Exacerbation of blood brain barrier dysfunction and brain injury after aneurysmal subarachnoid hemorrhage in endothelial specific S1PR1 knock out mice

ABSTRACT

Blood brain barrier (BBB) dysfunction upon ischemia and hypoxia has been implicated in the exacerbation of neuronal injury in stroke. Despite the therapeutic potential of the cerebrovascular endothelium, the limited understanding of the endothelial signaling pathways governing BBB function restricts progress towards developing novel therapeutic approaches specifically targeting the endothelium in stroke. Sphingosine-1-phosphate (S1P) is a potent modulator of endothelial function via its receptors (S1PR). Recent human and mouse studies indicate that anti-inflammatory endothelial S1P signaling via S1PR1 is limiting in cardiovascular and inflammatory diseases. Herein, we aimed to investigate the expression of S1PR1 in the mouse and human brain and the role of endothelial-specific S1PR1 signaling in BBB function, neuronal injury and neurological outcomes in a mouse model of aneurysmal subarachnoid hemorrhage (SAH), the most devastating type of stroke. We found that S1PR1 is the most abundant S1PR transcript in the mouse brain and in mouse and human brain endothelial cells (20-100 mRNA copies per cell). S1PR1 transcripts were significantly enriched (~6 fold) in mouse cortical microvessels compared to total brain. Using the S1PR1-eGFP knock in mouse, we found that S1PR1-eGFP is abundantly expressed in the cerebrovascular endothelium and neurons in the mouse brain. A similar pattern of expression was observed in human brain autopsy samples. Endothelial specific deletion of S1PR1, induced in adult mice (S1pr1 ^flox/flox × Cdh5-Cre^ERT2, referred to as S1pr1^iECKO) by tamoxifen administration, resulted in exacerbation of brain edema, neuronal injury as well as worsened neurological outcomes upon SAH compared to S1pr1 ^flox/flox littermates. No differences in the SAH grade, hemostasis in the tail bleeding assay or cerebral blood flow changes during and after SAH were found between groups. Mechanistically, S1pr1^iECKO exhibited aggravated BBB dysfunction and increased phosphorylation of myosin light chain (MLC) in isolated cortical microvessels, a downstream effector of the Rho-ROCK pathway implicated in endothelial inflammation and barrier dysfunction.

Our study indicates that inhibition of S1PR1 signaling specifically in the endothelium exacerbates BBB dysfunction upon ischemic-hypoxic injury and thereby heightens neuronal injury resulting in worsened neurological outcomes. Given the expression of S1PR1 in the cerebrovascular endothelium in humans and recent findings indicating that S1P-S1PR1 anti-inflammatory signaling in the endothelium could be limiting in cardiovascular disease and other inflammatory conditions, the therapeutic and diagnostic potential of the endothelial sphingosine-1-phosphate pathway in stroke is very high.

INTRODUCTION

The cerebrovascular endothelium, in coordination with pericytes ^{1, 2} and astrocytes ³ plays a critical role in the maintenance of the blood brain barrier (BBB). BBB dysfunction has been implicated in the exacerbation of neurovascular ischemic, hypoxic and inflammatory injury, which occurs after stroke, a leading cause of mortality and long-term disability worldwide⁴. For instance, the use and effectiveness of current therapies for ischemic stroke (i.e. tissue plasminogen activator, tPA, or mechanical recanalization) are severely limited by the vascular complications of reperfusion injury (i.e. blood brain barrier dysfunction leading to vasogenic edema and hemorrhagic transformation)^5–9. In hemorrhagic stroke, perihematoma edema volume expansion is associated with worsened outcomes ¹⁰. In subarachnoid hemorrhage (SAH), blood brain barrier breakdown occurs in the acute phase and correlates with more severe neurological deficits ^11–13. However, currently, there are no therapies specifically targeting the endothelium to promote BBB function and prevent exacerbation of neurovascular injury in stroke. A better understanding of the cellular and molecular mechanisms governing BBB function may help design novel therapies aimed at mitigating cerebral endothelial barrier dysfunction in stroke in order to prevent exacerbation of neurovascular injury and improve outcomes.

S1P, a bioactive lipid very abundant in plasma, is a potent modulator of endothelial and lymphocyte function. Plasma S1P originates from endothelial cells^{14, 15} and erythrocytes¹⁶ and it is bound to lipoproteins (e.g. HDL^{17, 18} via apolipoprotein M (ApoM)¹⁸), albumin and other chaperones. In mice, S1P signaling via S1PR1 is required for embryonic and postnatal vascular development and maturation in brain and other organs ^{19, 20,21}. In adult mice, S1PR1 promotes endothelial barrier function, vascular stabilization and inhibits endothelial inflammation ^22–24. These effects are dependent on the activation of anti-inflammatory signaling pathways (Gi-phosphatidylinositol-3-kinase leading to activation of Rac and Akt) which strengthen endothelial barrier function by promoting actin cytoskeleton dynamics (cortical actin assembly) and adherens junctions assembly to the cytoskeleton. In contrast, S1P binding to S1PR2 induces barrier dysfunction and activates pro-inflammatory signaling pathways (e.g. NFκB) in a Rho-Rho kinase dependent way. S1P-S1PR1 antiinflammatory signaling pathway is predominant in the endothelium when S1P is bound to HDL via ApoM ²⁴ (Reviewed in ²⁵). Indeed, a good part of the cardiovascular protective effects of HDL depend on its content of S1P ^26–28 and ApoM ²⁴, which also has been shown to protect plasma S1P from degradation ¹⁸. Recent studies indicate that S1P anti-inflammatory and vasoprotective signaling via S1PR1 could be limiting in human cardiovascular and inflammatory diseases ^28–31,25. For instance, in coronary artery disease, the content of ApoM or S1P in the HDL fraction is significantly reduced, and it inversely correlates with the severity of the disease^{28, 29}. In addition, in patients suffering inflammatory conditions such diabetes or sepsis, plasma ApoM levels are significantly decreased compared to control patients ^{32, 33}.

Interestingly, S1PR1 also plays a critical role in lymphocyte trafficking and function³⁴. Indeed, the immunosupressor FTY720, which has been approved by the FDA for multiple sclerosis ^{35, 36}, is a S1PR1 functional antagonist (i.e. a potent S1PR1 agonist which induces S1PR1 internalization and degradation ^{37, 38}). Recent studies attempting to understand the role of S1P in cerebral ischemia have relied on the use of the immunosuppressor FTY720 and other S1PR1 agonists ^{39, 40, 41,42}, which protected in experimental stroke, mainly due to their immunosuppressive effects⁴²(reviewed in²⁵). Thus, these studies could not determine the role of endothelial-specific S1P signaling in BBB function modulation and its impact on brain injury and stroke outcomes. Although immunosuppression is protective in experimental stroke, it is not clear that it could be a good therapeutic strategy in humans, since it increases the risk of developing infections and sepsis, which is an important cause of worsened outcomes and mortality in stroke patients ^{43, 44}. Hence, a better understanding of the role of endothelial S1P signaling in stroke will be important for future design of novel vasoprotective therapeutic agents targeting this pathway in the endothelium without compromising the immune response.

Given the pathophysiological relevance of the S1P-S1PR1 pathway in humans and in order to bridge this knowledge gap, in this study we aimed to investigate the expression of S1PR1 in the mouse and human brain as well as the role of endothelial-specific S1PR1 signaling in modulation of blood brain barrier function and its impact on brain injury upon subarachnoid hemorrhage, the most devastating type of stroke. Our results show the critical role of endothelial-specific S1PR1 signaling in BBB function and its impact on stroke outcomes. They also indicate direct causality between endothelial barrier dysfunction and the exacerbation of neuronal injury, highlighting the critical role of the endothelium in the pathophysiology of brain injury and its therapeutic potential.

METHODS

Mice

All animal experiments were approved by the Weill Cornell Institutional Animal Care and Use Committee. Endothelial cell specific S1pr1 knockout mice (S1pr1^flox/flox×Cdh5–Cre^ERT2; referred to as S1pr1^iECKO) were generated as we have described ⁴⁵. S1pr1^flox/flox mice ²⁰ were crossed to Cdh5–Cre^ERT2 mice ⁴⁶ to generate S1pr1^flox/flox Cdh5-Cre^ERT2 mice. Mice were treated with tamoxifen (Sigma-Aldrich) by oral gavage (75 mg kg⁻¹) for 3 days at the age of 8 weeks and used for the experiments 3-4 weeks after tamoxifen treatment. S1pr1^flox/flox littermates treated with tamoxifen were used as control mice. S1pr1-eGFP knock in mice ⁴⁷ weighing 26-30 g were used for the expression studies. All experiments were performed in male mice.

Endovascular perforation SAH surgery

SAH surgery was performed on C57/BL6/J, S1pr1^iECKO mice and their littermate S1pr1^flox/flox controls as we have previously described ⁴⁸. In brief, surgery was performed using a dissecting surgical microscope. Temperature was maintained at 36.5–37.5°C by using a thermostatic blanket (Harvard Apparatus, CMA 450 Animal Temperature Controller) throughout the procedure. Mice were anesthetized with isoflurane inhalation delivered by facemask in 100% of O₂. A 15 mm midline vertical incision was made on the skin in the head. The head was fixed in stereotactic frame and cerebral blood flow was measured by Laser-speckle imager (PeriCam PSI system, Perimed, Sweden). During surgery, mice were in supine position. A 10 mm midline vertical incision was made on the skin in the neck. The common carotid, external carotid and internal arteries were dissected from the adjacent tissue. The superior thyroid artery and the occipital artery were cauterized and cut. The external carotid artery was sutured with a dead knot and cauterized above the suture. A second suture loop was also placed in the external carotid artery just before the bifurcation of the common carotid artery. A slit-knot was placed around the common carotid artery. A small clip was applied to the internal carotid artery and the slip-knot around the common carotid artery was tightened temporally. A small incision was made in the external carotid artery stump. A 5-0 monofilament with a modified tip (0.28 mm diameter or 0.28 mm × 1.7 mm) was inserted into the incision and the knot around the external carotid artery was tightened to prevent bleeding. Then, the monofilament was advanced to the common carotid artery, the small clip on the internal carotid artery was removed and the monofilament was guided through the external carotid artery to the internal carotid artery. The knot around the common carotid artery was opened again and the monofilament was introduced to the bifurcation of the internal carotid artery. The monofilament was gently pushed ~1 mm further and then withdrawn to the external carotid artery. The knot around the external carotid artery was loosen and the monofilament was slowly removed. The external carotid artery was quickly ligated to prevent bleeding. The mouse was turned in prone position and induction of subarachnoid hemorrhage was confirmed by reduction of cerebral blood flow by Laser-speckle contrast imager. After the surgery, all animals were maintained in a small animal heated recovery chamber. Two different severities of SAH models were created by changing the tip shapes of a 5-0 monofilament; a rounded tip 0.3mm in diameter was used for mild SAH model, a tip 0.3 mm × 1.7 mm for severe SAH model. The surgeon and the investigator conducting the analysis were blinded to the genotype of the mice. Animals which did not exhibit a reduction in CBF upon endovascular rupture were excluded from the study.

RNA isolation, reverse transcription and quantitative PCR analysis (RT-qPCR)

Total RNA from mouse brains and cells was prepared using RNeasy Mini Kit (Qiagen, Valencia, CA) as instructed by the manufacturer. To generate cDNA, 100 ng of RNA was reverse transcribed using random primers and SuperScript II RT-polymerase (Invitrogen, Carlsbad, CA). Primers were designed using the Primer Express oligo design program software (Applied Biosystems, Foster City, CA). Real-time quantitative PCR was performed using the SYBR Green I assay on the ABI 7500 Sequence Detection System (Applied Biosystems). PCR reactions for each cDNA sample were performed in duplicate and copy numbers were calculated using standard curves generated from a master template as we previously described. The sequence of the primers used for qPCR are shown in Table 2.

Table 1. Physiological variables in S1pr1^flox/flox and S1pr1^iECKO mice.

Table 2. Sequences of the primers used for RT-qPCR

S1PR1 Immunohistochemistry

Human tissues were retrieved from Brigham and Women’s Department of Pathology archives; this work was approved by the Institutional Review Board (Protocol #2013P001431).

Immunohistochemistry for S1PR1 was performed on an automated stainer (Leica Bond III, Leica Biosystems, Buffalo Grove, IL) using an anti-human S1PR1 rabbit polyclonal antibody (Santa Cruz Biotechnology Inc.) at a final concentration of 1.3 ug/ml. The IHC technique for S1PR1 was validated as we have previously described ⁴⁹. 5μm formalin fixed paraffin embedded tissue sections of human frontal cortex were deparaffinized and processed using heat induced epitope retrieval with an EDTA-based buffer (Leica #AR9640) for 20 minutes and incubated with primary antibody for 30 minutes at room temperature. Secondary antibody (polymer) incubation (10 minutes) and diaminobenzidine-based signal generation (10 minutes) were performed per manufacturer’s instructions (Leica # DS9800). Pictures were taken using SPOT Insight Gigabit camera and SPOT Imaging Software (5.1).

Immunofluorescence staining

Under deep anesthesia, mice were perfused with cold PBS and subsequently with 4% PFA in PBS solution. The brains were removed, postfixed with 4% PFA for 24 h, transferred to 30% sucrose solution in PBS, embedded in OCT media and frozen. Coronal sections were cut (30 μm) in a cryostat and immunofluorescence was conducted in floating sections. Sections were preserved at −20°C in a PBS solution containing 30% polyethylene glycol and 30% sucrose. Sections were washed three times with Tris-buffered saline (TBS) and were then blocked with TBS-blocking solution (1 % bovine serum albumin, 0.2 % skim milk, and 0.3 % Triton X-100 in TBS) for 1 h and incubated with the specified primary antibodies in TBS-blocking solution overnight on a shaker at 4°C, followed by the appropriate secondary antibodies for 2 hours at room temperature. Brain slices were stained with 4’,6-diamidino-2-phenylindole (DAPI) for 7 minutes and were mounted onto slides. Samples were observed on a Zeiss LSM 510 Meta Confocal microscope (Carl Zeiss Microscopy, Thornwood, NY). Images were captured using Zen LE software (Carl Zeiss Microscopy).

Isolation of cortical microvessels

The brain microvessels were isolated as we have previously described ⁵⁰. All procedures were performed in a cold room. The brains were collected and rinsed in MCDB131 medium (Thermo Fisher Scientific) with 0.5% fatty acid-free BSA (Millipore Sigma). The leptomeninges, cerebellum, brainstem and white matter were removed on ice. The cortices were homogenized in 8 mL of MCDB131 medium with 0.5% fatty acid-free BSA using a 7-mL loose-fit Dounce tissue grinder (Sigma-Aldrich) with 10 strokes. The homogenates were centrifuged at 2,000 g for 5 min at 4 °C. The pellet was suspended in 15% dextran (molecular weight ~70,000 Da, Sigma-Aldrich) in PBS and centrifuged at 10,000 g for 15 min at 4 °C. The pellet was resuspended in MCDB131 with 0.5% fatty acid-free BSA and centrifuged at 5,000 g for 10 min at 4 °C. The pellet contained the brain microvessels.

Protein extraction from brain microvessels and western blotting

Brain microvascular fragments were lysed in HEPES-RIPA buffer (50 mM HEPES pH 7.5; 1% Triton; 0.5% sodium deoxycholate; 0.1% SDS; 500 mM NaCl; 10 mM MgCl2; 50 mM β-glycerophosphate) with 1x Protease inhibitor cocktail (CalBiochem), 1 mM Na3VO4 and 1 mM NaF and centrifuged at 15,000 r min⁻¹ for 15 min. Equal amount of proteins were mixed with SDS sample buffer, boiled and separated on a 4-15% polyacrylamide gel (Bio-Rad), transferred to PVDF membranes (Millipore Sigma), and blocked 5 % milk in 0.1% Tween-20 in TBS. Immunoblot analysis was performed with S1PR1 (1:250; Santa Cruz, cat. no. sc25489) and β-actin (1:1,000; Santa Cruz, cat. no. sc-1616 HRP) antibodies. Membrane were washed with 0.1% Tween-20 in TBS, incubated with anti-rabbit IgG secondary antibody conjugated to horseradish peroxidase (1:2,000; Cell Signaling), and protein bands were visualized with enhanced chemilumescent (ECL) reagent (Thermo Fisher Scientific) on a Protec OPTIMAX X-Ray Film Processor. Relative band intensities were obtained by densitometric analysis of images using ImageJ software.

Brain endothelial cell isolation and assessment of deletion efficiency of endothelial S1pr1 mRNA

Two weeks after tamoxifen treatment, mice were sacrificed and the brains were collected and rinsed in MCDB131 medium (Thermo Fisher Scientific) with 0.5% fatty acid-free BSA (Millipore Sigma). The cortices were homogenized in MCDB131 medium using a 7-mL loose-fit Dounce tissue grinder (Sigma-Aldrich) with 15 strokes. The homogenate was mixed with same amount of 30% dextran (molecular weight ~70,000 Da, Sigma-Aldrich) in PBS and centrifuged at 4,500 r min⁻¹ for 15 min at 4 °C. The pellet was resuspended in MCD131 medium and centrifuged at 2,400 r min⁻¹ for 10 min. The pellet was resuspended in Liberase TM solution (3.5 U; Roche) with DNaseI (0.4 U; AppliChem Inc) and digested at 37.0 °C for 90 min. The enzymatic reaction was stopped by adding 2 mM of EDTA and 2% of BSA. After centrifugation at 2,400 r min⁻¹ for 10 min, the pellet was incubated in purified Rat Anti-Mouse CD31antibody (MEC13.3) (1 : 100; BD Biosciences) with Dynabeads Sheep Anti-Rat IgG (Invitrogen, cat. no. 11035) for 35 min. CD31 positive endothelial cells were isolated by using DynaMag-2 Magnet (Thermo Fisher Scientific). Total RNA was extracted from the isolated endothelial cells using shredder (Qiagen) and RNeasy Mini Kit (Qiagen) with RNase-free DNase treatment (Qiagen) according to the manufacturer’s instructions. Reverse transcription was carried out using Verso cDNA Synthesis Kit (Thermo Fisher Scientific). Real-time PCR was performed on a real-time PCR system (Applied Biosystems, ABI 7500 Fast) by using PerfeCTa SYBR Green Fast Mix Low ROX. PCR primer sequences for target molecules are in Table 2.

Grading system for SAH

Blood volume in the subarachnoid space was assessed using the grading system previously reported ⁵¹. Mice were sacrificed under deep anesthesia 24 h after SAH induction and the brains were removed. Pictures of ventral surface of the brain depicting the basal cistern with the circle of Willis and the basilar artery were taken using a stereomicroscope (Olympus, SZX16) equipped with digital camera (Olympus, DP12). The basal cistern was divided into 6 segments and a grade from 0 to 3 was given to each segment: Grade 0, no subarachnoid blood; Grade 1, minimal subarachnoid blood; Grade 2, moderate blood clot with recognizable arteries; Grade 3, blood blot obliterating all arteries. Blood volume was evaluated by a total score ranging from 0 to 18 from six segments.

Tail bleeding assay

Tail bleeding time was determined as described previously⁵². A mouse was anesthetized with a mixture of ketamine and xylazine, and body weight was measured. The mouse was placed on a heating pad in prone position, the tail tip was about 4 cm blow the body horizon. A distal 5 mm segment of the tail was amputated, and the tail was immediately immersed in PBS pre-warmed at 37 °C. The time to complete arrest of bleeding was determined: complete arrest is no blood flow for 1 minute. The blood volume was determined by hemoglobin assay. Blood cells were separated by centrifuge at 4,000 r/min for 5 min at room temperature, and erythrocytes were resuspended in BD Pharm Lyse (BD Biosciences). After 10 min incubation in the buffer, the lysate was centrifuged at 10,000 rpm/min for 5 min. Hemoglobin concentrations were measured spectrophotometrically at 550 nm using a plate reader (Molecular Devices, SpectraMax M2e).

Mortality and neurological outcome

Mortality was assessed at 24, 48 and 72 hours after SAH induction. Gross neurological outcome was blindly evaluated before and at 24, 48 and 72 hours after surgery by sensorimotor scoring as described previously ^{53, 54}. Briefly, a motor score (0 to 12; spontaneous activity, limb symmetry, climbing, and balance) and a sensory score were used. Gross neurological outcome was evaluated by a total score of 4 to 24. Higher scores indicate better neurological outcome.

Brain water content

Brain edema was determined by using the wet/dry method as previously described ⁵⁵. Mice were sacrificed at 72 hours after surgery and the brains were quickly removed and separated into the left and right cerebral hemispheres and weighed (wet weight). The brain specimens were dried in an oven at 55°C for 72 hours and weighed again (dry weight). The percentage of water content was calculated as ([wet weight-dry weight]/wet weight) × 100%.

Cell death detection

DNA strand breakage during apoptosis after SAH was assessed by phospho-histon H2A.X (Ser 139) immunofluorescence as previously described ⁵⁶. Because phosphorylation of histon H2A.X at Ser 139 (γ-H2AX) is abundant, fast, and correlates well with each DNA strand breakage, it is the most sensitive marker that can be used to examine the DNA damage ⁵⁷. Mice were deeply anesthetized and perfused with cold PBS and subsequently with 4% PFA in PBS solution. The brains were removed, post-fixed with 4% PFA for 24 h and transferred to 30% sucrose solution. Frozen brains were cut with a 10 μm of thickness by a cryostat (Leica, CM3050 S). The brain slices were blocked with TBS-blocking solution (5% bovine serum albumin and 0.5% Tween 20 in PBS) for 1 hour at room temperature and incubated with phospho-histone H2A.X (Ser 139) (20E3) antibody (1:100; Cell Signaling) in blocking solution overnight on a shaker at 4 °C. Sections were washed three times with 0.5% Tween 20 in PBS and then incubated with goat anti-rabbit IgG Alexa-488 (1:200; Life Technologies, cat. no. A-11008). DAPI (40, 6-diamidino-2-phenylindole) staining was used as a counter staining. Sections were imaged by using an FluoView FV10i confocal microscope (Olympus, Japan) (original magnification, x 40). For quantification, the percentage of phospho-histon H2A.X positive cells per DAPI positive cells from three different fields in the ipsilateral cerebral cortex area per mouse (bregma −1.64 to −1.28 mm) were counted and the average values were plotted.

Assessment of blood brain barrier dysfunction

Small molecule (≤ 3 kDa) leakage from the blood into the brain parenchyma was assessed using dextran–tetramethylrhodamine (TMR) (3-kDa) dye (Thermo Fisher Scientific) Dextran-TMR (3-kDa) extravasation experiments were performed as described with modifications ^{2, 45, 58}. Mice were anesthetized and Dextran-TMR (3-kDa) (10 μg g⁻¹) was injected in the external jugular vein 22 hours after surgery. After 2 hours circulation, mice were deeply anesthetized and perfused with cold PBS to remove intravascular dye. The ipsilateral hemispheres were removed and homogenized in 1% Triton X-100 in PBS. The lysate was centrifuged at 15,000 r min⁻¹ twice for 15 min at 4 °C, and the supernatant was used to measure fluorescence (excitation/ emission = 540/590 nm, SpectraMax M2e, Molecular Devices). The relative fluorescence values (R.F.U.) were normalized by the brain weights.

To assess albumin extravasation, evans blue dye (EBD) was used because EBD binds to plasma albumin⁵⁹. Mice were anesthetized and 2% EBD (4 ml kg⁻¹) was injected in the external jugular vein 21 hours after surgery. After 3 hours circulation, mice were deeply anesthetized and perfused with cold PBS to remove intravascular dye. The ipsilateral hemispheres were removed and homogenized in 50% trichloroacetic acid in PBS. The lysate was centrifuged at 15,000 rpm min⁻¹ twice for 15 min at 4 °C, and the supernatant was used to measure fluorescence (excitation/ emission = 620/680 nm, SpectraMax M2e, Molecular Devices). The relative fluorescence units (R.F.U.) were normalized by the brain weights.

Statistical analysis

All results were expressed as mean ± SEM. Statistical analysis were performed with GraphPad Prism (GraphPad Software, Version 7.0c) by using two-tailed Student’s t-test, one-way analysis of variance (ANOVA) followed by Tukey’s test or two-way ANOVA followed by Bonferoni’s test. P values < 0.05 were considered statistically significant.

RESULTS

Abundant expression of S1PR1 in the mouse and human brain

We first investigated the expression of S1PR in brain, mouse primary neurons and mouse brain endothelial cells by reverse transcription and quantitative PCR analysis (RT-qPCR), as we have previously described ^{49, 60}. We found that S1PR1 is robustly expressed in the mouse brain in vivo, (Fig. 1A, 20.5± 2.4 copies/10⁶ 18S, which is equivalent to approximately 20.5± 2.4 S1PR1 mRNA copies/cell ⁶¹), as well as in the mouse brain endothelial cell line bEnd.3 ⁶² (Fig. 1B, 87.3±2.9 copies/cell), mouse primary neurons (Fig. 1C, 16.4±1.3 copies/cell) and mouse primary mixed glial cells (Fig. 1D, 8.3±0.15 copies/cell) in vitro. Then, we compared the expression of S1PR1 in cortical microvessels fragments⁴⁵ to its expression in total brain. Interestingly, we found that S1PR1 transcripts are highly enriched in cortical microvessels when compared to total brain (Fig. 1E, ~6 fold enrichment in microvessels vs whole brain).

Figure 1. S1PR1 mRNA is the most abundant S1PR transcript in brain, brain endothelial cells and neurons and it is highly enriched in brain microvessels.

S1PR mRNA levels in A) normal mouse brain, B) the mouse brain endothelial cell line bEnd.3, C) primary mouse neurons and D) primary mouse mixed glial cells. E) Comparison of S1PR1 mRNA levels in whole brain versus brain microvessels. Quantitative reverse transcription and polymerase chain reaction (RT-qPCR) demonstrates that S1PR1 transcript is predominant over the other S1PR and it is highly expressed in brain microvessels. B) Log scale. The individual values and the mean ± SEM are shown. N=3-6.*p<0.05.

Using the S1pr1-eGFP knock in mice, in which endogenous S1PR1 has been replaced by the S1PR1-eGFP fusion protein ⁴⁷, we found that S1PR1 is abundantly expressed in the cerebrovascular endothelium all throughout the brain (Fig. 2A, B, representative pictures from corpus callosum and cortex) as assessed by detection of S1PR1-eGFP and immunofluorescence (IF) analysis for the glucose transporter 1 (Glut-1). S1PR1 is also expressed in cortical (Supplementary Figure 1A) and hippocampal (Supplementary Figure 1B) neurons (co-staining with MAP-2) in vivo, both in the neuronal soma and the neuropil. In addition, S1PR1-eGFP is expressed in some GFAP positive cells, indicating that S1PR1 is also expressed in astrocytes (Supplementary Figure 1C, D). Similar pattern of expression was observed in the mouse brain when immunofluorescence analysis was conducted using a S1PR1 specific antibody previously validated in our laboratory ⁴⁹ (data not shown).

Figure 2. S1PR1 expression in the mouse brain in S1PR1-eGFP knock in mice.

S1PR1-eGFP fluorescence confocal analysis in the mouse brain. A-C) S1PR1-eGFP is expressed in the cerebrovascular endothelium. Immunofluorescence for Glut-1 (endothelial marker) is shown in the red channel. Two representative areas from the corpus callosum (A) and cortex (B, C) are shown.C) Selected region from B. Scale bar 20μm. Arrows point to endothelial cells with the brightest green signal. Representative pictures are shown. N=5-6.

In order to investigate the expression of S1PR1 in the human brain, we conducted IHC analysis in sections from the frontal lobe of a total of 5 brain autopsy samples. The S1PR1 antibody and IHC protocol used in human samples were previously validated in our laboratory ⁴⁹. We observed a very similar pattern of S1PR1 detection in these 5 samples. Representative pictures of the frontal cortex are shown in Figure 3. S1PR1 immuno positivity was observed both in the grey matter (GM) and white matter (WM) areas (Figure 3A, low magnification). S1PR1 was widely detected in the cerebrovascular endothelium of parenchymal vessels (in the grey matter and white matter, Fig. 3B, D-F) and pial vessels (Fig. 3C). In the grey matter, S1PR1 staining was observed in neuronal bodies and in the neuropil (Fig. 3B, C).

Figure 3. S1PR1 expression in the human brain.

Representative images of S1PR1 immunohistochemistry in 5 human brain autopsy samples. Formalin-fixed paraffin-embedded tissue sections were used for immmunohistochemical staining of S1PR1, as described in the Methods section. A) Expression of S1PR1 in the white and grey matter. Scale bar 500 μM. B) Representative picture showing S1PR1 immunopositivity in the neuropil, arterioles and capillaries of the grey matter. C) Detection of S1PR1 in the cerebrovascular endothelium of pial vessels. D-F) Representative pictures of the white matter. Notice S1PR1 positivity in parenchymal arterioles (D, F), venules (E) and capillaries (E,F). B-F) Scale bar 50 μM.

Altogether, our data indicate that, S1PR1 is the most abundant S1PR transcript in brain, brain endothelial cells and neurons. S1PR1 protein is also widely detected in the mouse and human brain, in the cerebrovascular endothelium and neurons. S1PR1 mRNA is highly enriched in mouse cerebral microvessels when compared to total brain.

Lack of S1pr1 in the endothelium results in worsened outcomes in the acute phase of SAH

In order to investigate the role of endothelial S1PR1 signaling in the pathophysiology of aneurysmal SAH, we used the endovascular rupture model of SAH⁴⁸, which is a well established and reproducible model that recapitulates key features of the pathophysiology of the acute phase of SAH. 24, 48 and 72 hours after SAH, cerebral microvessels were isolated from the mouse cortex of wild type mice to determine sphingosine 1-phosphate receptor 1 (S1PR1) mRNA and protein levels compared to sham mice. We found that S1PR1 mRNA levels in cortical microvessels did not significantly change compared to sham animals at 24, 48 or 72 h after SAH (Fig. 4A). Gfap mRNA levels in microvessels, which is induced in astrocytes upon inflammation, were significantly higher after SAH compared to sham (Fig. 4B, 3.07 ± 0,1, 4.05 ± 0.39 and 3.4 ± 0.56 fold induction at 24, 48 and 72 h respectively). The levels of endothelial (ZO-1, Fig. 4C), pericyte (CD13, Fig. 4D) or astrocyte end foot (Aqp4, Fig. 4E) markers did not significantly change suggesting similar cell composition among the microvessel preparations from these groups of mice. S1PR1 protein was widely detected by western blot assay in cerebral microvessels in sham animals (Supplementary Figure 2). Only a very modest increase in S1PR1 protein at 24 hours after SAH was observed (1.68 ± 0.19-fold). These data indicate that, in the acute phase of SAH, neurovascular inflammation is induced, assessed by increased GFAP expression (3-4 fold induction) but there were no significant changes in S1PR1 mRNA levels in cerebral microvessels. Only a modest increase in S1PR1 protein levels was observed 24 hours after SAH.

Figure 4. Increased mortality and worsened outcomes after SAH in endothelial specific S1pr1 null mice.

A-E) S1PR1, GFAP, ZO-1, CD13 and AQP4 mRNA levels in cerebral microvessels were quantified by RT-qPCR. F) S1pr1^flox/flox and S1pr1^iECKO mice were treated with tamoxifen (75 mg kg⁻¹) for the consecutive 3 days at the age of 8 weeks. S1pr1 mRNA expression levels were analyzed by qPCR in brain endothelial cells isolated from S1pr1^flox/flox and S1pr1^iECKO mice 2 weeks after tamoxifen treatment (n = 4). Deletion efficacy of endothelial specific S1pr1 in S1pr1^iECKO is shown relative to S1pr1^flox/flox mice. **P < 0.01, Student’s t-test. G) Survival curves after severe SAH surgery in S1pr1^flox/flox and S1pr1^iECKO mice (n = 10). H) Survival curves after mild SAH surgery S1pr1^flox/flox and S1pr1^iECKO mice (n = 13 or 19). G and H) Log-rank test, *P < 0.05. Individual values and mean ± s.e.m are shown. *P < 0.05, **P < 0.01, One-way ANOVA followed by Tukey’s test. Red line, S1pr1^iECKO; Blue line, S1pr1^flox/flox.

Given the abundant expression of S1PR1 in the cerebrovascular endothelium in the mouse and human brain, and the enrichment in S1PR1 mRNA in mouse cortical microvessels compared to total brain, we aimed to investigate the role of endothelial S1PR1 in early brain injury (EBI) after SAH. We generated endothelial-specific S1PR1 null mice (S1PR1 ^iECKO) by treating adult S1pr1^flox/flox×Cdh5–Cre^ERT2 mice (~2 month old) with 75 mg kg⁻¹ tamoxifen for 3 consecutive days (Fig.4F) as we have recently described⁴⁵. S1pr1^flox/flox littermate mice were also treated with tamoxifen and used as wild type control. We assessed the efficiency of deletion in S1PR1^iECKO by isolating brain endothelial cells from mice 2 weeks after tamoxifen treatment. RT-qPCR analysis demonstrated ~90% reduction of S1pr1 expression in S1pr1^iECKO mice compared to S1pr1^flox/flox littermate mice treated with tamoxifen in the same manner (Fig. 4F).

S1pr1^iECKO and S1pr1^flox/flox littermate mice were subjected to SAH surgery using a modified 5.0 suture (0.28 mm x 1.7 mm) to induce severe SAH as described in the method section ⁴⁸. The mortality rates were 10% in S1pr1^flox/flox mice at day 1, 2 and 3 after SAH. Interestingly, the mortality rates in endothelial-specific S1PR1 null mice (S1pr1^iECKO mice) were significantly higher compared to wild type (S1pr1^flox/flox,40% at day 1, and 60% at day 2 and 3, Fig. 4G).

Due to the higher mortality rates in S1pr1^iECKO mice, S1pr1^flox/flox and S1pr1^iECKO mice were subjected to a milder SAH model by changing the shape of the tip of the nylon suture to perforate the cerebral artery (0.28 mm diameter, as described in methods section) S1pr1^iECKO mice exhibited a trend towards a higher mortality rate (26.3% in day 1, 2 and 3) compared to S1pr1^flox/flox, which exhibited a mortality rate of 7.7% at day 1, 2 and 3(Fig. 4H), although it was not statistically significant. Given the high mortality rate in S1pr1^iECKO subjected to the severe SAH model (Fig. 4G), we used the milder SAH model for subsequent studies.

These data indicate that endothelial specific deletion of S1PR1 in mice results in a dramatic increase in mortality rates in the acute phase of SAH.

Genetic deletion of S1pr1 in the endothelium has no effect on hemostasis, blood volume in the subarachnoid space or CBF changes after SAH

Since the volume of subarachnoid blood directly correlates with worse outcomes after SAH ^63,51, we quantified the amount of subarachnoid blood upon SAH in S1pr1^iECKO and S1pr1^flox/flox mice by image analysis using a previously described SAH grading system ⁵¹. No significant differences were observed in the amount of subarachnoid blood between S1pr1^flox/flox and S1pr1^iECKO mice 24h after SAH (Fig. 5A, representative pictures). SAH grading was 11.50 ± 0.62 in S1pr1^flox/flox mice and 11.67 ± 0.42 in S1pr1^iECKO (Fig. 5B). In addition, we determined hemostasis using the tail bleeding assay ⁵². Consistent with the subarachnoid blood data, we did not find any significant differences in the bleeding times or blood volumes between S1pr1^flox/flox and S1pr1^iECKO mice in the tail bleeding assay. Bleeding times were 68.88 ± 4.98 seconds in S1pr1^flox/flox and 63.8 ± 4.14 seconds in S1pr1^iECKO (Fig. 5C). Hemoglobin contents, assessed by measuring the absorbance at 550 nm, were 0.73 ± 0.07 in S1pr1^iECKO mice compared to 0.70 ± 0.12 in S1pr1^flox/flox mice (Fig. 5D). Altogether these data indicate that hemostasis is not impaired in S1pr1^iECKO mice compared to wild type mice and that the amount of blood in the subarachnoidal space upon SAH is similar in wild type and S1pr1^iECKO mice.

Figure 5. Subarachnoid blood volume, hemostasis and CBF changes after SAH in S1pr1^flox/flox and S1pr1^iECKO mice.

A) Representative images of the ventral side of the brain depicting the basal cistern from S1pr1^flox/flox and S1pr1^iECKO mice at 24 hours after SAH. B) SAH grade was calculated by quantifying the blood in the subarachnoidal space by image analysis as described in methods section. (n = 6). C) and D) Tail bleeding assay for hemostasis: C) bleeding time and D) blood volume (n = 8). (B-D) Student’s t-test. Individual values and mean±SEM are shown. (E) CBF in the middle cerebral artery territory was measured by Laser-speckle contrast imager in in S1pr1^flox/flox and S1pr1^iECKO mice before (0 min.), 1 min, 10 min, 2 h, 4 h, 24 h after SAH induction. The relative CBF values (%) versus before SAH are shown. Data are mean ± SEM. (n = 6). Two-way ANOVA followed by Bonferoni’s test showed no statistically significant differences between wild type and S1pr1^iECKO. Red solid line, S1pr1^flox/flox ipsilateral side; blue solid, S1pr1^iECKO ipsilateral; light dotted line, S1pr1^flox/flox contralateral; light blue dotted line, S1pr1^iECKO contralateral. IL, ipsilateral side. CL, contralateral side.

We also determined cerebral blood flow (CBF) changes during SAH in both groups of mice by Laser-Speckle flowmetry. Cerebral blood flow in the middle cerebral artery (MCA) territory rapidly dropped ~1 minute after SAH in both the ipsilateral side (to 31.65 ± 3.46% of basal in S1pr1^iECKO and to 40.26 ± 4.09% of basal in S1pr1^flox/flox) and the contralateral side (to 82.42 ± 3.72% of basal in S1pr1^iECKO and to 77.32 ± 3.90% of basal in S1pr1^flox/flox). Then, CBF recovered to ~85% of basal 2 h after SAH in the ipsilateral side (to 87.46 ± 7.15% in S1pr1^iECKO versus to 82.83 ± 5.88% in S1pr1^flox/flox) and ~95 to 100% in the contralateral side (102.81 ± 1.97% in S1pr1^iECKO versus 95.34 ± 4.86% in S1pr1^flox/flox) both in S1pr1^iECKO and S1pr1^flox/flox mice (Fig. 5E). These data indicate that genetic deletion of S1pr1 in the endothelium did not significantly impact CBF changes during SAH. Finally, no significant differences were observed in arterial O₂ saturation, heart rate, pulse distention (a surrogate of pulse pressure) and respiratory rate between S1pr1^iECKO and S1pr1^flox/flox mice, before or after SAH (Table 1).

Altogether, these data indicate that endothelial-specific deletion of S1PR1 in adult mice did not have a significant impact on bleeding or clearance of subarachnoid blood, cerebral blood flow changes or systemic physiological parameters such as heart rate, respiratory rate, O₂ saturation or pulse distension.

Endothelial deletion of S1pr1 exacerbates brain edema and cell death after SAH leading to worsened neurological outcomes

In order to determine the impact of the lack of endothelial S1PR1signaling on brain injury after SAH, we assessed brain edema at 72 hours after mild SAH surgery by quantifying total brain water content. S1pr1^flox/flox mice exhibited a statistically significant increase in brain water content in the ipsilateral hemisphere (82.83 ± 0.53%) compared to sham (81.34 ± 0.20%) (Fig. 6A). Interestingly, in S1pr1^iECKO mice, total brain edema in the ipsilateral hemisphere was significantly exacerbated (84.39 ± 0.91%) compared to S1pr1^flox/flox (Fig. 6A). There was no increase in brain water content in contralateral hemisphere compared to sham.

Figure 6. Exacerbated cerebral edema and neuronal injury after SAH in endothelial specific S1pr1 null mice

A) 72 hours after SAH, brain edema was evaluated by brain water content in S1pr1^flox/flox and S1pr1^iECKO mice (n = 7 - 14). Brain water content was calculated as ([wet weight-dry weight]/wet weight) x 100. * p*0.05, one-way ANOVA followed by Tukey’s test. B) Immunofluorescence confocal analysis of phospho-histone H2A.X (DNA damage, green channel). Nuclear staining (DAPI) is shown in the blue channel. Representative images of ipsilateral hemisphere (cortex) from S1pr1^flox/flox and S1pr1^iECKO mice after SAH stained are shown. Scale bar, 50 μm. C) Quantification of phospho-histone H2A.X positive cells (%) (n = 7 or 8 mice). *P < 0.05, student’s t-test. The individual values and the mean ± SEM are shown. D) Neurological deficits after mild SAH surgery in S1pr1^flox/flox and S1pr1^iECKO mice (n = 12 or 14). From 4 to 24 points: 24 points (best), 4 points (worst). Data are mean ± s.e.m. *P < 0.05, **P < 0.01, One-way ANOVA followed by Tukey’s test. Red line, S1pr1^iECKO; Blue line, S1pr1^flox/flox.

Next, we analyzed cell death at 24 hours after SAH using phospho-histone H2A.X (Ser 139) immunostaining, a marker for DNA fragmentation resulted from apoptosis. We found that S1pr1^iECKO mice showed significantly higher number of phospho-histone H2A.X positive cells compared to S1pr1^flox/flox mice (50.85 ± 2.56% in S1pr1^iECKO versus 34.00 ± 2.98% in S1pr1^flox/flox) (Fig. 6B and C). No apoptotic cells were detected in sham animals.

Finally, we aimed to determine the impact of genetic deletion of S1PR1 in the endothelium on neurological outcomes after SAH. Neurological outcomes were determined by assessing motor and sensory function using a total scale of 4 to 24 (being 24 the best neurological outcome) as previously described^{53, 54}. Neurological outcomes at 24h and 48h after mild SAH surgery were worsened in S1pr1^iECKO mice (16.29 ± 1.37 and 17.14 ± 1.40, respectively) compared to S1pr1^flox/flox mice (20.91 ± 0.81 and 21.00 ± 0.83, respectively) (Fig. 6D).

These data indicate that genetic deletion of S1pr1 specifically in the endothelium significantly exacerbates total brain edema and cell death resulting in poorer neurological outcomes.

Endothelial specific deletion of S1pr1 in adult mice exacerbates blood brain dysfunction after SAH

Disruption of the BBB is an early event in the pathophysiology of neurovascular ischemic and hypoxic injury. Two distinct cellular mechanisms of BBB dysfunction have been described. Endothelial transcytosis ⁶⁴ (i.e. the transcellular pathway) ² allows the passage of high molecular weight molecules (e.g. albumin). In addition, small molecules can leak through the intercellular junctions (tight and adherens junction proteins, i.e. paracellular pathway) ^{58, 64–67}. In order to quantify BBB dysfunction upon SAH, we used two different tracers: Evans blue dye, which binds to plasma albumin (~66 kDa), and thus given its bigger size, it can leak into the brain parenchyma via transendothelial vesicle trafficking ⁶⁴ or after a severe loss of vascular integrity; tetramethylrhodamine (TMR)-labeled 3-kDa dextran which, due to its small molecular weight tracer can cross the BBB via the intercellular junctions (paracellular pathway) ^{2, 58}.

24 hours after SAH, albumin leakage, assessed by EBD extravasation assay, was significantly increased in S1pr1^flox/flox mice compared to sham animals (Fig 7A, 1.53 ± 0.06 fold in S1pr1^flox/flox SAH versus S1pr1^flox/flox sham). Albumin extravasation 24 after SAH was significantly higher in S1pr1^iECKO mice with regards to S1pr1^flox/flox (1.83 ± 0.08 in S1pr1^iECKO, Fig. 7A). Sham S1pr1^iECKO mice did not exhibit increased albumin leakage compared to sham wild type S1pr1^flox/flox litter mates (Fig. 7A). However, they did exhibit small molecule (3KDa TMR-dextran) BBB leakage (2.87 ± 0.45 fold in sham S1pr1^iECKO compared to sham S1pr1^flox/flox, Supplementary Fig. 3), consistent with our recently published studies ⁴⁵.

Figure 7. Endothelial specific S1pr1 null mice exhibit exacerbated BBB (albumin) leakage after SAH compared to wild type litter-mates.

A) Albumin BBB leakage, assessed by Evans Blue Dye extravasation, 24 hours after sham or SAH surgery in S1pr1^flox/flox and S1pr1^iECKO mice (n = 6 - 10). Evans blue dye was circulating for 3 hours. The individual values and mean ± SEM are shown. *P < 0.05, **P < 0.01, ***P < 0.001 (One-way ANOVA followed by Tukey’s test). y axis shows relative fluorescence units (R.F.U.) per gram of tissue. B) Increased phosphorylation of myosin light chain (p-MLC) in isolated cortical microvessels in S1pr1^iECKO mice compared to S1pr1^flox/flox. Cortical microvessels were isolated 24 h after SAH in S1pr1^iECKO and S1pr1^flox/flox mice. Western blot analysis for phospho-MLC (p-MLC) and β-actin was conducted. Individual values and mean ± SEM are shown. *P < 0.05 (t test).

Mechanistically, we found that exacerbated BBB dysfunction in endothelial-specific S1PR1 null mice, correlated with increased phosphorylation of myosin light chain (MLC) in isolated cortical microvessels (Fig. 6B, 6.96±1.6 fold in S1pr1^iECKO versus S1pr1^flox/flox), a downstream effector of the Rho-ROCK pathway implicated in endothelial inflammation and blood brain barrier dysfunction ^{58, 68–70}.

These data indicate that BBB dysfunction after SAH (albumin leakage) is heightened in mice lacking S1PR1 specifically in the endothelium compared to wild type and correlates with increased phosphorylation of MLC.

DISCUSSION

Increasing preclinical and clinical evidence indicate that BBB dysfunction correlates with exacerbated neuronal injury and plays a critical role in the pathophysiology of ischemic, hypoxic and inflammatory neurovascular injury in stroke ^5–13, traumatic brain injury^{71,72, 73} and other acute^{74, 75} and chronic ^76–79clinical conditions. Novel strategies to prevent BBB dysfunction may help minimize exacerbation of neuronal injury in these clinical settings. While the therapeutic potential of the endothelium is beginning to be recognized ^{25, 80, 81}, yet much research is needed to better understand the endothelial molecular mechanisms involved in the modulation of BBB function in these conditions.

Aneurysmal SAH, the most devastating type of stroke, occurs when an intracranial aneurysm ruptures. Compared to other types of stroke, SAH occurs earlier in life (40-60 years)⁸² and leads to higher mortality (50%) ⁸³. In addition, SAH survivors experience a high degree of disability and cognitive impairment (memory, language and executive function) ⁸⁴ with personal and societal consequences. SAH causes transient cerebral ischemia and accumulation of blood in the subarachnoid space, leading to brain injury, BBB dysfunction, cytotoxic and vasogenic cerebral edema in the acute phase ^{12, 85, 86}. Although numerous studies have reported the direct correlation between endothelial barrier dysfunction and worsened SAH outcomes in humans ^{87, 88} and mice ^{86, 89–94}, no direct causality regarding the contribution of BBB dysfunction to the exacerbation of neuronal injury has been shown. In this study, we investigated the role of the endothelium and BBB dysfunction in the aggravation of brain injury in the acute phase of SAH by using a well-established mouse model of aneurysmal SAH, the endovascular rupture model ⁴⁸. This experimental model faithfully recapitulates key features of the acute phase of SAH: upon rupture of the MCA, blood pours into the subarachnoid space, increasing intracranial pressure which gives rise to a brief period of transient (~3-5 minutes) cerebral ischemia ⁴⁸ (Fig. 5E, ~70%reduction of CBF in ipsilateral hemisphere). CBF slowly recovers over the time, but the brain remains hypoperfused in the acute phase of SAH (Fig. 5E, ~30% reduction vs basal). BBB breakdown, cerebral edema and neuronal injury ensue ^{89, 92}. In our study, we show that endothelial-specific genetic deletion of S1PR1 in adult mice exacerbates cerebral edema and neuronal injury, resulting in worsened outcomes in the acute phase of SAH. Thus, our study demonstrates the critical role of the endothelium in SAH outcomes and underscores the importance of a better understanding of the signaling pathways governing BBB function in the design of future therapeutic strategies targeting the endothelium to prevent the exacerbation of brain injury in this devastating condition.

S1P signaling via S1PR1 plays a critical role in endothelial function in humans and mice (reviewed in ²⁵). Activation of S1PR1 by S1P in the HDL fraction accounts for an important part of the cardiovascular protective effects of HDL^{24, 27–29}. S1PR1 also plays a critical role in lymphocyte function: mice lacking S1PR1 in hematopoietic cells are lymphopenic due to impaired egress from thymus and lymphoid organs³⁴. Interestingly, the immunosupressor FTY720-P, which has been approved by the FDA for multiple sclerosis ^{35, 36}, is a S1PR1 potent agonist. Given the phenotype of the mice lacking S1PR1 in hematopoietic cells³⁴ and in vitro studies which indicate that FTY720-P induces desensitization of S1PR1 ^{37, 38} it is accepted that FTY720-P, as well as other agonists ^{95, 96} and antagonists ⁹⁷ of S1PR1 lead to immunosuppression via inhibition of S1PR1 signaling. In experimental stroke, FTY720 and S1PR1 specific agonists, as other immunosuppressor drugs, have been shown to be protective^{39, 40,41, 42} and have even been tested in pilot stroke clinical trials ^98–100. Although the mechanisms are not completely clear, FTY720 protection has been attributed to its immunosuppressive effects ⁴² and its ability to desensitize S1PR1. Thus, the role of endothelial specific S1PR1 signaling in BBB function and stroke outcomes had remained elusive. We sought to address this knowledge gap by inducing the deletion of S1PR1 in adult mice, specifically in the endothelium ⁴⁶ and directly testing the role of endothelial S1PR1 signaling in BBB function and its impact on early brain injury upon SAH. Our data demonstrate that endothelial specific deletion of S1PR1 in adult mice exacerbates neuronal injury and worsens neurological outcomes in the acute phase of SAH, indicating that endothelial S1P signaling via S1PR1 is an endogenous protective signaling pathway that mitigates neurovascular ischemic/hypoxic/inflammatory injury.These data are consistent with previous studies from our laboratory and others which indicated that FTY720 exerts agonistic activity for S1PR1 in the endothelium and promotes endothelial barrier function ^23,25. Overall, our findings with the endothelial-specific S1PR1 mice highlight the therapeutic potential of this endothelial pathway in stroke. Novel future strategies to target this pathway specifically in the endothelium without compromising the immune response hold promise as novel vasoprotective therapeutic agents in stroke.

Mechanistically, we found that endothelial deletion of S1PR1 exacerbates BBB dysfunction (albumin leakage) upon SAH. BBB dysfunction heightens neurovascular injury by various mechanisms such as by allowing the entrance of neurotoxic plasma components into the brain parenchyma (e.g. albumin ¹⁰¹, fibrinogen ¹⁰²) and by compromising the ability of cerebral capillaries to deliver oxygen and nutrients to the neurons ^{7–9, 103}. In homeostatic conditions, the cerebrovascular endothelium, in cooperation with pericytes and astrocytes maintain a physical, metabolic and transport barrier to restrict the passage of molecules into the brain parenchyma ^{3, 104, 105}. In contrast to other organs, endothelial transcytosis (i.e. transcellular permeability) is very low in the brain¹⁰⁶. Crosstalk between the cerebrovascular endothelium and pericytes plays an important role in the inhibition of endothelial vesicular trafficking and expression of leukocyte adhesion molecules at the blood brain barrier ^2,107. In addition, the presence of tight junction proteins is also critical to maintain this physical barrier. Upon ischemic or hypoxic injury, BBB dysfunction starts early, primarily due to two cellular mechanisms, transcellular and paracellular permeability. The induction of gelatinase activity in cerebral microvessels ^{86, 92, 93, 108–111} leads to basal lamina degradation ^{7, 112}. In addition, pericyte and endothelial activation gives rise to increased endothelial transcytosis (transcellular permeability)^{2, 107}, allowing the passage of plasma proteins (e.g. albumin) into the brain parenchyma. Small plasma molecules can also leak via the intercellular junctions (i.e. paracellular permeability) ^{58, 64}, which become weakened by post-translational modifications (e.g. phosphorylation) and their disassembly from the actin cytoskeleton ¹¹³. With regards to molecular mediators and the signaling pathways governing these cellular processes leading to BBB dysfunction, generation of reactive oxygen species ¹¹⁴, NF-κB-mediated induction of MMP-9 ^{108, 115}, VEGF-induced increased vesicular trafficking and junctional destabilization ^{116, 117}, activation of Rho-ROCK pathway ^{58, 70} and increased angiopoietin 2/1 ratio ¹¹⁸ among others, have been proposed. We recently reported that endothelial deletion of S1PR1 in adult mice results in a defect in the assembly of tight junctions with the cytoskeleton in cerebral microvessels leading to BBB leakage of small molecules via the intercellular junctions (paracellular route) ⁴⁵. Interestingly, in healthy, non-challenged mice, this small molecule BBB leakage induced by endothelial deletion of S1PR1 in adult mice does not result in neuroinflammation or neurological deficits ⁴⁵. However, upon SAH challenge, BBB dysfunction (albumin leakage) and neurovascular injury were exacerbated in mice lacking S1PR1 specifically in the endothelium. At the molecular level, exacerbated BBB dysfunction in S1PR1 endothelial specific mice correlated with increased levels of phosphorylated MLC, a downstream effector of the Rho-ROCK pathway implicated in endothelial inflammation and blood brain barrier dysfunction ^{58, 68–70}. These data indicate that in endothelial S1pr1 null mice, increased p-MLC levels, possibly due to exacerbated Rho/ROCK signaling, correlates with the aggravation of BBB dysfunction and worsened outcomes upon SAH.

Other possible vascular mechanisms responsible for the worsened outcomes in endothelial specific S1pr1iECKO mice vs wild type could be differences in cerebral blood flow or in bleeding upon cerebral artery rupture. However, we did not observe differences in the CBF changes or physiological parameters during and after SAH between these two groups of mice. In addition, the amount of blood in the subarachnoidal space (SAH grade), which plays an important role in the severity of brain injury and SAH outcomes was similar in both groups of mice, indicating that there were no differences in bleeding or the clearance of the subarachnoidal blood. Furthermore, there were no differences in platelet function, assessed by the tail-bleeding assay, between wild type and S1pr1iECKO mice. Thus, the exacerbation of neuronal injury upon SAH in the endothelial specific s1pr1 null mice vs wild type cannot be attributed to differences in bleeding or in cerebral blood flow regulation. Altogether our data points to exacerbation of BBB dysfunction (albumin leakage) as the main cellular mechanism underlying the aggravation of brain injury upon endothelial deletion of S1PR1 in the adult mice, providing proof of concept of the critical role of endothelium and blood brain barrier dysfunction in the aggravation of brain injury in stroke.

Lastly, our study also underscores the pathophysiological relevance of the S1P pathway in humans. Using RT-qPCR in combination with mouse genetic approaches and IHC techniques previously validated in our laboratory ⁴⁹, we found that S1PR1 is the most abundant S1PR transcript in the mouse brain and mouse brain microvessels and widely detected in the human cerebrovascular endothelium and brain parenchyma. Our human data, together with our mouse studies using the endothelial-specific S1PR1 null mice, highlight the therapeutic and prognostic potential of the endothelial S1P pathway in stroke. Given recent human studies which indicate that S1P-S1PR1 vasoprotective signaling ^{24, 27} may be limiting in cardiovascular and inflammatory diseases ^{25, 28–31} our present data imply that patients with lower levels of ApoM or HDL-bound S1P and thus, having limiting endothelial S1PR1 antiinflammatory signaling, could be at risk of worsened outcomes upon stroke. In addition, our data could help explain the adverse effects reported in some subsets of multiple sclerosis patients in chronic treatment with FTY720, such as macular edema or posterior reversible encephalopathy syndrome, among others ^119–122.

In summary, our study demonstrates the critical role of the endothelial-specific S1PR1 signaling in BBB function and SAH outcomes in mice. Our human data strengthen the pathophysiological relevance of these findings and highlight the therapeutic and prognostic potential of the endothelium, more specifically the endothelial S1P signaling pathway in stroke. New strategies to modulate S1P signaling specifically in the endothelium to prevent exacerbation of brain injury without compromising the immune response ^{25, 123, 124} could hold promise as novel neurovascular protective therapies in stroke and other conditions associated with BBB dysfunction.

ACKNOWLEDGEMENTS

This work was supported by internal funds provided by the Departments of Surgery and Emergency Medicine, Beth Israel Deaconess Medical Center, the Department of Pathology and Laboratory Medicine, Weill Cornell Medicine, American Heart Association Grant-in-Aid 12GRNT12050110, NIH HL094465 and Leducq Foundation grants to TS.

We would like to thank Dan Li and Dr. Shou-ching Jaminet from the Multi-Gene Transcriptional Profiling Core Facility (Center for Vascular Biology Research, Beth Israel Deaconess Medical Center, Harvard Medical School) for the quantitative PCR analysis, Drs. Louise McCullough (University of Texas Health McGovern Medical School) and Tim Hla (Boston Children’s Hospital) for their input in the project.

The authors declare no competing financial or non-financial interests.