Introduction

Vascular networks are formed by an array of morphogenetic processes, such as sprout outgrowth, anastomosis and pruning, which depend on the tight coordination of dynamic endothelial cell behaviors (Betz et al., 2016; Ellertsdóttir et al., 2010; Phng, 2018; Schuermann et al., 2014). In order to transport blood, the newly formed blood vessels need to form a continuous lumen. Studies of lumen formation in mammalian and zebrafish embryos have revealed that the early blood vessels of the trunk are formed by different morphogenetic processes. The larger axial vessels, the dorsal aorta (DA) and posterior cardinal vein (PCV), form their lumens between coalescing endothelial cells, a process referred to as cord hollowing (Ellertsdóttir et al., 2010; Jin et al., 2005; Strilić et al., 2009; Zeeb et al., 2010). In vivo timelapse analyses of the formation of the intersomitic vessels (ISV) and the dorsal longitudinal anastomotic vessels (DLAV) in zebrafish have revealed the existence of two alternative mechanisms of lumenization. In one mechanism, called transcellular lumen formation, the luminal cell membrane invaginates into the cell. This cell hollowing process is driven by blood pressure and results in so-called seamless tubes, which contain a lumen surrounded by single cells (Betz et al., 2016; Gebala et al., 2016; Herwig et al., 2011 ; Lenard et al., 2013; Phng and Belting, 2021). Alternatively, local lumens, which are formed between individual cells within a sprout can be fused by cell rearrangements (Herwig et al., 2011; Sauteur et al., 2014). Endothelial cell rearrangements generate multicellular tubes (Betz et al., 2016; Phng and Belting, 2021). Studies examining lumenization of ISV and DLAV have shown that transcellular lumen formation requires blood pressure and that, in the absence of blood flow, endothelial cell rearrangements appear to be the default mechanism of vascular tube formation (Herwig et al., 2011 ; Lenard et al., 2013).

Endothelial cells are very motile - they can move within the angiogenic sprout and also in patent vessels in the presence of blood flow (Blum et al., 2008; Franco et al., 2016; Jakobsson et al., 2010). Here, a dynamic balance between inter-endothelial cell adhesion and plasticity allows angiogenic sprouting while maintaining the endothelial seal. Previous analysis on blood vessel formation and anastomosis in zebraflsh has shown that junctional remodeling is central to many aspects of morphogenetic endothelial cell-cell interactions. In particular, the adhesion molecule VE-cadherin (Cdh5) is essential for coordinated cell shape changes during multicellular tube formation, as loss of VE-cadherin was shown to inhibit cell rearrangements (Sauteur et al., 2014). Furthermore, rather than supporting a passive role of VE-cadherin, which is consistent with the maintenance of vascular integrity, an active force generating function for VE-cadherin in this process was suggested (Sauteur et al., 2014). By performing in vivo high-resolution time-lapse imaging and genetic analyses, we have shown that endothelial cells use oriented, oscillating junction-based lamellipodia (JBL) to move over each other and thus provide the physical means for cell rearrangements to drive multicellular tube formation (Paatero et al., 2018).

To better understand the cellular and molecular mechanisms of JBL function we have undertaken a detailed analysis of junctional and actomyosin dynamics. Taken together, our data show that JBL provide forward movement by generating pushing and pulling forces at the poles of junctional rings thereby elongating the cell and promoting cell rearrangements.

Results

Junction-based lamellipodia generate luminal connectivity

Lumenization of vascular sprouts is central to blood vessel formation. Although different morphogenetic mechanisms of vascular tube formation have been described, in vivo analysis in zebraflsh embryos has established cord hollowing as the default mechanism of blood vessel formation in the absence of blood flow (Herwig et al., 2011 ; Lenard et al., 2013). During the cord hollowing process neighboring cells form local lumens which become interconnected by cell rearrangements (Andrew and Ewald, 2010). In order to visualize the fusion of such luminal pockets we followed the localization GFP-Podxl, which labels the apical cell membrane, together with F-actin reporter UCHD-mRuby2, to delineate endothelial cell-cell junctions and F-actin base protrusions (Fig. 1). Time-lapse recordings revealed that fusion of apical membrane compartments occurs within a few hours upon initiation of DLAV formation and is associated with junctional rearrangements between the three associated cells (Fig. 1b, c and Supplementary movie 2). In agreement with our previous work, higher temporal resolution imaging showed that cell junctions formed transient F-actin-based protrusion (JBL) at the pole with respect to junctional ring elongation (Fig. 1c, d and Supplementary movie 1, and Supplementary movie 3) (Paatero et al., 2018).

Figure 1

JBL form distal junctions by de novo recruitment of VE-cadherin

We have previously shown that JBL are oscillating protrusions, which drive junction elongation and thereby cell rearrangements during tube formation. A salient feature of JBL is the formation of a distal junction in front of the JBL (Paatero et al., 2018).

To gain more insight into the dynamics of distal junction formation, we compared the distribution of Factin (Ruby-UCHD) with that of VE-cadherin (VE-cad-Venus) and ZO-1 (EGFP-ZO1), respectively (Fig. 2). Time-lapse experiments revealed distinct and dynamic localization of these proteins. During early phases of JBL formation VE-cadherin was diffusely distributed throughout the protrusion and accumulated in aggregates leading to a characteristic “double junction” pattern at the distal tip of the junctional ring (Fig. 2a and Supplementary movie 4). Compared to VE-cadherin and ZO-1 the early demarcation of F-actin at proximal and distal junction was less sharp (Fig. 2b, c and Supplementary movies 5, 6). However, early aggregates of VE-cadherin and ZO-1 were always positive for Ruby-UCHD, showing that these three proteins were present from the onset of distal junction formation. At subsequent stages, as the distal becomes more mature, ZO-1 and VE-cadherin at the proximal junction were maintained, while the Ruby-UCHD signal started to fade (Fig. 2c-e).

Figure 2

Formation of the distal junction may occur in different ways. One one hand, it may form by a local junctional disassembly, followed by a reaggregation at the distal end of the JBL. This would essentially consist of a junctional rearrangement process. On the other hand, the distal junction may form de novo by recruitment of VE-cadherin from pools outside of the proximal junction. In order to distinguish between these scenarios, we performed color conversion experiments (Supplementary Fig. 1 and Supplementary Movie 7). Pools of photoconvertible VE-cadherin (Cdh5-mClav2) were differentially labeled by exposure to UV light within a narrow region of interest at the pole of the junctional ring (Supplementary Fig. 1a), which resulted in efficient green-to-red conversion (Supplementary Fig. 1b). When we followed subsequent JBL dynamics over time, we noted that converted VE-cad was absent from the protrusion as well as the distal junction (Supplementary Fig. 1c and Supplementary movie 7. 5 experiments). Instead, the distal junction contained non-converted VE-cadherin, indicating that the distal junction was formed de novo by recruitment of VE-cadherin from pools outside the proximal junction.

Arp2/3 complex dynamics spatiotemporally correlates with JBL formation

Junctional protrusions are commonly seen in developing and newly formed blood vessels in the zebrafish embryo. Cell culture experiments, using human umbilical cord endothelial cells (HUVEC), have shown that such lamellipodial structures can control monolayer integrity and vascular permeability (Alves et al., 2018; Martinelli et al., 2013). In particular, so-called junction-associated intermittent lamellipodia (JAIL) are prominent in sub-confluent HUVEC, where they serve to repair junctional gaps (Taha et al., 2014). In contrast to JAIL JBL are highly polarized relative to the vessel axis and they drive cell rearrangements rather than maintaining vascular integrity (Paatero et al., 2018). However, both lamellipodial structures are similar in their emergence from and association with endothelial cell junctions.

JAIL formation appears to be triggered by a local reduction in junctional tension and requires the activation of the ARP2/3 complex. We used a transgenic zebraflsh line expressing Arpc1b-Venus in the vasculature to test whether Arp2/3 also regulates JBL formation. Time-lapse analysis showed a discrete spatiotemporal deployment of Arp2/3 at the leading edge of the forming JBL (Fig. 3). Imaging JBL over several oscillations revealed that Arp2/3 accumulation coincided with the initiation and the elongation of JBL, while it disappeared at the end of each JBL cycle (Fig. 3a and Supplementary movie 8). Comparison with ZO-1 showed that Arp2/3 localized distally with no overlap within the proximal junction (Fig. 3a). When we imaged Arp2/3 together with mRuby2-UCHD at higher temporal resolution (30s/frame), we observed that during JBL extension Arp2/3 was maintained at the leading edge (Fig. 3b, c and Supplementary movie 9). Taken together, these data show that Arp2/3 specifically marks the distal edge of the JBL during formation and elongation (Fig. 3d).

Figure 3

As the above findings suggest that Arp2/3 is involved in JBL formation, we interfered with Arp2/3 function by pharmacological inhibition using CK666 (Hetrick et al., 2013). We used acute treatments to avoid secondary effects and performed live-imaging on rearranging endothelial cell junctions. Application of 0.2 mM CK666 at 32 hpf completely disrupted junctional Arp2/3 localization within 30 min. (Supplementary Fig. 2). Analysis of F-actin organization revealed an increase of filopodia emanating from the junction in the area, which usually was occupied by JBL (Fig. 4a-c and Supplementary movies 10, 11). The abundance of these filopodia prevented the localization of JBL at the junctional poles consistent with previous cell culture studies (Henson et al., 2015). This suggests that filopodia are formed instead of lamellipodia in the absence of Arp2/3 activity. To test consequences of the loss of Arp2/3 activity on cell rearrangements, we measured the speed of junctional elongation in control and CK666 treated embryos. By imaging ZO-1 labeled junctions over 1 hour we found that the speed of junction elongation was strongly reduced from an average of 8µm/h to 3µm/h (Fig. 3d,e and Supplementary movies 12,13). Taken together the results show that JBL formation depends on Arp2/3 function and that the absence of Arp2/3 activity leads to a strong impairment of endothelial cell rearrangements during blood vessel formation.

Figure 4

Proximal and distal junctions resolve by junctional merging and fusion

The appearance of “double junctions” (consisting of a proximal and a distal junction) is a distinctive junctional feature observed in JBL. Their spatiotemporal dynamics suggest that they represent part of the mechanism by which JBL promote junction elongation. Live-imaging revealed a cyclic turnover between the two junctional states: single junction and double junction pattern. Time-lapse imaging of VE-cadherin-Venus at high temporal resolution showed that this turnover is mediated by the cyclic formation of the distal junction followed by the merging and fusion of the proximal and distal junctions (Fig. 5a and Supplementary movie 14).

Figure 5

Myosin light-chain localization correlates with junctional merging

The cyclic junctional pattern in JBL suggests that it may represent a physical ratchet mechanism, which drives junction elongation. Therefore, we wanted to explore whether junctional mergence could be driven by actomyosin contractility. To monitor the spatiotemporal distribution of actomyosin contractility during JBL cycle, we utilized myosin regulatory light chain (MLC) (Myl9a-GFP and Myl9b-Cherry) as reporters for non-muscular myosin II (NMII) activity (Lancino et al., 2018). Myl9 accumulated along the junctions but was strongly enriched in JBL at both ends of the junctional ring (yellow arrowheads in Fig. 5b).

During cell rearrangements JBL are formed at each pole of the junctional ring in a complementary fashion: each JBL is formed by one of the interacting cells corresponding to the forward orientation with respect to cell movement (Paatero et al., 2018)(see schematic in Fig. 5d). To test whether Myl9 accumulation corresponds to the forward orientation of the JBL, we analyzed rare mosaic situations in which Myl9a-GFP was expressed in only one of the partnering cells. Indeed, we observed high levels of Myl9a-GFP in JBL formed by the high-expressing cell pointing into the forward direction of the elongating junction (Fig. 5d, e).

Vascular tube formation by cell rearrangements has also been described in other vascular beds. We turned to the posterior cerebral vein (PCeV), which is generated by JBL-driven cell rearrangements converting the initial tube architecture from unicellular to multicellular (Lenard et al., 2013). When we compared Myl9a-EGFP distribution to mRuby2-UCHD, we found that the former localized within the JBL adjacent to the forming distal junction of the JBL (see yellow arrowheads in Fig. 5f and Supplementary movie 15).

In order to gain a better understanding how actomyosin contractility may drive junctional mergence, we imaged the distribution of Myl9 at high spatiotemporal resolution. We first used VE-cadherin-EGFP to label proximal and distal junctions as spatial landmarks and thus demarcate the area of interest for the quantification of Myl9b-mCherry in space and time (Fig. 6a, b, Supplementary Fig. 3, and Supplementary movies 16, 17, 18, 19, 20). We defined three areas – proximal junction, interjunctional and distal junction and quantified Myl9b-mCherry at two time-points (0” and 45”) at the onset and during junctional mergence, respectively (Fig. 6b). These quantifications reveal a redistribution Myl9 from the proximal and distal junctions towards the interjunctional space (Fig. 6c). This increase indicates a strong recruitment of Myl9 and a concomitant increase of actomyosin contractility within the area between the proximal and distal junctions.

Figure 6

Actomyosin contractility is required for junction elongation and junctional mergence

In order to assess the role of actomyosin contractility in junction elongation more directly, we used the Rock inhibitor Y-27632 (Uehata et al., 1997)and measured the length of junctional rings over 60 minutes. Rock inhibition strongly inhibited junction elongation compared to DMSO controls (Fig. 7a, b and Supplementary movies 21,22). To test whether the lack of junction elongation under Rock inhibition may be caused by defects in JBL function, we performed acute short-term Y-27632 treatment and performed live-imaging over 30 minutes to image JBL dynamics during reduction of actomyosin contractility – with a focus on the merging of proximal and distal junctions. To this end, we imaged 20 (Y-27632) and 13 (DMSO) events of junctional merging, respectively, and tracked individual interjunctional distances over time (Fig. 7c-e and Supplementary movies 23, 24). Inhibition of actomyosin contractility led to a strong inhibition of junctional mergence, ranging from a reduction in speed to a complete block in junctional mergence. This blockage in junctional mergence was reflected by the overall increased occurrence of “double junctions” under Rock inhibition (20 (Y-27632) vs 13 events (DMSO) in 18 videos). Taken together, these results show that high levels of actomyosin contractility reside within interjunctional space and that these contractile forces are responsible for junctional mergence. Furthermore, these data suggest that JBL formation together with junctional mergence are the driving force of junctional elongation and endothelial cell rearrangements during angiogenic blood vessel formation.

Figure 7

Discussion

Lumen formation is key in the formation of tubular organs such as blood vessels. Endothelial cells can use different morphogenetic pathways to generate a lumen, including membrane invagination, cord hollowing or lumen ensheathment (Eberlein et al., 2021; Phng and Belting, 2021). The dorsal longitudinal anastomotic vessel (DLAV) of the zebrafish embryo can be formed by either membrane invagination or cell rearrangements, which result in unicellular or multicellular tubes, respectively (Betz et al., 2016; Phng and Belting, 2021). Here, we have analyzed the cell rearrangements, which connect small luminal pockets within the initial endothelial rod, thereby generating a continuous lumen in a fully patent vascular tube.

The coalescence of local, extracellular lumens is commonly observed during the cord hollowing process of different biological tubes, including the notochord of Ciona intestinalis, the vertebrate dorsal aorta as well as the neural tube and intestine (Bagnat et al., 2007; Dong et al., 2009; Gladysheva et al., 2021; Jin et al., 2005; Strilić et al., 2009). In those tubular organs, in which lumen coalescence was analyzed, it was found that it is mainly caused by lumen expansion, which in turn is driven by directed liquid transport and hydrostatic pressure (Bagnat et al., 2007; Lowery and Sive, 2005; Munson et al., 2008; Navis and Bagnat, 2015). In contrast, the lumen coalescence observed in the zebrafish ISV, DLAV and PCeV is accomplished by active cell intercalation. Here, endothelial cells actively elongate their cell-cell junctions until they form novel cell-cell contacts, which allow the connection of the two adjacent lumens.

We have previously shown that endothelial junction elongation is driven by junction-based lamellipodia (JBL) (Paatero et al., 2018). Careful analysis of junctional and cytoskeletal reporters revealed that JBL dynamics can be subdivided into 4 discrete steps: 1) JBL formation, 2) formation of a new, distal junction, 3) junction conversion and 4) junction stabilization (Paatero et al., 2018). Taken together, these studies showed that JBL provide an oscillating ratchet-like mechanism, in which endothelial cells employ F-actin-based protrusions and VE-cadherin-mediated cell-cell contacts to move over each other. JBL thus provide the physical means for cell rearrangements to drive multicellular tube formation.

In this study we have further explored the junctional and actomyosin dynamics to discern how JBL generate the motive force required for junction elongation and cell rearrangements. In essence, our data show that JBL require ARP2/3 dependent F-actin polymerization for JBL formation as well as actomyosin contractility to merge the proximal and distal junctions at the end of the JBL cycle. Thus, by sequential deployment of pushing (F-actin polymerization) and pulling (junction merging) forces, JBL promote forward movement of endothelial cells (summarized in Fig. 8).

Figure 8

Oscillatory junctional protrusions of endothelial cells have also been described in cultured HUVECs (Cao et al., 2017; Taha et al., 2014). Here, so-called junction-associated intermittent lamellipodia (JAIL) form in similar intervals as JBL. However, differences in junctional dynamics suggest that JAIL and JBL represent related but different cellular activities. Notably and in contrast with JAIL, JBL form a distal junction at the leading edge of the protrusion. Our studies show that these “double junctions” are an integral part of the cellular mechanism of junction elongation. We have previously shown that JBL are most prominent during multicellular tube formation and become less prevalent and smaller thereafter (Paatero et al., 2018). It is therefore likely that JAIL and JBL have different functions during vascular development. While JBL are required for polarized, directed junctional remodeling, JAIL are involved in junctional maintenance and endothelial integrity. Lamellipodial structures resembling JBL, displaying double junctions, have recently been described in the oak leaf-shaped endothelial cells of dermal lymphatics in the mouse (Schoofs et al., 2025). Here, the double junctions are thought to regulate endothelial shape and in particular to stabilize the interendothelial cell overlap, which is important to control the interstitial fluid transport.

Our photoconversion experiments (see Supplementary Fig. 1) show that the distal junction is formed de novo from VE-cadherin pools originating from outside the existing (proximal) junction. During distal junction formation, we observe that VE-cadherin forms clusters resembling spot-junctions observed in the context of organ morphogenesis in other systems. Consistent with our observation such spotjunctions have been shown to be important in the formation of contractile F-actin fibers in these systems (Lecuit et al., 2011). While we are currently not able to monitor actin flow in our experimental in vivo system, our observation that the formation of the distal junction coincides with the formation of a distal F-actin belt is consistent with a clutch mechanism, in which transligation of VE-cadherin triggers reorganization of the cortical actin cytoskeleton similar to what has recently been described in cell culture studies (Noordstra et al., 2023). Furthermore, our previous findings from zebrafish cdh5 null mutants or embryos expressing a VE-cadherin protein (VE-cadΔCT) lacking the β-catenin binding site show that VE-cadherin mediated cell-cell interactions as well as VE-cad/F-actin interaction are important for JBL-driven junction elongation (Paatero et al., 2018; Sauteur et al., 2014).

The cell motility promoting activity of VE-cadherin during tube formation may be quite unique, considering that in most studies cadherins have been described to stabilize epithelia, limit cell motility and to regulate contact inhibition of locomotion(Campàs et al., 2024; Maître and Heisenberg, 2013; Noordstra et al., 2023) Cadherin-based cell interactions have been shown to be essential for dynamic cell movements in several morphogenetic processes (Friedl and Mayor, 2017; Mayor and Etienne-Manneville, 2016). In the case of collective border cell migration during Drosophila oogenesis and migration of primordial germ cells in zebrafish, cadherins have been shown to provide traction for collective and single cell migration, respectively (Blaser et al., 2006; Cai et al., 2014).

A salient feature of JBL is their appearance at both ends of each rearranging cell. This bipolar activity causes the cells to elongate in both directions and rearrange rather than to migrate. Furthermore, and in contrast to lamellipodia-driven mechanisms of cell migration, the domain of actomyosin contractility is confined within the interjunctional area (between distal and proximal junctions, see Figs. 6 and 8). This suggests that the restricted area of contractility is not only demarcated by the proximal and distal junctions, but that these junctions act as insulators to segregate compartments of different actomyosin tension along the cellular interface. We propose that this insulator function is essential to permit junction elongation.

In this study, we have investigated the cellular and molecular basis of force generation in JBL. Through detailed analysis of junctional, cytoskeletal, and regulatory proteins, we uncover a biphasic mechanism involving alternating protrusive and contractile forces. Notably, and in contrast to classical paradigms of cell migration, we show that actomyosin contractility is spatially restricted to a narrow zone at the junctional pole during endothelial cell movement. Together, our findings reveal a novel mechanism of cell rearrangement during vascular cord hollowing. JBL serve as localized modules that generate both protrusive and contractile forces during junctional remodeling and cell elongation. Moreover, JBL-localized junctions may act as insulators, spatially confining actomyosin tension to the junctional pole.

Material and Methods

Lead contact

Further information and request for resources and reagents should be directed and will be fulfilled by the Lead Contact; Heinz-Georg Belting (heinz-georg.belting@unibas.ch).

Materials availability

Materials and zebrafish lines generated and used in this study are available upon request to the lead contact.

Fish maintenance and stocks:

Zebrafish (Danio rerio) were maintained according to FELASA guidelines (Aleström et al., 2020). All experiments were performed in accordance with federal guidelines and were approved by the Kantonales Veterinäramt of Kanton Basel-Stadt (1027H, 1014HE2, 1014 G).

The following zebrafish lines were used in this study:

Sex was not considered in this study because zebrafish embryos used for the experiments were at developmental stages prior to sex differentiation. Therefore, the sex of the embryos was not determined or relevant to the outcomes.

Live imaging of zebrafish embryos

In DLAV experiments, zebrafish embryos between 30 to 32 hpf were dechorionated and anaesthetized with 0.16 mg/ml (1×) tricaine methanesulfonate (Sigma). The embryos were selected for presence of fluorescence and mounted into microwell dishes within 0.55% low-melting-point agarose (ROTI) and covered with E3 buffer containing 1× tricaine. In PceV experiments, transgenic embryos selected for presence of fluorescence were anaesthetized in 1× tricaine (0.08%) and mounted in a 35 mm glassbottom Petri dish (0.17 mm, MatTek), using 0.7% low melting agarose (Sigma) containing 0.08% tricaine and 0.003% 1-phenyl-2-thiourea (PTU; Sigma-Aldrich). The embryos were mounted into microwell dishes within 0.55% low-melting-point agarose (ROTI) and covered with E3 buffer containing 1× tricaine.

A Leica SP5 confocal microscope with a 40× water immersion objective was used for live-imaging experiments. High spatiotemporal resolution time-lapse images were acquired with an Olympus SpinSR spinning disc microscope using a 60X silicon oil (NA = 1.3) objective, with a z-stack step size of 0.6 µm at different frequencies depending on the experiment.

Photoconversion experiments

Photoconversion experiments were performed using a Leica SP5 confocal microscope with a 40× water immersion objective (NA = 1.1). Photoconversions were applied to manually selected regions of interest (ROIs) with a 405 nm laser for 10–20 seconds to induce conversion, monitored by simultaneous imaging at 488 nm to ensure complete conversion. All the imaging was performed at 28.5°C.

Junction elongation assay

Junctional elongation was assessed by measuring elongation speed of isolated junctional rings during DLAV formation between 30 and 32 hpf. Inhibitor treatments CK666 (200 µM), Y-27632 (75 µM), or DMSO (1%) were applied 1h before mounting of embryos into 0.55% low-melting point agarose. The same concentrations of chemicals were applied to the low-melting-point agarose mounting medium and the E3 medium on top of it before imaging and imaging the junctions for 60–90 min on a Leica SP5 (Y-27632) Olympus SpinSR spinning disc microscope or Olympus SpinSR spinning disc (CK666) microscope.

Junctional merging tracking

Speed of junctional merge was evaluated by monitoring isolated junctional rings during DLAV formation. Inhibitor treatment Y-27632 (75 µM) or DMSO (1%) were applied 30 min before mounting. The same concentrations of chemicals were applied to the low-melting-point agarose mounting medium and the E3 medium on top of it before imaging and imaging the junctions for 10–15 min on Olympus SpinSR spinning disc microscope. Upon distal junction appearance, distances were measured on the Fiji software, drawing a line between proximal and distal junction, and measuring the length of the lines.

Immunostaining

Zebrafish embryos of 30 to 32 hpf were fixed after dechorionation using Glyofixx (Thermo-Fisher Scientific) and incubated at 4°C overnight (ON) in 1.5 ml Eppendorf tubes (max. 20 embryos per tube). After fixation, embryos were washed four times for 5 minutes in 1× PBST at room temperature (RT) on a rocking platform. Embryos were then permeabilized using 0.5–1% Triton-X100 in PBST for 30 min at RT. They were then blocked with 2% BSA and 5% goat serum in PBST overnight with continuous shaking at 4°C. After blocking, the primary antibody solution was added. The primary antibodies were either diluted in blocking solution or Pierce Immunostain Enhancer Solution (Thermo Fisher Scientific). Primary antibody incubation was performed at 4°C for 3 days. After incubation, the solution was removed, and the embryos were washed at least four times for 30 minutes each with PBST at 4°C, or overnight. Subsequently, the embryos were incubated with secondary antibodies, diluted 1:2000 in blocking solution, and incubated at 4°C.

Image preparation and analysis

Image analysis and measurements were performed using ImageJ 2.9.0. Z-stacks were flattened by maximum slice intensity projections. Where needed, noise was reduced using Gaussian filtering (radius 0.7) and/or background subtracted (rolling ball radius 50) using ImageJ. Contrast and brightness of images were linearly adjusted. Image panels were created using the Open Microscopy Environment (OMERO). ARP2/3-Venus and mRuby2-UCHD localization were evaluated on a 4pixel-wide rectangle, and the average over length, corrected for background (evaluated as average of average intensity in 5 rectangles on the cell cytoplasm) and displayed in a graph using graphpad prism. Proximal, distal, and interjunctional space domains were defined using VE-cad distribution, My9-mcherry distribution was evaluated as average intensity in the distinct domains The intensities are corrected for background (which has been evaluated as average of average intensities in 5 rectangles in the cell cytoplasm) and displayed in a graph using graphpad prism.

Supplementary figures

Supplementary Figure 1:

Supplementary Figure 2:

Supplementary Figure 3:

Acknowledgements

We thank Kumuthini Kulendra and Andre Rodriguez for fish care and the Imaging Core Facility of the Biozentrum (Universität Basel) for microscopy support. We thank Anne Schmid (Institute Pasteur, Paris, France) for providing Tg(kdrl: Myl9a-GFP)ip5Tg and Michel Bagnat (Duke University, USA) for providing TgKI(tjp1a-tdTomato)pd1224. This work has been supported by the Kantons Basel-Stadt andBasel-Land and by grants from the Swiss National Science Foundation (310030_200701 and 310030B_176400) to M.A..

Additional information

Data availability

The data that support the findings of this study are available in the supporting information of this article.

Statistical analysis

GraphPad Prism 10.4.1 was used to perform statistical analyses. For every experiment and analysis, sample size is specified in the figure legend. Sample sizes were not pre-determined using statistical methods, and all data meeting the quality control criteria were included in the analyses. Randomization was not applied to the experiments. Unless specified otherwise, default settings were used for all software tools in the analyses. Whenever a p-value is presented in the text or figures, the corresponding statistical test is indicated.

Author contributions

L.M., J.Y., C.W., I.P., M.A. and H.-G.B. designed the experiments. L.M., J.Y., C.W. and I.P. performed experiments and analyzed the data. J.M. and C.H. generated Tg(fli1a:Arpc1b-mVenus-p2A-Arpc3-mTurq2)^mr25. L.M. and H.-G.B. wrote the manuscript. All authors reviewed the manuscript.

Additional files

Video 1. JBL zo1-gfp_uchd-mRuby2

Video 2. JBL drive local lumen fusion

Video 3. JBL adjacent to apical compartment

Video 4. Formation of the distal junction (VE-cad-Venus)

Video 5. Distal Junction Formation (mRuby-UCHD)

Video 6. Distal Junction Formation (mRuby-UCHD)

Video 7. The distal junctions forms de novo

Video 8. Arp2/3 oscillation at junctional pole

Video 9. Arp2/3 localization at the front end of the JBL

Video 10. Arp2/3 is required for JBL formation

Video 11. Arp2/3 is required for JBL formation

Video 12. Arp2/3 is required for junction elongation

Video 13. Arp2/3 is required for junction elongation

Video 14. Distal and proximal junctions merge

Video 15. MLC is enriched at the junctional poles

Video 16. Myosin light-chain dynamics correlate with junctional merging 1

Video 17. Myosin light-chain dynamics correlate with junctional merging 2

Video 18. Myosin light-chain dynamics correlate with junctional merging 3

Video 19. Myosin light-chain dynamics correlate with junctional merging 4

Video 20. Myosin light-chain dynamics correlate with junctional merging 5

Video 21. Actomyosin contractility drives junctional conversion (DMSO)

Video 22. Actomyosin contractility drives junctional conversion (Y-27632)

Video 23. Actomyosin contractility drives junctional conversion (DMSO)

Video 24. Actomyosin contractility drives junctional conversion (Y-27632)

Video 25. Molecular Mechanism of JBL function during endothelial junction elongation