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Suppression of transcytosis regulates zebrafish blood-brain barrier function

  1. Natasha M O'Brown
  2. Sean G Megason  Is a corresponding author
  3. Chenghua Gu  Is a corresponding author
  1. Harvard Medical School, United States
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Cite this article as: eLife 2019;8:e47326 doi: 10.7554/eLife.47326

Abstract

As an optically transparent model organism with an endothelial blood-brain barrier (BBB), zebrafish offer a powerful tool to study the vertebrate BBB. However, the precise developmental profile of functional zebrafish BBB acquisition and the subcellular and molecular mechanisms governing the zebrafish BBB remain poorly characterized. Here, we capture the dynamics of developmental BBB leakage using live imaging, revealing a combination of steady accumulation in the parenchyma and sporadic bursts of tracer leakage. Electron microscopy studies further reveal high levels of transcytosis in brain endothelium early in development that are suppressed later. The timing of this suppression of transcytosis coincides with the establishment of BBB function. Finally, we demonstrate a key mammalian BBB regulator Mfsd2a, which inhibits transcytosis, plays a conserved role in zebrafish, as mfsd2aa mutants display increased BBB permeability due to increased transcytosis. Our findings indicate a conserved developmental program of barrier acquisition between zebrafish and mice.

https://doi.org/10.7554/eLife.47326.001

Introduction

Blood vessels in the vertebrate brain are composed of a single layer of endothelial cells that possess distinct functional properties that allow the passage of necessary nutrients yet prevent unwanted entry of specific toxins and pathogens into the brain. This specialized endothelial layer forms the blood-brain barrier (BBB) and restricts the passage of substances between the blood and the brain parenchyma via two primary mechanisms: 1) specialized tight junction complexes between apposed endothelial cells to prevent intercellular transit (Reese and Karnovsky, 1967; Brightman and Reese, 1969) and 2) suppressing vesicular trafficking or transcytosis to prevent transcellular transit (Ben-Zvi et al., 2014; Andreone et al., 2015; Chow and Gu, 2017). BBB selectivity is further refined with the expression of substrate-specific transporters that dynamically regulate the influx of necessary nutrients and efflux of metabolic waste products (Sanchez-Covarrubias et al., 2014; Umans et al., 2017). While the BBB is comprised of endothelial cells, the surrounding perivascular cells including pericytes and astroglial cells, play a critical role in forming and maintaining barrier properties (Janzer and Raff, 1987; Armulik et al., 2010; Bell et al., 2010; Daneman et al., 2010; Wang et al., 2014). Collectively, endothelial cells and the surrounding perivascular cells form the neurovascular unit.

As the simplest genetic model organism with an endothelial BBB (Jeong et al., 2008), zebrafish offer a powerful tool to study the cellular and molecular properties of the vertebrate BBB (Xie et al., 2010; Vanhollebeke et al., 2015; Umans et al., 2017; O'Brown et al., 2018; Quiñonez-Silvero et al., 2019). Zebrafish have served as a great model system to study vascular biology due to their large clutch size, rapid and external development, and transparency for in vivo whole organism live-imaging (Lawson and Weinstein, 2002; Jin et al., 2005; Santoro et al., 2007; Armer et al., 2009; Herbert et al., 2009; Phng et al., 2009; Geudens et al., 2010; Herbert et al., 2012; Wilkinson and van Eeden, 2014; Franco et al., 2015; Vanhollebeke et al., 2015; Matsuoka et al., 2016; Ulrich et al., 2016; Venero Galanternik et al., 2017; Stratman et al., 2017; Geudens et al., 2018). Additionally, with the advent of CRISPR-Cas9 technology, zebrafish provide an efficient genetic toolkit for targeted mutagenesis (Hwang et al., 2013; Gagnon et al., 2014; Ablain et al., 2015; Varshney et al., 2015; Albadri et al., 2017; Hogan and Schulte-Merker, 2017). However, the subcellular and molecular mechanisms governing the formation and maintenance of the zebrafish BBB are only beginning to be characterized. Expanding our understanding of the zebrafish BBB can thus reveal the mechanistic similarities between the zebrafish and mammalian BBB to further evaluate the position of zebrafish as a model organism for studying the BBB.

Barrier properties of brain endothelial cells are induced by extrinsic signals from other cells in the surrounding microenvironment during development (Stewart and Wiley, 1981). In rodents, the BBB becomes functionally sealed in a spatiotemporal gradient, with the hindbrain and midbrain barriers becoming functional before the cortical barrier (Daneman et al., 2010; Ben-Zvi et al., 2014; Sohet et al., 2015). Within the cortex, barrier function is acquired along a ventral-lateral to dorsal-medial gradient (Ben-Zvi et al., 2014). In the zebrafish, existing studies have disagreed over the timing of zebrafish barrier formation, with some suggesting that BBB maturation occurs concurrently with angiogenesis (Jeong et al., 2008; Xie et al., 2010; Umans et al., 2017) and others providing a wide range beginning at 3 dpf and extending to 10 dpf (Fleming et al., 2013). These conflicting reports may be due to regional developmental gradients of barrier acquisition, differences in definitions of BBB development, which range from functional tracer restriction (Jeong et al., 2008; Fleming et al., 2013) to gene expression of BBB-specific selective transporters, such as glut1b (Umans et al., 2017), or variations in the experimental approaches used to assess BBB permeability such as the molecular weight and properties of tracers and circulation time. Therefore, we aim to resolve some of these discrepancies through detailed regional and subcellular characterizations of functional barrier acquisition in zebrafish.

Recent work in the mammalian blood-retinal barrier has indicated that the suppression of transcytosis governs functional barrier development (Chow and Gu, 2017). Interestingly, endothelial cells at the leaky neonatal angiogenic front possess functional tight junction complexes halting the intercellular passage of the tracer protein Horseradish Peroxidase (HRP) at the so-called ‘kissing points’. In contrast, these endothelial cells exhibit high levels of HRP-filled vesicles compared to functionally sealed proximal vessels. Moreover, these areas of elevated vesicular trafficking continue to correspond with barrier permeability at the angiogenic front until the barrier seals (Chow and Gu, 2017). Work in the mouse BBB has also demonstrated the importance of suppressing transcytosis in determining barrier permeability. Mice lacking the major facilitator super family domain containing 2a (Mfsd2a) lipid transporter exhibit increased levels of caveolae vesicles in CNS endothelial cells, resulting in increased barrier permeability (Ben-Zvi et al., 2014; Andreone et al., 2017). Whether this suppression of transcytosis also determines BBB function in zebrafish remains unknown.

Here in zebrafish, we find a spatiotemporal gradient of barrier acquisition, and capture the dynamics of BBB leakage as it matures during development using time lapse live imaging. We further find a conserved role for transcytosis suppression in determining barrier function, both during normal development and in mfsd2aa mutants.

Results

Posterior-anterior gradient of zebrafish BBB development

To determine when and how the zebrafish BBB becomes functional in different brain regions, we performed intracardiac injections of fluorescently conjugated tracers (1 kDa NHS and 10 kDa Dextran) simultaneously at different developmental stages and imaged live fish after 1 hr of tracer circulation (Figure 1A and B). NHS is widely used to assess mouse BBB permeability (Sohet et al., 2015; Chow and Gu, 2017; O'Brown et al., 2018). Additionally, NHS has been used successfully to assess junctional defects in occludin- and claudin-deficient animals (Chen et al., 1997; Furuse et al., 2002; Nitta et al., 2003), and was previously shown to be restricted within the adult zebrafish cerebral vasculature (Jeong et al., 2008). We used a combination of different molecular weight tracers to tease apart potential avenues of leakage, as tight junctional defects result specifically in the leakage of low-molecular-weight tracers 1 kDa and below into the brain parenchyma (Nitta et al., 2003; Campbell et al., 2008; Sohet et al., 2015; Yanagida et al., 2017). To assess BBB permeability, we measured tracer fluorescence intensity in the brain parenchyma and normalized to circulating levels of each tracer, that is to the fluorescence intensity of tracer within brain blood vessels, to account for between fish variation in tracer injections or circulation (details in the Materials and methods section). At 3 dpf, we observed the presence of both NHS and Dextran throughout the brain parenchyma (average of 8.5 ± 0.3 Tracer Intensity in the midbrain and average of 7.4 ± 0.4 NHS and 5.5 ± 0.6 Dextran Intensity in the hindbrain; Figure 1; Figure 1—figure supplement 1). These leakage assays revealed that the injected tracers were able to permeate into the brain parenchyma, suggesting that the BBB was not functionally sealed. In addition to the use of exogenous injected fluorescent tracers, we also assayed BBB permeability with an endogenous transgenic serum DBP-EGFP fusion protein (Tg(l-fabp:DBP-EGFP)) to account for injection artifacts (Xie et al., 2010). At 3 dpf, we observed similar leakage patterns with the transgenic serum protein as we did with the injected tracers (average of 8.9 ± 0.8 DBP-EGFP Intensity in the midbrain and average of 6.1 ± 0.4 DBP-EGFP Intensity in the hindbrain; Figure 1C and D; Figure 1—figure supplement 1). At 4 dpf, we observed a significant decrease in tracer intensity in the hindbrain (average of 4.1 ± 0.2 NHS Intensity, 3.9 ± 0.4 Dextran Intensity and 4.0 ± 0.4 DBP-EGFP Intensity; p<0.0001, p=0.0115 and p=0.0034 compared to 3 dpf for NHS, Dextran and DBP-EGFP by two-way ANOVA; Figure 1—figure supplement 1). However, the midbrain barrier remains leaky (average of 7.6 ± 0.5 NHS Intensity, 7.5 ± 0.8 Dextran Intensity and 6.8 ± 0.9 DBP-EGFP Intensity; p=0.66, p=0.89 and p=0.079 compared to 3 dpf for NHS, Dextran and DBP-EGFP by two-way ANOVA; Figure 1C and D). At 5 dpf, the midbrain parenchymal tracer intensities are dramatically reduced (average of 3.3 ± 0.3 NHS Intensity, 3.6 ± 0.2 Dextran Intensity and 3.2 ± 0.3 DBP-EGFP Intensity; p<0.0001, two-way ANOVA; Figure 1B and C) and no significant changes were observed in the hindbrain (Figure 1—figure supplement 1). No significant change was observed in midbrain tracer uptake from 5 to 6 dpf or from 5 to 10 dpf (Figure 1C and D), indicating that the midbrain barrier becomes sealed at 5 dpf. Of note, all three tracers showed nearly indistinguishable patterns of uptake, with similar parenchymal tracer intensity levels (Figure 1; Figure 1—figure supplement 1), suggesting that the leakage is not due to tight junctional defects but may rather be due to an increase in vesicular trafficking.

Figure 1 with 1 supplement see all
The midbrain BBB becomes functional at 5 dpf.

(A) Diagram of the tracer leakage assay. Fluorescently conjugated tracers (turquoise) were injected intracardially into transgenic fish that express mCherry in the vasculature (magenta; Tg(kdrl:mCherry)) and allowed to circulate for 1 hr before imaging. (B) Dorsal view maximum intensity projection of the larval brain vasculature at 3 dpf. Left image is pseudo-colored to demarcate the midbrain (violet) and the hindbrain (gold) vasculature. Right image shows the NHS tracer (turquoise) in the entire larval brain, with a large number of tracer-filled parenchymal cells in the midbrain. (C) Representative dorsal view maximum intensity projections of larval zebrafish midbrains at different developmental stages reveal increased permeability at 3 and 4 dpf compared to 5, 6 and 10 dpf. The increased early permeability was observed with two injected tracers of different sizes, a 1 kDa NHS (turquoise) and a 10 kDa Dextran (white), as well with an 80 kDa transgenic serum protein DBP-EGFP (green). Parenchymal tracer intensity outside of the vasculature (magenta) was measured and normalized to the blood vessel tracer intensity in each fish. Scale bars represent 50 µm. (D) Quantification of normalized parenchymal tracer intensity in the midbrain between 3 and 10 dpf reveals a significant decrease in tracer leakage at five dpf. There was no difference observed between different tracers at any time point. There was no significant change from 3 to 4 dpf or from 5 to 10 dpf, suggesting that the midbrain barrier seals around 5 dpf. N = 14–21 fish, each represented as a single dot on the plot. The mean and the standard error are drawn in black for each tracer and stage. ****p<0.0001, ns is not significant by two-way ANOVA.

https://doi.org/10.7554/eLife.47326.002

Time lapse imaging reveals two modes of developmental leakage

The vast majority of studies interrogating BBB permeability have relied on observing leakage in fixed static images from mutant mice, which does not reveal the dynamic nature of the BBB. A key advantage of zebrafish is the ability to examine biological processes in vivo in real-time, providing us with the unique opportunity to observe the process of functional barrier maturation (Figure 1). To examine the developmental dynamics of barrier leakage, we injected larvae at 3 dpf with fluorescently labeled 10 kDa Dextran and performed time lapse live imaging for 1 hr following injection, with time 0 being an average of 8 min post-injection. We observed a gradual and diffuse increase in extravascular Dextran intensity over time within the brain parenchyma (slope of 0.13 intensity/minute; Figure 2—videos 1 and 2; Figure 2A, C and E). In addition to the time-dependent increase in overall Dextran intensity in the brain parenchyma, we also observed parenchymal cells taking up tracer, both directly adjacent to blood vessels and at a small distance away (Figure 2A and C). To validate that the Dextran was in fact being taken up by cells rather than filling the extracellular space surrounding cells, we performed these same tracer injections in a transgenic line with fluorescently labeled cell membranes (Tg(actb2:MA-Citrine)) at 3 dpf and observed several tracer-filled cells, that is tracer confined within cell membranes outside of the vasculature (Figure 2—figure supplement 1). Similarly, when we performed these Dextran injections in a nuclear labeled transgenic line (Tg(actb2:Hsa.H2B-Scarlet)) at 3 dpf, we also observed Dextran co-localization with nuclear signal (Figure 2—figure supplement 1), further indicating that the injected tracers are taken up by scavenger parenchymal cells in addition to permeating the extracellular spaces.

Figure 2 with 4 supplements see all
Dynamic tracer leakage in the developing BBB via live imaging.

(A) Time course stills from Figure 2—Video 1 of tracer leakage at 3 dpf reveal an increase in parenchymal cells absorbing the Dextran tracer (outlined by dashed white lines) as well as a general increase in overall Dextran (green) intensity outside of the vasculature (magenta). An angiogenic tip cell becomes apparent at 33:23 and is demarcated by an asterisk (*). This tip cell produces two separate bursts of leakage observed in Figure 2—Video 2. (B) Time course stills from Figure 2—Video 3 of Dextran tracer dynamics at 5 dpf reveals a mature BBB, with reduced overall Dextran extravasation into the brain parenchyma. The scale bars represent 10 µm. (C and D) Dorsal maximum intensity projection of the midbrain at 3 dpf (C) and 5 dpf (D) at the first and last time point examined. While there is a large increase in overall parenchymal Dextran intensity over time at 3 dpf, the 5 dpf midbrain parenchyma appears relatively unaltered after an hour of Dextran circulation. Boxed regions are representative of the six areas per fish used for analysis in E. The scale bars represent 50 µm. (E) Quantification of Dextran intensity in the brain parenchyma over time at 3 dpf (green) and 5 dpf (black) shows a significant difference in tracer leakage dynamics (p<0.0001, Mann Whitney U test), with both more total Dextran accumulation and a faster rate of Dextran accumulation in the brain parenchyma at 3 dpf. N = 6 fish with 6 regions analyzed per fish.

https://doi.org/10.7554/eLife.47326.005

Interestingly, at 3 dpf, we observed an actively sprouting cell that extended away from the established vessel in a sporadic fashion (Figure 2—Video 2; Figure 2A). After 20 min of rambling migration, this sprout suddenly released a large bolus of Dextran that appeared to be taken up by a single parenchymal cell. Two min later, this same sprout released a second bolus of Dextran on the opposite side (Figure 2—Video 2). These rare large bursts of leakage were not unique to this sprout; they were also sporadically observed from established blood vessels (Figure 2—Video 1; Figure 2A). While this particular fish displayed three of these bursting events, defined as parenchymal Dextran intensity increasing to luminal levels in less than 1 min, only two of the other five measured 3 dpf fish displayed any bursts within the hour post tracer injection, one from a sprouting tip cell and one from a stable vessel, further indicating the infrequency of these large bursts. Cumulatively, this time lapse data revealed two discrete types of leakage occurring during this early developmental stage: steady and diffuse Dextran leakage into the parenchyma that makes up the vast majority of observed leakage and rare large bursts of leakage.

When we performed these same time lapse experiments at 5 dpf, we observed significantly less overall tracer accumulation in the brain parenchyma over the course of the hour in addition to reduced rates of tracer accumulation (slope of 6.833e-6 intensity/sec; Figure 2—Video 3; p<0.001, Mann-Whitney test; Figure 2B, D and E). Furthermore, 5 dpf fish never exhibited bursts of tracer leakage as observed at 3 dpf. These data suggest that at early stages, most tracer leakage occurs broadly through vessel walls via unrestricted transcytosis with an occasional transient rupture to endothelial integrity that leads to these large bursts of leakage. Once tracer is leaked into the intercellular space of the parenchyma it can be taken up by select parenchymal cells (Figure 2—figure supplement 1). As the BBB matures both sources of leakage sharply decrease.

Suppression of transcytosis coincides with functional BBB development

Given the observed differences in brain permeability during larval development, we next sought to determine the subcellular mechanisms underlying the development of a functional BBB. Therefore, we assessed BBB properties by performing intracardiac injections of electron-dense NHS-gold nanoparticles (5 nm) followed by transmission electron microscopy (TEM) at different developmental stages when the BBB is leaky (3 dpf) and when it is functionally sealed to circulating fluorescent tracers (5 and 7 dpf; Figure 3). All the blood vessels analyzed had a maximal diameter of 5 µm to enrich for capillaries and small veins. At 3 dpf, we found blood vessels in direct contact with neurons and pericytes, with the pericytes sharing the endothelial basement membrane (Figure 3A), as in the mammalian neurovascular unit. During this leaky developmental stage, the endothelial basement membrane was filled with the electron-dense gold nanoparticles (Figure 3A and B; turquoise arrowheads) with an average gold intensity of 1.29 ± 0.07 (luminal gold intensity normalized to 1.0; Figure 3G), further demonstrating that the BBB is functionally immature at 3 dpf. To decipher how the gold nanoparticles traverse from the lumen to the basement membrane, we first looked at the tight junctions to see if the nanoparticles passed between apposed endothelial cells. We consistently observed that the nanoparticles were halted at the ‘kissing points’ between endothelial cells, indicating that tight junctions were functional prior to formation of a functional BBB (49/49 functional tight junctions; Figure 3B; green arrowhead). The presence of functional tight junctions at this stage is in agreement with the previously reported expression of tight junction proteins ZO-1 and Claudin-5 in cerebral blood vessels at 3 dpf (Jeong et al., 2008; Xie et al., 2010). As transcytosis has been implicated in the maturation of the blood-retinal barrier (Chow and Gu, 2017), we next examined the levels of luminal and abluminal flask-shaped vesicles filled with gold nanoparticles as a means of assessing transcytosis. Quantification of gold-filled luminal and abluminal flask-shaped vesicles revealed an average of 0.21 ± 0.03 and 0.17 ± 0.02 vesicles/µm, respectively (Figure 3H and I). These data reveal that the basement membrane becomes filled with gold nanoparticles via vesicular transport rather than intercellular passage between immature tight junctions.

Suppression of transcytosis determines the timing of functional BBB formation.

(A, C, E) TEM images of individual blood vessel cross-sections after injection of electron-dense gold nanoparticles at 3 dpf (A), 5 dpf (C), and 7 dpf (E). Endothelial cells (EC) are pseudo-colored green, pericytes (P) are pseudo-colored purple and red blood cells (RBC) are pseudo-colored red when present in the lumen (L). Turquoise arrowheads highlight the gold-filled basement membrane at 3 dpf (A). (B, D, F) High magnification images (25000x) of the areas boxed in A, C, and E, respectively, with the endothelial cells outlined with white dashed lines. The images are oriented with the lumen (L) on top and the ablumen (A) on the bottom. Tight junctions are functional as early as 3 dpf (B), as seen by their ability to halt the gold nanoparticles at the so-called ‘kissing point’ (green arrowhead), and remain functional throughout development (D and F). Even though the tight junctions are functional at 3 dpf, the endothelial basement membrane is filled with electron-dense gold nanoparticles (B, turquoise arrowheads). This appears to be due to an elevated level of luminal and abluminal gold-filled vesicles (magenta arrows). The scale bars represent 200 nm. (G) Quantification of the endothelial basement membrane gold intensity normalized to luminal gold intensity. (H and I) Quantification of the vesicular densities both on the luminal (H) and abluminal (I) membrane of endothelial cells reveals a suppression of vesicular densities beginning at 5 dpf that remains constant at 7 dpf. N = 4 fish, each marked with a different color, with at least 10 blood vessels quantified for each fish and displayed as single points. ****p<0.0001, **p<0.01, ns is not significant by nested one-way ANOVA.

https://doi.org/10.7554/eLife.47326.011

When we repeated this assay at 5 dpf, when the barrier becomes less permeable to fluorescently conjugated tracers (Figures 1 and 2), we still observe close contacts between endothelial cells and pericytes (Figure 3C). At 5 dpf, the basement membrane was noticeably lacking gold particles as compared to 3 dpf (Figure 3D), with a reduced average gold intensity of 0.85 ± 0.02 (Figure 3G). Like at 3 dpf, the tight junctions remained functional at 5 dpf, based on their capacity to halt gold nanoparticles at the kissing points between neighboring endothelial cells (76/76 functional tight junctions; Figure 3D). However, the levels of vesicles both luminally and abluminally were notably decreased to 0.08 ± 0.01 and 0.07 ± 0.01 vesicles/µm, respectively (p<0.001 (luminal) and p<0.0001 (abluminal), nested two-way ANOVA; Figure 3H and I). At 7 dpf, the neurovascular cellular interactions remained constant with endothelial cells in close contact with pericytes and neurons, and with an unfilled basement membrane (average gold intensity of 0.91 ± 0.02; Figure 3E and G). Importantly, the tight junctions remained functional (51/51 functional tight junctions), and the low vesicular densities observed at 5 dpf remained comparably low at 7 dpf with 0.07 ± 0.01 vesicles/µm, both luminally and abluminally (Figure 3F, H and I). Taken together with the fluorescent tracer data, our data suggests that the zebrafish BBB becomes functionally impermeable at 5 dpf via the suppression of vesicular trafficking.

Conserved role of Mfsd2a in determining BBB function

Given the important role of suppressing transcytosis in determining the developmental maturation of the mouse BBB, and a similar correlation of the timing between the suppression of transcytosis and functional barrier formation in zebrafish, we wondered whether the key mammalian barrier regulator Mfsd2a, which suppresses caveolae mediated transcytosis (Ben-Zvi et al., 2014; Andreone et al., 2017), plays a conserved role in zebrafish. Zebrafish contain two paralogues of Mfsd2a, mfsd2aa and mfsd2ab, both of which are expressed in the developing zebrafish CNS (Guemez-Gamboa et al., 2015). In order to resolve the vascular expression of both paralogues, we performed fluorescent in situ hybridization (FISH) in kdrl:mCherry transgenic fish sections at 3 and 5 dpf and in adult brain sections. To control for off-target signal, we also performed FISH with a mouse Gfap probe, which should not bind to the zebrafish transcript (Figure 4). Vascular expression levels for mfsd2aa or mfsd2ab were then background corrected using the measurements from the Gfap probe. As a positive control, we also performed FISH for the BBB-specific vascular marker slc2a1a (also known as glut1b) (Umans et al., 2017; Quiñonez-Silvero et al., 2019), and observed consistent slc2a1a signal in cerebral vessels at all time points examined (Figure 4—figure supplement 1). FISH revealed undetectable levels of mfsd2aa expression in cerebral vessels at 3 dpf, higher levels at 5 dpf, when BBB endothelial cells suppress transcytosis (Figure 3), and intermediate levels in adult blood vessels (Figure 4). Mfsd2ab, on the other hand, is highly expressed in cerebral vessels at 3 and 5 dpf and lowly expressed in adult vessels (Figure 4). Mfsd2aa is 61% identical to human MFSD2A and 62% identical to mouse Mfsd2a (Figure 4—figure supplement 2A). Mfsd2ab is 64% identical to human MFSD2A and mouse Mfsd2a (Figure 4—figure supplement 2A). The two paralogues are only 68% identical to each other, but they both contain the lipid binding domain that is critical for governing barrier properties (Figure 4—figure supplement 2A; Andreone et al., 2017). Given the lack of a clear paralogue that most closely resembles Mfsd2a, we generated CRISPR mutants for both paralogues independently. Mfsd2aahm37/hm37 mutants have a 7 bp deletion in exon 2 (Figure 4—figure supplement 2B) that is predicted to lead to a premature stop codon at amino acid 82 (Figure 4—figure supplement 2A; black box) and exhibit reduced levels of mfsd2aa mRNA (Figure 4—figure supplement 2C). Homozygous mfsd2aa mutants are viable and fertile. While mfsd2aa mutants appear in Mendelian ratios at 5 dpf (105/408 fish), a reduced number of mfsd2aa mutants survive to adulthood (44/248 fish). Mfsd2abhm38/hm38 mutants have a 19 bp deletion in exon 5 (Figure 4—figure supplement 2B) that is predicted to lead to a premature stop codon at amino acid 175 (Figure 4—figure supplement 2A; black box) and exhibit reduced levels of mfsd2ab mRNA (Figure 4—figure supplement 2C). Homozygous mfsd2ab mutants are also viable and fertile and appear in Mendelian ratios both as larvae (51/215 fish) and in adulthood (33/124 fish). Neither mutant displayed obvious angiogenic defects, similar to the normal vasculature observed in mouse Mfsd2a mutants (Ben-Zvi et al., 2014). In addition to BBB defects, Mfsd2a knockout mice also exhibit microcephaly (Ben-Zvi et al., 2014). However, neither mutant displayed decreased larval brain size.

Figure 4 with 2 supplements see all
Fluorescent in situ hybridization (FISH) reveals vascular signal for both mfsd2aa and mfsd2ab at 5 dpf.

(A) FISH for mfsd2aa (left), mfsd2ab (middle) and mouse Gfap negative control (right) in Tg(kdrl:mCherry) 3 dpf, 5 dpf and adult brain tissue. FISH for mfsd2aa (left) reveals no vascular expression above background at 3 dpf, high levels at 5 dpf and low levels in adult blood vessels. FISH for mfsd2ab (middle) reveals high levels of vascular signal at 3 and 5 dpf and negligible vascular signal in adults. Neither Mfsd2a paralogue was expressed exclusively in the vasculature, outlined in white, at any time examined and mfsd2ab displayed higher overall expression throughout the larval brain than mfsd2aa. Scale bar represents 10 µm. (B) Quantification of the number of transcript puncta per blood vessel area in 3 dpf, 5 dpf and adult brain sections after background (mouse Gfap expression) subtraction. N = 6 sections from at least 3 different fish.

https://doi.org/10.7554/eLife.47326.013

To investigate whether either paralogue was necessary for barrier formation, we performed NHS tracer injection assays as we did to identify the developmental timeline of barrier formation and assayed for BBB leakage at 5 dpf. In addition to the use of the exogenous injected fluorescent tracer (1 kDa NHS), we also assayed the leakage of the endogenous transgenic serum DBP-EGFP fusion protein (Tg(l-fabp:DBP-EGFP)). Mfsd2aa mutants displayed at least a two-fold increase in midbrain and hindbrain parenchymal tracer leakage, both for the injected 1 kDa NHS and the endogenous serum transgene 80 kDa DBP-EGFP, at 5 dpf compared to wild-type sibling controls (Figure 5). These data suggest that mfsd2aa plays a similar role to mouse Mfsd2a in determining barrier properties. Conversely, mfsd2ab mutants displayed similarly low levels of parenchymal tracer intensity to wild-type controls at 5 dpf (Figure 5—figure supplement 1), indicating that mfsd2ab is dispensable for functional barrier formation. Similarly, when we injected different guides to target either mfsd2aa or mfsd2ab and assessed BBB permeability in F0 crispants, we observed increased BBB permeability in mfsd2aa crispants but not mfsd2ab crispants (Figure 5—figure supplement 2), confirming the mutant phenotypes. To see if the loss of both paralogues resulted in increased barrier permeability, we investigated mfsd2aahm37/hm37; mfsd2abhm38/hm38 double mutants for BBB permeability using the endogenous serum DBP-EGFP fusion protein (Figure 5—figure supplement 3). While mfsd2aa single mutants displayed the previously observed increase in barrier permeability and mfsd2ab single mutants did not, mfsd2aahm37/hm37; mfsd2abhm38/hm38 double mutants displayed similar levels of increased barrier permeability to the mfsd2aa single mutants (Figure 5—figure supplement 3). These data correspond with previously reported mfsd2aa morpholino knockdown leading to increased BBB permeability (Guemez-Gamboa et al., 2015). However, neither our mfsd2aa or mfsd2ab mutants display brain hemorrhage or increased embryonic lethality, as reported in the morphants (Guemez-Gamboa et al., 2015). Taken together, these data suggest that the two paralogues, while structurally similar, play different roles, with zebrafish mfsd2aa and mammalian Mfsd2a sharing a conserved role in the BBB.

Figure 5 with 3 supplements see all
Mfsd2aa mutants exhibit increased BBB permeability.

(A) Representative maximum intensity projection images of the brain of wild-type and mfsd2aa mutants injected with a fluorescent 1 kDa NHS tracer (turquoise) at 5 dpf. Mfsd2aa mutants have increased levels of NHS in the midbrain and hindbrain parenchyma outside of the vasculature (magenta; Tg(kdrl:mCherry)) compared to wild-type siblings. (B) Representative maximum intensity projection images of the brain of wild-type and mfsd2aa mutants expressing the fluorescently labeled 80 kDa transgenic serum protein DBP-EGFP (green) at 5 dpf. Mfsd2aa mutants have increased levels of DBP-EGFP in the midbrain and hindbrain parenchyma compared to wild-type siblings. The scale bar represents 50 µm. (C) Quantification of normalized parenchymal tracer intensity in the midbrain and hindbrain of wild-type (black) and mfsd2aa mutants (magenta) reveals that mfsd2aa mutants have significantly increased levels of tracer permeability, both for the injected NHS (A) and the endogenous transgene DBP-EGFP (B). Parenchymal tracer intensity outside of the vasculature was measured and normalized to the blood vessel tracer intensity for both the midbrain and hindbrain in each fish and displayed as a single point. The mean and the standard error are drawn as black lines. ****p<0.0001, ***p<0.001 by t test.

https://doi.org/10.7554/eLife.47326.017

Since mfsd2aa mutants contain a leaky BBB (Figure 5), we wanted to further investigate the dynamic nature of the leakage in mfsd2aa mutants using time lapse live imaging to determine how tracers enter the mutant brain and whether or not this would mirror the leakage observed in larval fish (Figure 6). In 9 mfsd2aa heterozygote control fish (Figure 6—Videos 1 and 2), we observed similarly low rates of Dextran accumulation to that observed in wild-type 5 dpf fish (Figure 2). In 9 mfsd2aa mutant fish (Figure 6—Videos 3 and 4), we also never observed any bursting directly from blood vessels as sporadically observed in 3 dpf fish (Figure 2). However, we uncovered a significantly increased rate of diffuse Dextran accumulation in the brain parenchyma in mfsd2aa mutants (slope of 0.098 intensity/min) compared to heterozygote controls (slope of 0.019 intensity/min; Figure 6). Interestingly, the increased Dextran permeability observed in mfsd2aa mutants closely resembles the rate and overall levels of Dextran permeability of the immature 3 dpf larvae (Figure 2).

Figure 6 with 4 supplements see all
Dynamic tracer leakage in mfsd2aa mutants via live imaging.

(A) Time course stills of Dextran tracer dynamics at 5 dpf in mfsd2aa heterozygote controls reveals a sealed BBB. (B) Time course stills of Dextran tracer dynamics in mfsd2aa mutants at 5 dpf reveals a leaky BBB, with increased overall Dextran extravasation into the brain parenchyma. The scale bar represents 20 µm. (C and D) Representative dorsal maximum intensity projection of the midbrain of mfsd2aa heterozygotes (C) and mfsd2aa mutants (D) at the first and last time point examined. While the heterozygotes restrict the Dextran within the cerebral blood vessels at 5 dpf, mfsd2aa mutants exhibit a large increase in overall parenchymal Dextran intensity over the course of 1 hr. Boxed regions are representative of the 6 areas per fish used for analysis in E. The scale bar represents 50 µm. (E) Quantification of Dextran intensity in the brain parenchyma over time in heterozygote controls (black) and mfsd2aa mutants (magenta) shows a significant difference in tracer leakage dynamics (p<0.0001, Mann Whitney U test), with both more total Dextran accumulation and a faster rate of Dextran accumulation in the mutant brain parenchyma than heterozygote controls. N = 9 fish with 6 regions analyzed and averaged per fish and normalized to Dextran intensity in circulation.

https://doi.org/10.7554/eLife.47326.022

Given the leakage phenotype in mfsd2aa mutants at 5 dpf, we next wanted to examine whether the leakage phenotype persisted into adulthood. To address this, we assessed the localization of the serum DBP-EGFP fusion protein in adult brains sections of mfsd2aa mutant and wild-type controls. As expected, the wild-type siblings retained the DBP-EGFP within their blood vessels throughout the brain (Figure 7). However, mfsd2aa mutants exhibited DBP-EGFP extravasation into the brain parenchyma (Figure 7), suggesting that the leakage phenotype was not limited to larval fish. Finally, to determine whether this increased permeability was due to increased transcytosis as in Mfsd2a knockout mice, we measured vesicular density in capillaries with luminal diameters less than 5 µm from adult mutant and wild-type siblings using TEM. The mfsd2aa mutant blood vessels appeared morphologically normal by TEM, composed of a thin single layer of endothelial cells in close contact with pericytes, as observed in their wild-type siblings (Figure 8A and B). A closer examination of the endothelial cells revealed electron-dense tight junction complexes between all apposed endothelial cells in both wild-type and mfsd2aa mutant fish (Figure 8C and D). However, while wild-type fish display similarly low levels of luminal (0.1 vesicles/µm) and abluminal (0.11 vesicles/µm) vesicular densities to those observed at 7 dpf (Figure 8C–F), mfsd2aa mutant fish display a significant increase in luminal and abluminal vesicular densities (0.29 and 0.27 vesicles/µm, respectively) compared to wild-type siblings (Figure 8C–F). Interestingly this increase in vesicular abundance is even higher than that observed during early barrier development at 3 dpf (Figure 3). These data also suggest that the increased tracer leakage observed in mfsd2aa mutants results from an increase in vesicular trafficking across the BBB, further supporting a conserved role for mfsd2aa in determining barrier properties. In contrast to mfsd2aa mutants, mfsd2ab mutants displayed similar levels of abluminal and luminal vesicular pit density to wild-type siblings (Figure 8—figure supplement 1), further demonstrating that the paralogue mfsd2ab does not play a conserved role in determining barrier properties in zebrafish.

Adult mfsd2aa mutants display increased BBB permeability.

(A and B) Sagittal sections of the adult brain of wild-type (A) and mfsd2aa mutants (B) reveals increased extravasation of the transgenic DBP-EGFP serum fusion protein (green) in the forebrain (FB), midbrain (MB) and hindbrain (HB) of mfsd2aa mutants compared to wild-type siblings. The scale bar represents 100 µm. (C) Quantification of normalized parenchymal DBP-EGFP tracer intensity in the forebrain, midbrain and hindbrain of wild-type (black) and mfsd2aa mutants (magenta) reveals that mfsd2aa mutants have significantly increased levels of DBP-EGFP permeability throughout the adult brain. Parenchymal tracer intensity outside of the vasculature was measured in 6 distinct areas, averaged and normalized to the blood vessel DBP-EGFP intensity for each region (FB, MB and HB) and displayed as a single point for each fish. The mean and the standard error are drawn as black lines. ****p<0.0001 by t test.

https://doi.org/10.7554/eLife.47326.028
Figure 8 with 1 supplement see all
Mfsd2aa mutants exhibit increased transcytosis.

(A and B) TEM images of individual blood vessel cross-sections of adult wild-type (A) and mfsd2aa mutant fish (B). Endothelial cells (EC) are pseudo-colored green, pericytes (P) are pseudo-colored purple and red blood cells (RBC) are pseudo-colored red. (C and D) High-magnification images of endothelial cells outlined with white dashed lines of wild-type (C) and mfsd2aa mutants (D). The images are oriented with the lumen (L) on top and the ablumen (A) on the bottom. Mfsd2aa mutants appear to have normal tight junctions (green arrowhead) but elevated levels of luminal and abluminal vesicles (magenta arrows). The scale bars represent 200 nm. (E and F) Quantification of the vesicular densities both on the luminal (E) and abluminal (F) side of the endothelial cells reveals that mfsd2aa mutants have increased vesicular densities. N = 4 fish, each marked with a different color, with 15 blood vessels quantified for each fish and displayed as single points. ****p<0.0001, **p<0.01 by nested t test.

https://doi.org/10.7554/eLife.47326.030

Discussion

One of the major advantages to studying the BBB in zebrafish is the ability to perform live imaging of tracer permeability dynamics. We provide some of the first data on the dynamics of immature barrier leakage during early development and of barrier leakage seen in a genetic mutant fish at the time when the BBB should be functionally mature. At 3 dpf, we observed two types of leakage, the gradual overall increase in the parenchyma and rare transient bursts, both from a migrating sprout and established blood vessels. We attribute some of the gradual diffuse increase in parenchymal tracer uptake to the high levels of vesicular trafficking observed in our TEM analyses in which the endothelial basement membrane became filled with gold nanoparticles. Time lapse analysis of mfsd2aa mutants revealed similar diffuse tracer leakage levels and rates to that observed in the leaky 3 dpf larvae. Since mfsd2aa mutant fish have normal angiogenesis but elevated endothelial transcytosis levels, the Dextran leakage at 5 dpf, when their siblings have a functional BBB, provides a direct measure of the in vivo rates and mode of BBB leakage due to unsuppressed transcytosis. The rarer bursts of leakage, on the other hand, could be due to fleeting tight junctional ruptures between neighboring endothelial cells rather than increased vesicular trafficking. As these events were extremely scarce during our time lapse imaging, it would be nearly impossible to capture these potential junctional breaches by TEM to confirm this hypothesis. However, with continuing advances in the resolution of fluorescence microscopy, we could use zebrafish to resolve whether these two types of leakage, gradual and bursting, proceed through the same or different subcellular routes into the brain parenchyma, either paracellularly or transcellularly.

While the BBB constitutes the largest interface for blood-brain exchange in adults (Nag and Begley, 2005), and represents the most well-studied cellular barrier, it is not the only brain barrier. The blood-cerebrospinal fluid (CSF) barrier is created by the polarized epithelial cells of the choroid plexus (CP) that projects directly into the brain ventricle and creates a barrier between the fenestrated capillaries of the CP and the CSF (Wolburg and Paulus, 2010). During development, these CP epithelial cells become selectively less permissive by expressing tight junction proteins, including Claudin-5 and Occludin, and nutrient transporters, including Mrp1, Mdr1 and Oat3 (Henson et al., 2014; van Leeuwen et al., 2018). In zebrafish, this blood-CSF barrier has been shown to become completely impermeable by 4 dpf to various molecular weight tracers including 3 kDa fluorescein, 10 kDa and 40 kDa Dextrans (Henson et al., 2014). However, while CP epithelial cells are initially permeable to 3 kDa fluorescein and 10 kDa Dextran at 2 dpf, they never allow the passage of the larger 40 kDa Dextran (Henson et al., 2014). Since we saw comparable BBB permeability for all tracers used during the developmental time course, including the 80 kDa DBP-EGFP serum transgene, this strongly suggests that the tracers entered the brain predominantly through leaky BBB vessels rather than through an immature blood-CSF barrier. The avascular arachnoid epithelium directly beneath the dura that completely ensheaths the CNS acts as a third barrier between the extracellular fluids of the CNS and the rest of the body (Abbott et al., 2006). Previous HRP tracer injections have shown that the zebrafish arachnoid barrier matures after 9 dpf (Jeong et al., 2008). Since we did not observe any differences in tracer impermeability between 5 and 10 dpf, when the arachnoid barrier is still leaky, this suggests that the immature arachnoid barrier does not significantly contribute to the observed tracer extravasation into the brain earlier in development. Finally, the similarities between the observed amounts and dynamics of leakage in 3 dpf immature brains and mfsd2aa mutant brains at 5 dpf, which contain BBB-specific endothelial transcytosis defects, further suggests that the observed tracer leakage at 3 dpf is predominantly arising through the BBB. Taken together, while alternative routes for tracer entry into the brain exist, including an immature blood-CSF or arachnoid barrier, our methods allowed us to specifically evaluate BBB functionality, and other routes are not significantly contributing to the tracer leakage we observed in the larval brain.

Among the various clathrin-independent transcytotic pathways (Tuma and Hubbard, 2003; Sandvig et al., 2018), caveolae are particularly abundant in vascular endothelial cells (Frank et al., 2003). Caveolae are caveolin-coated 50–100 nm flask-shaped invaginations of the plasma membrane (Palade, 1953; Palade, 1961). Furthermore, the suppression of caveolae-mediated transcytosis regulates the development and function of both the mouse blood-retinal barrier and the BBB (Andreone et al., 2017; Chow and Gu, 2017). Taken together, this suggests that the increased transcytosis during early larval stages is most likely caveolae-mediated. Interestingly, the linear profile of Dextran uptake in the midbrain at 3 dpf closely resembles the caveolae-mediated uptake of a fluorescently conjugated aminopeptidase P (APP) antibody in the mouse lung (Oh et al., 2007). While similarly linear, the scale is on the order of minutes in the larval zebrafish brain versus seconds in the mouse lung. This discrepancy in timing is most likely due to the large difference in the vesicular densities between the immature BBB endothelium (average of 2 vesicles/µm2) and the continuous endothelium of peripheral tissues, which ranges from 30 to 98 vesicles/µm2 on the luminal membrane of diaphragm and myocardial endothelium (Simionescu et al., 1974). Since loss of caveolin 1 (Cav1) is sufficient to precociously seal the mouse blood-retinal barrier at the angiogenic front (Chow and Gu, 2017), it would be interesting in future work to examine whether zebrafish cav1 mutants (Cao et al., 2016), which are viable as homozygotes, also exhibit an earlier onset of BBB maturation.

BBB permeability is tightly regulated by endothelial cell interactions with pericytes and astroglial cells. For the first time, we visualized their locations under TEM in zebrafish throughout barrier development. Pericytes have been shown to be essential for establishing the mammalian BBB (Armulik et al., 2010; Bell et al., 2010; Daneman et al., 2010). Similarly, pericyte-deficient notch3fh332 fish display increased BBB permeability in a tight junction independent manner like pericyte-deficient mice (Wang et al., 2014). Our TEM data show that zebrafish pericytes are in close contact with brain endothelial cells (Figures 2 and 5), even at the earliest stage examined (3 dpf; Figure 2), and are embedded within the endothelial basement membrane as in mammals. Our subcellular localization data is in line with a growing body of evidence for the conserved role and molecular profile of pericytes in the zebrafish BBB (Wang et al., 2014; Ando et al., 2016; Lei et al., 2017; Vanlandewijck et al., 2018; Ando et al., 2019).

Taken together this study provides a thorough characterization of the development of the zebrafish BBB, highlighting regional differences in timing of functional maturation and capturing the dynamics of the immature BBB. Furthermore, our developmental TEM series demonstrates the role of vesicular trafficking in regulating zebrafish BBB function during development. We have also shown that this down-regulation of vesicular trafficking is necessary for functional BBB formation, as mfsd2aa mutants display increased barrier permeability due to unsuppressed transcytosis. Finally, our time lapse imaging of mfsd2aa mutants provides the first direct measure of tracer permeability in vivo that results from increased transcytosis. We hope that this work will serve as a launching pad for future studies using zebrafish to understand the molecular regulators of BBB development and homeostasis in vertebrates.

Materials and methods

Zebrafish strains and maintenance

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Zebrafish were maintained at 28.5°C following standard protocols (Westerfield, 1993). All zebrafish work was approved by the Harvard Medical Area Standing Committee on Animals under protocol number 04487. Adult fish were maintained on a standard light-dark cycle from 9 am to 11 pm. Adult fish, age 3 months to 2 years, were crossed to produce embryos and larvae. For imaging mutant larvae, 0.003% phenylthiourea (PTU) was used beginning at 1 dpf to inhibit melanin production. These studies used the AB and Casper (mitfaw2/w2; mpv17a9/a9) (White et al., 2008) wild-type strains and the transgenic reporter strains Tg(l-fabp:DBP-EGFP)Iri500 (Xie et al., 2010), Tg(kdrl:HRAS-mCherry)s896 (Chi et al., 2008), abbreviated as Tg(kdrl:mCherry) in the text, Tg(actb2:MA-Citrine)hm26 (Mosaliganti et al., 2012) and Tg(actb2:Hsa.H2B-Scarlet)hm38.

Tracer injections

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Larvae were immobilized with tricaine and placed in an agarose injection mold with their hearts facing upwards. 2.3 nl of Alexa Fluor 405 NHS Ester (Thermo Fisher: A30000) or Alexa Fluor 647 10 kDa Dextran (Thermo Fisher: D22914) fluorescently conjugated tracers (10 mg/ml) were injected into the cardiac sac using Nanoject II (Drummond Scientific, Broomall, PA). Larvae were then mounted with 1.5% low gelling agarose (Sigma: A9414) in embryo water on 0.17 mm coverslips and imaged live within 2 hr post-injection. For developmental electron microscopy experiments, 2.3 nl of 5 nm NHS-activated gold nanoparticles (Cytodiagnostics: CGN5K-5–1, ~1.114 particles/ml in PBS) were injected into the cardiac sac just as for the fluorescently conjugated tracers. After 5 min of circulation, the fish were fixed for electron microscopy. The brains of transgenic DBP-EGFP adults were fixed in 4% paraformaldehyde in PBS overnight at 4°C. Following fixation the brains were washed three times and sagittally sectioned with a cryostat (30 µm) to reveal DBP-EGFP localization within the brain.

Transmission electron microscopy (TEM)

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Fish were anesthetized with tricaine and initially fixed by immersion in 4% paraformaldehyde (VWR:15713 s)/0.1M sodium-cacodylate (VWR:11653). Following this initial fixation, the larval fish and adults with exposed brains were further fixed for 7–14 days in 2% glutaraldehyde (Electron Microscopy Sciences: 16320)/4% paraformaldehyde/0.1M sodium-cacodylate at room temperature. Following fixation, larvae or dissected brains were washed overnight in 0.1M sodium-cacodylate. Entire larval heads or coronal vibratome free-floating sections of adult brains (50 µm) were post-fixed in 1% osmium tetroxide and 1.5% potassium ferrocyanide, dehydrated, and embedded in epoxy resin. Ultrathin sections of 80 nm were then cut from the block surface and collected on copper grids. The adult sections were counter-stained with Reynold’s lead citrate prior to imaging.

CRISPR mutants

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Mfsd2aa mutant fish were generated by injection of Cas9 RNA and a single guide RNA (5’-GGTGTGTTTTGCGATCGGAG-3’) targeting exon 3 into single-cell fertilized AB wild-type embryos. Mfsd2ab mutant fish were generated by injection of Cas9 protein and a single guide RNA (5’-TGAGAGCAGAGTAGGGCACG-3’) targeting exon 5 into single-cell fertilized double transgenic Tg(l-fabp:DBP-EGFP; kdrl:mCherry) embryos. F0 injected fish were raised, outcrossed to wild-type fish and screened for potential mutant founders by PCR and sequencing. Founders were outcrossed to wild-type fish at least three times before leakage analyses. Heterozygous fish were intercrossed for all leakage assays, with the exception of the mfsd2aa time lapse microscopy, where a heterozygous fish was crossed to a homozygous mutant. All larvae were imaged prior to genotyping to identify wild-type and mutant fish. The stable mfsd2aa mutant line was genotyped using 5’-AAATCACCTCTTCCAGTGAGGA-3’ and 5’-ATAGTAACAAAACGATGCTGAGCC-3’ primers followed by a Pvu1 restriction digest, which does not digest the mutant DNA. Mfsd2ab mutants were genotyped using 5’-GTCTACTCCATTTGCTGTACTTTGC-3’ and 5’-AGAGCAGAGTAGGGCACGTGGAAGC-3’ primers for the wild-type allele and 5’-GTCTACTCCATTTGCTGTACTTTGC-3’ and 5’-GTGCTGATGAAGCACTGATGAAGCA-3’ for the mutant allele.

Mfsd2aa crispant fish were generated by injection of Cas9 protein and two new guide RNAs (5’- GCTCACTAAAAACCCAACGG-3’ and 5’-TTGTAGAAAGAAGCCCAGGG-3’) targeting exons 2 and 3 into single-cell fertilized double transgenic Tg(l-fabp:DBP-EGFP; kdrl:mCherry) embryos. Similarly, mfsd2ab crispants were generated by injection of Cas9 protein and two new guide RNAs (5’-AAGCGAGATGGCAAAAGGAG-3’ and 5’- ACCCACAAACAAGATGATGG-3’) targeting exons 1 and 3 into single-cell fertilized double transgenic Tg(l-fabp:DBP-EGFP; kdrl:mCherry) embryos. F0 crispants were analyzed for leakage at 5 dpf.

qPCR

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qPCR was performed on cDNA obtained from wild-type, mfsd2aahm37, mfsd2abhm38, mfsd2aa and mfsd2ab F0 crispant 5 dpf whole larvae mRNA. An Eppendorf Realplex2 Real-Time PCR system was used for qPCR experiments and gene expression levels were normalized relative to that of a reference gene, 18s using primers 5’-TCGCTAGTTGGCATCGTTTATG-3’ and 5’-CGGAGGTTCGAAGACGATCA-3’. Levels of mfsd2aa were measured using 5’-CTCTTCACTTCGCTAGCCTTCATG-3’ and 5’-CGATGTAAACAGCAGTCTTTTTCCC-3’ primers and mfsd2ab levels were measured with 5’-TCTCGACTCTTAGTCTTGATTTCGC3’ and 5’-GAGTCCGTTTCTGAATCCATCTCG-3’. All reactions were performed in technical duplicates, and the results represent four independent biological samples including the SEM.

HCR fluorescent in situ hybridization (FISH)

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HCR RNA in situ hybridization (Molecular Instruments) experiments on 14 µm cryosections of fixed 3 and 5 dpf larvae and adult brains were performed as previously described (Choi et al., 2018). Briefly, sections were air dried and re-fixed in 4% paraformaldehyde in PBS for 10 min at room temperature. Following fixation, sections were further permeabilized using 1 µg/ml Proteinase K (ThermoFisher) for 5 min, followed by PBS washes. Slides were then put through an ethanol dehydration series of 50%, 70% and two rounds of 100% ethanol and air dried for 5 min. Dried slides were put into pre-hyb solution for at least 10 min at 37°C. Subsequently, the probes for mfsd2aa, mfsd2ab, slc2a1a and mouse Gfap were added to the slides at a final concentration of 4 nM to hybridize overnight at 37°C. The next day, slides were washed with a series of wash buffer to 5x SSCT (5x SSC with 0.1% Tween 20). Excess liquid was then removed and samples were immersed in amplification buffer at room temperature for 30 min prior to hairpin amplification, which occurred overnight at room temperature. The next day, slides were washed in 5x SSCT and mounted with Fluoromount-G (Electron Microscopy Sciences).

Imaging

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All imaging of tracer permeability was performed on a Leica SP8 laser scanning confocal microscope using the same acquisition settings using a 25x water immersion objective. Time lapse imaging was performed on the SP8 using a resonance scanner. FISH sections were also imaged with the Leica SP8 using a 63x oil immersion objective. A 1200EX electron microscope (JOEL) equipped with a 2 k CCD digital camera (AMT) was used for all TEM studies. Images were visualized and quantified using ImageJ (NIH) and Adobe Photoshop. Time lapse videos were visualized as 3D reconstructions and cropped to highlight particular blood vessels of interest using Imaris software (Bitplane).

Tracer permeability quantification

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All quantifications were performed on blinded image sets. For static images, parenchymal fluorescent tracer intensity was measured using ImageJ in the entire regional parenchyma outside of the vasculature in 30 µm thick maximum intensity projections of the larval brains. These projections began on average 15 µm below the mesencephalic vein to reduce the effects of potential leakage diffusion from the surface vessels and had the vasculature masked and removed for intensity quantification. These parenchymal tracer intensity values were then normalized to the tracer intensity within the vasculature to account for differential amounts of circulating tracer. For time lapse imaging, Dextran intensity was measured in 6 parenchymal regions of average intensity projections of the time lapse videos and averaged as a single value per fish. These values were normalized to the average fluorescence intensity in the lumen at each time point. Z-stacks were collected with 1 µm z-steps using a 25x water immersion objective on a Leica SP8 laser scanning confocal microscope.

TEM quantification

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For all TEM quantifications, vesicular density values were calculated from the number of non-clathrin coated flask-shaped vesicles per µm of endothelial luminal or abluminal membrane for each image collected. Larval basement membrane gold intensity was measured in 3 regions per endothelial cell and normalized to luminal gold intensity values within individual images using ImageJ. All images for analysis were collected at 12,000 x magnification on the JOEL 1200EX electron microscope. 10–15 vessels were quantified for each fish, with each color representing a different fish.

FISH quantification

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For all FISH experiments, we quantified the number of gene expression puncta within the vasculature of maximum intensity projections using ImageJ. These puncta were normalized to total blood vessel area within the image. Non-specific background levels of mouse Gfap probe binding were similarly assessed at all three time points. These background levels were then subtracted from mfsd2aa and mfsd2ab expression levels and negative values were set to 0.

Statistical analysis

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All statistical analyses were performed using Prism 8 (GraphPad Software). Two group comparisons were analyzed using an unpaired two-tailed t test. Nested t tests were employed for all electron microscopy comparisons to account for the actual N versus vessels analyzed. Multiple group comparisons were analyzed with two-way ANOVA, followed by a post hoc Tukey’s multiple comparison test. The time lapse leakage dynamics were analyzed with a Mann-Whitney U test to discriminate whether the two patterns of leakage accumulation were different. Sample size for all experiments was determined empirically using standards generally employed by the field, and no data was excluded when performing statistical analysis. Standard error of the mean was calculated for all experiments and displayed as errors bars in graphs. Statistical details for specific experiments, including exact n values and what n represents, precision measures, statistical tests used, and definitions of significance can be found in the Figure Legends.

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Decision letter

  1. Didier Y Stainier
    Senior Editor; Max Planck Institute for Heart and Lung Research, Germany
  2. Richard Daneman
    Reviewing Editor; University of California, San Diego, United States
  3. Benoit Vanhollebeke
    Reviewer; Université libre de Bruxelles, Belgium
  4. Michael R Taylor
    Reviewer; University of Wisconsin-Madison, United States
  5. Kelly Monk
    Reviewer; Vollum Institute, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Suppression of transcytosis regulates zebrafish blood-brain barrier development" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a guest Reviewing Editor and Didier Stainier as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Benoit Vanhollebeke (Reviewer #1); Michael R Taylor (Reviewer #2); Kelly Monk (Reviewer #3).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

All of the reviewers thought that the work was of potential interest, but that there were a number of major concerns that need to be addressed.

Based on the discussion the reviewers specific points included that: "there were several concerns from all reviewers regarding interpretation of data and terminology that should be addressed both in the major and minor comments" and "a more careful job with the analysis of the new mutants they have generated is required." Furthermore, all reviewers agreed that counting cells with tracer as a proxy for BBB leakage was concerning as this could be a neural intrinsic feature and BBB-independent process, and that the dynamic range of this assay remains unclear as changing the molar amounts of tracer does not lead to changes in cell counts.

The authors should address each of the concerns raised by the reviewers.

Reviewer #1:

This study by Brown et al. investigates BBB maturation in zebrafish larvae combining time-lapse confocal imaging of tracer extravasation, genetics and electron microscopy. By using different tracers, the authors assess the time-point at which developmental BBB maturity is reached in a spatiotemporal fashion. The decrease in leakage in the midbrain at 5 days post fertilization was further explored by TEM and shown to correlate with a decrease in intracellular transcytosis. The sealing of the blood brain barrier, correlating with a decrease in transcytosis, was further explored by studying the conserved function of Mfsd2a in this process.

The paper is of interest as it further documents the similarities between the zebrafish and the mammalian BBB structures. The TEM studies, combined with mfsd2aa mutants convincingly show that transcytosis suppression contributes to BBB maturation in zebrafish, as it does in the mouse.

The authors convincingly report two apparently independent tracer accumulation modes in the zebrafish larval brain. The first one, previously observed by other investigators, results in diffuse tracer accumulation across the parenchyma, follows near-linear kinetics and, in the context of this study, has been analyzed in the midbrain after at 3 and 5 dpf. The other mode is interesting as so far unreported and results in focal tracer accumulation secondary to perivascular "bursts" that can be captured by time-lapse confocal microscopy.

The authors propose that the steady and diffuse accumulation is secondary to transcytosis across the immature BBB endothelium and is controlled developmentally in the midbrain by mfsd2aa, while the other would result from sporadic vessel instability and uptake through what are referred to as "parenchymal cells". The number of such tracer-filled cells is then quantified as a proxy for tracer leakage across the BBB to infer both BBB closure and asses genetic mutants.

While overall interesting and timely, some of the leakage assay methodology central to the message of this manuscript might suffer from technical caveats that could be addressed before publication.

Major comments:

First. While the confocal time-lapse microscopy evidence is strong enough to support the idea that the perivascular bursts are secondary to leakage through the BBB endothelium, there is no direct evidence that the more distant focal accumulation, as well as the disuse accumulation throughout the parenchyma, results from leakage through the immature BBB. When dyes are injected in the zebrafish cardiovascular system, they can reach the brain parenchyma through multiple routes, with only one of them being the BBB. Jeong et al., 2008 reported that several mesencephalic vessels (that are not intraneural BBB vessels) are leaky at 3 dpf. These vessels are therefore a likely alternative tracer entry point to the nearby midbrain. Similarly, upon intracardial injection, tracers of any kind accumulate in the CSF within ventricular spaces, and could therefore reach the brain through the ependymal cell layer. The poorly characterized meningeal barriers (arachnoid, pia mater, glia limitans) constitute yet another portal to the brain. Of note, these latter were shown to close at much later stages (after 9 dpf) by Jeong et al., 2008. Therefore, tracer accumulation in the larval zebrafish brain does not necessarily imply leakage through the BBB. Of note, this challenge is particularly acute due the small size of the larval zebrafish brain, that in contrast to the mammalian structures, poorly allows for anatomical leakage inferences. Admittedly, addressing this issue experimentally is challenging and well beyond the scope of what can be achieved at the revision step of this particular study. In this reviewer's opinion, these anatomical considerations should however be carefully discussed by the authors as they impact directly several of the main conclusions of the manuscript.

Overall, we recommend rephrasing the manuscript in order to:

i) shift away from a BBB-centric view and give more space to the anatomical complexity of the multiple brain barriers. We would for instance suggest rephrasing the first section heading from ("P-A gradient of zebrafish BBB development" to "P-A gradient of zebrafish brain barriers development").

ii) Importantly, avoid any firm causality inferences between BBB phenotypes (like the presence or absence of vesicular transport) and at distance tracer accumulation. For instance, in the Abstract the sentence "Electron microscopy studies further reveal that this steady accumulation results from high levels of transcytosis that are eventually suppressed, sealing the BBB" should be rephrased or removed. Of note, this is not to say that the authors shouldn't discuss on the developmental correlation of transcytosis suppression and reduced tracer accumulation in the Discussion section. Similarly, the HRP leakage in the adult mfsd2aa mutants are convincing.

Second. While this reviewer agrees that it seems reasonable that the focal tracer accumulation indeed reflects intracellular accumulation in "scavenger " parenchymal cells, the authors fail to present evidence to support this claim. When considering carefully the Videos 1 and 2, tracer extravasation and "cellular" accumulation appear as a synchronous and perhaps unique process, which is puzzling. Moreover, the still images and the video recording these sporadic bursts do not allow discriminating with the alternative scenario of focal accumulation in extracellular spaces, or labelling of apoptotic cells or cell debris. As the authors use these "cells" as a "proxy of tracer leakage", this requires clarification. We suggest approaching this question by confocal imaging with counterstained nuclei and cell membranes.

Third. The authors propose a model in which unrestricted transcytosis triggers diffuse tracer accumulation, a process negatively regulated by mfsd2aa and quantified in WT animals in Figure 2E. Surprisingly, in mfsd2aa mutants, quantification of this mode of leakage was not performed. This is essential to support the model and should be performed.

Fourth. Instead, "parenchymal cells" were monitored (see comment #2). We were not convinced that monitoring the number of "cells" is predictive to leakage extent. It would only be so if (i) the total number of cells is high, stable during development and unaltered in genetic mutants and if (ii) the amount of substrate tracer is limiting so that the more tracer reaches the parenchyma, the more cells are detectable. None of these conditions seem established in light of the data presented. For instance, as the nature of the presumed cells is unknown, their total number (labeled or not) cannot be established and therefore different time points and genetic conditions cannot be compared (Figure 1D, Figure 1—figure supplement 1, Figure 4C, Figure 4—figure supplement 2, 3). How can the authors rule out that the total number of this presumptive cell type is not merely increased in mfsd2aa mutants? According to previous reports mfsd2aa is widely expressed throughout the zebrafish embryonic and larval brain. Similarly, how can the authors exclude that the number of these cells drops over time in the midbrain? Instead of counting "cells", the mean intensity of the "cells" could be analyzed (alternatively, asses the parenchymal leak, see third comment). If transcytosis (or more broadly tracer leakage) is increased, the anticipated result is an increase in the average fluorescence intensity of cell-associated label (rather than an increase in the actual number of cells).

Assessing the number of cells as a readout for transcytosis seems moreover also puzzling from cellular standpoints. Indeed, at least some of these "cells" were shown to be filled secondary to sudden rupture of endothelial integrity (Figure 2E) and therefore would not be a faithful readout for transcytosis upregulation. Final sentence in subsection “Posterior-Anterior Gradient of Zebrafish BBB Development” are unclear in that respect as well. We would agree with the authors that tight junctional defects would likely exhibit size-selective leakage properties (although that remains to be shown in zebrafish). Consequently, the observed labelling of the "parenchymal cells" with tracers of all tested sizes suggest an alternative leakage mode is at play (if we assume this is secondary to BBB defects, see above). It does however not discriminate vessel rupture from transcytosis. So in absence of vessel rupture quantification in mfsd2aa mutants versus WT, interpretation of the cell counts is ambiguous.

Reviewer #2:

1) This reviewer believes that the title of this manuscript overstates the conclusions of the study. To conclude that BBB development is regulated by suppression of transcytosis, implies that other characteristics of BBB development are also controlled by this cellular process. As BBB development requires the acquisition of many properties (e.g. formation of tight junctions, expression of multiple types of transporters, establishment of cellular interactions, etc), including the suppression of transcytosis as described here, please consider a title that more accurately reflects the results of the study.

2) The conclusions of the manuscript rely significantly upon the spatiotemporal expression of mfsd2aa (and mfsd2ab to a lesser extent). Based upon the results of this study, it would be predicted that mfsd2aa is not expressed in brain endothelial cells at 3 dpf, which allows for transcytosis of tracers into the brain parenchyma. Furthermore, it would be predicted that mfsd2aa is expressed in brain endothelial cells by 5 dpf, which then suppresses transcytosis of tracers into the brain parenchyma. However, the authors do not provide any expression data for either mfsd2aa or mfsd2ab.

In previous studies, Guemez-Gamboa et al. used whole-mount in situ hybridization to show that both zebrafish mfsd2aa or mfsd2ab transcripts are expressed throughout the nervous system in zebrafish at 24, 48, and 96 hpf. However, it is not clear from this data whether brain endothelial cells express either transcript. In addition, Thisse and Thisse showed basal expression of mfsd2aa in spinal cord neurons as early as 20-25 somites to Prim-5 stage, no spatial restriction at the Prim-15 to Prim-25 and High-pec to Long-pec stages, and neurocranium expression at 5 dpf (https://zfin.org/ZDB-GENE-041114-166/expression). Thus, it remains to be determined whether zebrafish mfsd2aa or mfsd2ab is expressed in brain endothelial cells.

Please provide expression data that demonstrates mfsd2aa expression (or lack of expression) in zebrafish brain endothelial cells at the appropriate developmental time points (i.e. 3 dpf, 5 dpf, and adult). Perhaps another valid strategy would be to rescue mfsd2aa mutants by cell autonomous expression of wild-type mfsd2aa in endothelial cells.

Guemez-Gamboa et al. (2015) Nature Genetics 47:809-813

Thisse and Thisse, (2004) ZFIN Direct Data Submission (http://zfin.org).

3) To quantify "leakiness", this study focused on the number of parenchymal cells that accumulated tracer in the midbrain and demonstrated that larvae at 3 dpf are leakier than 5 dpf. The authors also indicated that parenchymal cells in the hindbrain did not accumulate tracer at a significant level at either developmental time point. However, in the mfsd2aa mutants, the authors did not examine midbrain leakiness at 3 dpf or in the hindbrain at either 3 dpf or 5 dpf. Do the mfsd2aa mutants at 3 dpf exhibit increased leakiness compared to wild-types at 3 dpf? Also, if mfsd2aa suppresses transcytosis in all brain endothelial cells, then is the hindbrain leakier in mfsd2aa mutants at 3 dpf and 5 dpf? If not, please explain.

Reviewer #3:

This manuscript from O'Brown and colleagues represents the most careful analysis of zebrafish blood brain barrier (BBB) development performed to date. Given the genetic tractability of zebrafish, the unparalleled ability to perform in vivo imaging, and the amenability to small molecule screens, zebrafish is an ideal vertebrate model system for BBB study. Previous work raised a controversy as to whether the BBB forms in zebrafish by 3 days of age (Jeong et al., 2008; Umans et al., 2017) or between 3-10 days of age (Fleming et al., 2013). It is critically important to resolve this controversy and to define how the BBB forms in zebrafish to set the stage for future mechanistic, genetic, and drug discovery studies. For example, one can imagine small molecule screens in zebrafish larvae to find compounds that selectively open the BBB for therapeutic drug delivery. Without fully understanding how the BBB forms in zebrafish, however, these studies cannot be performed. O'Brown et al. begin to tackle this important problem by thoroughly characterizing BBB formation in zebrafish between 3-6 days of age. Using tracer injections, live imaging, and electron microscopy, they show that the hindbrain BBB is closed by 3 days, whereas the midbrain closes between 3-5 days. This developmental gradient is similar to mouse. Given that the lipid transporter Mfsd2a is required for proper BBB formation in mammals, O'Brown and colleagues also generate and analyze new mfsd2a zebrafish mutants. Mutant analysis indicates that the function of Mfsd2a is conserved from zebrafish to mammals.

There are some minor experimental additions that could be performed to strengthen the study's conclusions and impact.

1) Given that antibodies to mark tight junctions exist that work in zebrafish, it would be useful to add this analysis to the time course of BBB formation.

2) Given the controversy surrounding the timing of BBB closure in zebrafish, it is important to extend the time course to 10 days to be consistent with the Fleming et al. study. Additionally, the hindbrain at 3 days shows ~2 cells/embryo taking up tracer whereas the authors consider the midbrain BBB closed at 5-6 days with ~8 cells/larva taking up tracer. Thus, it is important to test if the number of cells taking up tracer further reduces with time.

3) The standard in the field is to rescue a new mutant with wild-type gene product to control for off-target CRISPR/Cas9 effects. Similarly, qPCR for mfsd2aa and mfsd2ab are important controls for the new mutants, especially since phenotypes were not observed in mfsd2ab mutants.

4) Not performing in vivo imaging of the mfsd2aa mutants seems like a missed opportunity. Such investigation could also reveal if both types of leakage observed in the time course are affected in mfsd2aa mutants. The analysis presented in Figure 2E would also be helpful to show for the mutants.

Overall this is an important descriptive study for both zebrafish development and BBB fields that will be foundational for future work in many laboratories.

https://doi.org/10.7554/eLife.47326.035

Author response

The authors should address each of the concerns raised by the reviewers.

Reviewer #1:

[…] Overall, we recommend rephrasing the manuscript in order to:

i) shift away from a BBB-centric view and give more space to the anatomical complexity of the multiple brain barriers. We would for instance suggest rephrasing the first section heading from ("P-A gradient of zebrafish BBB development" to "P-A gradient of zebrafish brain barriers development").

ii) Importantly, avoid any firm causality inferences between BBB phenotypes (like the presence or absence of vesicular transport) and at distance tracer accumulation. For instance, in the Abstract the sentence "Electron microscopy studies further reveal that this steady accumulation results from high levels of transcytosis that are eventually suppressed, sealing the BBB" should be rephrased or removed. Of note, this is not to say that the authors shouldn't discuss on the developmental correlation of transcytosis suppression and reduced tracer accumulation in the Discussion section. Similarly, the HRP leakage in the adult mfsd2aa mutants are convincing.

We thank the reviewer for his thoughtful remarks. We closely examined the literature about alternative routes into the brain during development and have added a paragraph to the discussion talking about the other brain barriers, their developmental timing and restrictiveness during development, as well as potential contribution to the leakage during early development. Based on this thorough literature review and our cumulative findings, we conclude that Figure 1 is mostly representing BBB leakage as discussed in subsection “Conserved Role of Mfsd2a in Determining BBB Function” as well as in the Discussion section. Overall, we have tempered our interpretations of the causality of tracer leakage between different experiments throughout the manuscript. For example, we have altered the referred to sentence in the Abstract to “Electron microscopy studies further reveal high levels of transcytosis in brain endothelium early in development that are suppressed later. The timing of this suppression of transcytosis coincides with the establishment of BBB function.”

Second. While this reviewer agrees that it seems reasonable that the focal tracer accumulation indeed reflects intracellular accumulation in "scavenger " parenchymal cells, the authors fail to present evidence to support this claim. When considering carefully the Videos 1 and 2, tracer extravasation and "cellular" accumulation appear as a synchronous and perhaps unique process, which is puzzling. Moreover, the still images and the video recording these sporadic bursts do not allow discriminating with the alternative scenario of focal accumulation in extracellular spaces, or labelling of apoptotic cells or cell debris. As the authors use these "cells" as a "proxy of tracer leakage", this requires clarification. We suggest approaching this question by confocal imaging with counterstained nuclei and cell membranes.

We agree with this reviewer, and also reviewer 2, that the “scavenger” parenchymal cells should be better characterized. In order to address these concerns, we have performed these same tracer leakage assays in membrane labelled fish and have observed both the diffuse accumulation in the extracellular spaces and the cellular accumulation within cell membranes and have included this data as a supplement (Figure 2—figure supplement 1). In addition, we have also performed these tracer leakage assays in nuclear labelled transgenic fish and have observed co-localization between the injected Dextran and the nuclear signal, further supporting that the injected tracers can be taken up by “scavenger” parenchymal cells (Figure 2—figure supplement 1).

Third. The authors propose a model in which unrestricted transcytosis triggers diffuse tracer accumulation, a process negatively regulated by mfsd2aa and quantified in WT animals in Figure 2E. Surprisingly, in mfsd2aa mutants, quantification of this mode of leakage was not performed. This is essential to support the model and should be performed.

We thank the reviewer for this comment and wholeheartedly agree that this is a crucial experiment to support the notion of increased transcytosis altering BBB leakage. We have performed the suggested time lapse leakage assays in mfsd2aa mutant and heterozygote control fish to observe the dynamics of leakage in a system with BBB-specific increased transcytosis. We observed similar diffuse leakage into the brain parenchyma in mfsd2aa mutants like we did at 3 dpf (Figure 6), both with similar overall levels and rates of leakage into the brain parenchyma.

Fourth. Instead, "parenchymal cells" were monitored (see comment #2). We were not convinced that monitoring the number of "cells" is predictive to leakage extent. It would only be so if (i) the total number of cells is high, stable during development and unaltered in genetic mutants and if (ii) the amount of substrate tracer is limiting so that the more tracer reaches the parenchyma, the more cells are detectable. None of these conditions seem established in light of the data presented. For instance, as the nature of the presumed cells is unknown, their total number (labeled or not) cannot be established and therefore different time points and genetic conditions cannot be compared (Figure 1D, Figure 1—figure supplement 1, Figure 4C, Figure 4—figure supplement 2, 3). How can the authors rule out that the total number of this presumptive cell type is not merely increased in mfsd2aa mutants? According to previous reports mfsd2aa is widely expressed throughout the zebrafish embryonic and larval brain. Similarly, how can the authors exclude that the number of these cells drops over time in the midbrain? Instead of counting "cells", the mean intensity of the "cells" could be analyzed (alternatively, asses the parenchymal leak, see third comment). If transcytosis (or more broadly tracer leakage) is increased, the anticipated result is an increase in the average fluorescence intensity of cell-associated label (rather than an increase in the actual number of cells).

Assessing the number of cells as a readout for transcytosis seems moreover also puzzling from cellular standpoints. Indeed, at least some of these "cells" were shown to be filled secondary to sudden rupture of endothelial integrity (Figure 2E) and therefore would not be a faithful readout for transcytosis upregulation. Final sentence in subsection “Posterior-Anterior Gradient of Zebrafish BBB Development” are unclear in that respect as well. We would agree with the authors that tight junctional defects would likely exhibit size-selective leakage properties (although that remains to be shown in zebrafish). Consequently, the observed labelling of the "parenchymal cells" with tracers of all tested sizes suggest an alternative leakage mode is at play (if we assume this is secondary to BBB defects, see above). It does however not discriminate vessel rupture from transcytosis. So in absence of vessel rupture quantification in mfsd2aa mutants versus WT, interpretation of the cell counts is ambiguous.

We truly thank this reviewer (and reviewer 2) for this comment and we have now analyzed the data in an alternative way as shown in Figure 1 and described in the Materials and methods section. We now measure the overall tracer intensity in the brain parenchyma normalized to tracer intensity within circulation, as we had previously done for the time lapse imaging, to quantify leakage across developmental stages and mutant lines. Interestingly, this analysis revealed high tracer permeability throughout the larval brain at 3 dpf, whereas our previous cell counting method showed the hindbrain barrier had been sealed at this time point based the lack of “scavenger” cells in the hindbrain. This difference could be due to the sensitivity of two quantification methods or the lack of these particular scavenger cells in the hindbrain. However, this has not altered our overall assessment of a posterior to anterior gradient of barrier sealing, as at 4 dpf, we observed a sealed hindbrain barrier but a leaky midbrain barrier and at 5 dpf, everything appeared sealed. The combination of both methods for leakage quantification strengthened this study.

Reviewer #2:

1) This reviewer believes that the title of this manuscript overstates the conclusions of the study. To conclude that BBB development is regulated by suppression of transcytosis, implies that other characteristics of BBB development are also controlled by this cellular process. As BBB development requires the acquisition of many properties (e.g. formation of tight junctions, expression of multiple types of transporters, establishment of cellular interactions, etc), including the suppression of transcytosis as described here, please consider a title that more accurately reflects the results of the study.

We thank the reviewer for pointing out the overgeneralization of the term BBB development, and have amended the title to reflect more specifically what we have measured: “Suppression of transcytosis regulates zebrafish blood-brain barrier function”.

2) The conclusions of the manuscript rely significantly upon the spatiotemporal expression of mfsd2aa (and mfsd2ab to a lesser extent). Based upon the results of this study, it would be predicted that mfsd2aa is not expressed in brain endothelial cells at 3 dpf, which allows for transcytosis of tracers into the brain parenchyma. Furthermore, it would be predicted that mfsd2aa is expressed in brain endothelial cells by 5 dpf, which then suppresses transcytosis of tracers into the brain parenchyma. However, the authors do not provide any expression data for either mfsd2aa or mfsd2ab.

In previous studies, Guemez-Gamboa et al. used whole-mount in situ hybridization to show that both zebrafish mfsd2aa or mfsd2ab transcripts are expressed throughout the nervous system in zebrafish at 24, 48, and 96 hpf. However, it is not clear from this data whether brain endothelial cells express either transcript. In addition, Thisse and Thisse showed basal expression of mfsd2aa in spinal cord neurons as early as 20-25 somites to Prim-5 stage, no spatial restriction at the Prim-15 to Prim-25 and High-pec to Long-pec stages, and neurocranium expression at 5 dpf (https://zfin.org/ZDB-GENE-041114-166/expression). Thus, it remains to be determined whether zebrafish mfsd2aa or mfsd2ab is expressed in brain endothelial cells.

Please provide expression data that demonstrates mfsd2aa expression (or lack of expression) in zebrafish brain endothelial cells at the appropriate developmental time points (i.e. 3 dpf, 5 dpf, and adult). Perhaps another valid strategy would be to rescue mfsd2aa mutants by cell autonomous expression of wild type mfsd2aa in endothelial cells.

Guemez-Gamboa et al. (2015) Nature Genetics 47:809-813

Thisse and Thisse, (2004) ZFIN Direct Data Submission (http://zfin.org).

To address this reviewer and reviewer 1’s concern, we have performed fluorescent in situ hybridization (FISH) on sections of 3 and 5 dpf larvae and adult brains from kdrl:mCherry transgenic fish, which fluorescently labels endothelial cells. These FISH experiments have shown that both mfsd2aa and mfsd2ab are most highly expressed in brain endothelial cells at 5 dpf, While mfsd2aa was not expressed above background levels at 3 dpf, mfsd2ab was highly expressed at both 3 and 5 dpf (Figure 4). Both paralogues are expressed in the adult brain tissue with higher levels of mfsd2aa than mfsd2ab. In addition, previously published RNA-seq from FAC-sorted endothelial cells at 48 hpf similarly revealed barely detectable expression of mfsd2aa and high levels of mfsd2ab (Bower et al., Nature Neuroscience, 2014). Taken together, these FISH experiments have nicely supported our hypothesis that the suppression of transcytosis coincides with the onset of mfsd2aa expression in the vasculature.

3) To quantify "leakiness", this study focused on the number of parenchymal cells that accumulated tracer in the midbrain and demonstrated that larvae at 3 dpf are leakier than 5 dpf. The authors also indicated that parenchymal cells in the hindbrain did not accumulate tracer at a significant level at either developmental time point. However, in the mfsd2aa mutants, the authors did not examine midbrain leakiness at 3 dpf or in the hindbrain at either 3 dpf or 5 dpf. Do the mfsd2aa mutants at 3 dpf exhibit increased leakiness compared to wild types at 3 dpf? Also, if mfsd2aa suppresses transcytosis in all brain endothelial cells, then is the hindbrain leakier in mfsd2aa mutants at 3 dpf and 5 dpf? If not, please explain.

As stated above, we have altered our method of quantifying leakage throughout the manuscript in response to multiple reviewers’ suggestions. This quantification method has been widely used in mice (Nitta et al., 2003; Bell et al., 2010; Daneman et al., 2010; Andreone et al., 2017; Chow et al., 2017). With this method of normalized brain parenchyma tracer intensity, we have now quantified leakage in mfsd2aa and mfsd2ab mutants both in the midbrain and hindbrain at 5 dpf to address potential regional differences, or lack thereof. This data is now included in Figure 5. When we examined DBP-EGFP tracer permeability in mfsd2aa mutant and wild-type fish at 3 dpf, we did not observe an increase in BBB permeability in mfsd2aa mutants compared to the already elevated levels in wild-type fish (data not shown), as expected from the lack of mfsd2aa expression at this time (Figure 4).

Reviewer #3:

[…] There are some minor experimental additions that could be performed to strengthen the study's conclusions and impact.

1) Given that antibodies to mark tight junctions exist that work in zebrafish, it would be useful to add this analysis to the time course of BBB formation.

We thank the reviewer for her kind remarks. We feel that this information is already available in the Jeong et al., 2008 and the Xie et al., 2010 papers, and therefore do not believe it needs to be repeated. However, we have further highlighted this fact in the Results section.

2) Given the controversy surrounding the timing of BBB closure in zebrafish, it is important to extend the time course to 10 days to be consistent with the Fleming et al. study. Additionally, the hindbrain at 3 days shows ~2 cells/embryo taking up tracer whereas the authors consider the midbrain BBB closed at 5-6 days with ~8 cells/larva taking up tracer. Thus, it is important to test if the number of cells taking up tracer further reduces with time.

We have now repeated the leakage assays at 10 dpf and included that data in Figure 1. Additionally, we have altered our metric of quantifying leakage to be less restricted to the number of parenchymal cells and are measuring tracer intensity in the brain parenchyma instead. This new quantification has been applied to the 10 dpf data.

3) The standard in the field is to rescue a new mutant with wild-type gene product to control for off-target CRISPR/Cas9 effects. Similarly, qPCR for mfsd2aa and mfsd2ab are important controls for the new mutants, especially since phenotypes were not observed in mfsd2ab mutants.

Given the late phenotype (5 dpf) we do not believe that the use of RNA to rescue the mfsd2aa leakage will work. While we have not rescued the phenotype, we have generated multiple F0 crispants for both mfsd2aa and mfsd2ab using new guides to target each gene, and observed similar leakage phenotypes in mfsd2aacrispants and did not observe any leakage phenotypes in mfsd2abcrispants, suggesting that the observed phenotypes are in fact due to the loss of the target gene (Figure 5—figure supplement 2). Both stable mutant lines have also been backcrossed to wild-type fish for at least 5 generations, and the leakage phenotype in mfsd2aa mutants still persists. We have now performed the requested qPCR for mfsd2aa and mfsd2ab in both mutant backgrounds to ensure reduced transcript levels (Figure 4—figure supplement 1).

4) Not performing in vivo imaging of the mfsd2aa mutants seems like a missed opportunity. Such investigation could also reveal if both types of leakage observed in the time course are affected in mfsd2aa mutants. The analysis presented in Figure 2E would also be helpful to show for the mutants.

We completely agree with the reviewer that these are critical experiments, and have now performed time lapse leakage assays in mfsd2aa mutant and heterozygous siblings, revealing the dynamics of increased transcytosis in the BBB (Figure 6).

https://doi.org/10.7554/eLife.47326.036

Article and author information

Author details

  1. Natasha M O'Brown

    Department of Neurobiology, Harvard Medical School, Boston, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0718-7037
  2. Sean G Megason

    Department of Systems Biology, Harvard Medical School, Boston, United States
    Contribution
    Conceptualization, Supervision, Methodology, Writing—original draft, Writing—review and editing
    For correspondence
    Sean_Megason@hms.harvard.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-9330-2934
  3. Chenghua Gu

    Department of Neurobiology, Harvard Medical School, Boston, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Writing—original draft, Project administration, Writing—review and editing
    For correspondence
    Chenghua_Gu@hms.harvard.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4212-7232

Funding

Damon Runyon Cancer Research Foundation (Postdoctoral Research Fellowship)

  • Natasha M O'Brown

Fidelity Biosciences

  • Chenghua Gu

National Institutes of Health (National Institutes of Health Director's Pioneer Award DP1 NS092473)

  • Chenghua Gu

Harvard Medical School (Mahoney Postdoctoral Fellowship)

  • Natasha M O'Brown

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

Acknowledgements

We thank members of the Gu and Megason laboratories for data discussion and comments on the manuscript; Dr. Zach O’Brown for discussions and comments on the manuscript; Dr. Bela Anand-Apte (Cleveland Clinic) for providing the transgenic Tg(l-fabp:DBP-EGFP) fish line (Xie et al., 2010) and Dr. Leonard Zon for providing the transgenic Tg(kdrl:HRAS-mCherry) line; and the HMS Electron Microscopy Core Facility, with special thanks to Louise Trakimas for all of her assistance in troubleshooting and preparing the TEM samples. This work was supported by the Damon Runyon Cancer Research Foundation (NMO), the Mahoney Postdoctoral Fellowship (NMO), the Fidelity Biosciences Research Initiative (CG), and the National Institutes of Health Pioneer Award DP1 NS092473 (CG). The research of CG was also supported in part by a Faculty Scholar grant from the Howard Hughes Medical Institute.

Ethics

Animal experimentation: This study was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the NIH. All animals were handled according to approved institutional animal care and use committee (IACUC) protocols (protocol 04487) of Harvard Medical School. All work was approved by the Harvard Medical Area Standing Committee on Animals.

Senior Editor

  1. Didier Y Stainier, Max Planck Institute for Heart and Lung Research, Germany

Reviewing Editor

  1. Richard Daneman, University of California, San Diego, United States

Reviewers

  1. Benoit Vanhollebeke, Université libre de Bruxelles, Belgium
  2. Michael R Taylor, University of Wisconsin-Madison, United States
  3. Kelly Monk, Vollum Institute, United States

Publication history

  1. Received: April 2, 2019
  2. Accepted: August 19, 2019
  3. Accepted Manuscript published: August 20, 2019 (version 1)
  4. Version of Record published: September 4, 2019 (version 2)

Copyright

© 2019, O'Brown et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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