1. Developmental Biology

Progressive mural cell deficiencies across the lifespan in a foxf2 model of cerebral small vessel disease

  1. Merry Faye E Graff
  2. Emma EM Heeg
  3. David A Elliott
  4. Sarah J Childs  Is a corresponding author
  1. Alberta Children’s Hospital Research Institute, University of Calgary, Canada
  2. Department of Biochemistry and Molecular Biology, University of Calgary, Canada
  3. Hotchkiss Brain Institute Advanced Microscopy Platform, University of Calgary, Canada
https://doi.org/10.7554/eLife.106720.3
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  1. Merry Faye E Graff
  2. Emma EM Heeg
  3. David A Elliott
  4. Sarah J Childs
(2026)
Progressive mural cell deficiencies across the lifespan in a foxf2 model of cerebral small vessel disease
eLife 14:RP106720.
https://doi.org/10.7554/eLife.106720.3
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eLife Assessment

This study provides important insights into mural cell dynamics and vascular pathology using a zebrafish model of cerebral small vessel disease. The authors present convincing evidence that partial loss of foxf2 function results in progressive, cell-autonomous defects in pericytes accompanied by endothelial abnormalities across the lifespan. By leveraging advanced in vivo imaging and genetic approaches, the work establishes zebrafish as a powerful and relevant model for dissecting the cellular mechanisms underlying cerebral small vessel disease.

https://doi.org/10.7554/eLife.106720.3.sa0

Abstract

Cerebral small vessel disease (SVD) is a leading cause of stroke and dementia and yet is often an incidental finding in aged patients due to the inaccessibility of brain vasculature to imaging. Animal models are important for modelling the development and progression of SVD across the lifespan. In humans, reduced FOXF2 is associated with an increased stroke risk and SVD prevalence in humans. In the zebrafish, foxf2 is expressed in pericytes and vascular smooth muscle cells and is involved in vascular stability. We use partial foxf2 loss of function (foxf2a-/-) to model the lifespan effect of reduced Foxf2 on small vessel biology. We find that the initial pool of pericytes in developing foxf2a mutants is strongly reduced. The few brain pericytes present in mutants have strikingly longer processes and enlarged soma. foxf2a mutant pericytes can partially repopulate the brain after ablation, suggesting some recovery is possible. Despite this capacity, adult foxf2a mutant brains show regional heterogeneity, with some areas of normality and others with severe pericyte depletion. Taken together, foxf2a mutants fail to generate a sufficient initial population of pericytes. The pericytes that remain have abnormal cell morphology. Over the lifespan, initial pericyte deficits are not repaired and lead to severely abnormal cerebrovasculature in adults. This work opens new avenues for modeling progressive genetic forms of human cerebral small vessel disease.

eLife digest

Every time you pause to think, remember a name, or read a sentence, the blood in your brain is quickly rerouted to the neurons doing the work. This redistribution depends on a vast network of blood vessels, from large arteries to microscopic capillaries, which deliver oxygen and energy directly to active brain cells.

For this system to function properly, the smallest blood vessels, the capillaries, must be able to regulate blood flow precisely. This control is provided by support cells on the outside of the capillary, such as pericytes or smooth muscle cells, which relax to open the vessel. When these cells fail, brain regions may no longer receive enough blood, even if larger vessels remain intact.

A breakdown of these cells is observed in cerebral small vessel disease, a leading cause of stroke and dementia. Unlike other types of strokes, this disease originates in the smallest blood vessels of the brain. However, it remains unclear whether it begins only in old age or much earlier in life. Understanding when and how this disease progresses is important because identifying its earliest mechanisms may offer opportunities to delay damage.

Graff et al. studied a zebrafish model carrying a mutation in foxf2a, which is linked to cerebral small-vessel disease in older humans. They found that the condition may not be exclusively age-related. When zebrafish had foxf2a levels reduced to about 50% of normal - similar to the reduction observed in humans with variants linked to cerebral small vessel disease - the fish developed blood vessel absormalities from the earlierst stages of life that persisted into adulthood. They also had fewer pericytes. Although pericytes could regenerate to some extent, blood vessel damage remained and worsened over the lifespan in this zebrafish model.

More detailed analyses revealed that pericytes showed signs of stress, which caused higher rates of cell death compared to zebrafish with normal foxf2a levels. In other words, although blood vessel damage could be partly repaired, it tended to deteriorate when foxf2a was absent.

These findings suggest that cerebral small vessel disease should may be better understood as a lifelong, progressive condition, where damage accumulates over time. Although approximately 20% of the population may carry genetic risk factors for this kind of disease, ongoing blood vessel damage and repair are common. Population-wide screening for individuals at risk of cerebral small vessel disease early in life, combined with targeted lifestyle and cardiovascular interventions, could greatly reduce the disease burden in the elderly.

Introduction

Brain microvessels supply billions of neurons and other brain cells with the oxygen and nutrients they need. Pathologies affecting brain microvasculature progress slowly and silently over a lifetime but have devastating consequences from decreased perfusion and vascular destabilization. Cerebral small vessel disease (CSVD) encompasses progressive heterogeneous changes in brain microvessels; it is the most common cause of vascular dementia and a significant contributor to stroke and cognitive decline (Østergaard et al., 2016). 25% of all strokes are the result of CSVD, yet effective targeted treatments remain elusive (Østergaard et al., 2016). This is in part due to the inability to both detect and assess progressive damage in the brain.

While there are several genes implicated in familial CSVD (i.e. NOTCH3, HTRA1, FOXC1, COL4A1, and COL4A2), there is a lack of suitable in vivo models for studying disease development and progression. Research has predominantly focused on NOTCH3, but over the last decade, FOXF2 has emerged as a risk locus for CSVD (Chauhan et al., 2016; Duperron et al., 2023). SNPs in the intergenic region between FOXF2 and FOXQ1 decrease FOXF2 expression and significantly increase stroke risk due to the variant decreasing the efficiency of ETS1 binding to a novel FOXF2 enhancer (Ryu et al., 2022).

Foxf2 promotes mural cell differentiation and vascular stability in the zebrafish brain and is expressed highly in brain pericytes (Ahuja et al., 2024 #1591); (Chauhan et al., 2016 #1403); (Reyahi et al., 2015 #1372); (Ryu et al., 2022 #1590). Brain pericytes interact closely with endothelial cells, contributing to extracellular matrix (ECM) deposition and blood-brain barrier (BBB) formation, in addition to providing vasoactivity and stability (Bahrami and Childs, 2020; Daneman et al., 2010; Dave et al., 2018; Stratman et al., 2009). In animal models, an absence of brain pericytes results in hemorrhages and accelerates vascular-mediated neurodegeneration (Bell et al., 2010; Wang et al., 2014). Foxf2 is clearly important for vascular stability across species, as loss of Foxf2 in mice and zebrafish leads to increased brain hemorrhage and alterations in brain pericyte numbers and differentiation (Chauhan et al., 2016; Reyahi et al., 2015; Ryu et al., 2022). Brain tissue from patients with aging-related dementias (i.e. post-stroke dementia, vascular dementia, Alzheimer’s disease) has reduced deep white matter pericytes and associated BBB disruption (Ding et al., 2020), suggesting that pericytes should be examined as mediators of CSVD progression in patients with FOXF2 deficiency.

We previously showed that complete loss of foxf2 in foxf2aca71; foxf2bca21 double-homozygous mutants in late embryogenesis leads to reduced brain pericyte numbers (Ryu et al., 2022). However, stroke susceptibility in humans is associated with reduced, but not absent, FOXF2 expression. Genome-wide association (GWA) indicates that carrying a minor allele of an SNP in a FOXF2 enhancer leads to reduced, but not absent, FOXF2 and is associated with stroke (Ryu et al., 2022). For this reason, we model CSVD using a zebrafish with reduced Foxf2 dosage using single homozygous foxf2a mutants. Zebrafish foxf2a and foxf2b genes are the result of genome duplication in zebrafish ~430 million years ago and have similar gene expression (Arnold et al., 2015; Chauhan et al., 2016). We have detected no difference in function between the two genes and, therefore, foxf2a loss of function may be similar to human heterozygous loss of FOXF2 function, a state that is observed in the population in GnomAD (Chen et al., 2024).

Strikingly, while pericytes in embryonic foxf2 mutants are clearly affected, foxf2 mutants can survive until adulthood, albeit with a reduced lifespan. How pericytes change across the lifespan while CSVD progresses is unknown. Here, we find that foxf2a mutants have significantly reduced brain pericyte numbers as embryos that do not recover over time. Pericytes in mutant embryos and larvae exhibit morphological abnormalities, including increased soma size, longer processes, and degeneration. We show that processes and soma in the adults are also abnormal, though their morphology differs over the lifespan. Although the initial pool of pericytes is smaller, mutants can regenerate pericytes after ablation. Our analysis suggests that foxf2 is required within pericytes to modulate numbers but also has a strong effect on morphology. We show that brain pericytes may contribute to the pathological progression of genetic CSVD, starting in embryonic development and continuing across the lifespan. Understanding the early developmental aspects of late-onset vascular conditions like CSVD will aid in the development of effective therapeutic strategies.

Results

Pericyte number is consistently lower in foxf2 mutant embryos and larvae

Embryonic phenotypes can lead to lifelong consequences. We have previously studied foxf2a;foxf2b double mutants only at a single embryonic stage at 3 days post-fertilization (dpf). To understand how a pericyte and cerebrovascular phenotype evolves and/or resolves over development, we conducted serial imaging of individual brains of foxf2a mutants at embryonic stages (3 and 5 dpf), and at larval stages (7 and 10 dpf). Mutants were live imaged using endothelial (kdrl:mCherry) and pericyte (pdgfrβ:Gal4, UAS:GFP) transgenic lines.

Embryonically, brain pericytes have a thin-strand morphology and are closely associated with, and extend processes over, the endothelium in the midbrain and hindbrain of zebrafish (Figure 1A–A’’). In wild-type embryos, the number of pericytes increases progressively from 3 through 10 dpf (Figure 1B). However, foxf2a mutants show significantly fewer pericytes on brain vessels at 3 dpf (Figure 1C; mean 21 in wild-type and 10 in mutants), and this pericyte deficiency persists through 5, 7, and 10 dpf (Figure 1C–D). The reduction in pericyte numbers shows variable penetrance, with some foxf2a mutants having pericyte numbers only slightly reduced from wild-type, and others that are severely diminished. The same pattern of pericyte reduction is seen with foxf2a mutants from a homozygous or heterozygous incross, suggesting there is no maternal effect (Figure 1E–F). Serial imaging of mutants with regional absence in earlier stages shows that defects in pericyte coverage persist into later stages, suggesting that the size of the initial pericyte population is a key determinant of later coverage (Figure 1—figure supplement 1). Double foxf2a;foxf2b mutants have a fully penetrant phenotype with significantly fewer brain pericytes in mutants than wild-type at every stage (Figure 1—figure supplement 2; mean 19 in the wild-type and 9 in mutants at 3 dpf). Incomplete penetrance in foxf2a single mutants could be due to genetic compensation from foxf2b. While foxf2b is not significantly upregulated in foxf2a mutants on average, individual embryos have highly variable foxf2b expression (Figure 1—figure supplement 3).

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Brain pericyte number is consistently lower and does not recover in foxf2a mutant larvae.

(A) Zebrafish brains were imaged using endothelial (red; Tg(kdrl:mCherry)) and pericyte (light blue; Tg(pdgfrβ:Gal4, UAS:GFP)) transgenic lines (arrows: brain pericytes). (A’-A’’) Brain pericyte soma (white arrows) and processes (yellow arrows) are closely associated with the endothelium. (B) Serially imaged wild-type and foxf2a mutant brains at 3, 5, 7, and 10 dpf. (C) Total brain pericyte numbers at 3, 5, 7, and 10 dpf. (D) Individual brain pericyte trajectories of serially imaged embryos over the same period. (E) Dorsal images of embryos for the indicated genotypes from a foxf2a heterozygous incross at 75 hpf. (F) Total brain pericytes at 75 hpf. Statistical analysis was conducted using multiple Mann-Whitney tests (C) and one-way ANOVA with Tukey’s test (F). Scale bars, 50 µm (A–B, E).

Figure 1—source data 1

All raw quantitative data underlying Figure 1 and supplements.

https://cdn.elifesciences.org/articles/106720/elife-106720-fig1-data1-v1.xlsx

Since Foxf2 conditional knockout mice show reduced expression of the pericyte marker Pdgfrβ, we tested whether pdgfrβ expression is reduced in foxf2a mutants, as this might introduce inaccuracies in pericyte counting. We used two methods, quantitative hybridization chain reaction (HCR) in situ hybridization and integrated density of the pdgfrβ transgene expression in wild-types and mutants carrying only a single copy of the transgene. We show that pdgfrβ mRNA nor transgene expression is not reduced in foxf2a mutants (Figure 1—figure supplement 3). Furthermore, we show that pdgfrβ has complete overlap in expression with other brain pericyte markers, such as ndufa4l2 and foxf2b using HCR (Figure 2A–D). Thus, pdgfrβ transgene or mRNA expression can reliably be used to count zebrafish brain pericytes in foxf2a mutants.

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Loss of foxf2 affects embryonic pericyte numbers, but not endothelial cell pattern.

(A) foxf2a expression in wild-type brains at 72 hpf using hybridization chain reaction (HCR) shows co-expression with pericyte marker ndufa4l2a. foxf2a is also lowly co-expressed in the endothelium with kdrl. Arrows show overlapping expression. (B) foxf2b is co-expressed with pericyte marker pdgfrβ, also lowly expressed in the endothelium (kdrl). (C) foxf2b and pdgfrβ are expressed in a similar, overlapping pattern in pericytes of wild-type and foxf2a mutants. (D) Pericyte marker nduf4al2a and pdgfrβ are expressed in a similar, overlapping pattern in pericytes in wild-type and foxf2a mutants. (E) Image of endothelium used to generate the total blood vessel network length. (F) Total vessel network length from Vessel Metrics software. (G) Scatter plot of hindbrain CtA diameters. (H) Scatter plot of pericyte density and pericyte coverage (I). Statistical analysis was conducted using one-way ANOVA with Tukey’s test. Scale bars, 10 µm (A–D), 50 µm (F).

Figure 2—source data 1

All raw quantitative data underlying Figure 2 and supplements.

https://cdn.elifesciences.org/articles/106720/elife-106720-fig2-data1-v1.xlsx

Although we focus on mural cells, Foxf2 is expressed in adult mouse brain endothelium (Vanlandewijck et al., 2018). We assessed foxf2a and foxf2b mRNA expression patterns using HCR in situ hybridization. At 3 dpf, foxf2a is co-localized with the pericyte marker ndufa4l2a in brain pericytes and shows low-level co-localized expression with the blood vessel marker kdrl in brain endothelium (Figure 2A). The expression pattern of foxf2b is similar. It is expressed in pericytes (pdgfrβ) and only weakly in endothelial cells (kdrl) (Figure 2B). This is supported by the DanioCell atlas of single-cell sequencing of multiple embryonic stages that shows expression of both genes is strongest in mural cells, pericytes, and vascular smooth muscle cells and low in endothelial cells (Sur et al., 2023; Figure 2—figure supplement 1). Thus, while foxf2a and foxf2b are principally expressed in pericytes, they are lowly expressed in endothelium during brain development.

Pericyte loss or impairment leads to alterations in vascular patterning in the mouse retina (Eilken et al., 2017). To assess if the endothelial network is affected in foxf2a mutants, we employed a Python workflow using Vessel Metrics (McGarry et al., 2024; Figure 2E). We found no statistical difference in total network length between wild-type and mutants at 3 dpf (Figure 2F) or hindbrain central artery diameter (Figure 2G). However, pericyte density (number of pericytes divided by the total network length) is reduced by 40% in foxf2a mutants (Figure 2H), reflecting the loss of pericytes with no change in vessel network length. Similarly, pericyte coverage of vessels (total process coverage from brain pericytes divided by the total network length) is reduced by 39% in mutants (Figure 2I). Our data suggest that during early development, foxf2a depletion primarily affects pericytes.

Early defects in brain vessel development have lifelong consequences

foxf2 mutant animals can survive to adulthood, albeit with a reduced lifespan (~1 year vs. >2 years). Are early pericyte deficiencies repaired, or is loss of pericytes unimportant to survival to adulthood? To understand how brain pericyte phenotypes evolve over the lifespan, we dissected adult wild-type and foxf2a mutant brains on pericyte and endothelial double transgenic backgrounds and imaged after iDISCO clearing. Gross measurements of standard length of the fish (snout to tail) show no significant differences between wild-type and mutants, except that female mutant brain length and width are significantly smaller at 11 months post fertilization (mpf) (Figure 2—figure supplement 2). However, there is no significant difference in the proportional brain length/standard length ratio in foxf2a mutants vs. wild-type.

Projected 3D views of light sheet images of the whole brain show striking defects in pericyte density, coverage, and vascular pattern in foxf2a mutants vs. wild-type adults (Videos 1 and 2; Figure 3A–B). Pericyte distribution is irregular, and blood vessel density is visibly reduced in mutant brains (Figure 3C–D). Using a machine learning workflow (Figure 3—figure supplement 1, Figure 3—source data 1), we segmented pericyte soma, or the vessel backbone for the entire adult brain, to count the total pericyte number in comparison to the total endothelial network length. We find a significant reduction in pericyte numbers in 3 mpf foxf2a mutant brains, which have only 45% of the pericytes of a wild-type brain (average of 24,567 pericytes/brain vs. wild-type 54,833 pericytes/brain; n=3 of each genotype; Figure 3E). However, at this stage, the vessel network length is not statistically different (Figure 3F). The density of pericytes on vessels is significantly decreased from 0.01 pericyte/µm to 0.006 pericytes/µm in foxf2a mutants showing a clear deficit (Figure 3G). In contrast, the mean vessel diameter across vessel segments is not significantly different at 3 mpf when all diameters are considered, nor when vessel diameters are grouped in 5 µm bins (n=522,037 wild-type and 381,544 foxf2a mutant diameters; Figure 3H–I).

Video 1
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Rotating view of a cleared wild-type brain at 3 mpf with pdgfrβ (blue) and kdrl (red).
Figure 3 with 5 supplements see all
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foxf2a mutants show strong brain vascular defects as adults.

(A–B) 3D projections of iDISCO-cleared immunostained whole wild-type and foxf2a-/- brains at 3 mpf, viewed ventrally. (C–D) Wild-type and foxf2a mutant 2 brain regions, viewed dorsally (arrows = defects in coverage). (E) Number of brain pericytes in three individual wild-type and mutant brains at 3 mpf detected using Imaris’ spot tool and machine learning. (F) Total vessel network length in three individual wild-type and mutant brains at 3 mpf using Ilastik and Imaris’ filament tool and machine learning. (G) Brain pericyte density calculated using number of brain pericytes per meter of vessel length. (H) Vessel diameter in three individual wild-type and mutant brains at 3 mpf using Imaris’ filament tool and machine learning. (I) Percentage of vessel segments in wild-type and mutant brains at 3 mpf segregated by vessel diameter (in 5 μm bins). (J) CUBIC-cleared wild-type and foxf2a mutant midbrain at 11 mpf (arrows: individual pericyte soma). C=caudal, D=dorsal, R=rostral, V=ventral. Statistical analysis was conducted using unpaired t-tests (E–H) and ANOVA with Dunnett’s post hoc test (I). Scale bars, 500 μm (A–B), 200 μm (C–D), 50 μm (J).

Figure 3—source data 1

All raw quantitative data underlying Figure 3 and supplements.

https://cdn.elifesciences.org/articles/106720/elife-106720-fig3-data1-v1.xlsx
Video 2
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Rotating view of a cleared foxf2a mutant brain at 3 mpf with pdgfrβ (blue) and kdrl (red).

Similarly, cleared dissected brains at 11 mpf were imaged and showed similarly striking vascular defects (Figure 3—figure supplement 2). Adult pericytes have a clear, oblong cell body with long, slender primary processes that extend from the cytoplasm and secondary processes that wrap around the circumference of the blood vessel (Figure 3J). These pericytes form a continuous network of processes to cover the blood vessels in the brain. In foxf2a 11 mpf brains, pericyte somas lose their oblong shape, and cell bodies cannot be easily distinguished from processes. Mutant cells also extend thickened linear processes, with no secondary processes encircling the vessels. Mutant pericytes do not form an extensive network with other neighbouring pericytes. We note that there is variable phenotypic penetrance in foxf2a adults at 3 mpf that is reflective of the incomplete penetrance in foxf2a embryos (Figure 3—figure supplement 2).

To view cellular morphology, we sectioned adult brains and immunolabeled pericytes and endothelium. In wild-type adult brains, we identified three subtypes of pericytes: ensheathing, mesh, and thin-strand, previously characterized in murine models (Berthiaume et al., 2018b; Figure 3—figure supplement 3). In comparing brain sections from both wild-type and foxf2a mutants, on smaller vessels, mutant pericytes exhibit more linear processes with barely distinguishable soma, markedly differing from the characteristic appearance of wild-type adult pericytes (Figure 3—figure supplement 4). Similarly, pdgfrβ-expressing cells on large-calibre vessels (mural cells, likely vascular smooth muscle cells (vSMCs)) show alterations in morphology and coverage (Figure 3—figure supplement 4). Although mutant embryos do not exhibit apparent abnormalities in their endothelium, large aneurysm-like structures are evident in the adult brain (Figure 3—figure supplement 5). These structures also appear to have decreased kdrl expression. Thus, both whole-mount and sectioned tissue show that brain vascular mural cell number and morphology are severely impacted in adult foxf2a-/- mutant brains, with increasing involvement of the endothelium, suggesting a worsening phenotype throughout life.

As foxf2a is expressed in vSMCs, we tested the effect of loss of foxf2a on vSMCs using confocal imaging of larvae and light sheet microscopy of cleared dissected 6 mpf adult wild-type and foxf2a mutant brains stained for transgenic endothelial (kdrl) and vSMC (acta2) markers. acta2-positive vSMCs are present on brain vessels around the Circle of Willis at embryonic (5 dpf) and larval (10 dpf) stages in both mutant and wild-type (Figure 4A and C). Furthermore, the number of vSMCs does not differ between mutant and wild-type (Figure 4B and D; Figure 4—source data 1). In adult brains, we find no significant difference in the total length of vSMCs in the brain, or in vSMC coverage (proportion of vessels covered by vSMCs) (n=3 of each genotype; Videos 3 and 4; Figure 4E–H). We note that at 6 mpf, there is a significant decrease in vessel network length, however (Figure 4G).

Figure 4
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Loss of foxf2a has no impact on acta2-expressing brain vascular smooth muscle cells.

(A) foxf2a+/++ and foxf2a-/- larvae at 5 dpf showing vSMC coverage in the brain using endothelial (red; Tg(kdrl:mCherry)) and vascular smooth muscle cell (vSMC) (light blue; Tg(acta2:GFP)) transgenic lines. (B) Scatter plot of total vSMCs at 5 dpf. (C) foxf2a+/++and foxf2a-/- larvae at 10 dpf showing vSMC coverage in the brain. (D) Scatter plot of total vSMCs at 10 dpf. (E) 3D projections of iDISCO-cleared immunostained whole wild-type and foxf2a-/- brains at 6 mpf, viewed ventrally. (F) Total vSMC length at 6 mpf using Imaris’ filament tool and machine learning. (G) Total vessel network length at 6 mpf Ilastic and Imaris’ filament tool and machine learning. (H) vSMC coverage per total blood vessel network length at 6 mpf. C=caudal, D=dorsal, R=rostral, V=ventral. Statistical analysis was conducted using unpaired t-tests. Scale bars, 20 µm (A, C), 200 μm (E).

Figure 4—source data 1

All raw quantitative data underlying Figure 4.

https://cdn.elifesciences.org/articles/106720/elife-106720-fig4-data1-v1.xlsx
Video 3
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Rotating view of a cleared wild-type brain at 6 mpf with acta2 (blue) and kdrl (red).
Video 4
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Rotating view of a cleared foxf2a mutant brain at 6 mpf with acta2 (blue) and kdrl (red).

Morphological abnormalities emerge in the larval brain of mutants

Considering the altered pericyte morphology of adult mutants, we revisited developmental stages to identify when these abnormalities first arise. We analyzed morphology at 3 and 10 dpf using in vivo confocal imaging to understand defects in the cell body (soma) and processes (Figure 5A). We found no significant difference in the soma area at 3 dpf, but at 10 dpf, there is a significant increase in mutant pericyte soma area compared to wild-type (Figure 5B). In parallel, wild-type pericytes undergo a slight reduction in soma size from 3 to 10 dpf.

Figure 5
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foxf2a mutant brain pericytes show increased soma size and process length.

(A) Wild-type and foxf2a-/- mutant brain pericytes at 3 and 10 dpf with tracings of individual pericytes (indicated by arrows). (B) Brain pericyte soma area at 3 and 10 dpf. (C) Multispectral Zebrabow labelling reveals pericyte-process interactions in the larval brain. (arrows: pericyte interaction points). (D) Total process length per pericyte at 3 and 10 dpf. (E) Varying pericyte-pericyte interactions at 10 dpf (arrows: interaction points). (F) Number of each type of interaction at 10 dpf. (G) Length of overlap when process interaction occurs. Statistical analysis was conducted using multiple Mann-Whitney tests in B, a one-way ANOVA with Tukey’s test at 3 dpf and a Kruskal-Wallis test with Dunn’s multiple comparisons test at 10 dpf in D. Scale bars, 25 µm (A), 20 µm (C), 5 µm (E).

Figure 5—source data 1

All raw quantitative data underlying Figure 5.

https://cdn.elifesciences.org/articles/106720/elife-106720-fig5-data1-v1.xlsx

The low number of brain pericytes in foxf2a mutants means that distinct pericyte processes can be distinguished and measured. However, in wild-types, pericyte processes are not easily distinguished. For accurate process length measurements in wild-type, we used multispectral labelling with a pericyte-specific Zebrabow transgenic line (pdgfrβ:Gal4, UAS:Zebrabow) (Pan et al., 2013; Whitesell et al., 2019). Cre mRNA was injected at the one-cell stage to activate random recombination, allowing us to visualize individual neighbouring pericytes (Figure 5C). Wild-type processes have a mean length of 81.3 µm vs 102.9 in the mutant at 3 dpf and 96.3 µm vs 147.6 in the mutant at 10 dpf (Figure 5D). Thus, the pericyte process lengths are significantly increased in foxf2a mutants at 3 and 10 dpf (Figure 5D).

Through the study of wild-type processes using multispectral labelling, we observed some differences in the behaviour of adjacent pericyte processes from that published in the adult mouse brain (Berthiaume et al., 2018a). While direct contact with no overlap between pericytes is the most common interaction in developing zebrafish, similar to mouse adult brain pericytes (Figure 5F), we also see some overlap (Figure 5E). The average overlap between wild-type pericyte processes at 10 dpf was 5.9 µm (Figure 5G). Together, our data show overlap in brain pericyte processes in wild-type animals.

To further explore mutant pericyte behaviour, we conducted serial imaging during larval development. Over time, some mutant pericyte processes form disconnected bead-like blebs with cell bodies that disappear over time (Figure 6A). This pericyte phenotype can occur on vessels with full patency and is not associated with endothelial regression. The bead-like remnants are highly prevalent during larval development in foxf2a-/- mutants but not present in wild-type (Figure 6B). To visualize process degeneration in real-time, we time-lapse imaged from 4 to 5 dpf (Figure 6C). The pericyte can undergo a process reminiscent of cell death with soma blebbing and process degeneration phenotypically resembling neural dendrite degeneration or pruning (Figure 6D).

Figure 6
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foxf2a mutant pericytes degenerate.

(A) foxf2a-/- mutant pericyte at 10 and 13 dpf with the degenerating process and cell body with a wild-type control from the same brain region (arrows: individual pericyte). (B) Bar graph with process blebbing phenotype penetrance in wild-type and mutant brains (n=total samples examined). (C) Time-lapse of a foxf2a-/- mutant midbrain from 4 to 5 dpf (arrows: individual pericyte). (D) Inset of mutant pericyte undergoing degeneration (arrows: blebbing). Scale bars, 20 µm (A, C).

Figure 6—source data 1

All raw quantitative data underlying Figure 6.

https://cdn.elifesciences.org/articles/106720/elife-106720-fig6-data1-v1.xlsx

In summary, larval foxf2a mutant pericytes show reduced numbers, increased soma size, and elongated processes with evidence of process degeneration. In addition, we make the novel observation of overlapping pericyte processes during zebrafish development.

Foxf2a mutants do not have an impaired capacity to repopulate brain pericytes

Zebrafish have regenerative capacity in various tissues (Becker et al., 1997; Lepilina et al., 2006; Otteson and Hitchcock, 2003), yet foxf2a mutant embryos that are pericyte-deficient maintain strong brain pericyte defects through aging, which suggests that either they may not be able to replenish absent/damaged pericytes, or that the smaller size of the initial pool in embryogenesis limits repair such that numbers never can catch up to wild-type. To differentiate these hypotheses, we tested whether foxf2a mutants lack the capacity to regenerate pericytes. We reduced pericyte numbers in a foxf2a heterozygous incross using a cell ablation strategy. Zebrafish expressing pdgfrβ:Gal4, UAS:NTR-mCherry, and flk:GFP transgenes were treated with 5 mM metronidazole (MTZ) at 50 hpf for 1 hr which ablates most pericytes. MTZ is a prodrug substrate that elicits cell death in nitroreductase (NTR)-expressing cells due to its cytotoxic derivatives. We then imaged and counted brain pericytes at 3 dpf (a day after treatment) and 10 dpf (recovery).

At 3 dpf, in the vehicle (DMSO) control group, there is the expected significant difference between wild-type and mutant pericyte numbers (Figure 7A and B; Figure 7—source data 1). When treated with metronidazole, both mutants and wild-types both have a similar, severe reduction in pericytes at 3 dpf post-ablation and are not significantly different from each other (Figure 7B). By 10 dpf, pericytes are partially repopulated in both wild-type and mutant ablated groups (Figure 7C). Surprisingly, given the initial pericyte number defects in foxf2a mutant embryos, we find no significant difference between any groups (Figure 7D; mean 70 in wild-type DMSO treated, 53 in mutant DMSO treated, 43 in wild-type MTZ treated, 33 in mutant MTZ treated). This shows that foxf2a mutants retain their ability to repopulate pericytes following ablation. This data is important to distinguish the timing and mechanism of pericyte reduction in foxf2a mutants. While foxf2a mutant pericytes can regenerate following induced catastrophic loss, it cannot compensate for the initially smaller pool of pericytes during embryogenesis. This data suggests that foxf2a plays a critical role in the establishment of a properly sized initial pericyte pool during early development. Regeneration mechanisms, although intact, cannot fully compensate for this initial reduction in pericytes.

Figure 7
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foxf2a mutants regenerate brain pericytes normally after genetic ablation.

Zebrafish brains were imaged using endothelial (red; Tg(kdrl:GFP)) and pericyte (light blue; Tg(pdgfrβ:Gal4, UAS:NTR-mCherry)) transgenic lines. (A) Wild-type and mutant brains at 3 dpf in control (DMSO) and treated (MTZ) groups. (B) Total brain pericytes at 3 dpf. (C) Wild-type and mutant brains at 10 dpf. (D) Total brain pericytes at 10 dpf. Statistical analysis was conducted using a one-way ANOVA (B) or Kruskal-Wallis test with Dunn’s multiple comparisons test (D). Scale bars, 50 μm (A, C).

Figure 7—source data 1

All raw quantitative data underlying Figure 7.

https://cdn.elifesciences.org/articles/106720/elife-106720-fig7-data1-v1.xlsx

Discussion

Our reduced dosage model of Foxf2 demonstrates disease processes at the cellular level in intact animals, giving insight into pathological changes that occur during CSVD that have not been observed in other models or humans. Our data supports a developmental origin for this type of CSVD, which then progresses and evolves across the lifespan (Figure 8).

Figure 8
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Model of foxf2a mutant brain pericyte defects over the lifespan.

Wild-type pericytes develop normally in the embryo and establish extensive, continuous coverage over vessels by adulthood. foxf2a mutant pericytes exhibit abnormal morphology during development that worsens over the lifespan, with mutant vessels developing atypical morphology and discontinuous coverage.

Dosage sensitivity of Foxf2

The logic for using a reduced dosage of Foxf2 in our studies is to better match the effect of common population variants leading to CSVD. We previously modelled the effect of a high-risk CSVD and stroke-associated SNV on the expression of FOXF2 in human cells, showing that it can reduce expression by ~50% (Ryu et al., 2022). Since zebrafish have two FOXF2 orthologs (foxf2a and foxf2b), foxf2a homozygotes have equivalent FOXF2 dosage as human heterozygotes, assuming that foxf2a and foxf2b not only exhibit similar expression but also share similar functions. Foxf2 is a dosage-sensitive gene, as zebrafish foxf2a heterozygotes show a significant reduction in brain pericytes. Similarly, mouse Foxf2 is also dosage sensitive (Reyahi et al., 2015). We find that foxf2a mutants are variably penetrant, while foxf2a;foxf2b double mutants are fully penetrant. Genetic compensation is common with gene duplication (Kok et al., 2015; Rouf et al., 2023) and would explain variability in disease phenotypes in different individuals with the same mutation.

Impaired embryonic pericyte coverage in Foxf2 deficiency

We show that numbers of brain pericytes are reduced at multiple developmental and adult stages when foxf2a and/or foxf2a;foxf2b are lost. The deficiency is significant at every stage, including 3 dpf, the earliest time point that pericytes are robustly observed in development. Deficient numbers could be due to a reduction in the pericyte precursor population (i.e. nkx3.1 positive cells Ahuja et al., 2024), or to an inability of foxf2a-deficient cells to differentiate into pdgfrβ-positive pericytes. Our data does not allow us to distinguish these, although pdgfrβ expression in foxf2a mutants is transcriptionally unchanged, suggesting that pericyte number in early development is the primary phenotype. Additional evidence for pericytes being the primary cells affected is that there is no difference in total vessel network length or hindbrain CtA diameter at 3 dpf in foxf2a heterozygotes or mutants when pericyte numbers are reduced. A reduction in pericytes is expected to have early consequences, as pericytes provide signals to the endothelium for quiescence and arterial-venous identity (Mäe et al., 2021 #1750), as well as allowing contractility of cerebral blood vessels in early development (Bahrami and Childs, 2020). Fewer pericytes distributed on a normal-sized endothelial network length result in reduced vessel coverage. It is intriguing, therefore, that pericyte process length is increased at both 3 and 10 dpf, potentially to compensate for low pericyte density. A similar elongated pericyte phenotype and reduced coverage is seen in mice with constitutive Pdgfβret knockout (Mäe et al., 2021). The convergent phenotypes after manipulation of two genes (foxf2a in fish and Pdgfβ ret in mice) that reduce pericyte number suggest that the intrinsic pericyte response to depletion of pericyte density is to elongate, perhaps to attempt to cover ‘naked’ vessels. We note that previous mouse Foxf2 knockouts using a conditional Wnt1-Cre driver saw contrasting results to ours, with increased pericyte numbers, although a similar loss of vascular stability is seen in both models (Reyahi et al., 2015). The reason for the difference in the direction in pericyte number may be experimental due to the different knockout technique (conditional in mice which only removes neural crest-derived pericytes vs. full knockout which removes mesodermal, neural crest-derived and endothelial foxf2a in fish), or the difference could reflect a species-specific difference.

Insights on fundamental pericyte process properties

The reasons behind the hypergrowth of pericyte processes in foxf2a-/- mutant brains is unclear. Berthiaume et al. (Berthiaume et al., 2018b) propose that repulsive interactions among pericytes in the adult brain establish boundaries between adjacent pericytes, preventing their processes from overlapping. In this case, one might hypothesize that foxf2a mutant pericytes, which are less dense along vessels, would lack feedback from adjacent pericytes, leading to uncontrolled growth.

In mice, adult brain pericytes form a non-overlapping network along capillaries, although their processes occasionally approach each other without overlapping (Berthiaume et al., 2018a; Hill et al., 2015). Overlap between pericyte processes have not been well studied in the developing brain, but pdgfrb+ve cells in developing zebrafish fin also overlap (Leonard et al., 2022). Strikingly, using multispectral (Zebrabow) labeling, we find processes overlap in the developing zebrafish brain. Overlaps occur on capillary segments and at branch points. Previous studies in mice were conducted in the adult brain, and it is possible that overlapping processes in the developing brain are transient, zebrafish-specific, or were potentially overlooked in the mouse experiments.

We have also observed forked extensions at the tips of some pericyte projections at 10 dpf in wild-type embryos, where pericytes are not in contact. Whether these unique structures are involved in attractive/repulsive signals between pericytes or help determine the direction and extent of process growth is unclear, but understanding the fundamental mechanisms governing pericyte-pericyte process interactions would yield valuable insights into development and disease, as well as into how pericyte depletion results in abnormally long processes.

Morphological abnormalities in pericyte soma in foxf2a mutants

A second morphological abnormality of foxf2a-deficient pericytes is that their soma size is increased at 10 dpf, which may also be a compensatory change. Pathological changes in soma size have not been observed in brain pericytes. However, increased neuronal soma size is observed in Amyotrophic Lateral Sclerosis or Lhermitte-Duclos (Dukkipati et al., 2018; Kwon et al., 2001) and is linked to mTor signaling (Kwon et al., 2001). In neuronal populations, moderate soma swelling can be an adaptation for survival, while rapid, drastic swelling indicates imminent death (Rousseau et al., 1999). It is, therefore, not surprising that we observe degeneration of pre-existing pericytes in foxf2a mutant but not wild-type animals. This degeneration phenotypically resembles neuronal dendrite remodelling and pruning during development (Fukui et al., 2012; Greenwood et al., 2007). Further work to test whether pericytes share similar mechanisms of degeneration in response to stress or cellular damage may provide further insight into phenotypic progression.

Changes in foxf2a mutants across the lifespan

For the first time, we have examined how vascular phenotypes change in the adult brain using iDISCO clearing. We find that adult soma take on very unusual, ‘stiff’ shapes and become almost indistinguishable from processes. The adult processes are shorter and more linear than the embryonic processes. Hypertrophic embryonic pericytes either do not survive to adulthood or undergo morphological changes over time. Given the limited knowledge in this area, the underlying factors driving this shift in pericyte morphology are unclear. Actin is present near the plasma membrane of cells, where it provides structural and mechanical support, enabling motility and determining cell structure. Given the abnormal shapes of adult pericytes, alterations in actin may contribute to their irregular morphology.

In the adult foxf2a mutant brain, abnormal pdgfrβ-expressing cells are observed on large-calibre vessels and likely represent vSMCs. We observed large-calibre aneurysm-like vessels, suggesting that vSMCs may lose functionality over time, thereby increasing the vulnerability of underlying vessels to dilation and weakening. Loss of acta2 and, subsequently, vSMCs has been associated with conditions, such as thoracic aortic aneurysms and dissections (Guo et al., 2009). However, we found no significant change in either vSMC cell number in embryonic development or in network length of vSMCs in the adult brain, suggesting that the primary phenotype in foxf2a mutants is in pericyte cells and that any changes to vSMCs are likely secondary.

Similarly, brain vessel diameter and network length are not significantly altered in embryonic foxf2 mutants, but in 6-month-old adults, the vessel network length is significantly decreased. Our data suggest that foxf2 deficiency contributes to cumulative vascular defects. Adult endothelial defects may be secondary to pericyte defects, although foxf2a is expressed in both pericytes and endothelial cells, and both cell types may be affected autonomously.

The initial pericyte pool is critical for lifelong vascular health

To understand the mechanism by which foxf2a influences pericyte numbers, we needed to distinguish between its role in early development and later roles. Our lab recently demonstrated that pericyte precursors express nkx3.1 and foxf2a before pericyte differentiation and prior to expression of canonical markers, such as pdgfrβ (Ahuja et al., 2024). Here, we show that the pericyte pool in foxf2a mutants is reduced at the earliest embryonic stage that it can be measured. Zebrafish are remarkably regenerative and able to repair cardiac, retinal, spinal cord, and pancreatic damage among other tissues (Poss and Tanaka, 2024). We, therefore, anticipated that foxf2a mutants might be able to regenerate new pericytes to replace lost pericytes. This was tested using genetic ablation of pericytes. Ablation is very efficient in mutants and wild-type. Surprisingly, 7 days after ablation, foxf2a mutants show recovery of pericyte numbers. Thus, when placed under extreme stress, embryonic foxf2a mutants can regenerate pericytes; yet, under baseline conditions, foxf2a mutant pericytes do not replenish. This experiment is important to distinguish an underlying mechanism of the foxf2a pericyte phenotype. Our data suggest that the earliest and most important difference in foxf2a mutant pericytes is their low initial number. Even though some repair is possible, it is not enough to compensate. Reduced pericyte density is associated with elongated processes, enlarged soma, and degeneration in mutants, which are likely signs of cellular stress. Over a lifetime, the deficiency leads to progressive damage to the vascular network. Determining that there is a critical embryonic window for developing a robust pericyte precursor pool helps us focus on early interventions to mitigate these deficits before damage over the lifespan worsens. The necessity of Foxf2 during development for the establishment of proper brain vasculature was also seen in a mouse Foxf2 knockout (Reyahi et al., 2015). Further research into the early role of foxf2a in establishing initial pericyte specification/differentiation, and the impact of early vascular defects on disease progression, will be crucial for developing strategies to prevent and treat cerebrovascular conditions.

Materials and methods

Zebrafish husbandry and strains

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All procedures were conducted in compliance with the Canadian Council on Animal Care, and ethics approval was granted by the University of Calgary Animal Care Committee (AC21-0169). Embryos were maintained in E3 medium (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4, pH = 7.2) (Westerfield, 1995) at 28 °C. Larvae up to 10 dpf were maintained in an incubator with a light cycle (14 hr light, 10 hr darkness), with daily water changes and feeding. As zebrafish sex is not determined until 28 dpf, sex is not considered in embryo and larval experiments. For adult experiments, results were compared by sex where sufficient n’s were available. Results from the progeny of het in-crosses were blinded in that they were quantified before genotyping. All reagents and strains are listed in Key resources table. All experiments included wild-type as a comparison group, and developmental stages, n’s, and genotypes are noted for each experiment source data table for each figure.

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Strain, strain background (D. rerio)foxf2aca71Ryu et al., 2022ZDB-ALT-230214–10
Strain, strain background (D. rerio)foxf2bca22Chauhan et al., 2016ZDB-ALT-160809–1
Strain, strain background (D. rerio)Tg(acta2:GFP)ca7Whitesell et al., 2014ZDB-ALT-120508–1
Strain, strain background (D. rerio)Tg(4xUAS:Zebrabow-B)a133Pan et al., 2013ZFIN: ZDB-ALT-130816–3
Strain, strain background (D. rerio)Tg(flk:GFP)la116Choi et al., 2007ZDB-TGCONSTRCT-070529–1
Strain, strain background (D. rerio)Tg(kdrl:mCherry)ci5Proulx et al., 2010ZFIN: ZDB-ALT-110131–57
Strain, strain background (D. rerio)Tg(UAS:NTR-mCherry)c264Davison et al., 2007ZDB-ALT-070316–1
Strain, strain background (D. rerio)TgBAC(4xUAS:EGFP)mpn100TgDeMaria et al., 2013ZFIN: ZDB-TGCONSTRCT-140812–1
Strain, strain background (D. rerio)TgBAC(pdgfrβ:GAL4FF)ca42Whitesell et al., 2019ZFIN: ZDB-ALT-200102–2
Strain, strain background (D. rerio)TgBAC(pdgfrβ:EGFP)ca41TgWhitesell et al., 2019ZDB-TGCONSTRCT-160609–1
Antibodyanti mCherry, rat monoclonalThermo Fisher, M11217RRID:AB_25366111 in 500
Antibodyanti Green Fluorescent Protein, mouse monoclonalClontech, 3 P 632380RRID:AB_100134271 in 500
AntibodyDonkey anti mouse 488Thermo Fisher, A-21202RRID:AB_1416071 in 500
AntibodyGoat anti rat 555Thermo Fisher, A-21434RRID:AB_25358551 in 500
Commercial assay or kitFluoromount-G Mounting Medium with DAPIInvitrogenE141201
Commercial assay or kitFluoromount-G Mounting MediumThermo Fisher00-4958-02
Commercial assay or kitTaqman SNP genotyping kit for foxf2bca22Applied BiosystemsANAACEC
Commercial assay or kitKAPA2G Fast Hotstart Genotyping MixRocheKK5621
Chemical compound, drugDimethylsulfoxideSigmaD8418
Chemical compound, drugPhenylthioureaSigmaP7629
Chemical compound, drugUltraPure AgaroseInvitrogen16520–050
Chemical compound, drugMetronidazoleSigmaM3761
Software, algorithmImaris 10.3Oxford InstrumentsRRID:SCR_007370
Software, algorithmIlastichttps://www.ilastik.org/RRID:SCR_015246
software, algorithmFiji (ImageJ)Schindelin et al., 2012RRID:SCR_002285
Software, algorithmGraphPad Prism 10GraphpadRRID:SCR_002798
Software, algorithmAdobe PhotoshopAdobeRRID:SCR_014199
Software, algorithmVesselMetricsMcGarry et al., 2024 Microvasc Res, 2024
Commercial assay or kitHCR Probe (v3.0) ndu4al2aMolecular Instruments
Commercial assay or kitHCR Probe (v3.0) kdrlaMolecular Instruments
Commercial assay or kitHCR Probe (v3.0) pdgfrβMolecular Instruments
Commercial assay or kitHCR Probe (v3.0) foxf2aMolecular Instruments
Commercial assay or kitHCR Probe (v3.0) foxf2bMolecular Instruments
Sequence-based reagentfoxf2a-genotyping-forwardIDTATG CAC TCG GCT CTC CAA AA
Sequence-based reagentfoxf2a-genotyping-reverseIDTGAT CGC CAT GAC TAT CGG GG

Genotyping

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Adult fish were anesthetized in 0.4% Tricaine (Sigma) and placed on a cutting surface. A small portion of the tip of the fin was clipped using a razor blade, and the fish were returned to the system to recover. For developmental DNA isolation, whole embryos, or larvae (up to 10 dpf) were anesthetized in 0.4% Tricaine prior to sampling.

Genomic DNA (gDNA) was extracted using the HotSHOT DNA isolation protocol, adapted from Meeker et al., 2007. To isolate gDNA, tissue or embryo was placed in 50 μL of Base Solution and incubated at 95 °C for 30 min. 5 μL of Neutralization Solution was added to neutralize the reaction. Samples were spun down in a mini centrifuge, and DNA was sampled from the top portion of the solution to avoid undigested samples.

Zebrafish were genotyped for target genes or the presence of transgenes using the KAPA2G Fast HotStart Genotyping Mix as per the manufacturer’s instructions. foxf2 mutants were genotyped using foxf2a primers (Key resources table). A custom TaqMan probe for foxf2bca22 (ANAACEC) was obtained from Applied Biosystems. Samples were genotyped using the QuantStudio 6 Flex Real-Time PCR System (Applied Biosystems) with the wild-type allele reported in FAM and the mutant allele reported in VIC.

In situ hybridization

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Custom probes for foxf2a (PRD069), nduf4al2 (RTD146), kdrl (PRI089), pdgfrβ (PRA654), and foxf2b (PRF462) were obtained from Molecular Instruments (Los Angeles, CA) for Hybridization Chain Reaction (HCR) in situ hybridization. In situ staining was performed according to the manufacturer’s instructions. Samples were permeabilized with proteinase K (1 mg/mL stock; Invitrogen, 4333793) at various times and concentrations, depending on their age.

Integrated intensity

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pdgfrβ and foxf2b intensity was obtained using ImageJ from in-situ stained embryos at 3 dpf. Mean fluorescent intensity was measured using the freehand selection tool to circle each pericyte. The background mean intensity was subtracted from the pericyte mean intensity to standardize each measurement, and the intensities of each fish were averaged. The same protocol was used to measure the intensity of the pdgfrβ transgene in 3 dpf live embryos.

Total RNA extraction and cDNA synthesis

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Total RNA was extracted using the Research RNA Clean & Concentrator-5 Kit (Zymo Research, R1015) with some modifications. 50 dechorionated embryos were collected at 48 hpf and homogenized in TRIZOL (Ambion, 15596026) reagent with a syringe needle. Briefly, samples were centrifuged at 16,000 rpm for 2 min to remove pigment. After separating the solution from the precipitate, chloroform was added, and the mixture was centrifuged at 12,000 rpm for 15 min at 4 °C. The aqueous layer containing nucleic acids was removed, and an equal volume of 95–100% ethanol was added, then mixed well. The rest of the protocol followed the manufacturer’s instructions using the Zymo-spin IC column. After eluting RNA with RNase/DNase-free water, samples were quantified using a Nanodrop to evaluate RNA quality. For cDNA synthesis, qSCRIPT cDNA Supermix (QuantaBio, 95048–100) was used according to the manufacturer’s instructions and stored at –20 °C.

RT-qPCR

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To assess relative gene expression, real-time quantitative PCR (RT-qPCR) was carried out using custom-designed primers. PowerUP SYBR Green Master Mix (Applied Biosystems, A25742) was utilized on a QuantStudio 6 Flex Real-Time PCR System (Applied Biosystems). Cycle conditions followed the protocol of a PowerUP SYBR Green standard reaction: 50 °C for 2 min, 95 °C for 2 min, and 40 cycles of 95 °C for 15 seconds and 60 °C for 60 s. To calculate the fold change in gene expression, mutants were compared to wild-type using ΔΔCT calculations (Livak and Schmittgen, 2001).

Brain dissection

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Zebrafish were first euthanized in Tricaine and mounted on a Sylgard gel plate with dissection pins to stabilize them. Scissors were used to sever the brain stem entirely at the base of the head, posterior to the hindbrain. The eyes were removed with forceps/micro-scissors, and the optic stalks were cut. An incision was then made at the posterior portion of the skull plate anteriorly. Forceps were used to pull back the skull plate and orbital bones. Any nerve connections were severed, and the brains were removed and immediately fixed in ice-cold 4% PFA overnight at 4 °C.

Brain tissue clearing

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Whole zebrafish brains were cleared by either CUBIC (Susaki et al., 2015), or a modified iDISCO+ (Renier et al., 2016) protocol. For CUBIC, samples were incubated in a 1:1 ratio of Reagent 1 (25% Quadrol (Sigma, 122262), 25% Urea, and 15% Triton X-100):H2O overnight at 37 °C. Next, brains were incubated in 100% Reagent 1 at 37 °C until the tissue was transparent. Finally, brains were rinsed in PBS, mounted in 2% low melting point agarose and re-placed in 100% Reagent 1 at 37 °C until the tissue was transparent.

For iDISCO, the samples were first dehydrated in methanol:H2O dilution series for 30 min each, then chilled at 4 °C. Next, samples were incubated overnight in a 1:3 ratio of dichloromethane (DCM; Sigma, 270997):methanol at room temperature. The following day, samples were washed in methanol and then chilled at 4 °C before bleaching in fresh 5% H2O2 in methanol overnight at 4 °C. Then, samples were rehydrated in methanol:H2O dilution series for 30 min each, then washed in PTx.2 (0.2% Triton X-100 in PBS) twice over 2 hr at room temperature. Next, samples were incubated in permeabilization solution (0.3 M glycine, 20% DMSO in 400 mL PTx.2) at 37 °C for up to two days, after which the samples were rinsed in PBS twice over 1 hr. Samples were then washed three times over 2 hr in 0.5 mM SDS/PBS at 37 °C for three days before being incubated in primary antibody (Key resources table) in 0.5 mM SDS/PBS at 37 °C for two more days. The primary antibodies were refreshed in PTx.2 and left for another 4 days before overnight washing in PTwH (10 μg/ml heparin and 0.2% Tween-20 in PBS) at 37 °C. Samples were left in secondary antibodies (Key resources table) in PTwH/3% NSS at 37 °C for 3 days, refreshed and left for another 3 days before washing overnight in PTwH. After immunolabeling, brains were mounted in 2% low melting agarose (LMA) and dehydrated in methanol:H2O dilution series for 30 min each and left overnight at room temperature. Next, brains were incubated in a 1:3 ratio of DCM: methanol at room temperature for 3 hr before washing in 100% DCM for 15 min twice. Finally, brains were incubated in ethyl cinnamate (Sigma, NSC6773) for 2 hr before replacing the solution and incubating overnight at room temperature.

Tissue sectioning

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For vibratome sections, brains were fixed in 4% PFA, washed, and mounted in 4% LMA (Invitrogen, 16520–050) before sectioning with a vibratome (Leica, VT1000S) to obtain a series of transverse sections at 50 µm.

Cre mRNA injections

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Wild-type fish on a Zebrabow transgenic background (Pan et al., 2013) were injected with 1 pg of Cre mRNA at the one-cell stage. Embryos were kept in E3 at 28 °C until imaging.

Immunofluorescence

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Sections for immunofluorescence were briefly washed in PBS, followed by 2% Triton X-100 in PBS for permeabilization. Sections were moved from the permeabilization solution into blocking buffer (5% goat serum, 3% BSA, 0.2% Triton X-100 in PBS) for 1 hr before incubation in primary antibody. Sections were then washed and left in secondary antibodies for 3 hr at room temperature. After incubation, sections were washed, mounted, and cover-slipped with Fluoromount-G Mounting Medium, with DAPI or without counterstain.

Drug treatments

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Embryos were dechorionated prior to treatment, and all drug treatments included DMSO at an equivalent concentration to the drug solution as a control (Sigma, D8418). Treatments were performed in a 24-well plate with approximately 15 embryos per well. For pericyte ablation experiments, 5 mM Metronidazole (MTZ; Sigma, M3761) was applied at 50 hpf for 1 hr with a DMSO control.

Microscopy

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Imaging fluorescent transgene expression in live embryos, antibody staining, and fluorescent staining were completed using an inverted laser scanning confocal microscope (LSM900; Zeiss) with a 10 X (0.25 NA), 20 X (0.8 NA), 40 X water (1.1 NA), or 60 X (1.4 NA) oil objectives. Laser wavelengths included blue (405 nm), green (488 nm), red (561 nm), and far red (640 nm). Embryos were maintained in phenylthiourea (PTU) from 24 hpf onwards to prevent pigment development, anesthetized in 0.4% tricaine and mounted in 0.8% LMA dissolved in E3, on a clear imaging dish. In some cases, live samples were retrieved from the agarose following imaging for further imaging at later time points, genotyping, or other data collection. Images were processed using Zen Blue and Fiji (Schindelin et al., 2012) software.

Imaging of whole adult brains was completed using the Light-sheet microscope (Ultramicroscope II equipped with a SuperPlan module; Miltenyi Biotec) with a 4 X (0.35 NA) objective at 1.0 zoom. Laser wavelengths included green (488 nm) and red (561 nm). Images were processed using Zen Blue, Fiji, and Imaris software.

Vessel network quantitation

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Confocal images from embryos were analyzed using the Python-based software tool Vessel Metrics (McGarry et al., 2024). The total vessel network length was measured starting from below the middle cerebral vein and dorsal longitudinal vein until the emergence of the basal communicating artery. Vessels included in measurements are as follows: middle mesencephalic central arteries, posterior mesencephalic central arteries, the primordial hindbrain channels, and the posterior region of the basilar artery. The forebrain vessels (i.e. anterior cerebral veins) were not included. Blood vessel diameter was restricted to the horizontal hindbrain central arteries, which were comparable between all images.

Adult vessel network length was measured using Ilastik (Berg et al., 2019) and Imaris (version 10.2.0; Bitplane AG, Oxford Instruments) software. Vessels were annotated in Ilastik to obtain a probability map. The probability map was imported into Imaris as a new channel for each image. The surface tool with machine learning was then used to annotate the vessels a second time, using the Ilastik map as the training guide. A mask was created from this surface. The filament tool was then used to create a 3D network of the vessel, using the surface mask as a training guide. The total network length and the diameter of each vessel segment were exported from the filament statistics tab.

Smooth muscle cell coverage was measured using the same pipeline; however, the raw channel was used for the surface tool annotation without using Ilastik. Total network length was exported from the filament statistics tab.

Pericyte quantitation

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Embryonic pericytes were manually counted using the Fiji counting or tracing tool on flattened Z-stacks from confocal images of the whole zebrafish head. Pericyte counting and analysis were restricted to the mid and hindbrain regions for all metrics. For pericyte process lengths, individual processes extending from a single soma were measured and summed to determine the total process length per pericyte (µm). In instances where two processes appeared to merge or cover the same blood vessel, half of the total length was added to each pericyte. For Zebrabow images, only pericytes with processes distinguishable from neighbouring pericytes were measured. For soma size, individual cell bodies were traced, and the area was determined by the software in µm2.

Adult pericytes were counted using the spot tool in Imaris. Using the raw channel, the spot tool was trained to identify pericyte cell bodies. The total spot number, equivalent to the number of pericyte cell bodies, was exported from the spot statistics tab. A mask of the spot tool was created to better visualize pericytes.

Statistics

All statistical analyses were performed using GraphPad Prism 10, with significance determined by p<0.05. Statistical tests conducted are included in figure captions. If no significance is indicated, it is not significant. The D’Agostino-Pearson test was used to assess the normality of data sets, and in cases where the data did not pass the normality test, non-parametric statistics were used. Experimental N’s are reported in source data accompanying each figure. Results are expressed on graphs as mean ± standard deviation (SD). Only significant p-values are indicated.

Materials availability

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All materials used in this study are freely available on request.

Data availability

Numerical data for all experiments is included for each figure.

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Article and author information

Author details

  1. Merry Faye E Graff

    1. Alberta Children’s Hospital Research Institute, University of Calgary, Calgary, Canada
    2. Department of Biochemistry and Molecular Biology, University of Calgary, Calgary, Canada
    Contribution
    Conceptualization, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – original draft, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0009-0006-6861-5759

  2. Emma EM Heeg

    1. Alberta Children’s Hospital Research Institute, University of Calgary, Calgary, Canada
    2. Department of Biochemistry and Molecular Biology, University of Calgary, Calgary, Canada
    Contribution
    Conceptualization, Formal analysis, Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0009-0002-0879-133X

  3. David A Elliott

    Hotchkiss Brain Institute Advanced Microscopy Platform, University of Calgary, Calgary, Canada
    Contribution
    Resources, Software, Methodology
    Competing interests
    No competing interests declared

  4. Sarah J Childs

    1. Alberta Children’s Hospital Research Institute, University of Calgary, Calgary, Canada
    2. Department of Biochemistry and Molecular Biology, University of Calgary, Calgary, Canada
    Contribution
    Conceptualization, Resources, Supervision, Writing – original draft, Project administration, Writing – review and editing
    For correspondence
    schilds@ucalgary.ca
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2261-580X

Funding

Canadian Institutes of Health Research (PJT-183631)

  • Sarah J Childs

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

Acknowledgements

This work was funded by the Canadian Institutes of Health Research PJT-183631. MFG received a Canada Graduate Scholarship Master’s from the Canadian Institutes of Health Research, the Alberta Graduate Excellence Scholarship (AGES) for Master’s Research from the Province of Alberta, and a Biochemistry and Molecular Biology Department scholarship from the University of Calgary. EH received the Alvin Libin Graduate Scholarship in Cardiovascular Research, the Alberta Children’s Hospital Research Institute (ACHRI) Graduate Scholarship for Master’s Research, the ACHRI Graduate Scholarship for Doctoral Research, and the Alberta Graduate Excellence Scholarship (AGES) for Doctoral Research from the Province of Alberta. We acknowledge the Alberta Children’s Hospital Research Institute and Hotchkiss Brain Institute Imaging facilities for microscopes and technical support.

Ethics

This study was performed in strict accordance with the recommendations CCAC guidelines: Zebrafish and other small, warm-water laboratory fish from the Canadian Council on Animal Care. All animals were handled according to the approved institutional University of Calgary Animal Care Committee (AC21-0169).

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  1. Merry Faye E Graff
  2. Emma EM Heeg
  3. David A Elliott
  4. Sarah J Childs
(2026)
Progressive mural cell deficiencies across the lifespan in a foxf2 model of cerebral small vessel disease
eLife 14:RP106720.
https://doi.org/10.7554/eLife.106720.3

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