Introduction
The mammalian brain is submerged in cerebrospinal fluid (CSF), the majority of which is continuously produced by the choroid plexus, a monolayered epithelial structure that protrudes from the ventricular wall [
1,
2]. The CSF serves crucial roles such as cushioning the brain and thus protecting it from mechanical insult [
3] in addition to acting as the transport route for delivery/disposal of nutrients, metabolites, and immune modulators [
4]. Dysregulation of CSF dynamics occurs in a range of neuropathologies, among others posthemorrhagic hydrocephalus (PHH) [
5‐
7], in which water accumulates in the brain after the hemorrhagic event, causing ventriculomegaly and elevation of the intracranial pressure (ICP). PHH may arise secondary to spontaneous or traumatic intracranial hemorrhage in all age groups [
8,
9], particularly frequently following hemorrhage in the subarachnoid and ventricular compartments. Such pathological posthemorrhagic brain water accumulation is generally attributed to impaired clearance of CSF caused by obstruction of CSF flow pathways. Currently, the only available treatment relies on drainage of the excess fluid from the brain via surgical CSF diversion through external ventricular drainage, ventricular shunt placement or ventriculostomy [
10,
11]. These surgical procedures associate with frequent side effects, i.e. infections and shunt malfunctions, which necessitate alternative treatment options. However, pharmacological interventions have proven suboptimal [
12,
13], partly due to the knowledge gap concerning the molecular mechanisms of CSF secretion and their regulatory properties in health and disease.
The serum lipid lysophosphatidic acid (LPA) [
14] enters the brain with hemorrhagic events, such as during traumatic brain injury and a murine model thereof [
15]. LPA appears to worsen the outcome in the process [
15], as it does in neonatal mice models [
16,
17]. These mouse models of (embryonic or postnatal) PHH displayed a range of LPA-mediated morphological changes in the week(s) following intracranial LPA administration, such as ventricular enlargements, 3rd ventricular occlusion, elevated ICP, thinning of cortical layers, and cilia loss along the lateral ventricular walls [
16,
17]. LPA effectuated these features via their G protein-coupled LPA receptors expressed in the brain tissue (LPA
R1-LPA
R6) [
16‐
18] and thus appear to act as a key factor in the etiology of PHH.
Emerging evidence suggests a component of CSF hypersecretion in some forms of hydrocephalus in patients and experimental animal models, i.e. choroid plexus hyperplasia, choroid plexus tumours, or PHH [
5,
6,
19]. In the rodent experimental model of intraventricular hemorrhage (IVH), the PHH was demonstrated to occur following CSF hypersecretion [
6]. The molecular coupling between the hemorrhagic event and the hydrocephalus formation was proposed to rely on the associated inflammatory response activating the Toll-like Receptor 4 (TLR4) signalling pathway, which ultimately promoted hyperactivity of the choroidal Na
+/K
+/2Cl
− cotransporter NKCC1 [
6], that serves as a key contributor to CSF formation by rodent and canine choroid plexus [
6,
20‐
22]. Hemorrhage-induced CSF hypersecretion may, in addition, be promoted by the LPA entrance occurring with the brain insult.
The choroid plexus epithelial cells abundantly express the Ca
2+-permeable nonselective cation channel transient receptor potential vanilloid 4 (TRPV4) [
23,
24]. TRPV4 is polymodal in a sense of its several distinct manners of activation, one of which being lipid-mediated modulation of channel activity [
25]. TRPV4 has been implicated in CSF secretion following demonstration of its ability to modulate transepithelial ion flux in choroid plexus cell lines [
26,
27] and of TRPV4 antagonists effectively alleviating ventriculomegaly in a genetic animal model of hydrocephalus [
28]. TRPV4 could thus act as the molecular link coupling the hemorrhage-mediated LPA elevation to the CSF hypersecretion and ensuing hydrocephalus.
Here, we demonstrate that LPA is indeed elevated in patients with subarachnoid haemorrhage (SAH) and in an animal model of IVH and reveal its ability to directly modulate the ion channel TRPV4, which subsequently promotes NKCC1-mediated CSF hypersecretion and ventriculomegaly.
Methods
Patients
CSF samples were collected between June 2019 and September 2021 from 12 patients (mean age: 63 y, range: 40–77 y, 7 F/ 5 M) with acute SAH admitted and treated for the condition at Department of Neurosurgery at Rigshospitalet, Copenhagen, Denmark. CSF samples were obtained within 48 h of ictus (n = 12, see Additional file
1: A for LPA levels as a function of time from ictus to CSF sampling) through an external ventricular drain (EVD) inserted on clinical indications. To exclude that measured CSF parameters could be affected by neuroinfections requiring antibiotic treatment, patients with no signs of neuroinfection at admission or during their treatment were selected. All included patients later received a permanent ventriculo-peritoneal shunt because of continued need for CSF diversion. As control subjects, 14 patients undergoing preventive surgery for unruptured aneurysms (vascular clipping) were enrolled (mean age: 61 y, range: 39–71 y, 8 F/6 M), and CSF was collected from the basal cisterns during surgery prior to clipping of the aneurysm. CSF was collected in polypropylene tubes (Sarstedt), centrifuged at 2000×
g for 10 min at 4 °C, split into aliquots, and stored at − 80 °C no later than 2 h after collection and until quantification of LPA (see below). Written informed consent were obtained from all patients or next of kin depending on the capacity of the patients and the study was approved by the Ethics committee of the Capital Region of Denmark (H-19001474 /69197/H-17011472).
Animals
All animal experiments complied with the relevant ethical regulations and conformed to European guidelines. The Danish Animal Experiments Inspectorate approved the experiments (permission no. 2016-15-0201-00944 and 2018-15-0201-01595). Adult male Sprague Dawley rats (Janvier Labs) of 9 weeks used for the animal experimentation were housed with a 12:12 light cycle and access to water and food ad libitum accordingly to the ARRIVE guidelines. Animals were ordered weekly from the supplier for a specific set of experiments and randomized within the group and evenly distributed to each of the experimental conditions.
Anaesthesia and physiological parameters
The experimental animals were anaesthetized with intraperitoneal (i.p.) injection with 6 mg ml− 1 xylazine + 60 mg ml− 1 ketamine (ScanVet) in sterile water (0.17 ml per 100 g body weight, pre-heated to 37 °C). Animals were re-dosed with half ketamine dose as required to sustain anesthesia. Animals were excluded if they did not respond to the anaesthesia regimen immediately (for the live imaging of the CSF secretion experiments, where the animals were not ventilated) or within the first three re-dosings (for the ventriculo-cisternal perfusion assay (see below) and ICP measurements, where the animals were mechanically ventilated). Such events occurred rarely. Isofluorane (1000 mg g−1, ScanVet) was employed for survival procedures (mixed with 1.8 1 min− 1 air/0.1 1 min− 1 O2 (Attene vet), 5% to induce anesthesia and 1–2.5% to sustain anesthesia. During magnetic resonance scanning of the animals, anesthesia was maintained at 1–1.5% isoflurane in a 1:1 mixture of air:O2. The body temperature of the anesthetized rats was maintained at 37 °C by a homeothermic monitoring system (Harvard Apparatus). Mechanical ventilation was included for anesthetic protocols longer than 30 min to ensure stable respiratory partial pressure of carbon dioxide and arterial oxygen saturation and thus stable plasma pH and electrolyte content. A surgical tracheotomy was performed and the ventilation controlled by the VentElite system (Harvard Apparatus) by 0.9 l min− 1 humidified air mixed with 0.1 l min− 1 O2 adjusted with approximately 3 ml per breath, 80 breath min− 1, a Positive End-Expiratory Pressure (PEEP) at 2 cm, and 10% sigh for a ~ 400 g rat. The ventilation settings were optimized for each animal using a capnograph (Type 340, Harvard Apparatus) and a pulse oximeter (MouseOx® Plus, Starr Life Sciences) after system calibration with respiratory pCO2 (4.5–5 kPa), pO2 (13.3–17.3 kPa), and arterial oxygen saturation (98.8–99.4%) (ABL90, Radiometer). For survival procedures, the rats were preoperatively given the analgesics buprenorphine p.o. (0.4 mg kg− 1, Sandoz) and carprofen subcutaneously (5 mg kg− 1, Norbrook). The former was re-administered 24 h postoperatively.
Solutions and chemicals
The majority of the experiments were conducted in HCO3−-containing artificial cerebrospinal fluid (aCSF; (in mM) 120 NaCl, 2.5 KCl, 2.5 CaCl2, 1.3 MgSO4, 1 NaH2PO4, 10 glucose, 25 NaHCO3, pH adjusted with 95% O2/5% CO2). In experiments where the solution could not be equilibrated with 95% O2/5% CO2, during the experimental procedure (intracranial pressure monitoring and RNA sequencing), the solution was instead buffered by HEPES (HEPES-aCSF; (in mM) 120 NaCl, 2.5 KCl, 2.5 CaCl2, 1.3 MgSO4, 1 NaH2PO4, 10 glucose, 17 Na-HEPES, adjusted to pH 7.4 with NaOH). Pharmacological inhibitors were dissolved in DMSO and kept as stock solutions at − 20 °C. These were either purchased from Sigma (bumetanide: B3023, GSK1016790A: G0798, RN-1734: R0658, lysophosphatidic acid: L7260, Closantel: 34093) or MedChemExpress (WNK463: HY-100,828). All control solutions contained appropriate concentrations of vehicle (DMSO, D8418, Sigma), which amounted to 0.05–0.1% DMSO.
Experimental intraventricular hemorrhage (IVH) in rats
The surgery was performed on rats anesthetized with isoflurane under aseptic conditions with body temperature maintained at 37 °C using a rectal probe and feedback-controlled heating pad (Harvard Apparatus). Rats were positioned in a stereotaxic frame (Harvard Apparatus) and the skull exposed with a midline incision. A cranial burr hole was drilled above the right lateral ventricle (0.6 mm posterior and 1.6 mm lateral to bregma), after which the rats were removed from the stereotaxic frame, the femoral artery catheterized, and approximately 300 µl blood was collected (the control rats underwent sham operation). Immediately thereafter, 200 µl of this autologous blood sample (or saline) was manually injected over the course of 15 min via a 27-gauge needle inserted stereotaxically into the burr hole in the right lateral ventricle (4.5 mm ventral) [
29] and expected to spread throughout the ventricular system containing approximately 180 µl CSF in rats of the chosen species and age (181 ± 18 µl, n = 3, quantified by MRI, see below). The needle was kept in place for 5 min before retraction to prevent backflow. The skin incisions were closed with sutures and the rats were allowed to recover before returning to the housing facility.
CSF extraction and alpha-LISA
24 h post-IVH surgery, the rats were anesthetized and placed in a stereotaxic frame. CSF was sampled from cisterna magna with a glass capillary (30-0067, Harvard Apparatus pulled by a Brown Micropipette puller, Model P-97, Sutter Instruments) placed at a 5° angle (7.5 mm distal to the occipital bone and 1.5 mm lateral to the muscle-midline). CSF was collected in polypropylene tubes (Sarstedt), centrifuged at 2000×g for 10 min at 4 °C, divided into aliquots, and stored at − 80 °C. The animals were excluded from the study if blood was detected in the CSF obtained from the saline control animals (occurred in one rat). The LPA content of the CSF samples was determined with alpha-LISA by use of a ready-to-use microwell kit designed to detect native LPA in either rats (MBS774994, MyBioSource) or humans (MBS707296, MyBioSource). The CSF samples were added to wells pre-coated with LPA antibody, followed by addition of streptavidin-horseradish peroxidase to form an immune complex. Upon a wash step, chromogen substrate solutions were added and plate reading conducted in a microplate photometer (Synery™ Neo2 Multi-mode Microplate Reader; BioTek Instruments) according to the manufacturer’s instructions.
Determination of brain water content following intraventricular LPA exposure
25 µl LPA (100 µM) was delivered intraventricularly into anesthetized rats (sham operated rats were used as comparison) as described for the blood delivery in the IVH procedure. This concentration was chosen based on the expected (97%) loss of LPA following lipid interactions with pipettes, tubes and syringe surfaces in the absence of a lipid-carrier protein [
16]. In addition, a single bolus injection into one lateral ventricle will be diluted in the entire ventricular system (of around 180 µl) and gradually washed away from the choroid plexus surface by the sustained CSF secretion in the 24 h interval between injection and experiment. We thereby expect a substantially lower LPA concentration at the surface of the choroid plexus over the 24 h period. 24 h post-injection, the rat was sacrificed and its brain swiftly removed, placed in a pre-weighed porcelain evaporating beaker (Witeg) and weighed within 1 min after brain isolation. The brain tissue was homogenized with a steel pestle and dried at 100 °C for 72 h to a constant mass. The dry brain was weighed, and the brain water content determined in ml/gram dry weight using the equation: (wet weight - dry weight)/dry weight. The weighing was done in a randomized and blinded fashion.
Magnetic resonance imaging (MRI)
Anesthetized rats underwent MRI in a 9.4 Tesla preclinical horizontal bore scanner (BioSpec 94/30 USR, Bruker BioSpin) equipped with a 240 mT/m gradient coil (BGA-12 S, Bruker) at the Preclinical MRI Core Facility, University of Copenhagen. The scanner was interfaced to a Bruker Avance III console and controlled by Paravision 6.1 software (Bruker). Imaging was performed with an 86 mm-inner-diameter volume resonator and a 4-channel surface quadrature array receiver coil. The animal body temperature was maintained at 37 ± 0.5 °C with a thermostatically controlled waterbed and its respiratory rate monitored by an MR-compatible monitoring system (SA Instruments). The imaging protocol consisted of T
2-weighted 2D rapid acquisition with relaxation enhancement (2D-RARE) for reference spatial planning with the following settings: repetition time (TR) = 4000 ms, effective echo time (TE) = 60 ms, number of averaging (NA) = 4, RareFactor = 4, slice thickness = 500 μm, in-plane resolution = 137 × 273 μm, 25 coronal slices, total acquisition time (TA) = 8.5 min. For obtaining high resolution CSF volumetry, a 3D constructive interference steady-state sequence (3D-CISS) [
30] image was calculated as a maximum intensity projection (MIP) from 4 realigned 3D-TrueFISP volumes with 4 orthogonal phase encoding directions (TR = 4.6 ms, TE = 2.3 ms, NA = 1, Repetitions = 2, Flip angle = 50°, 3D spatial resolution 100 × 100 × 100 μm, RF phase advance 0, 180, 90, 270°, TA = 28 min). To obtain optimal spatial uniformity, all acquired 3D-TrueFISP volumes were motion-corrected before calculation as MIP, and the image bias field was removed with Advanced Normalization Tools (ANTs) [
31,
32]. For each brain sample, the total brain volume was automatically segmented by using region growing with ITK-snap (version 3.8.0) [
33]. In addition, the pixel intensity factorized semi-automatic thresholding was performed to segment the lateral ventricle in each hemisphere. The volume measurement of the whole brain and lateral ventricles were performed in ITK-snap. The analysis was carried out in a blinded fashion.
ICP monitoring
A burr hole was drilled above the right lateral ventricle (using the coordinates 1.3 mm posterior to Bregma, 1.8 mm lateral to the midline, and 0.6 mm ventral through the skull) of anesthetized and ventilated rats placed in a stereotaxic frame, in which a 4 mm brain infusion cannula (Brain infusion kit 2, Alzet) was placed. On the contralateral side of the skull, an epidural probe (PlasticsOne, C313G) was placed in a cranial window (4 mm diameter) and secured with dental resin cement (Panavia SA Cement, Kuraray Noritake Dental Inc.). The cannula was pre-filled with HEPES-aCSF and connected to a pressure transducer (APT300) and a transducer amplifier module TAM-A (Hugo Sachs Elektronik). The pressure signal was visualized and recorded with a sample rate of 1 kHz using BDAS Basic Data Acquisition Software (Harvard Apparatus, Hugo Sachs Elektronik). Jugular compression was applied in the beginning and at the end of the experiment to confirm proper ICP recording. Pre-heated aCSF (containing DMSO or 1.8 mM RN1734; expected ventricular concentration 50 µM) was slowly infused (0.5 µl min− 1) into the lateral ventricle.
CSF production rate
The CSF production rate was determined with the ventriculo-cisternal perfusion (VCP) technique. An infusion cannula (Brain infusion kit 2, Alzet) was stereotaxically placed in the right lateral ventricle of an anesthetized and ventilated rat (using the same coordinates as for ICP), through which a pre-heated (37 °C, SF-28, Warner Instruments) dextran-containing solution (HCO
3−-aCSF containing 1 mg ml
− 1 TRITC-dextran (tetramethylrhodamine isothiocyanate-dextran, MW = 155 kDa; T1287, Sigma) was perfused at 9 µl min
− 1. CSF was sampled at 5 min intervals from cisterna magna as described above for CSF extraction. The cisterna magna puncture and associated continuous fluid sampling prevents elevation of the ICP during the procedure. The dilution of the infused solution is assigned to endogenously secreted CSF, irrespective of origin of this fluid [
20]. 60 min into the experiment, the control test solution was replaced with one containing either TRPV4 activator (GSK, 500 nM) or TRPV4 inhibitor (RN1734, 50 µM), with an expected 2-fold ventricular dilution (more in the contralateral ventricle) followed by continuous ‘wash-away’ from the choroid plexus surface by the sustained CSF secretion. Continuous (24 h) exposure to high concentrations of GSK have been reported to be toxic to cultured cells [
34], and we therefore verified cell viability and morphological integrity of acutely excised choroid plexus exposed to 500 nM GSK for the 1 h that mimics the VCP experimental procedure (Additional file
1: B). The fluorescent content was measured in a microplate photometer (545 nm, Synery™ Neo2 Multi-mode Microplate Reader; BioTek Instruments), and the production rate of CSF was calculated from the equation:
$$Vp=ri* \frac{Ci-Co}{Co}$$
where
Vp = CSF production rate (µl min
− 1),
ri = infusion rate (µl min
− 1),
Ci = fluorescence of inflow solution,
Co = fluorescence of outflow solution, calculated based on stable time intervals from 50 to 65 min and 100–120 min.
Live imaging of CSF movement
Through a burr hole in the skull of the anaesthetized rat (same coordinates as for ICP and VCP), a Hamilton syringe (RN 0.40, G27, a20, Agntho’s) was placed (4 mm deep) with 15 µl HCO3−-aCSF containing vehicle (DMSO) or drug of interest (TRPV4 activator, 1 µM GSK; TRPV4 inhibitor, 100 µM RN1734, expected to be immediately diluted in ∼180 µl native CSF). The content was injected during 10 s to target both lateral ventricles. This first injection was intended to allow the inhibitors to act on their target transporters prior to introduction of the fluorescent dye. The procedure was repeated 5 min later with identical syringe content, but with addition of the carboxylate dye (MW = 1,091, IRDye 800 CW, P/N 929-08972, LI-COR Biosciences) to the syringe content (10 µM of carboxylate dye). The rat was swiftly placed in a Pearl Trilogy Small Animal Imaging System (LI-COR Biosciences) and within 1 min after last ventricular injection, images were obtained at 30 s intervals (800 nm channel, 85 μm resolution, for 5 min). A white field image was taken at the termination of each experiment, after which the rat was sacrificed. The isolated brain was split into the two hemispheres and placed on a coverslip to record a final micrograph to ensure proper targeting of the ventricular compartment (800 nm channel). Images were analysed using LI-COR Image Studio 5.2 (LI-COR Biosciences) and data presented as fluorescence intensity in a region of interest placed in line with lambda, normalized to signals in the first image. Analyses were done in a randomized and blinded fashion.
RNASeq of rat choroid plexus
Isolated rat choroid plexus (lateral and 4th) were stored in RNAlater (R0901, Sigma) at − 80 °C prior to RNA extraction and library preparation (performed by Novogene Company Limited with NEB Next® Ultra™ RNA Library Prep Kit (NEB)) prior to RNA sequencing (paired-end 150 bp, with 12 Gb output) on an Illumina NovaSeq 6000 (Illumina). Quality control and adapter removal was done with Novogene. The 150 base paired-end reads were mapped to rat reference genome Rnor_6.0.103 (
Rattus norvegicus) using Spliced Transcripts Alignment to a Reference (STAR) RNA-seq aligner (v.2.7.2a) [
35]. The mapped alignment generated by STAR was normalized to transcripts per million (TPM) [
36] with RSEM (v 1.3.3) [
37]. The RNAseq data obtained from this choroid plexus tissue has also been employed for other analysis [
38]. Gene information was gathered with mygene 3.1.0 python library [
39,
40] (
http://mygene.info) where GO terms [
41‐
43] for cellular components were extracted. The list of genes annotated as “Voltage-gated ion channels”, “Ligand-gated ion channels”, and “Other Ion channel” was obtained from the Guide to Pharmacology webpage (
https://www.guidetopharmacology.org/download.jsp) [
44] and employed to generate a list of plasma membrane ion channel proteins (in which ion channels in intracellular membranes were excluded). The list of rat kinases was obtained from the Kyoto Encyclopedia of Genes and Genomes (KEGG) database (
https://www.kegg.jp/kegg/genes.html) [
45], with organism specific filtering “rno” (Rattus norvegicus), for entries of ‘EC 2.7.10.2’ (non-specific protein-tyrosine kinase), ‘EC 2.7.12’ (Dual-specificity kinases) with the two sub-categories, and ‘EC 2.7.11’ (Protein-serine/threonine kinases) with the 33 sub-categories. Scripts and program parameters can be found at
https://github.com/Sorennorge/MacAulayLab-RNAseq1. Network analysis was carried out with the string database (
string-db.org) [
46] with the search criteria: Multiple proteins in organism ‘Rattus norvegicus’: TRPV4, SLC12A2 (NKCC1), WNK1, WNK3, WNK4, Stk39 (SPAK).
Tissue preparation and immunohistochemistry
Anesthetized rats were perfusion fixed with 4% paraformaldehyde and the brains removed for post-fixation in the same fixative overnight, cryoprotected in 25% sucrose, and frozen on solid CO2. Saggital and coronal sections (16 μm) were cut and stored at − 20 °C. Blockage was done with 10% swine serum diluted in PBS + 1%Tween-20 (PBST), and the sections were immunolabeled with primary rabbit anti-NKCC1 (Abcam #AB59791, 1:400) and mouse anti-TRPV4 (BD BioSciences #AB53079, 1:500) overnight at 4 °C, and Alexa FluorTM488 and Alexa FluorTM647 (ThermoFisher Scientific 1:500) for 2 h at room temperature. Finally, the sections were mounted with ProLong Gold DAPI mounting medium (Dako). The confocal images were acquired with a Zeiss LSM710 point laser (Argon Lasos RMC781272) scanning confocal microscope with a Zeiss Plan-Apochromat 63×/numerical aperture (NA) 1.4 oil objective (Carl Zeiss, Oberkochen). All micrographs were sampled in a frame scan mode.
Proximity ligation assay
Proximity ligation assay (PLA) by indirect detection was done using Duolink reagents and instructions (Sigma-Aldrich) on rat brain cryosections (16 μm) (see ‘Tissue preparation’ section). After incubation in Duolink blocking solution, the tissue sections were incubated with anti-TRPV4 (LS-Bio #LS-C94498, 1:400) and anti-NKCC1 (Dundee University #S022D, 2 µg/ml) overnight at 4 °C. The secondary antibodies were conjugated to a MINUS and a PLUS PLA probe, and were detectable as fluorescent speckles only upon close contact [
47]. A technical control omitting both of the primary antibodies was included. Ligation and amplification of the samples were done according to the Duolink protocol, the sections were mounted with coverslips using the Duolink In Situ Mounting Medium with DAPI (Sigma-Aldrich). Micrographs were acquired with a Zeiss LSM700 point laser (Argon Lasos RMC781272) scanning confocal microscope with a Zeiss Plan-Apochromat 20×/numerical aperture (NA) 1.6 oil objective (Carl Zeiss, Oberkochen). All micrographs were sampled in a frame scan mode.
86Rb+efflux experiments
Acutely isolated lateral choroid plexuses were placed in cold HCO
3--aCSF but allowed to recover at 37 °C for 5–10 min before the experiment. Choroidal isotope accumulation was performed by a 10 min incubation in gas-equilibrated HCO
3--aCSF with 1 µCi ml
−1 86Rb
+ (NEZ07200, Polatom) and 4 µCi ml
−1 3H-mannitol (NET101, extracellular marker, PerkinElmer). The choroid plexus was briefly washed (15 s) prior to incubation in 0.5 ml efflux medium (HCO
3--aCSF containing the indicated modulators; 20 µM bumetanide (NKCC1 inhibitor), 100 nM GSK (TRPV4 agonist), 50 µM RN (TRPV4 inhibitor), 20 µM Closantel (SPAK kinase-inhibitor), 20 nM WNK463 (WNK kinase-inhibitor), 25 µM LPA, or vehicle (DMSO). 0.2 ml of the efflux medium was collected into scintillation vials at 10 s time intervals and replaced with fresh HCO
3--aCSF. Upon termination of the experiment, the choroid plexuses were dissolved in 1 ml Solvable (6NE9100, PerkinElmer) and the isotope content determined by liquid scintillation counting with Ultima Gold
™ XR scintillation liquid (6,013,119, PerkinElmer) in a Tri-Carb 2900TR Liquid Scintillation Analyzer (Packard). The choroid plexus
86Rb
+ content corrected for
3H-mannitol content (extracellular background) was calculated for each time point, and the natural logarithm of the choroid plexus content
At/
A0 was plotted against time. Slopes indicating the
86Rb
+ efflux rate constants (min
− 1) were determined from linear regression analysis [
20,
48].
Electrophysiological recordings in heterologously expressing Xenopus laevis oocytes
Defolliculated oocytes obtained from
Xenopus laevis were purchased from Ecocyte Bioscience. cRNA (TRPV4) was prepared from linearized plasmids (pXOOM containing cDNA encoding TRPV4) using the mMESSAGE mMACHINE T7 kit (Ambion) and extracted with MEGAclear (Ambion), according to the manufacturer’s instructions. The oocytes were microinjected with 4 ng cRNA per oocyte with a Nanoject microinjector (Drummond Scientific Company). The oocytes were kept at 19 °C for 3 days in Kulori medium prior to experiments. Oocytes were kept in Kulori medium (90 mM NaCl, 1 mM KCl, 1 mM CaCl
2, 1 mM MgCl
2, 5 mM HEPES (pH 7.4)) with inclusion of ruthenium red until the day of experiments (100 μm; Sigma Aldrich, R-2751) to suppress TRPV4 activity and ensuing cell death [
49]. Conventional two-electrode voltage-clamp was performed using a DAGAN CA‐1B High Performance oocyte clamp (DAGAN, Minneapolis, MN, USA) with a Digidata 1440 A interface controlled by pCLAMP software, version 10.5 (Molecular Devices, Burlingame, CA, USA). Borosilicate glass capillaries were used to pull electrodes (PIP5; HEKA Elektronik, Lambrecht, Germany) with a resistance of 1.5–3 MΩ when filled with 1 M KCl. Oocytes were placed in an experimental recording chamber and perfused with a test solution containing 100 mM NaCl, 2 mM KCl, 1 mM MgCl
2, 1 mM CaCl
2 and 10 mM HEPES (Tris buffered pH 7.4, 213 mosmol l
-1). Current traces were obtained by stepping the holding potential of -20 mV to test potentials ranging from − 130 mV to + 50 mV in increments of 15 mV in pulses of 200 msec. Recordings were low pass‐filtered at 500 Hz, sampled at 1 kHz and the steady‐state current activity was analysed at 160–180 ms after applying the test pulse.
Discussion
Here, we reveal a molecular coupling between a brain hemorrhagic event and the ensuing ventricular enlargement signifying PHH. The origin of hydrocephalus formation is generally sought in blockage of the CSF exit routes [
56‐
59]. In conditions with no discernable exit route blockage, ventriculomegaly could be caused by a component of CSF hypersecretion [
6,
60‐
63]. Hypersecretion of CSF may occur particularly in hydrocephalus etiologies where inflammation could be a pathogenetic factor [
6,
60,
63]. However, here we report that the mere presence of a blood lipid, LPA, appears to contribute to the CSF hypersecretion that may lead to ventriculomegaly in experimental rats. We detected an elevation of the phospholipid LPA in the CSF from patients with SAH and from rats following experimentally-induced IVH. Such LPA elevation has previously been reported for patients and mice experiencing traumatic brain injury [
15], indicative of brain entrance of the serum lipid LPA [
14] along with the hemorrhagic event. Such ventricular LPA elevation is able to mimic the hemorrhagic event and cause ventriculomegaly and elevated ICP in the week(s) following intracranial LPA administration [
16,
17]. These LPA-mediated morphological changes were effectuated via the G protein-coupled LPA receptors expressed in the brain tissue (LPA
R1-LPA
R6) [
16‐
18]. Here, we demonstrate an additional acute (24 h) effect of intraventricularly-delivered LPA causing ventriculomegaly and elevated brain fluid content in adult rats by its ability to act as an agonist of the TRPV4 channel. TRPV4 is expressed in various tissues and cell types throughout the body, for review see [
25], but in the brain it is highly concentrated in the choroid plexus [
26,
28,
64], in which we reveal it as the 6th highest-expressed ion channel, localized to the luminal membrane (this study and [
24]). Inhibition of TRPV4 modulates transepithelial ion flux in immortalized choroid plexus cell lines [
26,
27] and abolishes ventriculomegaly in a genetic rat model of hydrocephalus [
28]. Accordingly, we demonstrated an acute reduction of ICP in healthy rats exposed to an intraventricularly-delivered TRPV4 inhibitor. The reduction in ICP came about by TRPV4’s ability to directly modulate the rate of CSF secretion. TRPV4 activity thus modulates the CSF secretion rate in healthy rats and thereby may contribute to governing of the ICP. Notably, with its luminal localization on the choroid plexus, TRPV4 inhibitors may fail to reach the target if non-cell permeable inhibitors are delivered i.p., therefore the lack of effect on CSF secretion upon inclusion of TRPV4 inhibitor in an earlier study [
65].
TRPV4 can be directly activated by various lipid compounds [
25], and we here add LPA to the list of lipid agonists of TRPV4. LPA exposure elevated the TRPV4-mediated current in TRPV4-expressing
Xenopus laevis oocytes, an elevation that was absent in the presence of a TRPV4 inhibitor and in control oocytes lacking TRPV4 expression. Such a reduced experimental system allows for determination of direct functional interactions and promoted LPA as an agonist of TRPV4. The molecular coupling between TRPV4 activation and elevated activity of a transport mechanism implicated in CSF secretion originated in the TRPV4-mediated Ca
2+ influx, which activated the WNK/SPAK signaling pathway as also observed in salivary glands [
54]. These kinases are highly expressed in choroid plexus, with SPAK ranking as the highest expressed kinase (this study), and are well-established regulators of NKCC1 activity via modulation of the transporter phosphorylation status [
6,
66,
67]. Activation of this pathway, with either LPA or a well-established synthetic lipid agonist of TRPV4, terminated in NKCC1 hyperactivity and a resultant elevation of the CSF secretion rate (see schematic in Fig.
5). NKCC1 is a key contributor to the CSF secretion in mice, rats, and dogs [
6,
20,
21] as revealed with intraventricular delivery of the NKCC1 inhibitor bumetanide. With alternative delivery routes (i.v. or i.p.), bumetanide fails to reach its target on the luminal surface of choroid plexus in sufficient concentrations [
6,
65]. A coupling between the SPAK/WNK signaling cascade and NKCC1-mediated CSF hypersecretion has previously been described in an experimental IVH rat model [
6]. That study, however, promoted the immune receptor TLR4 and NFκβ as the molecular links between the hemorrhagic event and the SPAK-induced NKCC1-mediated CSF hypersecretion [
6]. These two PHH-related molecular pathways both promoting NKCC1 hyperactivity could well occur in parallel on different time scales with the LPA/TRPV4-mediated effects discernible within minutes, followed by a slower route (perhaps minutes–hours) through a hemorrhage/TLR4-mediated path through NFκβ-mediated transcription events leading to activation of SPAK [
6]. On a more prolonged time scale (days–weeks) LPA may act on its various G protein-coupled receptors and promote the ciliopathy, 3rd ventricular occlusion [
16,
17], and potentially other micro-blockages not discernible on brain imaging, together contributing to the ventriculomegaly characteristic of PHH. Notably the LPA receptor expression in brain tissue is developmentally regulated [
68] and it remains, at present, unresolved if the LPA-mediated pathological morphological changes observed in the neonatal mice [
16,
17] would occur in the adult. PHH may thus occur by various mechanisms and in distinct structures in the brain at different time points following the hemorrhagic event [
60]. Importantly, it appears that PHH arises, in part, from a component of CSF hypersecretion (this study and [
6,
63]), which may well occur in other brain pathologies with disturbed brain fluid dynamics. Delineation of the molecular underpinnings governing this (and other) CSF secretory disturbances may open new avenues for the pharmacological therapy wanting for these conditions. Although more could arise with future studies, potential targets could be TRPV4 [
28], the LPA signalling pathways [
15], the NKCC1 [
6], or the regulatory kinases, SPAK [
69,
70] and the serum/glucocorticoid-regulated kinase 1 (SGK1) [
71] inhibition of which may reduce hydrocephalus or lesion size in different cerebral pathological insults.
Limitations to the study include CSF sampling from cisterna magna in the rodent experimentation, from the ventricular compartment in the patients with SAH, and from the basal cisterns in the control patients. The different patient sample sites were dictated by ethical limitations in invasive CSF sampling, but could influence our results if the CSF composition differs between these locations. The control samples may contain blood contamination from the surgical opening. Such contamination is expected to raise the LPA levels in the control CSF. Our results would thus be expected to show even further differences between the two CSF groups, if one could test control CSF samples with no traces of blood. In the rat IVH model, the control rats received a saline injection (instead of the orthologous blood injection) and were therefore not naïve at the point of CSF sampling 24 h later. We cannot rule out that this manipulation could promote an inflammatory response, which in itself could affect the CSF LPA levels. However, the human CSF samples were obtained from patients undergoing preventive surgical clipping of unruptured aneurisms and therefore had to such surgical intervention prior to the sampling event. Of note, with the invasive nature of experimental approaches towards determination of agonist-induced modulation of intracranial pressure, cerebrospinal fluid secretion rates, and choroidal transport mechanisms, these parameters were obtained from the rodent (IVH) animal model. Such findings may not fully capture the human SAH condition. With the expected lipid-plastic interaction during the experimental procedures, most (approximately 97% [
16]) of the LPA is lost in the process. The concentration of free LPA in our experimental solutions is therefore an estimate and may not exactly match the concentrations detected in the CSF of humans and rats. Lastly, with the expected interplay with different transport mechanisms and regulatory factors present in the choroid plexus, our data may just reveal one segment of hemorrhage-related hydrocephalus formation. TRPV4 activation or LPA elevation may well act on other transporters or processes in the CSF secreting tissue and/or on the ventricular lining or various drainage pathways.
In conclusion, we demonstrate that the serum lipid LPA, entering the ventricular system during a hemorrhagic event, acts directly on TRPV4, and thus serves as a novel lipid agonist of this choroidal ion channel. Activation of TRPV4 causes the NKCC1 hyperactivity underlying the TRPV4-mediated elevated CSF secretion rate seemingly contributing to the ensuing ventriculomegaly signifying PHH. Future studies aimed at elucidating the pathophysiological changes in brain fluid dynamics occurring with diverse neuropathologies, ideally, should take into account formation rates of CSF, its path through the ventricular system, as well as its drainage, as most of these pathologies are likely to affect more than one of these aspects.
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