous rab5c mutant line was generated by injections of rab5c specific guide RNAs (gRNAs) and Cas9 protein, growth to adulthood, identification of germline-transmitting adults, growth of the F1 generation, and identification of the mutation. Detailed genomic analysis revealed that the resulting rab5c mutant (designated rab5c^mu229) contained a premature stop codon, thus terminating protein synthesis at 29 (instead of 221) amino acids (Fig. S5D,E). We then analyzed vascular development in MO-injected embryos and in the homozygous mutant offspring of rab5c^+/mu229 parents. Similar to the DN-Rab5C-expressing embryos, the homozygous mutants died during later stages of development. We observed mispatterning of the ISVs and impaired dorsal migration in individual ISV sprouts in embryos injected with rab5c MOs (compared to Control MOs), as well as in rab5c^mu229/mu229 embryos (compared to rab5c^+/+ siblings) (Fig. 4c,d), thus confirming that Rab5c is required for tip cell formation and normal ISV migration in vivo. Additionally, abnormal sprouting angiogenesis was also revealed by live imaging of ISV development using time-lapse microscopy (Movies S1,S2 and Fig. S6A).

We also investigated the levels of zebrafish Vegfr2 (Kdrl) protein in embryos injected with Control MO or rab5c MO by Western blotting (Fig. 5a). Similar to what we observed in HUVECs, a significant reduction in Kdrl protein levels was apparent in Rab5c-depleted embryos (Fig. 5a,b). Consistently, tip cells in these embryos formed shorter filopodia, as revealed by Rab5c knockdown in Tg(fli1a:lifeact-EGFP)^mu240 embryos, which express LifeAct-eGFP in the vasculature to visualize the actin cytoskeleton (Movies S3,S4 and Fig. 5c) [48]. Vegf-induced tip cell specification in zebrafish requires initial activation of Notch signaling in tip cells, which subsequently directs them into developing arteries [49, 50]. Since we found an impairment of tip cell morphology and behavior in Rab5c-depleted embryos, we also addressed whether Notch signaling was affected. For this purpose we used the Tg(TP1bglob:VenusPEST)^s940 transgenic line to image the highly dynamic expression of a very short-lived Venus protein (Venus-PEST), that is driven by the TP1 promoter element and thus reports activation of Notch signaling [51]. Notch activation was visible in arteries, as well as in some tip cells and arterial ISVs (Fig. 5d), in addition to a number of non-vascular mesenchymal cells. As expected from our morphological data, a strong reduction of Venus-PEST expression in rab5c MO-injected embryos was observed, indicating that activation of Notch signaling was reduced and further supporting our hypothesis that Rab5c is required for efficient generation of functional tip cells (Fig. 5d).

Altogether, these data show that in line with our observations in HUVECs (Figs. 1–3), Rab5c is crucial for the maintenance of Vegfr2 levels in zebrafish, and thereby regulates tip cell formation and sprouting angiogenesis in vivo.

The previous sections have shown that activation of endothelial Rab5C is required for sprouting angiogenesis. Because Rabs are activated by GEFs, we next investigated which of the eight known GEFs for Rab5 GTPases are expressed in endothelial cells, using publicly available genome-wide mRNA expression profiles. This analysis suggested that HUVECs express at least seven different Rab5 GEFs, while RINL was not represented on the used arrays (Fig. 6a and Table S2). To investigate which of these GEFs is involved in the Rab5C-dependent effects on VEGFR2 and angiogenic sprouting, we silenced the expression of each individual GEF, including RINL, and assessed VEGFR2 cell-surface levels by flow cytometry. Of all investigated GEFs, only the depletion of Ras and Rab interactor (RIN) 2 significantly reduced the surface levels of VEGFR2, to the same extent as the depletion of Rab5C (Fig. 6b). Furthermore, total protein levels of VEGFR2 were also reduced by RIN2 depletion, as assessed by Western blotting (Fig. 6c, d). As in the case of Rab5C depletion, the reduction of VEGFR2 levels was associated with an increase in VEGF-induced VEGFR2 degradation (Fig. 6e). Finally, we performed a sprouting assay using RIN2-depleted cells. As expected, both the number of formed sprouts as well as the total network length were suppressed by RIN2 depletion (Fig. 6f–i).

Collectively, our data show that knockdown of RIN2 expression recapitulates the effects of Rab5C depletion, and strongly suggest that RIN2-mediated Rab5C activation prevents VEGFR2 degradation to promote sprouting angiogenesis.

We next tested whether forced Rab5C activation can circumvent the requirement for RIN2 to drive sprouting. For this purpose, we generated a constitutively active (CA) mutant (Q80L) of human Rab5C, which blocks GTP hydrolysis, fused to mScarlet (Fig. S6B). Expression of this mutant into HUVECs caused enlargement of EEs, indicative of constitutive Rab5C activation (Fig. S6C) [52]. We then expressed mScarlet-CA-Rab5C into RIN2-depleted cells, which slightly lowered endogenous Rab5C expression, whereas RIN2 knockdown by itself did not affect endogenous Rab5C levels or vice versa (Fig. 7a; Fig. S6D). Importantly, expression of CA-Rab5C in RIN2-depleted HUVECs rescued the sprouting defects caused by RIN2 deficiency, and thus bypassed the requirement for RIN2 (Fig. 7b). These data confirm that 1) Rab5C activation promotes sprouting angiogenesis, and 2) the primary role of RIN2 in sprouting angiogenesis is to activate Rab5C.

To assess the effects of RIN2 knockdown on Rab5C function in vivo, we generated a splice-blocking MO (e3i3) against the zebrafish rin2 gene (Fig. S7A,B), which was injected into Tg(fli1a:mCherry-WT-hRAB5C)^mu227;Tg(kdrl:GFP)^s843 zebrafish embryos to examine Rab5C localization. Depletion of zebrafish Rin2 protein resulted in a partial failure of Rab5C to localize in vesicular compartments (Fig. 7c), indicating that the recruitment of Rab5C requires Rin2 and hence, that the Rin2-Rab5c interaction is functionally conserved in zebrafish.

We also addressed the functional role of zebrafish Rin2 in sprouting angiogenesis in Tg(kdrl:GFP)^s843 embryos. Knockdown of Rin2 resulted in impaired endothelial cell migration in the ISVs, similar to that induced by deficiency of Rab5c (Fig. 7d, e). The migration defect was again accompanied by reduced activation of Notch signaling, as analyzed by Tp1:Venus-Pest expression (Fig. S7C).

In summary, these results indicate that Rin2 is crucial for Rab5C activation and recruitment in endothelial cells, which regulates Vegf/Notch signaling, gene expression, tip/stalk cell specification, and sprouting angiogenesis (Fig. 7f).

The following antibodies were used: anti-VEGFR2 (R&D Systems, AF357), anti-α-tubulin (Sigma-Aldrich, T6199), anti-AKT (Cell signaling technology, #9272), anti-p(S473)AKT (Cell signaling technology, #4060S), anti-VEGFR1 (Abcam, ab32152), anti-p(Y204)ERK-1/2 (Santa Cruz Biotechnology, sc-7383), anti-ERK-1/2 (Santa Cruz Biotechnology, sc-153), and anti-RIN2 (Sigma-Aldrich, HPA034641). HRP-conjugated secondary antibodies (#P0447, #P0399 and #P0449) were from DAKO, Alexa Fluor-conjugated secondary antibodies (A21445, A21200, and A21467) were from Thermo Fisher. Recombinant human VEGF-A (VEGF165) was purchased from R&D Systems (293-VE), fibronectin (#F1141) and thrombin (#T1063) were from Sigma-Aldrich, human fibrinogen (Haemocomplettan P, B02BB01) was from CSL Behring. Bafilomycin (#tlrl-baf1) was from InvivoGen, Leupeptin (#4041) and Pepstatin (#4397) were from PeptaNova, Hoechst 33,342 (#H3570), phalloidin (#T7471), and CellTracker Fluorescent Probes (CMFDA green, #C2925 and CMTPX red, #C34552) were from Thermo Fisher Scientific. All used primers were purchased from Integrated DNA Technologies, Inc. (Table S3). The human WT and DN RAB5C open reading frames (ORFs) were synthesized, cloned into pDONR221, and sequence-verified by Thermo Fisher Scientific for gateway cloning. For the zebrafish experiments, the RAB5C ORFs were cloned into a modified version of pTol2-fli1ep:CherryDest (a kind gift from Nathan Lawson; Addgene plasmid #73,493). To generate this destination vector, the mCherry sequence was PCR-amplified without stop codon using oligos extended with restriction sites (fwd: 5′-GACGATGCTAGCGCCACCATGGTGAGCAAGGGCGAG-3′ rev: 5′-GACGATTACGTAGATATCCTTGTACAGCTCGTCCATG-3′). This product was cloned into pTol2-fli1ep:CherryDest using the NheI and SnaBI restriction sites. Next, a Gateway Reading Frame Cassette A containing the appropriate attR1/2 sites was cloned into the EcoRV site, 3′ of the mCherry ORF, creating pTol2-fli1ep:CherryDestV2. Finally, the human WT and DN RAB5C ORFs in pDONR221 were cloned into pTol2-fli1ep:CherryDestV2 using LR Clonase II enzyme mix (Invitrogen), creating the 4 pTol2-fli1ep:Cherry-RAB5 plasmids. For lentiviral transduction of the RAB5 ORFs into HUVECs, we modified pLenti6.3/V5-Dest (Invitrogen) by removing an attR1/2 cassette by digestion with EcoRI and EcoRV, and introducing an NheI site using 2 oligos (5′-AATTCGCTAGCGACGACGAT-3′ and 5′-ATCGTCGTCGCTAGCG-3′). Next, the NheI/SnaBI fragment from pTol2-fli1ep:Cherry-RAB was cloned into the NheI/EcoRV sites of pLenti6.3/V5-Dest. To generate the mScarlet-Rab5C-Q80L construct, site-directed mutagenesis was performed on an mScarlet-Rab5C-wt construct using primer 5′-GTTTGAGATCTGGGACACAGCTGGACTGGAGCGGTATCACAGC-3′ and its reverse complement, in which CAG, coding for Q (Glutamine) was replaced with CTG coding for L (Leucine). All cloned plasmids used in experiments were sequenced with at least double coverage.

For analysis of relative mRNA expression levels in transduced HUVECs, total RNA was isolated using the RNeasy kit (Qiagen) according to the manufacturer’s instructions. After reverse transcription to cDNA using the SuperScript III First-Strand Synthesis System (Thermo Fisher Scientific), qPCR was performed with the SensiFAST SYBR No-ROX kit (Bioline) and the indicated primers (Table S3), either on a LightCycler PCR system (Roche) or a StepOnePlus system (Applied Biosystems). Duplicate reactions were performed for each gene and expression was normalized to that of β-actin.

HUVECs were treated as indicated, detached with accutase (Sigma-Aldrich) for 2 min, and washed in 2% FCS in PBS. Cells were stained with primary antibodies for 1 h on ice, washed twice, followed by incubation with secondary antibodies for 45 min on ice and washed twice. Cells were analyzed on a Canto-II flow cytometer (BD Immunocytometry Systems) equipped with FACSDiva software.

Expression of Rab5 GEFs in HUVECs (Table S2) was determined by analysis of Affymetrix U133 Plus 2.0 genome-wide mRNA expression profiles in the public domain using the NCBI Gene Expression Omnibus (GEO: GSE7307) website [58, 59]. GEO was first searched for studies on low-passage, non-recombinant, non-stimulated HUVECs with data normalization using the MAS5.0 algorithm (Affymetrix Inc., Santa Barbara, CA). This resulted in a total of 11 published studies comprising 29 separate arrays, 17 of which were listed with present call analysis, that were queried for the expression of all known human Rab5 GEFs [20]. RINL was not represented on the Affymetrix arrays and was not further analyzed. Array data were analyzed as described using R2; an in-house developed Affymetrix analysis and visualization platform (http://​r2.​amc.​nl). Probes were ranked depending on high expression values and widespread expression (% of samples with significant expression for that gene) in the data collections tested.

Cells were washed in ice-cold PBS and lysed on ice in NP40 lysis buffer (50 mM Tris–HCl, pH 7.4, 100 mM NaCl, 10 mM MgCl₂, 1% NP-40, 10% Glycerol), supplemented with protease inhibitor cocktail (Sigma-Aldrich). Cell lysates were cleared by centrifugation, heated at 95ºC in SDS sample buffer (50 mM Tris–HCl pH 6.8, 2% SDS, 10% glycerol, 1% β-mercaptoethanol, 12.5 mM EDTA, 0.02% bromophenol blue), and proteins were resolved by SDS-PAGE. Thereafter, proteins were transferred to nitrocellulose membranes (GE Healthcare Amersham) and aspecific binding was blocked using 5% (w/v) milk (Campina) in TBST (150 mM NaCl, 10 mM Tris, 0.1% Tween 20, pH 8.0) for 30 min. Immunoblotting was performed with the indicated antibodies by incubation with primary antibodies o/n at 4ºC and with secondary antibodies for 1 h at RT. Membranes were washed 3 × with TBST after each step. Proteins were visualized using ECL chemiluminescence reagent (Thermo Fisher Scientific), light sensitive films (Fuji Film) and a film processor (Konica Minolta, SRX-101A). Quantification of bands was performed by densitometry using ImageJ. Band intensity of the protein of interest was corrected to that of α-tubulin.

For VEGF signaling and VEGFR2 degradation assays, HUVECs in 6-well plates were grown to confluence. Then, cells were starved overnight and subsequently either left untreated or stimulated with 50 ng/ml VEGF for the indicated time-points, whereafter the cells were processed for Western blotting. Inhibitors were used as indicated.

Sprouting assays were performed according to previously established protocols with minor modifications [42, 43]. In brief, HUVECs were incubated with collagen-coated microcarrier beads (Sigma-Aldrich, C3275). The next day, the beads were detached by washing and transferred to 48-well plates containing fibrinogen in PBS (2 mg/ml) supplemented with 0.15 U/ml aprotinin and 6.25 U/ml thrombin. EGM-2 medium was added on top of the gels. Beads were imaged by time-lapse microscopy on a Widefield system using a 10 × dry lens objective (Carl Zeiss MicroImaging, Inc.). Images were recorded at 15-min intervals during 48 h. For each condition, 20 beads were analyzed per experiment. Sprouting was assessed as the average number of large (length of sprout > than the bead diameter) sprouts per bead. Alternatively, beads were embedded in fibrin gels in ‘half-area’ glass-bottom 96-well imaging plates (Corning, #4580). After 48 h, beads were fixed, stained with Hoechst and phalloidin, and visualized by generating z-stacks on a confocal microscope. Maximum projections of z-stacks were created using Fiji/ImageJ (version 1.52e), de-speckled once, and the contrast was enhanced with 0.3% and analyzed using the Angiogenesis plugin [60]. The phalloidin staining was used for correct segmentation of the sprouts. For the sprouts stained with CellTracker, the number of tip cells per condition was counted and normalized to the total number of cells in that condition.

Zebrafish (Danio rerio) husbandry and embryo maintenance were carried out under standard conditions at 28.5 ℃ [61]. Embryonic developmental stages were determined as described [62]. Transgenic lines used in this study were Tg(kdrl:GFP)^s843, Tg(kdrl:Hras-mCherry)^s896, Tg(TP1bglob:VenusPEST)^s940, Tg(fli1a:lifeact-EGFP)^mu240 [48, 51, 63, 64]. All animal experiments were performed in compliance with the relevant laws and institutional guidelines, were approved by local animal ethics committees and were conducted at the University of Münster and the MPI for Molecular Biomedicine with permissions granted by the Landesamt für Natur, Umwelt und Verbraucherschutz (LANUV) of North Rhine-Westphalia.

The transgenic lines Tg(fli1a:mCherry-WT-hRAB5C)^mu227 and Tg(fli1a:mCherry-DN-hRAB5C)^mu228 zebrafish lines were newly generated using Tol2-mediated DNA integration [65]. The rab5c^mu229 mutation was newly generated by CRISPR/Cas9. Annealed template oligonucleotides were transcribed into gRNAs using MEGAshortscript T7 Kit (Ambion) as described [66]. The following oligos were used for gRNA generation: rab5c 5′-AAAGCACCGACTCGGTGCCACTTTTTCAAGTTGATAACGGACTAGCCTTATTTTAACTTGCTATTTCTAGCTCTAAAACCCGTCGGGAACAAAATCTGCCTATAGTGAGTCGTATTACGC-3′ (rab5c target sequence: 5′-GCAGATTTTGTTCCCGACGGGGG). A total of 1 nl of 300 ng/ml gRNA and 3.22 μg/μl Cas9 NLS protein (M0646T, New England Biolabs) were injected at the single-cell stage. Efficiency of the CRISPR-targeting as well as genotyping were performed by PCR, using fwd 5′-GCCTCTCCATCCTTTTCTCA-3′ and rev 5′-TCACGAACACACTCCAGCTC-3′ and subsequent restriction enzyme digest with Hpy99I.

For MO-mediated gene knockdown, embryos were injected at the one-cell stage with 3 ng control MO (Gene Tools), 5′-CCTCTTACCTCAGTTACAATTTATA-3′, 3 ng rab5c MO, 5′-CGCTGGTCCACCTCGCCCCGCCATG-3′ 3 ng rin2 MO, 5′-GCAGTGTGTTTTAACTCTCACCTTA-3′. Sequence of the rab5c ATG MO was as previously described [37]. The rab5c and rin2 splice-site MOs were generated as shown (Fig. S6A,S8A), and validated by RT-PCR. For mosaic overexpression of mCherry-WT-Rab5C or mCherry-DN-Rab5C, embryos were injected at the single-cell stage with 100 pg plasmid and 250 pg Tol2 mRNA per embryo. Effects on vascular development were analyzed at 30–32 hpf.

Total RNAs were prepared from 36 hpf embryos using TRIzol reagent (Invitrogen), and reverse-transcribed by random hexamer primers using Superscript III (Invitrogen) according to the manufacturer’s instruction. PCR was performed using gene-specific primers as below. Efficiency of the knockdown was assessed by PCR using the primers fwd: AGAGCTGCGTACATCCAACC and rev: TGCAAGCCAAAATTCAAAGA.

After removing chorionic membrane and yolk sac, embryos injected with either control MO or rab5c MO were directly lysed in 1 × SDS sample buffer, and subjected to Western blot analysis with anti-Kdrl antibody (Kerafast, ES1003) and anti-alpha-Actin antibody (Santa Cruz Biotechnology, sc-47778) as described previously [67].

HUVECs on glass coverslips were fixed with 4% paraformaldehyde (Merck) in PBS containing 1 mM CaCl₂ and 0.5 mM MgCl₂ (PBS + +) for 10 min, and permeabilized with 0.4% Triton X-100 (Sigma-Aldrich) in PBS +  + for 5 min. Aspecific antibody binding was prevented by blocking with 2% BSA Fraction V (Sigma-Aldrich) in PBS +  + for 15 min. Following incubation with the indicated primary antibodies, coverslips were washed with 0.5% BSA Fraction V in PBS +  + and antibody binding was visualized using secondary antibodies. Coverslips were washed and subsequently mounted in 10% Mowiol, 2.5% Dabco, 25% glycerol, pH 8.5. Image acquisition was performed on a Leica SP8 confocal microscope using a 63 × oil immersion objective, after which images were processed using Leica Application Suite X and Fiji/ImageJ (version 1.52e) software.

Sprouting was visualized using a 25 × long-distance water objective on a Leica SP5 confocal set-up. z-stacks were created of 1.5 μm per slice and the different channels were sequentially scanned between stacks.

Confocal microscopy of zebrafish was performed using an LSM780 (Carl Zeiss Microscopy GmbH; objective lens: Plan Achromat 20/0.8, 20x; LD C-Apochromat 40x/1.1 W Korr M27). Living zebrafish embryos were embedded in 1% agarose (dissolved in E3 medium containing Tricaine) and analyzed using Imaris 9 software (Bitplane) as previously described [68].

For time-lapse analysis in zebrafish, a temperature of 28.5 °C was maintained using a heating chamber. Images were collected every 6 min during 1 h (for imaging of filopodia) or every 15 min during 6 h (for imaging of ISV sprouting). Assembly of confocal stacks and time-lapse movies was conducted using Imaris 8/9 software (Bitplane).

sh_Ctrl, sh_Rab5A or sh_Rab5C cells were seeded in 10 cm dishes and grown to confluence. Cells were starved overnight, whereafter they were either left untreated or stimulated with 50 ng/ml VEGF for 1 h or 4 h. Subsequently, cells were lysed using RLT buffer (Qiagen) with 1% β-mercapto-ethanol and stored at −80 °C until analysis. RNA-seq libraries were prepared from samples of 2 independent experiments with the mRNA KAPA HyperPrep Kit (Illumina) using Truseq Illumina LT adapters. Sequencing was performed on an Illumina Hiseq4000 system (single-read 50 bp), 10 samples per lane.

Reads were subjected to quality control (FastQC, dupRadar, Picard Tools), trimmed using Trimmomatic v0.36, and aligned to the hg38 genome using HISAT2 v2.1.0. Counts were obtained with HTSeq v0.11.0. Statistical analyses were performed using the edgeR and limma/voom packages [69, 70]. Genes with no counts in any of the samples were removed while genes with more than 2 reads in at least 3 of the samples were kept. Count data were transformed to log2-counts per million (logCPM), normalized by applying the trimmed mean of M-values method and precision-weighted using voom. Differential expression was assessed using an empirical Bayes’ moderated t-test within limma’s linear model framework including the precision weights. Resulting p values were corrected for multiple testing using the Benjamini–Hochberg false discovery rate. Additional gene annotation was retrieved from Ensembl (release 94) using the biomaRt/Bioconductor package. Geneset enrichment was performed using CAMERA with a preset value of 0.01 for the inter-gene correlation using collections H, C1, C2, C3, C5, C6, and C7 retrieved from the Molecular Signatures Database (MSigDB v6.2; Entrez Gene ID version), complemented with a user-defined geneset containing the genes constituting the observed VEGF transcriptome. Data analysis were performed using R (v3.5.2) and Bioconductor (v3.8). Sequencing data have been made available in the GEO under accession number: GSE134947.

Statistical analysis was performed using one-way ANOVA for multiple comparisons and unpaired t-tests for comparisons between two conditions, unless stated otherwise. Throughout the paper, statistically significant differences are indicated by * (p < 0.05), ** (p < 0.01), *** (p < 0.001), or **** (p < 0.0001).