New Results

Sodium nitroprusside prevents the detrimental effects of glucose on the neurovascular unit and behaviour in zebrafish

K. Chhabria, A. Vouros, C. Gray, R.B. MacDonald, Z. Jiang, R.N. Wilkinson, K Plant, E. Vasilaki, C. Howarth, T.J.A. Chico
doi: https://doi.org/10.1101/576942

Abstract

Diabetes is associated with dysfunction of the neurovascular unit, although the mechanisms of this are incompletely understood, and currently no treatment exists to prevent these negative effects. We previously found that the NO donor sodium nitroprusside (SNP) prevents the detrimental effect of glucose on neurovascular coupling in zebrafish. We therefore sought to establish the wider effects of glucose exposure on both the neurovascular unit and on behaviour in zebrafish and the ability of SNP to prevent these.

We incubated 4 days post fertilisation (dpf) zebrafish embryos in 20mM glucose or mannitol for five days until 9dpf, with or without 0.1mM SNP co-treatment for 24h (8-9dpf), and quantified vascular nitric oxide reactivity, vascular mural cell number, expression of a klf2a reporter, glial fibrillary acidic protein (GFAP) and TRPV4, as well as spontaneous neuronal activation at 9dpf, all in the optic tectum. We also assessed the effect on light/dark preference and locomotory characteristics during free-swimming studies.

We find that glucose exposure significantly reduced nitric oxide reactivity, klf2a reporter expression, vascular mural cell number and TRPV4 expression, while significantly increasing spontaneous neuronal activation and GFAP expression (all in the optic tectum). Furthermore, when we examined larval behaviour we found glucose exposure significantly altered light/dark preference and high and low speed locomotion while in light. Co-treatment with SNP reversed all these molecular and behavioural effects of glucose exposure.

Our findings comprehensively describe the negative effects of glucose exposure on the vascular anatomy, molecular phenotype, and function of the optic tectum and on whole organism behaviour. We also show that SNP or other NO donors may represent a therapeutic strategy to ameliorate the complications of diabetes on the neurovascular unit.

Introduction

The prevalence of diabetes has quadrupled in the previous two decades, incurring an enormous burden of morbidity and healthcare expenditure worldwide (Zhang et al., 2010; Scully, 2012). Diabetes is a risk factor for both macrovascular (such as myocardial infarction and stroke) and microvascular (causing renal impairment and retinopathy) disease (Chase et al., 1989; Jorgensen et al., 1994; Miettinen et al., 1998; Stratton et al., 2000). Diabetes is also associated with neurological disorders including cognitive impairment (dementia) (Stewart & Liolitsa, 1999; Areosa & Grimley, 2002; MacKnight et al., 2002; Ciudin et al., 2017; Simo et al., 2017; Groeneveld et al., 2018). The mechanisms underlying this association are incompletely understood, and no specific therapies have been identified to prevent or reverse the effects of diabetes on neurological function.

Both diabetes and neurological diseases are associated with dysfunction of the neurovascular unit (NVU) (Zlokovic, 2010; Mogi & Horiuchi, 2011; Gardner & Davila, 2017). The NVU comprises neurons, astrocytes, myocytes, pericytes, endothelial cells (ECs) and extracellular matrix. Interactions between these ensures neuronal energy demands are met through increased local blood flow via neurovascular coupling (NVC) (Roy & Sherrington, 1890; Attwell et al., 2010). Recent evidence suggests that ECs are crucial to NVU function (Toth et al., 2015; Guerra et al., 2018) as they release vasoactive substances such as nitric oxide (NO) (Ignarro et al., 1987; Palmer et al., 1987). NO production is regulated by various endothelial genes including the Kruppel-like family of transcription factors (KLFs)(Gracia-Sancho et al., 2011) particularly KLF2 which is regulated by changes in flow and inflammation (Dekker et al., 2002; SenBanerjee et al., 2004b). ECs share a common basement membrane with pericytes that aid EC development (Gerhardt & Betsholtz, 2003). Pericyte coverage, essential to both blood brain barrier integrity and NVC, is affected in various neuropathologies (Tilton et al., 1985; Frank et al., 1990; Peppiatt et al., 2006; Pfister et al., 2008; Vates et al., 2010; Sagare et al., 2013; Hall et al., 2014). Astrocytes are the predominant glial cell in the brain and perform several functions including release of vasoactive factors (Zonta et al., 2003; Takano et al., 2006). Astrocytes also express glutamine synthetase (GS), an enzyme involved in the recycling of glutamate released by active neurons (Bergles et al., 1999; Bringmann et al., 2013). Mammalian studies show that astrocytes sense changes in vascular tone through activation of the mechanosensor TRPV4 (Vallinoid transient receptor potential) (Filosa et al., 2013), which is also expressed in ECs. Together all these cell types maintain the functional NVU.

The zebrafish is increasingly used as a model of human disease (Dooley & Zon, 2000; Lieschke & Currie, 2007). This has a number of advantages over existing mammalian models, particularly ease of in vivo cellular imaging and the ability to test the effect of drugs by immersion. Although most zebrafish studies attempting to model human disease have examined the anatomic or molecular effects of genetic or other manipulation (Patton & Zon, 2001; Lieschke & Currie, 2007), a range of behavioural assays have been applied to study more complex “whole organism” phenotypes, such as memory or aggression (Blaser & Gerlai, 2006; Oliveira et al., 2011). The zebrafish has previously been used as a model of diabetes, either by exposure to medium containing glucose (Capiotti et al., 2014), genetic manipulation (Kimmel et al., 2015) or ablation of the beta-cells of the pancreas (Pisharath et al., 2007).

We recently established a novel zebrafish larval model of NVC in which incubation of larvae in 20mM glucose impaired NVC. We found the NO donor sodium nitroprusside (SNP) rescued this effect (Chhabria et al., 2018) although the mechanism for this is unclear. We therefore wish to better understand the mechanism and consequences of NVU dysfunction induced by glucose. In the present study we have now examined the effect of glucose exposure on NO production in the NVU, klf2a expression, mural cell number, glial fibrillary acidic protein (GFAP) expression, TRPV4 expression, spontaneous neuronal activation, light/dark preference and larval locomotory behaviour. We find that glucose exposure affects all these aspects of NVU function and behaviour, and that co-treatment with SNP completely prevents all the detrimental effects of glucose exposure. Our findings provide insight into the effect of hyperglycaemia on NVU function and further support for the possibility that NO donors represent plausible drug candidates to ameliorate the detrimental effects of hyperglycemia.

Materials and Methods

Transgenic zebrafish

All zebrafish studies were conducted in accordance with the Animals (Scientific Procedures) Act, 1986, United Kingdom and covered by Home Office Project Licence 70/8588 held by TC. Reporting of experimental outcomes were in compliance with ARRIVE (Animal Research: Reporting in Vivo Experiments) guidelines (Kilkenny et al., 2010).

Maintenance of adult zebrafish was conducted according to previously described husbandry standard protocols at 28°C with a 14:10 hours (h) light:dark cycle (Lawrence, 2007). The following zebrafish lines were used; Tg(kdrl:HRAS-mCherry)s916 labelling EC membrane (Hogan et al., 2009), Tg(klf2a:GFP) expressing GFP under control of the zebrafish klf2a promoter, Tg(sm22ab:nls-mcherry)sh480 labelling mural cells expressing a smooth muscle actin binding protein and Tg (nbt:GCaMP3) which allows quantification of neuronal calcium levels (Meza Santoscoy, 2014; Bergmann et al., 2018).

Glucose, mannitol and SNP treatment

Glucose, mannitol and SNP (Sigma) were prepared in E3 medium to final concentrations of 20mM (glucose and mannitol) or 0.1mM SNP (Chhabria et al., 2018). Zebrafish larvae for in vivo imaging or immunostaining were incubated in E3 medium containing glucose/mannitol from 4-9 days post fertilization (dpf) and SNP from 8-9 dpf (Chhabria et al., 2018).

Assessment of NO reactivity in the cerebral vessels

Larvae exposed to glucose/mannitol with/without SNP as above were incubated with 2.5 µM Diaminofluorescein-FM (DAF-FM) in DMSO (1%) at 9 dpf for 3 hours at 28°C in the incubator in the dark. Larvae were washed with glucose/mannitol solution to remove excess DAF-FM, and then imaged on the lightsheet microscope.

Glutamine synthetase, GFAP and TRPV4 Immunohistochemistry

Immunohistochemistry (IHC) was used to observe and quantify changes in TRPV4 (AB2241068; ThermoFisher) and the glial specific markers, GS (mab302; Millipore) and GFAP (zrf-1; Zebrafish International Resource Center, Oregon). The protocol was adapted from (Inoue and Wittbrodt 2011). Larvae for each different treatment (mannitol or glucose, with or without co-treatment with SNP) were fixed in 4% Paraformaldehyde (PFA) overnight at 4°C followed by a 5min wash with 1x PBS before resuspending in 100% Methanol (MeOH) for storage at −20°C. Samples were rehydrated from MeOH with 3x 10min washes with phosphate buffer saline + 0.1% tween (PBST), with gentle agitation on a rocker. Larvae were then suspended in 150mM Tris-HCl (pH 9) for 5 minutes, followed by heating at 70°C for 15 minutes. Larvae then underwent 2x 10min washes with PBST, followed by 5min washes with distilled water (dH2O). Larvae were permeabilized using ice cold acetone at −20°C for 20 minutes, followed by 2x 5min washes with dH2O, then equilibrated 2x 5min washes in PBST. Subsequently, larvae were incubated in blocking buffer (B-buffer: 10% sheep serum, 0.8% triton X100 and 1% bovine serum albumin in PBST) for 3 hours at 4°C. B-buffer was replaced with incubation buffer (I-buffer: 1% sheep serum, 0.8% triton X100 and 1% bovine serum albumin in PBST) containing the primary (1°) antibodies (Abs): 1:250 GS (mouse monoclonal), 1:100 GFAP (mouse monoclonal), 1:300 TRPV4 (rabbit polyclonal) and 1:1000 DAPI followed by incubation at 4°C for 3 days with gentle agitation on a rocker.

Residual 1° Abs were removed by 3x hourly washes in PBST (at room temperature (RT)). Larvae then underwent 2x 10 min washes with PBS + 1% triton X100, followed by 2x hourly equilibration washes in PBS-TS (PBS + 1% triton X100+10% sheep serum).

Larvae were incubated in I-buffer containing secondary antibodies (2° Abs): anti-rabbit, 488 nm (A28181, Invitrogen), anti-mouse, 647 nm (A-11011, Invitrogen) and anti-mouse, 561 nm (A-11034, Invitrogen), corresponding to respective 1° Abs, at 1:500 dilutions for 2.5 days, in the dark on a rocker at 4°C.

Prior to imaging on the lightsheet microscope, larvae were washed three times in PBS-TS (at RT), followed by two 1 hour washes with PBST, each at RT. Larvae were mounted in 1% low melting point agarose (LMP) (Sigma) and imaged for the glial patterning in the brain for different markers.

Lightsheet fluorescent imaging

Lightsheet fluorescent microscopy (LSFM) was performed on 9 dpf larvae on a Zeiss Lightsheet Z.1 microscope. Larvae were minimally anesthetised (using 4.2% v/v tricaine methanesulfonate) and embedded in 1% agarose in a glass capillary (inner diameter ∼ 1mm) while imaging. We acquired 3D z stacks with 800 x 600 pixels (1 pixel = 0.6 µm) in X-Y direction and a depth of 100 slices in Z directions (slice thickness = 1 µm).

For imaging spontaneous neuronal activity, we acquired time lapses of single ‘z’ plane optic tectum with our previous acquisition settings and frequency quantifications (Chhabria et al., 2018).

Image analysis: klf2a quantification

Acquired 3D image stacks were converted to 2D maximum intensity projections and the tectal vasculature was segmented out. Tectal vascular length was extracted (Chhabria et al., 2018). The segmented vasculature was used as a binary mask, followed by normalizing the total intensity of the green channel (klf2a:GFP) in the optic tectum to the tectal vascular length.

Image analysis: quantification of vascular mural cells

Acquired 3D image stacks were converted to 2D maximum intensity projections followed by segmenting the sm22absh480 nuclei (red channel) using intensity based thresholding similar to the method of RBC segmentation described in (Chhabria et al., 2018). Segmented cells in the optic tectum were enumerated (in a fixed vascular volume) using custom written MATLAB scripts used for all treatment groups.

Behavioural analysis

Light/Dark preference

Analysis of light/dark preference of the larval zebrafish was designed on a similar principle to that of the adult zebrafish light/dark preference test (Blaser and Penalosa 2011). A 12-well plate was modified by adhering three cellophane films (blue, green and yellow) to half of each well to create a ‘dark’ side that allowed the camera to track larvae movement by infrared (IR). Larvae from different treatment groups were placed on the light side of the well filled with 5 ml E3 (without methylene blue).

The plate was placed inside a Viewpoint-Zebrabox system (1% light intensity) and the tracking protocol was built allocating dark and light regions of each well prior to the start of imaging (to get x,y coordinates of dark/light regions separately for analysis). Speed thresholds were set as high speed > 6.4mm/s, low speed 3.3-6.3mm/s, and inactive < 3.3mm/s. Total experimental duration was one hour, inclusive of acclimatization (recording started immediately post adding larvae to individual well).

Automated locomotion analysis

Using previous rodent based methods developed for Morris water maze (Wolfer & Lipp, 2000; Graziano et al., 2003; Gehring et al., 2015; Illouz et al., 2016; Vouros et al., 2018), we quantified four different features from the swimming trajectories of zebrafish. Coordinates of the swimming trajectories were extracted from the Viewpoint-Zebrabox system and were segmented into smaller paths delimited by light/dark, followed by quantification of path features listed in Figure 1. Small path segments, with lengths lower than 1st percentile of segments, generated as an artefact of light/dark transitions were removed from further analysis. The path features eccentricity (ε) and mean point distance from ellipsoid (MPDE) quantify the spatial elongation of the locomotion trajectories and are used as measures of exploration in the field while mean point distance from centre (MPDC) of the well, represents thigmotaxis (preference of edge vs. centre of well). Additionally, we quantified the number of transitions between light and dark areas (Figure 1).

Figure 1:
Figure 1: Mathematical description of features calculated from the larval trajectories.

A: Schematic diagram of the well showing parameters calculated for various features. B: Table describing the formulae of calculated features for each coordinates of the trajectories. The minimum enclosing ellipsoid with centre (xe, ye) and major and minor axes of a and b is defined as the unique closed ellipse of smallest volume which enclose all points (xi, yi) of a path. Embedded Image is the Euclidean distance of every point of the path to centre of the well, (xa, ya). Embedded Image is the distance between the centre of minimum enclosing ellipsoid (xe, ye) and ith point of the path. For each feature, two distinctive numeric examples are provided.

Experimental design and statistical analysis

Experiments were designed using the NC3Rs experimental design analysis (EDA) tool. GraphPad Prism (La Jolla, CA®) was used for statistical comparisons. All intergroup comparisons were performed using two-way ANOVA with posthoc multiple comparison tests (Sidak’s test) where appropriate. P values <0.05 were considered to be statistically significant. Data are shown as mean ± standard error of mean (s.e.m.) unless specified.

Results

Glucose exposure reduces vascular nitric oxide reactivity which is prevented by co-treatment with SNP

Studies with diabetic patients have shown reduced bioavailability of NO in the ECs (Williams et al., 1996; Pieper, 1998). We therefore first examined whether glucose exposure reduces NO availability in the cerebral vessels. We used DAF-FM staining to visualise NO reactivity in 9dpf zebrafish embryos exposed to 20mM Glucose or mannitol (as osmotic control) with or without co-treatment with 0.1mM SNP. Representative micrographs of the vessels in the left optic tectum are shown in Figure 2A. We observed variable levels of NO reactivity that co-localised with the kdrl:HRAS-mcherry endothelial reporter in animals treated with mannitol. Co-treatment with mannitol and the NO donor, SNP did not alter the intensity of vascular NO reactivity compared to treatment with mannitol alone (Figure 2). Exposure to glucose significantly reduced vascular NO reactivity in the tectal vessels (Figure 2), in keeping with data from other models (Pieper, 1998; Du et al., 2001). Co-treatment with SNP prevented this reduction in vascular NO reactivity (Figure 2).

Figure 2:
Figure 2: Effect of mannitol/glucose treatment with/without SNP on NO reactivity, quantified by intensity of DAF-FM staining.

A: Representative micrographs of tectal vessels showing separate and merged channels (green: DAF-FM staining, red: kdrl:HRAS-mcherry) for 20mM mannitol or glucose exposed larvae co-treated with or without SNP. Scale bar represents 20 µm. B: Quantification of DAF-FM intensity in the tectal vessels (n=25, 24, 27 and 24 larvae for mannitol, mannitol + SNP, glucose and glucose + SNP, respectively). Data are mean ± s.e.m. *p<0.05.

Glucose exposure reduces endothelial klf2a which is prevented by SNP co-treatment

The shear-stress responsive transcription factor klf2a is protective against vascular disease (Dekker et al., 2002; Dekker et al., 2005; Chiu et al., 2009). To determine whether glucose exposure affects endothelial shear stress sensing and klf2a expression, we quantified the intensity of a Tg(klf2a:GFP) reporter in the cerebral vessels of zebrafish with glucose or mannitol with or without SNP co-treatment. Representative micrographs are shown in Figure 3A. Glucose exposure significantly reduced intensity of the klf2a:GFP reporter compared to mannitol alone (Figure 3B). Although SNP co-treatment with mannitol had no effect compared with mannitol alone (Figure 3), SNP co-treatment prevented the glucose-induced reduction in the klf2a reporter expression.

Figure 3:
Figure 3: Effect of mannitol/glucose treatment with/without SNP on expression of klf2a:GFP expression.

A: Representative micrographs of tectal vessels in Tg(klf2a:GFP) exposed to 20mM mannitol or glucose co-treated with or without SNP. Scale bar represents 20 µm. B: Quantification of the klf2a:GFP intensity in the tectal vessels (n=26 larvae/group). Data are mean ± s.e.m. *p<0.05.

Glucose exposure reduces the number of vascular mural cells on the tectal vessels which is prevented by SNP co-treatment

NO is required for mural cell function and can evoke hyperpolarization in mural cells (including pericyte and smooth muscle cell) causing vasodilation (Sakagami et al., 2001; Lee et al., 2005). Mural cell loss is a feature of diabetes (Pfister et al., 2008) but no therapy has been shown to reverse this. We therefore examined the effect of glucose on vascular mural cells. We used a sm22ab:nls-mCherry reporter to quantify the number of vascular mural cells present in the optic tectum. Representative micrographs are shown in Figure 4A. Glucose exposure induced a significant reduction in the number of vascular mural cells compared with either mannitol or mannitol plus SNP (Figure 4B). Co-treatment with SNP prevented the reduction of vascular mural cells induced by glucose exposure.

Figure 4:
Figure 4: Effect of mannitol/glucose treatment with/without SNP on mural cell number on the tectal vessels.

A: Representative micrographs of tectal vessels showing separate and merged channels (green: fli1:GFF:UAS:GCaMP6, red: sm222ab:nls-mcherry sh480) for 20mM mannitol or glucose exposed larvae co-treated with or without SNP. Scale bar represents 20 µm. White arrows indicate mural cell nuclei. B: Quantification of the number of sm22ab:nls-mcherrysh480 nuclei on the tectal vessels for 20mM mannitol or glucose exposed larvae co-treated with or without SNP (n=28 larvae/group). Data is mean ± s.e.m. *p<0.05, **p<0.01.

Glucose exposure increases GFAP expression in the optic tectum which is prevented by SNP co-treatment

Glial cells play major roles in maintenance of the blood brain barrier and NVU function (Janzer & Raff, 1987; Prat et al., 2001). Experimental studies have shown over-expression of GFAP (termed astrogliosis) in response to both hyperglycemia and type 1 diabetes (Coleman et al., 2004). We thus examined the effect of glucose or mannitol with or without SNP on GFAP expression. Representative micrographs of whole mount 9 dpf old zebrafish are shown in Figure 5A. Glucose exposure increased GFAP expression compared to mannitol treatment (Figure 5B), in keeping with astrogliosis in other diabetic models. This was prevented by co-treatment with SNP (Figure 5B).

Figure 5:
Figure 5: Effect of mannitol/glucose treatment with/without SNP on GFAP expression.

A: Representative micrographs showing the effect of mannitol/glucose exposure with/without SNP treatment on GFAP expression (Red channel represents GFAP and blue channel represents DAPI). Scale bar represents 20 µm. B: Quantification of GFAP expression in the optic tectum (n=16, 12, 18 and 20 larvae for larvae for mannitol, mannitol + SNP, glucose and glucose + SNP, respectively). Data are mean ± s.e.m. *p<0.05, ***p<0.001.

Glucose exposure reduces the expression of TRPV4 in the optic tectum which is prevented by SNP co-treatment

Since hyperglycemia downregulates TRPV4 in the ECs of the retinal microvasculature (Monaghan et al., 2015), we investigated TRPV4 expression by immunohistochemistry in 9 dpf old zebrafish larvae exposed to glucose or mannitol with or without SNP treatment. We performed immunohistochemistry for TRPV4 to first compare expression patterns in the optic tectum. Representative micrographs are shown in Figure 6. Glucose exposure decreased tectal TRPV4 expression (which includes radial glial, endothelial or neuronal expression of TRPV4) compared to mannitol. Co-treatment with SNP prevented the effect of glucose on TRPV4 expression (Figure 6).

Figure 6:
Figure 6: Effect of mannitol/glucose treatment ± SNP on TRPV4.

A: Representative micrographs of optic tectum showing the effect of mannitol/glucose treatment with/without SNP treatment on the expression of TRPV4. Scale bar represents 20 µm. B: Quantification of the TRPV4 intensity in the optic tectum in a fixed volume of the tissue (n=21, 20, 24 and 26 larvae for larvae for mannitol, mannitol + SNP, glucose and glucose + SNP, respectively). Data are mean ± s.e.m. **p<0.01, ****p<0.0001.

Glucose exposure reduces expression of glutamine synthetase in the optic tectum which is prevented by SNP co-treatment

Glutamine synthetase, a glial-specific enzyme involved in recycling of extracellular/extrasynaptic glutamate to glutamine, is reduced in neurological disorders and diabetes (Lieth et al., 2000; Burbaeva et al., 2003). We therefore examined expression of glutamine synthetase in the optic tectum. Glucose exposure induced a significant reduction in glutamine synthetase expression, which was prevented by SNP co-treatment (Figure 7).

Figure 7:
Figure 7: Effect of mannitol/glucose treatment with/without SNP on glutamine synthetase.

A: Representative micrographs of optic tectum showing the effect of mannitol/glucose treatment with/without SNP treatment on the expression of glutamine synthetase. Scale bar represents 20 µm. B: Quantification of the glutamine synthetase intensity in the optic tectum in a fixed volume of tissue (n=21, 20, 24 and 26 larvae for larvae for mannitol, mannitol + SNP, glucose and glucose + SNP, respectively). Data are mean ± s.e.m. *p<0.05, **p<0.01.

Glucose exposure increases frequency of spontaneous neuronal calcium transients which is prevented by SNP co-treatment

The previous results showed that glucose exposure affects both the anatomy of the NVU (vascular mural cell loss) and induces dysregulation of gene expression, including a reduction of glutamine synthetase which might be expected to cause neuronal hyperexcitability. We therefore quantified neuronal activity in our model. Representative time series of spontaneous neuronal activation (quantified as ΔF/Fo) in larvae exposed to mannitol or glucose with or without SNP are shown in Figure 8A. Glucose exposure increased neuronal activation compared to mannitol exposure (Figure 8B). Co-treatment with SNP prevented this glucose-induced increase in neuronal activation (Figure 8B), suggesting that diabetes-induced neuronal hyperexcitability may be mediated via reduced NO bioavailability.

Figure 8:
Figure 8: Effect of mannitol/glucose treatment with/without SNP on frequency of spontaneous neuronal calcium transients.

A: Time series of neuronal activation (ΔF/Fo in zebrafish (5 larvae/group) exposed to (left:top-to-bottom); mannitol, mannitol+SNP, glucose and glucose+SNP. Red arrowheads indicate the detected peaks. B: Quantification of frequency of neuronal calcium transients for each of the groups (n=28 larvae/group). Data are mean ± s.d. in A and mean ± s.e.m in B. *p<0.05

Glucose exposure alters light/dark preference and locomotion which is prevented by SNP co-treatment

Although the previous data clearly demonstrate molecular and functional defects of the NVU induced by glucose, if such disturbances are clinically relevant, they would be expected to manifest in overt behavioural or neurological consequence. We therefore assessed the effect of glucose exposure on free swimming behaviour in zebrafish larvae to attempt to identify behavioural consequences of glucose exposure. We first examined the effect of glucose exposure on light-dark preference. Figure 9A shows the trajectories of representative zebrafish larva for a period of 1 h in each treatment group. Red and green paths indicate high and low speed locomotion respectively. We tested light/dark preference using percentage of time spent in the light or dark side of the well. Mannitol exposed larvae at 9 dpf showed a preference for light, spending ∼80% time in the light. This was significantly reduced by glucose exposure (Figure 9B). However, co-treatment with SNP prevented the effect of glucose on light/dark preference (Figure 9B).

Figure 9:
Figure 9: Effect of mannitol/glucose treatment with/without SNP on larval zebrafish behaviour.

A: Representative trajectories of 9 dpf old zebrafish moving in half darkened wells (of a 12 well plate) as tracked by Viewpoint software for mannitol or glucose with or without SNP treatment. Red trajectories represent high speed (> 6.4mm/s), green low speed (3.3-6.3 mm/s), and black inactive (< 3.3mm/s) B: Percentage of time spent in light region of the well by larvae (n = 50, 45, 44 and 56 larvae for mannitol, mannitol+SNP, glucose and glucose+SNP, respectively). C: Quantification of percentage time spent in low and high speed locomotion in the light region by the same animals as in B. D: Quantification of percentage time spent in low and high speed locomotion in the dark region by the same animals as B-C. E: Quantification of number of transitions into the light/dark regions for same larvae as in B-D. Data are mean ± s.e.m. *p<0.05, **p<0.01 and ****p<0.0001.

To further characterise larval behaviour, we quantified the time spent in high speed (> 6.4mm/s), low speed (3.3-6.3 mm/s), and inactive < 3.3mm/s locomotion in the light (Figure 9C) or dark side of the wells (Figure 9D). Glucose exposure significantly increased the time larvae spent in both low and high speed locomotion while in the light and SNP prevented this effect (Figure 9C). In contrast, glucose did not induce any significant difference in the amount of time spent in either high or low speed locomotion while in the dark side of the well (Figure 9D). We quantified the number of transitions between light and dark areas of the well and found that glucose exposure induced a significant increase in the number of transitions that was prevented by co-treatment with SNP (Figure 9E).

We next investigated three further features of larval zebrafish locomotion; path eccentricity, mean point distance to ellipsoid (MPDE), and mean point distance to centre (MPDC) (Figure 1) in the light and dark sides of the wells. Glucose induced significant increases in path eccentricities, MPDE and MPDC in the light side of the well which was prevented by SNP (Figure 10A-C). When we quantified these behaviours in the dark side of the well, we saw a similar significant increase in path eccentricity and MPDE but not in MPDC (Figure 10D-F). Again, SNP prevented the effect of glucose on these aspects of behaviour.

Figure 10:
Figure 10: Effect of mannitol/glucose treatment with/without SNP on various features of zebrafish locomotion.

Quantification of mean frequency of path eccentricities (A, D), MPDE (B, E) and MPDC (C, F) (n= 50, 45, 44 and 56 larvae for mannitol, mannitol+SNP, glucose and glucose+SNP, respectively) in A-C: light and D-F: dark regions of the well. Data are mean ± s.e.m. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001

Discussion

We here describe a comprehensive molecular, anatomic, functional, and behavioural study of the effect of glucose exposure on the NVU in zebrafish. The zebrafish model has a range of advantages, coupling simplicity, speed, and cost-effectiveness with sophisticated in vivo imaging in a whole-organism setting. We show that a relatively short (5d) exposure to 20mM glucose (a concentration seen in the blood of poorly controlled human diabetics (Amiel et al., 1988; Boyle et al., 1988)) affects every constituent cell type of the NVU that we examined. Glucose exposure impaired both vascular NO production and vascular mural cell number, which builds on our previous work showing that tectal endothelial patterning is impaired by glucose exposure (Chhabria et al., 2018). These effects were also associated with reduced endothelial klf2a expression, which is known to promote vascular inflammatory gene expression, thrombosis, and atherosclerosis (SenBanerjee et al., 2004a; Lin et al., 2005). Developmental studies in klf2a(-/-) mice have shown reduced vascular mural cell recruitment suggesting the role of klf2a in maintaining endothelial-mural cell interactions (Wu et al., 2008; Fukuhara et al., 2009; Gaengel et al., 2009). Abnormal mural cell recruitment or migration is associated with various microangiopathies and is commonly observed in diabetes (Hammes et al., 2002; Gaengel et al., 2009).

In addition to the negative effects of glucose on the vascular component of the NVU, we found that glucose exposure induced upregulation of GFAP, indicating astrogliosis, with concomitant reductions in glutamine synthetase and TRPV4. In both rodents and zebrafish, TRPV4 channels are expressed on astrocytes, neurons and ECs (Vriens et al., 2005; Benfenati et al., 2007; Grant et al., 2007; Mangos et al., 2007; Marrelli et al., 2007; Amato et al., 2012). Interestingly studies have shown that TRPV4 in ECs can regulate eNOS (Sukumaran et al., 2013) and the presence of a feedback loop from eNOS to TRPV4 to modulate TRPV4 based Ca2+ signalling (Yin et al., 2008). Rodent models have shown that TRPV4 on cortical astrocyte endfeet can evoke changes in intracellular astrocyte calcium concentration, thereby modulating vascular tone and contributing to NVC (Dunn et al., 2013; Filosa et al., 2013). Rodent experiments support our observations by showing TRPV4 downregulation in streptozotocin induced diabetes (Monaghan et al., 2015).

Retinal studies with streptozotocin-induced hyperglycemia in rodents have suggested that hyperplasia of the Muller cells (retinal analog of astrocytes) could lead to an increase in GFAP (Newman & Reichenbach, 1996; Rungger-Brandle et al., 2000). Astrocytic glutamate clearance is also impaired under high glucose conditions (Coleman et al., 2004) making neurons susceptible to depolarization, a possible cause of neurotoxicity. This could result in accumulation of glutamate in the extrasynaptic space leading to recurrent neuronal depolarization. This is concordant with our observation of an increased number of spontaneous calcium peaks in the glucose treated larvae. Neuronal hyperexcitability and increased firing is known to be associated with seizures, commonly observed in diabetic patients (Martinez & Megias, 2009; Baviera et al., 2017). The various cellular markers shown here to be affected by glucose exposure could explain a predisposition of diabetics to seizures. Increased neuronal firing could also lead to abnormal and non – precise pre and post synaptic neuronal firing causing defects in the synaptic plasticity mechanisms necessary for cognition and memory. Further exploration of this could help define the relationship between diabetes and cognitive defects.

The anatomic and molecular effects of glucose exposure on the NVU were associated with altered embryonic behaviour, with a reduction in preference for light, which is a measure of unconditioned anxiety and related disorders in rodents and zebrafish (Kulesskaya & Voikar, 2014; Kysil et al., 2017). Unconditioned anxiety is influenced by environmental, emotional and cognitive factors (Arrant et al., 2013). It is based on an approach/avoidance conflict between the drive to explore a novel area and an aversion to brightly lit/completely dark open spaces in adult/larval zebrafish, respectively (Bourin & Hascoet, 2003; Arrant et al., 2013). The approach/avoidance conflict is well-studied in mammals and is known to have various neural substrates in the brain, including the limbic system, anterior cingulate cortex, ventral striatum and prefrontal cortex (Aupperle et al., 2015). Although zebrafish do not possess a cortex, their ventral and dorsal telencephalic area (Vd and Dm, respectively) are homologous to the mammalian amygdala and striatum (Maximino et al., 2013). Thus impaired light/dark preference as described in the present study could imply an abnormal circuitry in the zebrafish Vd/Dm. Anatomical studies with zebrafish have shown that Vd/Dm system project to the optic tectum and hence any defects could also affect the optic tectum (Scott & Baier, 2009; Nevin et al., 2010).

Various studies have described larval zebrafish behavioural differences with anxiolytic or anxiogenic treatments (Egan et al., 2009; Richendrfer et al., 2012). However, this is the first study characterizing the effect of hyperglycemia on geometrical and positional characteristics of zebrafish locomotion. Altered positional and geometric features with glucose exposure implies an altered behaviour such as increased exploration and thigmotaxis. Various zebrafish studies have shown increased exploration and thigmotaxis with anxiogenic drug treatments (Egan et al., 2009; Blaser et al., 2010). This further points to the association of glucose exposure and diabetes to anxiety related brain activation, which warrants further investigation. Although human diabetes is a complex disorder and our zebrafish model examines only the effect of hyperglycemia, our findings broadly reproduce those in other cell based and mammalian model (Williams et al., 1996; Li et al., 2002). Diabetes is well known to reduce vascular NO levels, and our work reproduces this. Although zebrafish are not known to possess endothelial nitric oxide synthase (eNOS) (Syeda et al., 2013), our results strongly indicate vascular NO production. Previous studies have linked hyperglycemia and pharmacologically induced diabetes to reductions in cerebral blood flow (Dandona et al., 1978; Stevens et al., 2000). A recent study demonstrated rescuing effects of the NO donor, SNP, on hyperglycemia induced neurovascular uncoupling (Chhabria et al., 2018). Using the same protocol as described in (Chhabria et al., 2018) to induce hyperglycemia in larval zebrafish, we have now described multiple effects of hyperglycemia on cellular markers of the NVU, essential for regulation of CBF and on zebrafish behaviour.

This is the first study to demonstrate that SNP reverses the negative consequences of hyperglycemia on neurovascular anatomy and behaviour. The ability of SNP to prevent all the observed anatomical, molecular and behavioural effects of glucose exposure is exciting as it may represent a possible treatment for diabetes-associated neurovascular dysfunction. Future studies are needed to assess whether these effects of glucose exposure and SNP or other NO donors are seen in mammalian models or humans. NO donors are already widely used clinically for angina and heart failure, and are very cheap and off-patent. Therefore, if mammalian pre-clinical studies support our findings clinical studies could rapidly be performed to examine the ability of NO donors to ameliorate or prevent the consequences of diabetes on neurological dysfunction.

Author contributions

C.H. and T.J.A.C conceived the work detailed, prepared the manuscript and wrote the grant funding the work. K.C. performed all experiments, performed all data analysis and prepared the manuscript. K.P. assisted with experiments. A.V. and E.V. performed and assisted with behavioural data analysis. C.G. generated the klf2a reporter line. Z.J. and R.N.W. generated the Tg(sm22ab:nls-mcherry)sh480 reporter line. R.B.M. provided the antibodies, protocols and assisted with the immunohistochemistry. All authors assisted with writing and editing the manuscript.

Funding Acknowledgements

This work was funded by a Project Grant from the National Centre for Replacement, Refinement, and Reduction of Animals in Research (NC3Rs) NC/P001173/1. C.H. is the recipient of a Sir Henry Dale Fellowship jointly funded by the Wellcome Trust and the Royal Society (Grant Number 105586/Z/14/Z). The Zeiss Z1 lightsheet microscope was funded via British Heart Foundation Infrastructure Award IG/15/1/31328 awarded to T.C.

Competing Interests

The authors declare no competing financial interests.

Acknowledgments

We are very grateful to the aquarium staff of the Bateson Centre for expert husbandry and advice. We are grateful to John P. Ashton and Sarah Baxendale for expert training and assistance with the behavioural experiments.

References

  1. ↵
    Amato V, Vina E, Calavia MG, Guerrera MC, Laura R, Navarro M, De Carlos F, Cobo J, Germana A & Vega JA. (2012). TRPV4 in the sensory organs of adult zebrafish. Microscopy research and technique 75, 89–96.
    OpenUrlPubMed
  2. ↵
    Amiel SA, Sherwin RS, Simonson DC & Tamborlane WV. (1988). Effect of intensive insulin therapy on glycemic thresholds for counterregulatory hormone release. Diabetes 37, 901–907.
    OpenUrlAbstract/FREE Full Text
  3. ↵
    Areosa SA & Grimley EV. (2002). Effect of the treatment of Type II diabetes mellitus on the development of cognitive impairment and dementia. The Cochrane database of systematic reviews, CD003804.
  4. ↵
    Arrant AE, Schramm-Sapyta NL & Kuhn CM. (2013). Use of the light/dark test for anxiety in adult and adolescent male rats. Behavioural brain research 256, 119–127.
    OpenUrl
  5. ↵
    Attwell D, Buchan AM, Charpak S, Lauritzen M, MacVicar BA & Newman EA. (2010). Glial and neuronal control of brain blood flow. Nature 468, 232–243.
    OpenUrlCrossRefPubMedWeb of Science
  6. ↵
    Aupperle RL, Melrose AJ, Francisco A, Paulus MP & Stein MB. (2015). Neural substrates of approach-avoidance conflict decision-making. Human brain mapping 36, 449–462.
    OpenUrlCrossRefPubMed
  7. ↵
    Baviera M, Roncaglioni MC, Tettamanti M, Vannini T, Fortino I, Bortolotti A, Merlino L & Beghi E. (2017). Diabetes mellitus: a risk factor for seizures in the elderly-a population-based study. Acta Diabetol 54, 863–870.
    OpenUrl
  8. ↵
    Benfenati V, Amiry-Moghaddam M, Caprini M, Mylonakou MN, Rapisarda C, Ottersen OP & Ferroni S. (2007). Expression and functional characterization of transient receptor potential vanilloid-related channel 4 (TRPV4) in rat cortical astrocytes. Neuroscience 148, 876–892.
    OpenUrlCrossRefPubMedWeb of Science
  9. ↵
    Bergles DE, Diamond JS & Jahr CE. (1999). Clearance of glutamate inside the synapse and beyond. Curr Opin Neurobiol 9, 293–298.
    OpenUrlCrossRefPubMedWeb of Science
  10. ↵
    Bergmann K, Meza Santoscoy P, Lygdas K, Nikolaeva Y, MacDonald RB, Cunliffe VT & Nikolaev A. (2018). Imaging Neuronal Activity in the Optic Tectum of Late Stage Larval Zebrafish. Journal of developmental biology 6.
  11. ↵
    Blaser R & Gerlai R. (2006). Behavioral phenotyping in zebrafish: comparison of three behavioral quantification methods. Behavior research methods 38, 456–469.
    OpenUrlCrossRefPubMedWeb of Science
  12. ↵
    Blaser RE, Chadwick L & McGinnis GC. (2010). Behavioral measures of anxiety in zebrafish (Danio rerio). Behavioural brain research 208, 56–62.
    OpenUrlCrossRefPubMedWeb of Science
  13. ↵
    Bourin M & Hascoet M. (2003). The mouse light/dark box test. European journal of pharmacology 463, 55–65.
    OpenUrlCrossRefPubMedWeb of Science
  14. ↵
    Boyle PJ, Schwartz NS, Shah SD, Clutter WE & Cryer PE. (1988). Plasma glucose concentrations at the onset of hypoglycemic symptoms in patients with poorly controlled diabetes and in nondiabetics. New England Journal of Medicine 318, 1487–1492.
    OpenUrlCrossRefPubMedWeb of Science
  15. ↵
    Bringmann A, Grosche A, Pannicke T & Reichenbach A. (2013). GABA and Glutamate Uptake and Metabolism in Retinal Glial (Muller) Cells. Frontiers in endocrinology 4, 48.
  16. ↵
    Burbaeva G, Boksha IS, Turishcheva MS, Vorobyeva EA, Savushkina OK & Tereshkina EB. (2003). Glutamine synthetase and glutamate dehydrogenase in the prefrontal cortex of patients with schizophrenia. Progress in neuro-psychopharmacology & biological psychiatry 27, 675–680.
    OpenUrlCrossRefPubMed
  17. ↵
    Capiotti KM, Antonioli R, Jr.., Kist LW, Bogo MR, Bonan CD & Da Silva RS. (2014). Persistent impaired glucose metabolism in a zebrafish hyperglycemia model. Comparative biochemistry and physiology Part B, Biochemistry & molecular biology 171, 58–65.
    OpenUrl
  18. ↵
    Chase HP, Jackson WE, Hoops SL, Cockerham RS, Archer PG & O’Brien D. (1989). Glucose control and the renal and retinal complications of insulin-dependent diabetes. Jama 261, 1155–1160.
    OpenUrlCrossRefPubMedWeb of Science
  19. ↵
    Chhabria K, Plant K, Bandmann O, Wilkinson RN, Martin C, Kugler E, Armitage PA, Santoscoy PL, Cunliffe VT, Huisken J, McGown A, Ramesh T, Chico TJ & Howarth C. (2018). The effect of hyperglycemia on neurovascular coupling and cerebrovascular patterning in zebrafish. Journal of cerebral blood flow and metabolism: official journal of the International Society of Cerebral Blood Flow and Metabolism, 271678X18810615.
  20. ↵
    Chiu JJ, Usami S & Chien S. (2009). Vascular endothelial responses to altered shear stress: pathologic implications for atherosclerosis. Annals of medicine 41, 19–28.
    OpenUrlCrossRefPubMedWeb of Science
  21. ↵
    Ciudin A, Espinosa A, Simo-Servat O, Ruiz A, Alegret M, Hernandez C, Boada M & Simo R. (2017). Type 2 diabetes is an independent risk factor for dementia conversion in patients with mild cognitive impairment. Journal of diabetes and its complications 31, 1272–1274.
    OpenUrl
  22. ↵
    Coleman E, Judd R, Hoe L, Dennis J & Posner P. (2004). Effects of diabetes mellitus on astrocyte GFAP and glutamate transporters in the CNS. Glia 48, 166–178.
    OpenUrlCrossRefPubMed
  23. ↵
    Dandona P, James IM, Newbury PA, Woollard ML & Beckett AG. (1978). Cerebral blood flow in diabetes mellitus: evidence of abnormal cerebrovascular reactivity. British medical journal 2, 325–326.
    OpenUrlAbstract/FREE Full Text
  24. ↵
    Dekker RJ, van Soest S, Fontijn RD, Salamanca S, de Groot PG, VanBavel E, Pannekoek H & Horrevoets AJG. (2002). Prolonged fluid shear stress induces a distinct set of endothelial cell genes, most specifically lung Kruppel-like factor (KLF2). Blood 100, 1689–1698.
    OpenUrlAbstract/FREE Full Text
  25. ↵
    Dekker RJ, van Thienen JV, Rohlena J, de Jager SC, Elderkamp YW, Seppen J, de Vries CJ, Biessen EA, van Berkel TJ, Pannekoek H & Horrevoets AJ. (2005). Endothelial KLF2 links local arterial shear stress levels to the expression of vascular tone-regulating genes. The American journal of pathology 167, 609–618.
    OpenUrlCrossRefPubMedWeb of Science
  26. ↵
    Dooley K & Zon LI. (2000). Zebrafish: a model system for the study of human disease. Current opinion in genetics & development 10, 252–256.
    OpenUrlCrossRefPubMedWeb of Science
  27. ↵
    Du XL, Edelstein D, Dimmeler S, Ju Q, Sui C & Brownlee M. (2001). Hyperglycemia inhibits endothelial nitric oxide synthase activity by posttranslational modification at the Akt site. J Clin Invest 108, 1341–1348.
    OpenUrlCrossRefPubMedWeb of Science
  28. ↵
    Dunn KM, Hill-Eubanks DC, Liedtke WB & Nelson MT. (2013). TRPV4 channels stimulate Ca2+- induced Ca2+ release in astrocytic endfeet and amplify neurovascular coupling responses. Proceedings of the National Academy of Sciences 110, 6157–6162.
    OpenUrlAbstract/FREE Full Text
  29. ↵
    Egan RJ, Bergner CL, Hart PC, Cachat JM, Canavello PR, Elegante MF, Elkhayat SI, Bartels BK, Tien AK, Tien DH, Mohnot S, Beeson E, Glasgow E, Amri H, Zukowska Z & Kalueff AV. (2009). Understanding behavioral and physiological phenotypes of stress and anxiety in zebrafish. Behavioural brain research 205, 38–44.
    OpenUrlCrossRefPubMedWeb of Science
  30. ↵
    Filosa JA, Yao X & Rath G. (2013). TRPV4 and the regulation of vascular tone. Journal of cardiovascular pharmacology 61, 113–119.
    OpenUrlCrossRefPubMed
  31. ↵
    Frank RN, Turczyn TJ & Das A. (1990). Pericyte coverage of retinal and cerebral capillaries. Investigative ophthalmology & visual science 31, 999–1007.
    OpenUrlAbstract/FREE Full Text
  32. ↵
    Fukuhara S, Sako K, Noda K, Nagao K, Miura K & Mochizuki N. (2009). Tie2 is tied at the cell-cell contacts and to extracellular matrix by angiopoietin-1. Experimental & molecular medicine 41, 133.
    OpenUrl
  33. ↵
    Gaengel K, Genove G, Armulik A & Betsholtz C. (2009). Endothelial-mural cell signaling in vascular development and angiogenesis. Arterioscler Thromb Vasc Biol 29, 630–638.
    OpenUrlAbstract/FREE Full Text
  34. ↵
    Gardner TW & Davila JR. (2017). The neurovascular unit and the pathophysiologic basis of diabetic retinopathy. Graefe’s archive for clinical and experimental ophthalmology = Albrecht von Graefes Archiv fur klinische und experimentelle Ophthalmologie 255, 1–6.
    OpenUrl
  35. ↵
    Gehring TV, Luksys G, Sandi C & Vasilaki E. (2015). Detailed classification of swimming paths in the Morris Water Maze: multiple strategies within one trial. Sci Rep 5, 14562.
  36. ↵
    Gerhardt H & Betsholtz C. (2003). Endothelial-pericyte interactions in angiogenesis. Cell and tissue research 314, 15–23.
    OpenUrlCrossRefPubMedWeb of Science
  37. ↵
    Gracia-Sancho J, Russo L, Garcia-Caldero H, Garcia-Pagan JC, Garcia-Cardena G & Bosch J. (2011). Endothelial expression of transcription factor Kruppel-like factor 2 and its vasoprotective target genes in the normal and cirrhotic rat liver. Gut 60, 517–524.
    OpenUrlAbstract/FREE Full Text
  38. ↵
    Grant AD, Cottrell GS, Amadesi S, Trevisani M, Nicoletti P, Materazzi S, Altier C, Cenac N, Zamponi GW, Bautista-Cruz F, Lopez CB, Joseph EK, Levine JD, Liedtke W, Vanner S, Vergnolle N, Geppetti P & Bunnett NW. (2007). Protease-activated receptor 2 sensitizes the transient receptor potential vanilloid 4 ion channel to cause mechanical hyperalgesia in mice. J Physiol 578, 715–733.
    OpenUrlCrossRefPubMedWeb of Science
  39. ↵
    Graziano A, Petrosini L & Bartoletti A. (2003). Automatic recognition of explorative strategies in the Morris water maze. J Neurosci Methods 130, 33–44.
    OpenUrlCrossRefPubMedWeb of Science
  40. ↵
    Groeneveld O, Reijmer Y, Heinen R, Kuijf H, Koekkoek P, Janssen J, Rutten G, Kappelle L, Biessels G & group C-Is. (2018). Brain imaging correlates of mild cognitive impairment and early dementia in patients with type 2 diabetes mellitus. Nutrition, metabolism, and cardiovascular diseases: NMCD 28, 1253–1260.
    OpenUrl
  41. ↵
    Guerra G, Lucariello A, Perna A, Botta L, De Luca A & Moccia F. (2018). The Role of Endothelial Ca(2+) Signaling in Neurovascular Coupling: A View from the Lumen. International journal of molecular sciences 19.
  42. ↵
    Hall CN, Reynell C, Gesslein B, Hamilton NB, Mishra A, Sutherland BA, O’Farrell FM, Buchan AM, Lauritzen M & Attwell D. (2014). Capillary pericytes regulate cerebral blood flow in health and disease. Nature 508, 55–60.
    OpenUrlCrossRefPubMedWeb of Science
  43. ↵
    Hammes HP, Lin J, Renner O, Shani M, Lundqvist A, Betsholtz C, Brownlee M & Deutsch U. (2002). Pericytes and the pathogenesis of diabetic retinopathy. Diabetes 51, 3107–3112.
    OpenUrlAbstract/FREE Full Text
  44. ↵
    Hogan BM, Herpers R, Witte M, Heloterä H, Alitalo K, Duckers HJ & Schulte-Merker S. (2009). Vegfc/Flt4 signalling is suppressed by Dll4 in developing zebrafish intersegmental arteries. Development 136, 4001–4009.
    OpenUrlAbstract/FREE Full Text
  45. ↵
    Ignarro LJ, Buga GM, Wood KS, Byrns RE & Chaudhuri G. (1987). Endothelium-derived relaxing factor produced and released from artery and vein is nitric oxide. Proceedings of the National Academy of Sciences 84, 9265–9269.
    OpenUrlAbstract/FREE Full Text
  46. ↵
    Illouz T, Madar R, Louzon Y, Griffioen KJ & Okun E. (2016). Unraveling cognitive traits using the Morris water maze unbiased strategy classification (MUST-C) algorithm. Brain, behavior, and immunity 52, 132–144.
    OpenUrl
  47. ↵
    Janzer RC & Raff MC. (1987). Astrocytes induce blood-brain barrier properties in endothelial cells. Nature 325, 253–257.
    OpenUrlCrossRefPubMedWeb of Science
  48. ↵
    Jorgensen HS, Nakayama H, Raaschou HO & Olsen TS. (1994). Effect of blood pressure and diabetes on stroke in progression. Lancet 344, 156–159.
    OpenUrlCrossRefPubMedWeb of Science
  49. ↵
    Kilkenny C, Browne W, Cuthill IC, Emerson M, Altman DG & Group NCRRGW. (2010). Animal research: reporting in vivo experiments: the ARRIVE guidelines. British journal of pharmacology 160, 1577–1579.
    OpenUrlCrossRefPubMedWeb of Science
  50. ↵
    Kimmel RA, Dobler S, Schmitner N, Walsen T, Freudenblum J & Meyer D. (2015). Diabetic pdx1-mutant zebrafish show conserved responses to nutrient overload and anti-glycemic treatment. Scientific Reports 5.
  51. ↵
    Kulesskaya N & Voikar V. (2014). Assessment of mouse anxiety-like behavior in the light-dark box and open-field arena: role of equipment and procedure. Physiology & behavior 133, 30–38.
    OpenUrl
  52. ↵
    Kysil EV, Meshalkina DA, Frick EE, Echevarria DJ, Rosemberg DB, Maximino C, Lima MG, Abreu MS, Giacomini AC, Barcellos LJG, Song C & Kalueff AV. (2017). Comparative Analyses of Zebrafish Anxiety-Like Behavior Using Conflict-Based Novelty Tests. Zebrafish 14, 197–208.
    OpenUrlCrossRef
  53. ↵
    Lawrence C. (2007). The husbandry of zebrafish (Danio rerio): a review. Aquaculture 269, 1–20.
    OpenUrlCrossRefWeb of Science
  54. ↵
    Lee JS, Kang Decker N, Chatterjee S, Yao J, Friedman S & Shah V. (2005). Mechanisms of nitric oxide interplay with Rho GTPase family members in modulation of actin membrane dynamics in pericytes and fibroblasts. The American journal of pathology 166, 1861–1870.
    OpenUrlCrossRefPubMedWeb of Science
  55. ↵
    Li Q, Zemel E, Miller B & Perlman I. (2002). Early retinal damage in experimental diabetes: electroretinographical and morphological observations. Exp Eye Res 74, 615–625.
    OpenUrlCrossRefPubMedWeb of Science
  56. ↵
    Lieschke GJ & Currie PD. (2007). Animal models of human disease: zebrafish swim into view. Nature Reviews Genetics 8, 353–367.
    OpenUrlCrossRefPubMedWeb of Science
  57. ↵
    Lieth E, LaNoue KF, Antonetti DA, Ratz M & Group PSRR. (2000). Diabetes reduces glutamate oxidation and glutamine synthesis in the retina. Experimental eye research 70, 723–730.
    OpenUrlCrossRefPubMedWeb of Science
  58. ↵
    Lin Z, Kumar A, SenBanerjee S, Staniszewski K, Parmar K, Vaughan DE, Gimbrone MA, Jr.., Balasubramanian V, Garcia-Cardena G & Jain MK. (2005). Kruppel-like factor 2 (KLF2) regulates endothelial thrombotic function. Circ Res 96, e48–57.
    OpenUrlAbstract/FREE Full Text
  59. ↵
    MacKnight C, Rockwood K, Awalt E & McDowell I. (2002). Diabetes mellitus and the risk of dementia, Alzheimer’s disease and vascular cognitive impairment in the Canadian Study of Health and Aging. Dementia and geriatric cognitive disorders 14, 77–83.
    OpenUrlCrossRefPubMedWeb of Science
  60. ↵
    Mangos S, Liu Y & Drummond IA. (2007). Dynamic expression of the osmosensory channel trpv4 in multiple developing organs in zebrafish. Gene expression patterns: GEP 7, 480–484.
    OpenUrl
  61. ↵
    Marrelli SP, O’Neil RG, Brown RC & Bryan RM, Jr... (2007). PLA2 and TRPV4 channels regulate endothelial calcium in cerebral arteries. American journal of physiology Heart and circulatory physiology 292, H1390–1397.
    OpenUrlCrossRefPubMedWeb of Science
  62. ↵
    Martinez MD & Megias SM. (2009). [Occipital seizures with electroencephalographic alterations as the initial manifestation of diabetes mellitus]. Endocrinologia y nutricion: organo de la Sociedad Espanola de Endocrinologia y Nutricion 56, 458–460.
    OpenUrl
  63. ↵
    Maximino C, Lima MG, Oliveira KR, Batista Ede J & Herculano AM. (2013). “Limbic associative” and “autonomic” amygdala in teleosts: a review of the evidence. Journal of chemical neuroanatomy 48-49, 1–13.
    OpenUrlCrossRefPubMed
  64. ↵
    Meza Santoscoy PL. (2014). Analysis of the transcriptional and behavioural responses to seizure onset in a zebrafish model of epilepsy. University of Sheffield.
  65. ↵
    Miettinen H, Lehto S, Salomaa V, Mahonen M, Niemela M, Haffner SM, Pyorala K & Tuomilehto J. (1998). Impact of diabetes on mortality after the first myocardial infarction. The FINMONICA Myocardial Infarction Register Study Group. Diabetes Care 21, 69–75.
    OpenUrlAbstract/FREE Full Text
  66. ↵
    Mogi M & Horiuchi M. (2011). Neurovascular coupling in cognitive impairment associated with diabetes mellitus. Circulation Journal 75, 1042–1048.
    OpenUrl
  67. ↵
    Monaghan K, McNaughten J, McGahon MK, Kelly C, Kyle D, Yong PH, McGeown JG & Curtis TM. (2015). Hyperglycemia and Diabetes Downregulate the Functional Expression of TRPV4 Channels in Retinal Microvascular Endothelium. PLoS One 10, e0128359.
    OpenUrlCrossRefPubMed
  68. ↵
    Nevin LM, Robles E, Baier H & Scott EK. (2010). Focusing on optic tectum circuitry through the lens of genetics. BMC biology 8, 1.
  69. ↵
    Newman E & Reichenbach A. (1996). The Müller cell: a functional element of the retina. Trends in neurosciences 19, 307–312.
    OpenUrlCrossRefPubMedWeb of Science
  70. ↵
    Oliveira RF, Silva JF & Simoes JM. (2011). Fighting zebrafish: characterization of aggressive behavior and winner-loser effects. Zebrafish 8, 73–81.
    OpenUrlCrossRefPubMed
  71. ↵
    Palmer RM, Ferrige A & Moncada S. (1987). Nitric oxide release accounts for the biological activity of endothelium-derived relaxing factor.
  72. ↵
    Patton EE & Zon LI. (2001). The art and design of genetic screens: zebrafish. Nature reviews Genetics 2, 956–966.
    OpenUrlCrossRefPubMedWeb of Science
  73. ↵
    Peppiatt CM, Howarth C, Mobbs P & Attwell D. (2006). Bidirectional control of CNS capillary diameter by pericytes. Nature 443, 700–704.
    OpenUrlCrossRefPubMedWeb of Science
  74. ↵
    Pfister F, Feng Y, vom Hagen F, Hoffmann S, Molema G, Hillebrands JL, Shani M, Deutsch U & Hammes HP. (2008). Pericyte migration: a novel mechanism of pericyte loss in experimental diabetic retinopathy. Diabetes 57, 2495–2502.
    OpenUrlAbstract/FREE Full Text
  75. ↵
    Pieper GM. (1998). Review of alterations in endothelial nitric oxide production in diabetes: protective role of arginine on endothelial dysfunction. Hypertension 31, 1047–1060.
    OpenUrlCrossRef
  76. ↵
    Pisharath H, Rhee JM, Swanson MA, Leach SD & Parsons MJ. (2007). Targeted ablation of beta cells in the embryonic zebrafish pancreas using E. coli nitroreductase. Mechanisms of development 124, 218–229.
    OpenUrlCrossRefPubMedWeb of Science
  77. ↵
    Prat A, Biernacki K, Wosik K & Antel JP. (2001). Glial cell influence on the human blood-brain barrier. Glia 36, 145–155.
    OpenUrlCrossRefPubMedWeb of Science
  78. ↵
    Richendrfer H, Pelkowski SD, Colwill RM & Creton R. (2012). On the edge: pharmacological evidence for anxiety-related behavior in zebrafish larvae. Behavioural brain research 228, 99–106.
    OpenUrlCrossRefPubMed
  79. ↵
    Roy CS & Sherrington CS. (1890). On the regulation of the blood-supply of the brain. The Journal of physiology 11, 85.
    OpenUrlCrossRefPubMed
  80. ↵
    Rungger-Brandle E, Dosso AA & Leuenberger PM. (2000). Glial reactivity, an early feature of diabetic retinopathy. Invest Ophthalmol Vis Sci 41, 1971–1980.
    OpenUrlAbstract/FREE Full Text
  81. ↵
    Sagare AP, Bell RD, Zhao Z, Ma Q, Winkler EA, Ramanathan A & Zlokovic BV. (2013). Pericyte loss influences Alzheimer-like neurodegeneration in mice. Nat Commun 4, 2932.
    OpenUrlCrossRefPubMed
  82. ↵
    Sakagami K, Kawamura H, Wu DM & Puro DG. (2001). Nitric oxide/cGMP-induced inhibition of calcium and chloride currents in retinal pericytes. Microvasc Res 62, 196–203.
    OpenUrlCrossRefPubMedWeb of Science
  83. ↵
    Scott EK & Baier H. (2009). The cellular architecture of the larval zebrafish tectum, as revealed by gal4 enhancer trap lines. Frontiers in neural circuits 3, 13.
    OpenUrl
  84. ↵
    Scully T. (2012). Diabetes in numbers. Nature 485, S2–3.
    OpenUrlCrossRefPubMedWeb of Science
  85. ↵
    SenBanerjee S, Lin Z, Atkins GB, Greif DM, Rao RM, Kumar A, Feinberg MW, Chen Z, Simon DI & Luscinskas FW. (2004a). KLF2 Is a novel transcriptional regulator of endothelial proinflammatory activation. J Exp Med 199, 1305–1315.
    OpenUrlAbstract/FREE Full Text
  86. ↵
    SenBanerjee S, Lin ZY, Atkins GB, Greif DM, Rao RM, Kumar A, Feinberg MW, Chen ZP, Simon DI, Luscinskas FW, Michel TM, Gimbrone MA, Garcia-Cardena G & Jain MK. (2004b). KLF2 is a novel transcriptional regulator of endothelial proinflammatory activation. J Exp Med 199, 1305–1315.
    OpenUrlAbstract/FREE Full Text
  87. ↵
    Simo R, Ciudin A, Simo-Servat O & Hernandez C. (2017). Cognitive impairment and dementia: a new emerging complication of type 2 diabetes-The diabetologist’s perspective. Acta Diabetol 54, 417–424.
    OpenUrlPubMed
  88. ↵
    Stevens MJ, Obrosova I, Cao X, Van Huysen C & Greene DA. (2000). Effects of DL-alpha-lipoic acid on peripheral nerve conduction, blood flow, energy metabolism, and oxidative stress in experimental diabetic neuropathy. Diabetes 49, 1006–1015.
    OpenUrlAbstract
  89. ↵
    Stewart R & Liolitsa D. (1999). Type 2 diabetes mellitus, cognitive impairment and dementia. Diabetic medicine: a journal of the British Diabetic Association 16, 93–112.
    OpenUrl
  90. ↵
    Stratton IM, Adler AI, Neil HA, Matthews DR, Manley SE, Cull CA, Hadden D, Turner RC & Holman RR. (2000). Association of glycaemia with macrovascular and microvascular complications of type 2 diabetes (UKPDS 35): prospective observational study. Bmj 321, 405–412.
    OpenUrlAbstract/FREE Full Text
  91. ↵
    Sukumaran SV, Singh TU, Parida S, Narasimha Reddy Ch E, Thangamalai R, Kandasamy K, Singh V & Mishra SK. (2013). TRPV4 channel activation leads to endothelium-dependent relaxation mediated by nitric oxide and endothelium-derived hyperpolarizing factor in rat pulmonary artery. Pharmacological research 78, 18–27.
    OpenUrlCrossRefPubMed
  92. ↵
    Syeda F, Hauton D, Young S & Egginton S. (2013). How ubiquitous is endothelial NOS? Comp Biochem Phys A 166, 207–214.
    OpenUrl
  93. ↵
    Takano T, Tian G-F, Peng W, Lou N, Libionka W, Han X & Nedergaard M. (2006). Astrocyte-mediated control of cerebral blood flow. Nature neuroscience 9, 260–267.
    OpenUrlCrossRefPubMedWeb of Science
  94. ↵
    Tilton RG, Faller AM, Burkhardt JK, Hoffmann PL, Kilo C & Williamson JR. (1985). Pericyte degeneration and acellular capillaries are increased in the feet of human diabetic patients. Diabetologia 28, 895–900.
    OpenUrlCrossRefPubMedWeb of Science
  95. ↵
    Toth P, Tarantini S, Davila A, Valcarcel-Ares MN, Tucsek Z, Varamini B, Ballabh P, Sonntag WE, Baur JA, Csiszar A & Ungvari Z. (2015). Purinergic glio-endothelial coupling during neuronal activity: role of P2Y1 receptors and eNOS in functional hyperemia in the mouse somatosensory cortex. American journal of physiology Heart and circulatory physiology 309, H1837–1845.
    OpenUrlCrossRefPubMed
  96. ↵
    Vates GE, Takano T, Zlokovic B & Nedergaard M. (2010). Pericyte constriction after stroke: the jury is still out. Nature medicine 16, 959–959.
    OpenUrlCrossRefPubMed
  97. ↵
    Vouros A, Gehring TV, Szydlowska K, Janusz A, Tu Z, Croucher M, Lukasiuk K, Konopka W, Sandi C & Vasilaki E. (2018). A generalised framework for detailed classification of swimming paths inside the Morris Water Maze. Sci Rep 8, 15089.
    OpenUrl
  98. ↵
    Vriens J, Owsianik G, Fisslthaler B, Suzuki M, Janssens A, Voets T, Morisseau C, Hammock BD, Fleming I, Busse R & Nilius B. (2005). Modulation of the Ca2 permeable cation channel TRPV4 by cytochrome P450 epoxygenases in vascular endothelium. Circ Res 97, 908–915.
    OpenUrlAbstract/FREE Full Text
  99. ↵
    Williams SB, Cusco JA, Roddy MA, Johnstone MT & Creager MA. (1996). Impaired nitric oxide-mediated vasodilation in patients with non-insulin-dependent diabetes mellitus. Journal of the American College of Cardiology 27, 567–574.
    OpenUrlFREE Full Text
  100. ↵
    Wolfer DP & Lipp HP. (2000). Dissecting the behaviour of transgenic mice: is it the mutation, the genetic background, or the environment? Experimental physiology 85, 627–634.
    OpenUrlCrossRefPubMedWeb of Science
  101. ↵
    Wu J, Bohanan CS, Neumann JC & Lingrel JB. (2008). KLF2 transcription factor modulates blood vessel maturation through smooth muscle cell migration. Journal of Biological Chemistry 283, 3942–3950.
    OpenUrlAbstract/FREE Full Text
  102. ↵
    Yin J, Hoffmann J, Kaestle SM, Neye N, Wang L, Baeurle J, Liedtke W, Wu S, Kuppe H, Pries AR & Kuebler WM. (2008). Negative-feedback loop attenuates hydrostatic lung edema via a cGMP- dependent regulation of transient receptor potential vanilloid 4. Circ Res 102, 966–974.
    OpenUrlAbstract/FREE Full Text
  103. ↵
    Zhang P, Zhang X, Brown J, Vistisen D, Sicree R, Shaw J & Nichols G. (2010). Global healthcare expenditure on diabetes for 2010 and 2030. Diabetes research and clinical practice 87, 293–301.
    OpenUrlCrossRefPubMedWeb of Science
  104. ↵
    Zlokovic BV. (2010). Neurodegeneration and the neurovascular unit. Nature medicine 16, 1370–1371.
    OpenUrlCrossRefPubMed
  105. ↵
    Zonta M, Angulo MC, Gobbo S, Rosengarten B, Hossmann K-A, Pozzan T & Carmignoto G. (2003). Neuron-to-astrocyte signaling is central to the dynamic control of brain microcirculation. Nature neuroscience 6, 43–50.
    OpenUrlCrossRefPubMedWeb of Science
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Posted March 19, 2019.
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Sodium nitroprusside prevents the detrimental effects of glucose on the neurovascular unit and behaviour in zebrafish
K. Chhabria, A. Vouros, C. Gray, R.B. MacDonald, Z. Jiang, R.N. Wilkinson, K Plant, E. Vasilaki, C. Howarth, T.J.A. Chico
bioRxiv 576942; doi: https://doi.org/10.1101/576942
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