Abstract
The role of blood flow in vascular development is complex and context-dependent. In this study, we quantify the effect of the lack of blood flow on embryonic vascular development on two vascular beds, namely the cerebral and trunk vasculature in zebrafish. We perform this by analysing vascular topology, endothelial cell (EC) number, EC distribution, apoptosis, and inflammatory response in animals with normal blood flow or absent blood flow. We find that absent blood flow reduced vascular area and EC number significantly in both examined vascular beds, but the effect is more severe in the cerebral vasculature, and severity increases over time. Absent blood flow leads to an increase in non-EC-specific apoptosis without increasing tissue inflammation, as quantified by cerebral immune cell numbers and nitric oxide. Similarly, while stereotypic vascular patterning in the trunk is maintained, intra-cerebral vessels show altered patterning, which is likely to be due to vessels failing to initiate effective fusion and anastomosis rather than sprouting or path-seeking. In conclusion, blood flow is essential for cellular survival in both the trunk and cerebral vasculature, but particularly intra-cerebral vessels are affected by the lack of blood flow, suggesting that responses to blood flow differ between these two vascular beds.
Introduction
Endothelial cells (ECs) perform multiple functions during normal physiology including wound healing, tissue regeneration, immune response, menstruation, and pregnancy (1, 2, 3). Cerebral EC dysfunction is associated with neurodegenerative diseases, arteriovenous malformations, aneurysms, and stroke (4, 5). Increasing evidence suggests ECs display different molecular and functional properties according to their anatomical site, such as the cerebral or trunk vessels (6, 7, 8, 9, 10). This highlights the importance of studying the responses of multiple vascular in development and territories to experimental manipulations.
Zebrafish embryos are a frequently used model to study vascular development and disease (11, 12, 13). Fluorescent transgenic reporter lines allow cellular and sub-cellular visualization in vivo (14). Advanced microscopy, such as light sheet fluorescence microscopy (LSFM), acquires vascular information in great anatomical depth and over prolonged periods of time (15), allowing data acquisition to be rich in information and detail. Zebrafish embryonic transparency allows non-invasive and in vivo studies of different vascular beds in the same animal.
After 24 h post-fertilization (hpf), the zebrafish basic body plan is established and cardiac contraction starts. Within this timeframe, vasculogenesis forms the primary vessels (16, 17). Blood flow plays an important role in processes, such as EC polarization, vascular lumenization, pruning, and cardiac trabeculation (18, 19, 20, 21, 22, 23, 24), and directly impacts vascular architecture (25). Zebrafish embryos can survive for 7 days post fertilization (dpf) without blood flow via oxygen diffusion due to their small size (13, 26). This makes them well suited to examine the role of blood flow on vasculogenesis and angiogenesis.
One approach to study the effect of absent blood flow in zebrafish is the knockdown of cardiac troponin T2A (tnnt2a) using antisense morpholino oligonucleotides (MO). This prevents cardiac contraction and thus blood flow (27, 28). Contrary to recent debates on MO phenocopying mutant phenotypes, the tnnt2a MO phenocopies the silent heart (sih) mutation (28), making it a widely used MO-based approach to study the lack of blood flow. Additionally, a control MO group accounts for injection-induced developmental delays (29). Previous studies used this MO-based method to study the impact of blood flow on specific cerebral (30, 31) and trunk vessels (32), showing that EC numbers are reduced in the trunk when blood flow is absent (32, 33).
Even though these studies have provided invaluable insights into the role of blood flow, important questions remain unanswered about the role of blood flow in vascular development. These include whether absent blood flow has the same effects in different vascular beds, whether vessels of different identities respond differently, which steps of vasculature formation are altered, and whether effect sizes of absent blood flow differ in magnitude over time.
To examine these questions. we here use LSFM 3D in vivo imaging of the cerebral and trunk vasculature of 2–5dpf zebrafish embryos with and without blood flow induced by tnnt2a knockdown. Our results show that even though the gross vascular responses to blood flow are comparable in different territories, some differences between anatomical sites exist, suggesting vascular territories exhibit differential sensitivity to absent blood flow.
Materials and methods
Zebrafish husbandry
Experiments were performed according to the rules and guidelines of institutional and UK Home Office regulations under the Home Office Project Licence 70/8588 held by TC.
Maintenance of adult zebrafish was performed as described in standard husbandry protocols (34, 35). Embryos, obtained from controlled mating, were kept in E3 (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4) medium buffer with methylene blue and staged according to Kimmel et al. (36). The following transgenic reporter lines were used: Tg(kdrl:HRAS-mCherry)s916 (37) visualizes EC membrane, Tg(fli1a:eGFP)y1 (14) visualizes EC cytosol, Tg(flk1:nls-eGFP)zf109 (38) visualizes EC nuclei, Tg(mpx:GFP)i114 (39) visualizes neutrophils, and Tg(fms:GAL4.VP16)i186, Tg(UAS-E1b:nfsB.mCherry)il149 (40) visualizes macrophages. To assess cell death in vivo, we created the stable transgenic Tg(secAnnexinV:mVenus)SH632 in pDestCryaa:RFP, similar to (41) and the plasmid cloning of (42).
Morpholino injection
Development of functional heart contraction was inhibited via injection of tnnt2a ATG morpholino (1.56 ng final concentration), as described in (27, 28) (sequence 5’-CATGTTTGCTCT GATCTGACACGCA-3’). Control morpholino injections (5’-CCTCTTACCTCAGTTATTTATA-3’; no target sequence and little/no biological activity; Genetools, LLC) were performed with the above final concentration to study off-target effects of injections. Injections were conducted at a one-cell stage using phenol red as an injection tracer.
Chemical and histological stains
In vivo visualization of cell death was performed using 2 µg/mL solution of Acridine Orange (Sigma) in 1X E3 for 2 h in 3dpf embryos, followed by three washes in E3 before image acquisition (43).
In vivo visualization of inflammation via nitric oxide (NO) (44) was performed using 2.5 µM DAF-FM-DA (Molecular Probes; D23844) (45) for 6 h in 3dpf embryos. DMSO control was performed at the same concentration and duration.
Image acquisition settings and properties
Anaesthetized embryos were embedded in 2% LMP-agarose with 0.01% Tricaine in E3 (MS-222, Sigma). Data acquisition of the cranial and trunk vasculature was performed using a Zeiss Z.1 light sheet microscope, Plan-Apochromat 20×/1.0 Corr nd = 1.38 objective, dual-side illumination with online fusion, activated Pivot Scan, image acquisition chamber incubation at 28°C, and a scientific complementary metal-oxide-semiconductor (sCMOS) detection unit. The properties of acquired data were as follows: 16 bit image depth, 1920 × 1920 × 400–600 voxel (x,y,z; approximate voxel size of 0.33 × 0.33 × 0.5 µm, respectively). Multicolour images in double-transgenic embryos were acquired in sequential mode.
Image analysis
As tnnt2a can develop considerable oedema, we ensured comparability between samples by standardized image acquisition from the dorsal view, including the most dorsal vessel the dorsal longitudinal vein (DLV) to the more ventral basilar artery (BA; Supplementary Fig. 3A, see section on supplementary materials given at the end of this article). For image analysis, ROI selection was performed as previously described (Supplementary Fig. 9B) (46).
Automatic nuclei detection
To remove salt-and-pepper noise (47), a 6-by-6 neighbourhood median filter (48) was applied, while background removal was achieved with the rolling ball algorithm with size 200 (49). Following, 2D maximum intensity projection, and detection of local noise maxima using Fiji Software (50, 51). The Voronoi diagram was established following Otsu thresholding (52). Nuclei nearest neighbour distance (NND) was quantified in MIPs following pre-processing as above and Otsu thresholding, using the Fiji NND plugin (https://icme.hpc.msstate.edu/mediawiki/index.php/Nearest_Neighbor_Distances_Calculation_with_ImageJ.html).
Signal intensities
Signal intensity measurement of Acridine Orange and DAF-FM was conducted by creating 3D vascular masks following Sato filter for vascular enhancement and Otsu thresholding segmentation as previously described (46, 53, 54). Signal mean was quantified in ROIs in 2D MIPs. The signal-to-noise (SNR) ratio was quantified as the mean signal in ROI divided by the mean signal of the background (ROI placed outside the fish with a size of 10 × 10 μm). Acridine orange foci were detected in ROI following 2D median filtering using the detection of maxima with intensity over 10.
Manual analysis
All analysis was performed using Fiji (51).
Diameter of basilar artery (BA) was measured approximately 30 μm before bifurcating into posterior (caudal) communicating segments (PCS).
Posterior cerebral vein (PCeV) was measured approximately 20 μm before turning dorsally.
Posterior cardinal vein (PCV) and dorsal aorta (DA) diameters were measured above the cloaca, with three measurement points each in the same animal.
Intersegmental vessel (ISV) lengths of arterial (aISVs) and venous (vISVs) were measured above the cloaca, with three measurement points of the same vessels each in the same animal.
Vessel-specific nuclei measurements were conducted manually. First, nuclei were counted in aISVs and vISVs closest to the cloaca. Secondly, nuclei in the PCeV and BA were measured in an 80 μm long ROI positioned from the posterior end of the PCeV.
CtA sprouting and connectivity to BA were measured manually from the left PHBC.
Immune cells
Intracranial macrophages and neutrophils were quantified manually in 3D after ROI selection of the dorsal cerebral vasculature.
Availability of data and the code
Data and code will be made available upon request.
Statistical analysis
Gaussian distribution conformation was evaluated using the D’Agostino-Pearson omnibus test (21). Statistical analysis was performed using one-way ANOVA or paired Students t-test in GraphPad Prism Version 7 (GraphPad Software). Statistical significance was represented as: *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001. Graphs show mean values ± s.d. unless otherwise indicated. Image representation and visualization were done with Inkscape Version 0.48 (https://www.inkscape.org). Images were visualized as maximum intensity projections (MIPs). 3D rendering was performed using Arivis software (arivis AG, Munich, Germany).
Results
Cerebrovascular patterning is impaired by absent blood flow
We first examined whether cerebrovascular patterning is altered by lack of blood flow. Thus, we compared the cerebral vascular morphology of uninjected controls (Fig. 1A, B, C and D), control MO (Fig. 1E, F, G and H), and tnnt2a MO-injected embryos (Fig. 1I, J, K and L). When examining the global morphology, tnnt2a MO showed pericardial oedema, eye oedema, smaller eyes, and abnormal body curvature, which became more severe over time (Supplementary Fig. 1). The tnnt2a morphants were smaller with overtly abnormal cerebral vasculature, with effects becoming more severe from 2 to 5dpf. Intra-cerebral vessels, especially in the midbrain, were most severely affected showing altered growth patterns and an overall reduction of cerebral size was observed (Fig. 1L, dotted line). The primary head sinus (PHS), which extends laterally to the otic vesicle (OV), was enlarged, suggesting OV enlargement (Fig. 1L, filled arrowheads), while peripheral/perineural vessels, such as the primordial hindbrain channel (PHBC), formed relatively normally (Fig. 1L, unfilled arrowhead; schematic of cerebral vessels from 2 to 5dpf Supplementary Fig. 2). Our observations also confirm previous work in which vessels from the spinal cord and brain are still undergoing integration in the absence of blood flow (55).
The effect of tnnt2a morpholino knockdown on cerebral vessel development. (A, B, C and D) Cerebral vasculature from 2 to 5dpf in Tg(kdrl:HRAS-mCherry)s916 uninjected animals. (E, F, G and H) Cerebral vasculature in control MO-injected animals. (I, J, K and L) Cerebral vasculature in tnnt2a MO injected animals (n = 7–10; two experimental repeats). The lack of blood flow in tnnt2a morphants perturbs vascular development and worsens over time. Comparison between treatment groups shows that the midbrain vasculature is severely affected in tnnt2a MO (dotted lines) and the PCV surrounding the OV is enlarged (filled arrowhead), while the PHBC appears normal (unfilled arrowhead; d – dorsal, v – ventral; representative images colour-coded by depth).
Citation: Vascular Biology 3, 1; 10.1530/VB-21-0009
We next examined the distribution of EC in these animals (Fig. 2). This also suggested that peripheral vessels were less severely affected by absent blood flow, for example, when comparing nuclei distribution in the middle cerebral vein (MCeV, Fig. 2, unfilled arrowhead) to central arteries (Fig. 2, black arrowhead). This was confirmed when examining Voronoi diagrams of EC nuclei distribution (Fig. 2M, N and O). Voronoi tessellation, overlapping nuclei images, aided the visualization of EC nuclei density/distribution by partitioning MIPs into sub-regions based on nuclei position. Thus the Voronoi diagram lines and their colour indicate the spatial relationship of nuclei to each other, for example, larger tessellation and brighter colours show that a larger region is taken up by a nucleus. This showed EC distribution to be altered, particularly in the basilar artery (BA) or midbrain vessels.
Lack of blood flow impacts cerebral EC number. (A, B, C and D) Cerebral EC nuclei from 2 to 5dpf in Tg(flk1:nls-eGFP)zf109 uninjected control. (E, F, G and H) Cerebral EC nuclei in control MO. (I, J, K and L) Cerebral EC nuclei in tnnt2a MO (n = 7–10; two experimental repeats). Comparison of cerebral EC nuclei shows reduced cell numbers in tnnt2a MO with CtAs (filled arrowhead) being particularly affected (d – dorsal, v – ventral; representative images colour-coded by depth). (M, N and O) Voronoi (image partitioning based on nuclei position) diagrams of cerebral EC nuclei suggests nuclei numbers to be maintained in peripheral vessels such as the PHBC (white arrowheads), while EC density is reduced in the midbrain (grey arrowhead) and BA (unfilled arrowhead).
Citation: Vascular Biology 3, 1; 10.1530/VB-21-0009
This suggested that the effect of lack of blood flow becomes more severe over time and that central vessels are more impacted than peripheral vessels.
Vascular patterning in the trunk is preserved, but vascular morphology altered
We next examined vascular patterning in the trunk of the same animals to study whether the observed effects were conserved or different in these vascular beds (Fig. 3A, B, C, D, E, F, G, H, I, J, K and L). Vessels formed by vasculogenesis were observed and in contrast to the cerebral vessels, trunk vessel patterning was largely unaltered in the absence of flow. However, the morphology of the caudal vein plexus (CVP) was less defined upon lack of blood flow (Fig. 3, white arrowhead) and intussusceptive pillars were lacking (Fig. 3A′, E′ and I′). All our observations were consistent with previous studies that blood flow is required for plexus to caudal vein remodelling (56). Intussusception, and dorsal aorta (DA)-to-CVP segregation (57), but not intersegmental vessel (ISV) migration (17).
The effect of absent blood flow on trunk vessel development. (A, B, C and D) Trunk vasculature from 2 to 5dpf in Tg(kdrl:HRAS-mCherry)s916 uninjected control. (E, F, G and H) Trunk vasculature in control MO (n = 7–10; two experimental repeats). (I, J, K and L) Trunk vasculature in tnnt2a MO. Topology of the trunk vasculature is established in the absence of flow, but morphology of the CV is severely altered (black arrowheads) and intussusceptions are lacking (green arrowhead; d – dorsal, v – ventral; representative images colour-coded by depth).
Citation: Vascular Biology 3, 1; 10.1530/VB-21-0009
We next examined whether trunk EC numbers were changed. Examination of EC nuclei suggested EC numbers in the trunk were less severely impacted by lack of blood flow in comparison to cerebral vessels (Fig. 4), with EC distribution in animals without blood flow being similar in the CV but altered in ISVs (Fig. 4M, N and O).
Lack of blood flow impacts trunk EC number. (A, B, C and D) Trunk EC nuclei from 2 to 5dpf in Tg(flk1:nls-eGFP)zf109 uninjected control. (E, F, G and H) Trunk EC nuclei in control MO. (I, J, K and L) Trunk EC nuclei in tnnt2a MO (n = 7–10; two experimental repeats). Visual comparison of trunk EC nuclei suggests comparable numbers between treatment groups (d – dorsal, v – ventral; representative images colour-coded by depth). (M, N and O) Voronoi analysis of trunk EC nuclei shows EC distribution is maintained in the CV (black arrowhead), a decrease is observed in ISVs (red arrowhead), and significant changes are observed more anteriorly (unfilled arrowhead).
Citation: Vascular Biology 3, 1; 10.1530/VB-21-0009
Together, this suggested that vascular growth patterns in the trunk were unaltered, but remodelling, including intussusception, were lacking and that this is accompanied by changes in EC numbers and distribution.
Lack of blood flow affects vascular area and EC number
To elucidate the effect of absent blood flow over time, we quantified cerebrovascular area (vascular voxels in 2D (MIPs)). This was significantly reduced in tnnt2a morphants from 2 to 5dpf (Fig. 5A; uninjected control 2dpf P < 0.0001, 3dpf P < 0.0001, 4dpf P < 0.0001, 5dpf P < 0.0001), with similar results in craniovascular branch points (Supplementary Fig. 3A). Similarly, the number of cerebral EC nuclei was significantly reduced in tnnt2a morphants from 2 to 5dpf; while EC number almost doubled from 2 to 5dpf in controls, EC numbers were almost unaltered in fish without blood flow (Fig. 5B; uninjected control 2dpf P = 0.0409, 3dpf P < 0.0001, 4dpf P < 0.0001, 5dpf P < 0.0001). To examine the relationship between vascular area and EC nuclei number, vascular area-to-nuclei ratios were calculated (Fig. 5E), showing a decrease in uninjected controls (0.78-fold-change 2–5dpf) and control MO (0.87-fold-change 2–5dpf), while the ratio increased in tnnt2a MO (1.69-fold-change 2–5dpf). This showed that the number of nuclei-to-vasculature increased in controls but decreased in animals without blood flow.
We performed the same analysis on the trunk, and found that vascular area was significantly reduced by absent blood flow at 2dpf (P = 0.0074) and 3dpf (P = 0.0195), but not 4dpf (P = 0.1904) and 5dpf (P > 0.9999) in tnnt2a MO (Fig. 5C). Additionally, the number of nuclei was not significantly lower at 2dpf (P = 0.1514; Fig. 5D) but was significantly decreased at 3–5dpf (3dpf P = 0.0018, 4dpf P = 0.0013, 5dpf P < 0.0001). Similar results were observed when assessing trunk vascular branch points as co-metric (Supplementary Fig. 3B). Analysis of vascular-to-nuclei ratio showed that vascular area increased in all three groups (Fig. 5F; fold-change 2–5dpf: uninjected control 1.10, control MO 1.55, tnnt2a MO 2.51). Together, this suggested that cerebral vessels were more severely affected than trunk vessels by the absence of blood flow and that lack of blood flow increases the vascular-to-nuclei ratio in both the brain and trunk.
We next quantified the nuclei nearest neighbour distance (NND), finding it to be decreased in tnnt2a MO in both vascular beds (Fig. 5G and H), supporting the visual assessment of Voronoi diagrams and previous studies that suggest the less net distance between ECs in the absence of flow (56).
Quantification of vasculature and nuclei. (A) The cerebral vasculature is reduced in tnnt2a MO from 2 to 5dpf (n = 7– 10; two experimental repeats). (B) The number of cerebral EC nuclei is reduced in tnnt2a MO from 2 to 5dpf. (C) The trunk vasculature is reduced in tnnt2a MO at 2–3dpf, but not 4–5dpf (n = 7– 10; two experimental repeats). (D) The number of trunk EC nuclei is not altered in tnnt2a MO at 2dpf, but significantly reduced from 3 to 5dpf. (E) The ratio of cerebral vasculature-to-nuclei remains consistent in uninjected controls (green) and control MO (magenta), but increases over time in tnnt2a MO (blue). (F) The ratio of trunk vasculature-to-nuclei remains consistent in uninjected controls (green) and control MO (magenta), but increases over time in tnnt2a MO (blue). (G) Quantification of cerebral nuclei nearest neighbour distance (NND) showed nuclei distance to be decreased in tnnt2a MO in comparison to uninjected controls (P < 0.0001) and control MO (P < 0.0001; Kruskal–Wallis test). (H) Quantification of trunk nuclei NND showed nuclei distance to be decreased in tnnt2a MO in comparison to uninjected controls (P < 0.0001) and control MO (P < 0.0001; Kruskal–Wallis test).
Citation: Vascular Biology 3, 1; 10.1530/VB-21-0009
To assess how the observed reduced head size (Supplementary Fig. 4A) related to vascular and nuclei density, vasculature-to-ROI and nuclei-to-ROI ratios were quantified. This showed that for all three groups, vascular coverage was maintained at about 30% from 2 to 5dpf (Supplementary Fig. 4B), with ratios higher in tnnt2a MO at 5dpf in comparison to uninjected controls (P = 0.0074). Similarly, the nuclei-to-ROI ratios were maintained over time in all three groups (Supplementary Fig. 4C), but with higher ratios in tnnt2a MO than in controls.
Together, this showed that even though tnnt2a MO have a reduced head size, vasculature-to-ROI and nuclei-to-ROI ratios are maintained or even larger than in controls.
Arterial and venous responses to lack of blood flow are vascular bed dependent
To study whether vessels of different identities were differentially or similarly impacted by lack of blood, we quantified the diameter of selected arteries and veins in the brain and trunk 3dpf. Quantification of cerebral BA diameter showed no difference between tnnt2a MO and controls (Supplementary Fig. 5A; uninjected control P > 0.9999, control MO P > 0.9999), while the diameter of the posterior cerebral vein (PCeV) was reduced in animals without blood flow (Supplementary Fig. 5B; uninjected control P = 0.0175, control MO P = 0.1230). BA diameter was reduced by 2.5% (uninjected control 16.41 µm; tnnt2a MO 16.00 µm), while the PCeV diameter was reduced by 36.46% (uninjected control 11.52 µm; tnnt2a MO 7.32 µm). Quantifying trunk ISV diameters, aISV diameter (Supplementary Fig. 5C; uninjected control P = 0.0017, control MO P = 0.0121) and vISV diameter (Supplementary Fig. 5D; uninjected control P = 0.0010, control MO P = 0.0094) were both reduced in tnnt2a MO. The mean diameter of aISVs was reduced by 40.4% (uninjected control 10.52 µm; tnnt2a MO 6.27 µm), while the mean diameter of vISVs was reduced by 46.64% (uninjected control 11 µm; tnnt2a MO 5.87 µm). The diameter of the DA (Supplementary Fig. 5E; uninjected control P = 0.0008, control MO P = 0.0105) as well as the posterior cardinal vein (PCV) (Supplementary Fig. 5F; uninjected control P = 0.0159, control MO P = 0.0548) were reduced in tnnt2a MO. The mean diameter of the DA was reduced by 40.76% (uninjected control 25.86 µm; tnnt2a MO 15.32 µm), while the mean diameter of the PCV was reduced by 29.1% (uninjected control 27.46 µm; tnnt2a MO 19.47 µm). This suggested that vessel diameter is impacted by flow in the vessel of venous and arterial identity in the trunk, while arterial vessel calibre of the BA is unaltered without flow.
To further examine whether EC number was affected by vessel identity and/or vascular bed, EC nuclei per vessel were quantified. Analysis of nuclei of cerebral BA and PCeV in 80 μm ROI showed no significant difference between those vessels in the examined treatment groups and no significant change upon lack of blood flow (Supplementary Fig. 5G). Quantification of nuclei in trunk aISV and vISV showed no significant difference in EC numbers between vessels of different identity (Supplementary Fig. 5H). Upon lack of blood flow arterial EC number was not reduced (P = 0.5756), but venous EC number was significantly reduced (P = 0.0005).
Together, this suggests that vascular responses are vascular-bed as well as vascular-identity specific.
Non-EC-specific cell death is increased by absent blood flow
To examine whether cell death contributed to the reduced EC numbers observed in tnnt2a morphants, we quantified this using the transgenic Tg(secAnnexinV:mVenus)SH632 and the live dye Acridine Orange.
Visual inspection of the transgenic reporter line suggested an overall increase in cell death in tnnt2a MO compared to controls in the head (Fig. 6A, B and C) but not trunk (Fig. 6D, E and F), with specific foci found in the brain (white arrowhead). To examine whether the observed increase in annexin levels was vascular or non-vascular, we extracted non-vascular (Fig. 6A′, B′, C′, D′, E′ and F′) from vascular (Fig. 6A″, B″, C″, D″, E″ and F″) signal by producing vascular masks in 3D, finding that the observed cell death was not EC-specific. Quantification of tissue signal-to-noise ratio (SNR; Fig. 6G and H) showed an increase in the head in tnnt2a MO compared with uninjected controls (P = 0.0012, Fig. 6G) but not trunk (P > 0.9999, Fig. 6H). The number of annexin foci was not increased in tnnt2a MO (Fig. 6I; uninjected control P > 0.9999).
Cell death is increased by absent blood flow. (A, B and C) Cell death, visualized in the transgenic reporter line Tg(secAnnexinV:mVenus)SH632 (n = 13–15; three experimental repeats; 3dpf). (D, E and F) Annexin levels in the trunk vasculature appeared visually similar between groups. (G) Quantification of cerebral annexin SNR showed an increase in tnnt2a MO in comparison to uninjected controls (P = 0.0012) but not control MO (P = 0.072; Kruskal–Wallis test). (H) Quantification of trunk annexin SNR showed no increase in tnnt2a MO in comparison to uninjected controls (P > 0.9999) or control MO (P > 0.9999; Kruskal–Wallis test). (I) Quantification of cerebral annexin foci (white arrowheads) showed no increase in tnnt2a MO in comparison to uninjected controls (P > 0.9999) and control MO (P = 0.4542; Kruskal–Wallis test).
Citation: Vascular Biology 3, 1; 10.1530/VB-21-0009
To examine this further, the same experiments were conducted using the live dye Acridine orange, which showed similar results. However, cell death foci were clearly increased in fish without blood flow using the Acridine Orange assay (Supplementary Fig. 6).
Together, this suggests that overall cell death is increased in tnnt2a MO, not restricted to EC.
Inflammatory responses are not triggered by the absence of blood flow
We next examined whether immune cell numbers would be altered due to the observed cell death potentially triggering an inflammatory response, or local tissue ischemia due to lack of blood flow, equally triggering an inflammatory response (58). As the observed effects of lack of blood flow were more severe in the head vasculature, quantification was only performed in this vascular bed.
Quantification of the number of macrophages in controls and tnnt2a MO at 3dpf (Fig. 7A, B and C) showed no difference between groups (Fig. 7D; P = 0.2356). Similarly, no difference was found in intracranial neutrophil numbers at 3dpf when comparing tnnt2a MO to controls (Fig. 7E, F and G; P = 0.1708).
Lack of blood flow does not impact the number of cerebral innate immune cells. (A) Identification of immune cells was performed in 3D, allowing to discern non-specific signal (unfilled arrowhead) from immune cells (filled arrowhead). (B and C) Macrophages (magenta) were quantified examining the transgenic Tg(fli1a:eGFP)y1, Tg(fms:GAL4.VP16)i186, Tg(UAS-E1b:nfsb.mCherry)il149. (D) Number of macrophages was not changed upon blood flow loss (P = 0.2356; n = 12; 3dpf; Mann–Whitney U test). (E and F) Neutrophils (green) were examined in the transgenic reporter line Tg(mpx:GFP)i114, Tg(fms:GAL4.VP16)i186, Tg(UAS-E1b:nfsb.mCherry)il149. (G) Number of neutrophils was not changed upon blood flow loss (P = 0.1708; n = 12; two experimental repeats; 3dpf; Mann–Whitney U test).
Citation: Vascular Biology 3, 1; 10.1530/VB-21-0009
To further examine whether the observed cell death was associated with altered tissue inflammation in the absence of altered macrophage and neutrophil numbers, we visualised the inflammatory mediator nitric oxide (NO) using the live dye DAF-FM. The visual assessment showed no difference in DAF-FM levels in the head (Fig. 8A, B and C) and trunk (Fig. D, E and F). This was confirmed by SNR quantification in the head (Fig. 8G; P = 0.7967) and trunk (Fig. 8H; P = 0.9371) which showed no differences comparing samples with blood flow to samples without blood flow.
Inflammation is not increased by absent blood flow. (A, B and C) Examining nitric oxide (NO; visualized by DAF-FM) as an inflammatory marker, showed similar levels of NO in the head of uninjected controls, control MO, and tnnt2a MO (uninjected control n = 11, control MO n = 12, tnnt2a MO n = 11; three experimental repeats; 3dpf). (D, E and F) DAF-FM levels in the trunk vasculature appeared visually similar between groups. (G) Quantification of cerebral DAF-FM SNR showed no difference between tnnt2a MO and uninjected controls (P > 0.9999) or control MO (P > 0.9999; Kruskal–Wallis test). (H) Quantification of trunk DAF-FM SNR showed no difference between tnnt2a MO and uninjected controls (P = 0.1391) or control MO (P = 0.1216; Kruskal–Wallis test).
Citation: Vascular Biology 3, 1; 10.1530/VB-21-0009
Interestingly, we observed that the DAF-FM signal, previously described to be localized in the bulbus arteriosus after 2dpf (59), was absent in fish without blood flow (Supplementary Fig. 7), suggesting bulbus arteriosus NO expression is blood flow-dependent.
Together, our data suggested that the lack of blood flow does increase cell death but not tissue inflammation or immune cell recruitment at the investigated time-point.
Sprouting angiogenesis is altered but not halted by lack of blood flow
We next examined whether the observed changes in vascular patterning were due to changes in angiogenic sprouting. To examine this, we first examined trunk ISV sprouting and subsequent dorsal longitudinal anastomotic vessel (DLAV) formation, which are known to develop in a highly stereotypic anterior-to-posterior growth pattern. Comparison of time-lapse data in controls and embryos without blood flow (Supplementary Fig. 8A), showed that ISVs are competent to sprout in fish with blood flow, as previously suggested (17). Also in agreement with previous studies, we did not observe inverse membrane blebbing to occur in the absence of flow (60). However,DLAV anastomosis is delayed in fish without flow. Following the initial delay, ISVs sprout laterally to form the DLAV upon anastomosis. While vessels in controls become subsequently perfused upon anastomosis, this is not the case in fish without blood flow.
We next examined cerebrovascular sprouting by examining CtA sprouting and numbers, due to their stereotypic sprouting pattern from the PHBC and anastomosis with the BA. This showed that CtA’s were able to sprout from the PHBC and merge with the BA (Supplementary Fig. 8B) and when quantifying the number of sprouts this showed no significant difference in sprout number in fish without blood flow (Supplementary Fig. 8C; uninjected controls P = 0.4737). However, quantification of CtA’s merged to the BA, were significantly reduced in the absence of blood flow (Supplementary Fig. 8D; uninjected controls P = 0.0005).
Importantly, for all examined vessels stereotypic patterning was observed in the absence of flow, suggesting that path-finding is dependent on factors other than blood flow.
Discussion
In this study, we present the first assessment of the impact of blood flow on zebrafish embryonic vascular development from 2 to 5dpf, and present the first comparison of the impact of flow in two vascular beds, namely the head and trunk. We show that though the overall response to lack of blood flow is similar in both vascular beds, the head vasculature is more severely impacted than trunk vasculature, with intra-cerebral vessels particularly being affected. Additionally, the effect of absent blood flow increases over time. Our data show that lack of blood flow significantly increases cell death without significant cell death in ECs, without evidence of a significant increase of tissue inflammation, as quantified by cerebral immune cell numbers and nitric oxide (Table 1).
Overview of vascular properties altered upon lack of blood flow.
| Without blood flow | ||
|---|---|---|
| Head | Trunk | |
| Vasculogenesis | Yes | Yes |
| Angiogenic sprouts | Yes | Yes |
| Anastomosis | Reduced | Delayed |
| Inverse membrane blebbing | N/A | No |
| Kugeln | No | N/A |
| Cell death | Increased | No |
| Macrophages and neutrophils | No | N/A |
| DAF-FM | No | No |
| Vascular area | Reduced | Reduced |
| EC number | Reduced | Reduced |
| Vasculature:nuclei ratio | Increased | Increased |
| NND | Reduced | Reduced |
| BP | Reduced | Reduced |
| Artery diameter | No | Reduced |
| Vein diameter | Reduced | Reduced |
| Artery nuclei | No | No |
| Vein nuclei | No | Reduced |
Our work complements previous work (33), which found blood flow cessation to induce EC apoptosis in zebrafish embryos at 30hpf, while our assessment of cerebral macrophage numbers extends the examinations of Xu et al. who limited their studies to macrophage of the tectum (61). Our finding that NO is lacking in the bulbus arteriosus at 3dpf in fish without blood flow is the first functional evidence of bulbus arteriosus NO to be blood flow-dependent (59).
Our data showed that even in the complete absence of flow, stereotypic vascular patterning is preserved in the trunk and peripheral cerebral vessels. Our findings show that sprouting angiogenesis occurs in both vascular beds in the absence of flow but that anastomosis is delayed in the trunk and reduced in the head. This shows that in both, the trunk and head vasculature, vascular sprouting and path-seeking can occur in the absence of flow and is therefore not solely dependent on pressure gradients but likely to be induced by molecular or biochemical cues. However, successful fusion and anastomosis are blood flow dependent in both examined vascular beds. In agreement with previous work, we find that vascular plexus remodelling (56, 57) and vascular lumenization are blood flow-dependent (60). While previous studies suggest that pruning is directly driven by blood flow (19), it was never shown that this is the case for vessels that have never experienced blood flow. Our data show that early vascular networks are established. We suggest that, particularly in the head, multiple mechanisms contribute to the altered vascularization (a) lacking vascular self-fusion and remodelling in the absence of flow could be the mechanism for the observed phenotype, as suggested for the subintestinal vein (23); (b) reduced vascular stabilization (62, 63); (c) potentially changed EC polarity and maintenance needs to be examined (18).
Critically, our work shows that vessel identity affects cell response to flow and that this is furthermore impacted by the vascular bed. Thus, we suggest that even though genetic factors establish vascular identity prior to blood flow (64, 65), the fine-tuning of vascular response is likely to be caused by extrinsic vascular-bed-specific signaling molecules or biochemical cues (7).
Our data show that EC nuclei net distance is reduced in the absence of flow by examination of NND and Voronoi tessellation. Our study suggests that this might be in part due to a reduced vascular area (i.e. vasculature and nuclei numbers are reduced; vascular-to-nuclei ratios are unaltered in early development but changed with age) rather than significantly altered cell movements as nuclei are observed in all formed vessels. However, it has been previously shown that ECs migrate against the flow, thus the lack of blood flow might also alter EC net distance via migration changes (66).
Our studies show that cerebral EC cell numbers double from 2 to 5dpf in controls but remain stationary in fish without blood flow. Even though we observe cell death, there is a high likelihood that decreased proliferation due to the lack of blood flow is a contributing factor (67).
Once established blood vessels are established, also their EC properties and functions rely on blood flow, as it was previously shown regarding maintenance (7, 62, 63, 68), polarity (66, 69), and kugeln (9).
The finding that central cerebral vessels show severely altered growth patterns, while peripheral vessels are less impacted, suggests that the response to blood flow is different in different vascular territories; whether this is due to a difference in vessel formation (e.g. peripheral vessels are formed from angiogenesis-derived clusters, while central vessels are formed by angiogenesis from primary vessels) (70, 71) or identity (e.g. perineural, extra-cerebral, vs intra-cerebral) (72) requires future investigation.
The mechanisms of the observed effects in response to absent blood flow still require future investigation. This could include examining the expression of different mechanoreceptors and EC properties to complement previous findings about the importance and impact of blood flow (32, 73, 74, 75, 76, 77). Similarly, molecular pathways and their context-dependent interpretation (such as VEGF, BMPs, Wnt (62, 78, 79, 80)) might play a role in encountered differences in the head and trunk vasculature.
Together, our findings emphasize the important role of blood flow in vascular patterning and development and highlight different responses in different vascular territories to mechanical stimuli.
Key points
-
We here use zebrafish as a model to quantitatively assess the impact of the lack of blood flow in development and compare its impact in two vascular beds, namely the cerebral to trunk vasculature.
-
In both vascular beds, vascular growth and endothelial cell number are reduced by lack of blood flow, with increasing effect size from 2–5 days post fertilization.
-
Examination of vascular patterning shows that while in the absence of flow vasculogenesis and sprouting occur, anastomosis is delayed in the trunk and reduced in the head.
-
Studying differential responses in vessel types, we found that vascular responses to absent blood flow are vascular-bed as well as vascular-identity specific.
-
We found non-EC-specific cell death to be increased in both vascular beds, with a larger effect size in the brain, but that this cell death occurs without triggering recruitment of immune cells (macrophages or neutrophils) or tissue inflammation.
Supplementary materials
This is linked to the online version of the paper at https://doi.org/10.1530/VAB-21-0009.
Declaration of interest
Timothy Chico is an Editorial Board Member of Vascular Biology. Timothy Chico was not involved in the review or editorial process for this paper, on which he is listed as an author. The other authors declare that they have no conflict of interest.
Funding
This work was supported by a University of Sheffield, Department of Infection, Immunity and Cardiovascular Disease, Imaging and Modelling Node Studentship and an Insigneo Institute for in silico Medicine Bridging fund awarded to E K. R S was funded by the Medical Research Council Discovery Medicine North Doctoral Training Partnership. G B was funded by NC3Rs and the British Heart Foundation. J S-C was funded by the British Heart Foundation FS/18/2/33221. N H is funded by Association Européenne contre les Leucodystrophies, Grant/Award Number: ELA 2016‐012F4. P C E was funded by NC3Rs and the British Heart Foundation RG/19/10/34506. The Zeiss Z1 light-sheet microscope was funded via British Heart Foundation Infrastructure Award IG/15/1/31328 awarded to T C.
Author contribution statement
Funding obtained by E K, P A, T C, J S-C, N H, and P C E; data acquisition, E K, R S, G B, and K P; investigation, validation, and data curation, E K; formal visualization and analysis, E K; resources, P A, T C, J S-C, N H, and P C E; project administration, E K and P A; writing – original draft, E K; writing – review and editing, all authors. T C and P A: joint senior authors.
Acknowledgements
The authors are grateful to Heba Ismail for feedback, Deepak Ailani for sharing chemicals, and Fiona Wright for technical support, as well as the Bateson Centre Zebrafish Facility Staff for support and advice. The authors thank all funders, including the University of Sheffield Department of Infection, Immunity and Cardiovascular Disease, Insigneo Institute for in silico Medicine, Medical Research Council, NC3Rs, and the British Heart Foundation. The authors also thank the reviewers for their in-depth invaluable feedback to improve the manuscript.
References
- 1↑
Demir R, Yaba A, Huppertz B. Vasculogenesis and angiogenesis in the endometrium during menstrual cycle and implantation. Acta Histochemica 2010 112 203–214. (https://doi.org/10.1016/j.acthis.2009.04.004)
- 2↑
Jung S, Kleinheinz J. Angiogenesis – the key to regeneration. In Regenerative Medicine and Tissue Engineering, pp. 443–472, 2013. (https://doi.org/10.5772/55542)
- 3↑
Singer AJ, Clark RAF. Cutaneous wound healing. New England Journal of Medicine 1999 341 738–746. (https://doi.org/10.1056/NEJM199909023411006)
- 4↑
Feigin VL, Norrving B, Mensah GA. Global burden of stroke. Circulation Research 2017 120 439–448. (https://doi.org/10.1161/CIRCRESAHA.116.308413)
- 5↑
Lackland DT, Weber MA. Global burden of cardiovascular disease and stroke: hypertension at the core. Canadian Journal of Cardiology 2015 31 569–571. (https://doi.org/10.1016/j.cjca.2015.01.009)
- 6↑
Abbott NJ, Patabendige AAK, Dolman DEM, Yusof SR, Begley DJ. Structure and function of the blood–brain barrier. Neurobiology of Disease 2010 37 13–25. (https://doi.org/10.1016/j.nbd.2009.07.030)
- 7↑
Chico TJA, Kugler EC. Cerebrovascular development: mechanisms and experimental approaches. Cellular and Molecular Life Sciences 2021 78 4377–4398. (https://doi.org/10.1007/s00018-021-03790-1)
- 8↑
Huntley MA, Bien-Ly N, Daneman R, Watts RJ. Dissecting gene expression at the blood-brain barrier. Frontiers in Neuroscience 2014 8 355. (https://doi.org/10.3389/fnins.2014.00355)
- 9↑
Kugler EC, van Lessen M, Daetwyler S, Chhabria K, Savage AM, Silva V, Plant K, MacDonald RB, Huisken J & Wilkinson RN et al.Cerebrovascular endothelial cells form transient Notch-dependent cystic structures in zebrafish. EMBO Reports 2019 20 e47047. (https://doi.org/10.15252/embr.201847047)
- 10↑
Vanlandewijck M, He L, Mäe MA, Andrae J, Ando K, Del Gaudio F, Nahar K, Lebouvier T, Laviña B & Gouveia L et al.A molecular atlas of cell types and zonation in the brain vasculature. Nature 2018 554 475–480. (https://doi.org/10.1038/nature25739)
- 11↑
Bakkers J Zebrafish as a model to study cardiac development and human cardiac disease. Cardiovascular Research 2011 91 279–288. (https://doi.org/10.1093/cvr/cvr098)
- 12↑
Chico TJA, Ingham PW, Crossman DC. Modeling cardiovascular disease in the zebrafish. Trends in Cardiovascular Medicine 2008 18 150–155. (https://doi.org/10.1016/j.tcm.2008.04.002)
- 13↑
Gut P, Reischauer S, Stainier DYR, Arnaout R. Little fish, big data: zebrafish as a model for cardiovascular and metabolic disease. Physiological Reviews 2017 97 889–938. (https://doi.org/10.1152/physrev.00038.2016)
- 14↑
Lawson ND, Weinstein BM. In vivo imaging of embryonic vascular development using transgenic zebrafish. Developmental Biology 2002 248 307–318. (https://doi.org/10.1006/dbio.2002.0711)
- 15↑
Huisken J, Swoger J, Del Bene F, Wittbrodt J, Stelzer EHK. Optical sectioning deep inside live embryos by selective plane illumination microscopy. Science 2004 305 1007–1009. (https://doi.org/10.1126/science.1100035)
- 16↑
Isogai S, Horiguchi M, Weinstein BM. The vascular anatomy of the developing zebrafish: an atlas of embryonic and early larval development. Developmental Biology 2001 230 278–301. (https://doi.org/10.1006/dbio.2000.9995)
- 17↑
Isogai S, Lawson ND, Torrealday S, Horiguchi M, Weinstein BM. Angiogenic network formation in the developing vertebrate trunk. Development 2003 130 5281–5290. (https://doi.org/10.1242/dev.00733)
- 18↑
Campinho P, Vilfan A, Vermot J. Blood flow forces in shaping the vascular system: a focus on endothelial cell behavior. Frontiers in Physiology 2020 11 552. (https://doi.org/10.3389/fphys.2020.00552)
- 19↑
Chen Q, Jiang L, Li C, Hu D, Bu JW, Cai D, Du JL. Haemodynamics-driven developmental pruning of brain vasculature in zebrafish. PLoS Biology 2012 10 e1001374. (https://doi.org/10.1371/journal.pbio.1001374)
- 20↑
Hove JR, Koster RW, Forouhar AS, Acevedo-Bolton G, Fraser SE, Gharib M. Intracardiac fluid forces are an essential epigenetic factor for embryonic cardiogenesis. Nature 2003 421 172–177. (https://doi.org/10.1038/nature01282)
- 21↑
Kochhan E, Lenard A, Ellertsdottir E, Herwig L, Affolter M, Belting HG, Siekmann AF. Blood flow changes coincide with cellular rearrangements during blood vessel pruning in zebrafish embryos. PLoS ONE 2013 8 e75060. (https://doi.org/10.1371/journal.pone.0075060)
- 22↑
Lee J, Fei P, Packard RRS, Kang H, Xu H, Baek KI, Jen N, Chen J, Yen H & Kuo CC et al.4-Dimensional light-sheet microscopy to elucidate shear stress modulation of cardiac trabeculation. Journal of Clinical Investigation 2016 126 1679–1690. (https://doi.org/10.1172/JCI83496)
- 23↑
Lenard A, Daetwyler S, Betz C, Ellertsdottir E, Belting HG, Huisken J, Affolter M. Endothelial cell self-fusion during vascular pruning. PLoS Biology 2015 13 e1002126. (https://doi.org/10.1371/journal.pbio.1002126)
- 24↑
Ricard N, Simons M. When it is better to regress: dynamics of vascular pruning. PLoS Biology 2015 13 e1002148. (https://doi.org/10.1371/journal.pbio.1002148)
- 25↑
Hoefer IE, den Adel B, Daemen MJAP. Biomechanical factors as triggers of vascular growth. Cardiovascular Research 2013 99 276–283. (https://doi.org/10.1093/cvr/cvt089)
- 26↑
Stainier DY, Fishman MC. The zebrafish as a model system to study cardiovascular development. Trends in Cardiovascular Medicine 1994 4 207–212. (https://doi.org/10.1016/1050-1738(9490036-1)
- 27↑
Sehnert AJ, Stainier DYR. A window to the heart: can zebrafish mutants help us understand heart disease in humans? Trends in Genetics 2002 18 491–494. (https://doi.org/10.1016/s0168-9525(0202766-x)
- 28↑
Sehnert AJ, Huq A, Weinstein BM, Walker C, Fishman M, Stainier DYR. Cardiac troponin T is essential in sarcomere assembly and cardiac contractility. Nature Genetics 2002 31 106–110. (https://doi.org/10.1038/ng875)
- 29↑
Stainier DYR, Raz E, Lawson ND, Ekker SC, Burdine RD, Eisen JS, Ingham PW, Schulte-Merker S, Yelon D & Weinstein BM et al.Guidelines for morpholino use in zebrafish. PLoS Genetics 2017 13 e1007000. (https://doi.org/10.1371/journal.pgen.1007000)
- 30↑
Fortuna V, Pardanaud L, Brunet I, Ola R, Ristori E, Santoro MM, Nicoli S, Eichmann A. Vascular mural cells promote noradrenergic differentiation of embryonic sympathetic neurons. Cell Reports 2015 11 1786–1796. (https://doi.org/10.1016/j.celrep.2015.05.028)
- 31↑
Rödel CJ, Otten C, Donat S, Lourenço M, Fischer D, Kuropka B, Paolini A, Freund C, Abdelilah-Seyfried S. Blood flow suppresses vascular anomalies in a zebrafish model of cerebral cavernous malformations. Circulation Research 2019 125 e43–e54. (https://doi.org/10.1161/CIRCRESAHA.119.315076)
- 32↑
Watson O, Novodvorsky P, Gray C, Rothman AMK, Lawrie A, Crossman DC, Haase A, McMahon K, Gering M & Van Eeden FJ et al.Blood flow suppresses vascular Notch signalling via dll4 and is required for angiogenesis in response to hypoxic signalling. Cardiovascular Research 2013 100 252–261. (https://doi.org/10.1093/cvr/cvt170)
- 33↑
Serbanovic-Canic J, de Luca A, Warboys C, Ferreira PF, Luong LA, Hsiao S, Gauci I, Mahmoud M, Feng S & Souilhol C et al.Zebrafish model for functional screening of flow-responsive genes. Arteriosclerosis, Thrombosis, and Vascular Biology 2017 37 130–143. (https://doi.org/10.1161/ATVBAHA.116.308502)
- 34↑
Aleström P, D’Angelo L, Midtlyng PJ, Schorderet DF, Schulte-Merker S, Sohm F, Warner S. Zebrafish: housing and husbandry recommendations. Laboratory Animals 2020 54 213–224. (https://doi.org/10.1177/0023677219869037)
- 35↑
Westerfield M The Zebrafish Book: A Guide for Laboratory Use of Zebrafish (Brachydanio rerio). University of Oregon Press, 1993.
- 36↑
Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF. Stages of embryonic development of the zebrafish. Developmental Dynamics 1995 203 253–310. (https://doi.org/10.1002/aja.1002030302)
- 37↑
Chi NC, Shaw RM, De Val S, Kang G, Jan LY, Black BL, Stainier DYR. Foxn4 directly regulates tbx2b expression and atrioventricular canal formation. Genes and Development 2008 22 734–739. (https://doi.org/10.1101/gad.1629408)
- 38↑
Blum Y, Belting HG, Ellertsdottir E, Herwig L, Lüders F, Affolter M. Complex cell rearrangements during intersegmental vessel sprouting and vessel fusion in the zebrafish embryo. Developmental Biology 2008 316 312–322. (https://doi.org/10.1016/j.ydbio.2008.01.038)
- 39↑
Renshaw SA, Loynes CA, Trushell DMI, Elworthy S, Ingham PW, Whyte MKB. A transgenic zebrafish model of neutrophilic inflammation. Blood 2006 108 3976–3978. (https://doi.org/10.1182/blood-2006-05-024075)
- 40↑
Gray C, Loynes CA, Whyte MKB, Crossman DC, Renshaw SA, Chico TJA. Simultaneous intravital imaging of macrophage and neutrophil behaviour during inflammation using a novel transgenic zebrafish. Thrombosis and Haemostasis 2011 105 811–819. (https://doi.org/10.1160/TH10-08-0525)
- 41↑
Morsch M, Radford R, Lee A, Don EK, Badrock AP, Hall TE, Cole NJ, Chung R. In vivo characterization of microglial engulfment of dying neurons in the zebrafish spinal cord. Frontiers in Cellular Neuroscience 2015 9 321. (https://doi.org/10.3389/fncel.2015.00321)
- 42↑
Hamilton N, Rutherford HA, Petts JJ, Isles HM, Weber T, Henneke M, Gärtner J, Dunning MJ, Renshaw SA. The failure of microglia to digest developmental apoptotic cells contributes to the pathology of RNaseT2-deficient leukoencephalopathy. Glia 2020 68 1531–1545. (https://doi.org/10.1002/glia.23829)
- 43↑
Verduzco D, Amatruda JF. Analysis of cell proliferation, senescence and cell death in zebrafish embryos. Methods in Cell Biology 2011 101 19–38. (https://doi.org/10.1016/B978-0-12-387036-0.00002-5)
- 44↑
Zhang C, Li C, Jia X, Wang K, Tu Y, Wang R, Liu K, Lu T, He C. In vitro and in vivo anti-inflammatory effects of polyphyllin VII through downregulating MAPK and NF-κB pathways. Molecules 2019 24 875. (https://doi.org/10.3390/molecules24050875)
- 45↑
Kojima H, Nakatsubo N, Kikuchi K, Kawahara S, Kirino Y, Nagoshi H, Hirata Y, Nagano T. Detection and imaging of nitric oxide with novel fluorescent indicators: diaminofluoresceins. Analytical Chemistry 1998 70 2446–2453. (https://doi.org/10.1021/ac9801723)
- 46↑
Kugler E, Plant K, Chico T, Armitage P. Enhancement and segmentation workflow for the developing zebrafish vasculature. Journal of Imaging 2019 5 14. (https://doi.org/10.3390/jimaging5010014)
- 48↑
Lim J Two-Dimensional Signal and Image Processing, pp. 469–476. Englewood Cliffs, NJ: Prentice Hall, 1990.
- 49↑
Sternberg SR Biomedical image processing. Computer 1983 16 22–34. (https://doi.org/10.1109/MC.1983.1654163)
- 50↑
Burger W, Burge M. Digital Image Processing. London: Springer, 2008. (https://doi.org/10.1007/978-1-4471-6684-9)
- 51↑
Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S & Schmid B et al.Fiji – an Open Source platform for biological image analysis. Nature Methods 2012 9 676–682. (https://doi.org/10.1038/nmeth.2019)
- 52↑
Otsu N A threshold selection method from gray-level histograms. IEEE Transactions on Systems, Man, and Cybernetics 1979 9 62–66. (https://doi.org/10.1109/TSMC.1979.4310076)
- 53↑
Kugler E, Chico T, Armitage P. Image analysis in light sheet fluorescence microscopy images of transgenic zebrafish vascular development. In Medical Image Understanding and Analysis. MIUA 2018, pp. 343–353. Eds Nixon M, Mahmoodi S, Zwiggelaar R. Cham: Springer, 2018. (https://doi.org/10.1007/978-3-319-95921-4_32)
- 54↑
Kugler E, Chico T, Armitage PA. Validating segmentation of the zebrafish vasculature. In Medical Image Understanding and Analysis, pp. 270–281. Eds Zheng Y, Williams BM, Chen K. Cham: Springer International Publishing, 2020. (https://doi.org/10.1007/978-3-030-39343-4_23)
- 55↑
Kimura E, Isogai S, Hitomi J. Integration of vascular systems between the brain and spinal cord in zebrafish. Developmental Biology 2015 406 40–51. (https://doi.org/10.1016/j.ydbio.2015.07.015)
- 56↑
Goetz JG, Steed E, Ferreira RR, Roth S, Ramspacher C, Boselli F, Charvin G, Liebling M, Wyart C & Schwab Y et al.Endothelial cilia mediate low flow sensing during zebrafish vascular development. Cell Reports 2014 6 799–808. (https://doi.org/10.1016/j.celrep.2014.01.032)
- 57↑
Karthik S, Djukic T, Kim JD, Zuber B, Makanya A, Odriozola A, Hlushchuk R, Filipovic N, Jin SW, Djonov V. Synergistic interaction of sprouting and intussusceptive angiogenesis during zebrafish caudal vein plexus development. Scientific Reports 2018 8 9840. (https://doi.org/10.1038/s41598-018-27791-6)
- 58↑
Eltzschig HK, Carmeliet P. Hypoxia and inflammation. New England Journal of Medicine 2011 364 656–665. (https://doi.org/10.1056/NEJMra0910283)
- 59↑
Grimes AC, Stadt HA, Shepherd IT, Kirby ML. Solving an enigma: arterial pole development in the zebrafish heart. Developmental Biology 2006 290 265–276. (https://doi.org/10.1016/j.ydbio.2005.11.042)
- 60↑
Gebala V, Collins R, Geudens I, Phng LK, Gerhardt H. Blood flow drives lumen formation by inverse membrane blebbing during angiogenesis in vivo. Nature Cell Biology 2016 18 443–450. (https://doi.org/10.1038/ncb3320)
- 61↑
Xu J, Wang T, Wu Y, Jin W, Wen Z. Microglia colonization of developing zebrafish midbrain is promoted by apoptotic neuron and lysophosphatidylcholine. Developmental Cell 2016 38 214–222. (https://doi.org/10.1016/j.devcel.2016.06.018)
- 62↑
Bussmann J, Wolfe SA, Siekmann AF. Arterial-venous network formation during brain vascularization involves hemodynamic regulation of chemokine signaling. Development 2011 138 1717–1726. (https://doi.org/10.1242/dev.059881)21429983.
- 63↑
Packham IM, Gray C, Heath PR, Hellewell PG, Ingham PW, Crossman DC, Milo M, Chico TJA. Microarray profiling reveals CXCR4a is downregulated by blood flow in vivo and mediates collateral formation in zebrafish embryos. Physiological Genomics 2009 38 319–327. (https://doi.org/10.1152/physiolgenomics.00049.2009)
- 64↑
Geudens I, Coxam B, Alt S, Gebala V, Vion AC, Meier K, Rosa A, Gerhardt H. Artery-vein specification in the zebrafish trunk is pre-patterned by heterogeneous Notch activity and balanced by flow-mediated fine-tuning. Development 2019 146 dev181024. (https://doi.org/10.1242/dev.181024)
- 65↑
Weijts B, Gutierrez E, Saikin SK, Ablooglu AJ, Traver D, Groisman A, Tkachenko E. Blood flow-induced Notch activation and endothelial migration enable vascular remodeling in zebrafish embryos. Nature Communications 2018 9 5314. (https://doi.org/10.1038/s41467-018-07732-7)
- 66↑
Franco CA, Jones ML, Bernabeu MO, Vion AC, Barbacena P, Fan J, Mathivet T, Fonseca CG, Ragab A & Yamaguchi TP et al.Non-canonical Wnt signalling modulates the endothelial shear stress flow sensor in vascular remodelling. eLife 2016 5 e07727. (https://doi.org/10.7554/eLife.07727)
- 67↑
Bazmara H, Soltani M, Sefidgar M, Bazargan M, Mousavi Naeenian M, Rahmim A. The vital role of blood flow-induced proliferation and migration in capillary network formation in a multiscale model of angiogenesis. PLoS ONE 2015 10 e0128878. (https://doi.org/10.1371/journal.pone.0128878)
- 68↑
Neto F, Klaus-Bergmann A, Ong YT, Alt S, Vion AC, Szymborska A, Carvalho JR, Hollfinger I, Bartels-Klein E & Franco CA et al.YAP and TAZ regulate adherens junction dynamics and endothelial cell distribution during vascular development. eLife 2018 7 e31037. (https://doi.org/10.7554/eLife.31037)
- 69↑
Kwon HB, Wang S, Helker CSM, Rasouli SJ, Maischein HM, Offermanns S, Herzog W, Stainier DYR. In vivo modulation of endothelial polarization by apelin receptor signalling. Nature Communications 2016 7 11805. (https://doi.org/10.1038/ncomms11805)
- 70↑
Proulx K, Lu A, Sumanas S. Cranial vasculature in zebrafish forms by angioblast cluster-derived angiogenesis. Developmental Biology 2010 348 34–46. (https://doi.org/10.1016/j.ydbio.2010.08.036)
- 71↑
Siekmann AF, Standley C, Fogarty KE, Wolfe SA, Lawson ND. Chemokine signaling guides regional patterning of the first embryonic artery. Genes and Development 2009 23 2272–2277. (https://doi.org/10.1101/gad.1813509)
- 72↑
Vanhollebeke B, Stone OA, Bostaille N, Cho C, Zhou Y, Maquet E, Gauquier A, Cabochette P, Fukuhara S & Mochizuki N et al.Tip cell-specific requirement for an atypical Gpr124- and Reck-dependent Wnt/beta-catenin pathway during brain angiogenesis. eLife 2015 4 e06489. (https://doi.org/10.7554/eLife.06489)
- 73↑
Feng S, Bowden N, Fragiadaki M, Souilhol C, Hsiao S, Mahmoud M, Allen S, Pirri D, Ayllon BT & Akhtar S et al.Mechanical activation of hypoxia-inducible factor 1α drives endothelial dysfunction at atheroprone sites. Arteriosclerosis, Thrombosis, and Vascular Biology 2017 37 2087–2101. (https://doi.org/10.1161/ATVBAHA.117.309249)
- 74↑
Mahmoud MM, Serbanovic-Canic J, Feng S, Souilhol C, Xing R, Hsiao S, Mammoto A, Chen J, Ariaans M & Francis SE et al.Shear stress induces endothelial-to-mesenchymal transition via the transcription factor Snail. Scientific Reports 2017 7 3375. (https://doi.org/10.1038/s41598-017-03532-z)
- 75↑
Novodvorsky P, Chico TJA. The role of the transcription factor KLF2 in vascular development and disease. Progress in Molecular Biology and Translational Science 2014 124 155–188. (https://doi.org/10.1016/B978-0-12-386930-2.00007-0)
- 76↑
Serbanovic-Canic J, Souilhol C, Evans PC. Shear stress makes its mark on the endothelial genome. Cardiovascular Research 2019 115 1449–1451. (https://doi.org/10.1093/cvr/cvz088)
- 77↑
Souilhol C, Serbanovic-Canic J, Fragiadaki M, Chico TJ, Ridger V, Roddie H, Evans PC. Endothelial responses to shear stress in atherosclerosis: a novel role for developmental genes. Nature Reviews: Cardiology 2020 17 52–63. (https://doi.org/10.1038/s41569-019-0239-5)
- 78↑
Benz F, Wichitnaowarat V, Lehmann M, Germano RF, Mihova D, Macas J, Adams RH, Taketo MM, Plate KH & Guérit S et al.Low wnt/β-catenin signaling determines leaky vessels in the subfornical organ and affects water homeostasis in mice. eLife 2019 8 e43818. (https://doi.org/10.7554/eLife.43818)
- 79↑
Liang D, Chang JR, Chin AJ, Smith A, Kelly C, Weinberg ES, Ge R. The role of vascular endothelial growth factor (VEGF) in vasculogenesis, angiogenesis, and hematopoiesis in zebrafish development. Mechanisms of Development 2001 108 29–43. (https://doi.org/10.1016/s0925-4773(0100468-3)
- 80↑
Wiley DM, Jin SW. Bone morphogenetic protein functions as a context-dependent angiogenic cue in vertebrates. Seminars in Cell and Developmental Biology 2011 22 1012–1018. (https://doi.org/10.1016/j.semcdb.2011.10.005)
