Impaired Cerebrospinal Fluid Circulation and Cerebral Lymphatic Drainage in a Rat Model of Chronic Hydrocephalus

Impaired Cerebrospinal Fluid Circulation and Cerebral Lymphatic Drainage in a Rat Model of Chronic Hydrocephalus | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article Impaired Cerebrospinal Fluid Circulation and Cerebral Lymphatic Drainage in a Rat Model of Chronic Hydrocephalus Dong Bin Back, Bo-Ryoung Choi, Kyoung Ja Kwon, Dong-Hee Choi, and 3 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-3989278/v1 This work is licensed under a CC BY 4.0 License Status: Posted Version 1 posted You are reading this latest preprint version Abstract Background The cerebrospinal fluid (CSF) not only protects the brain but also maintains homeostasis by removing metabolic waste produced by brain activity. This study hypothesizes that chronic CSF circulatory dysfunction, such as chronic hydrocephalus or normal pressure hydrocephalus (NPH), may be a critical condition in neurodegenerative diseases associated with metabolic waste accumulation. Methods To investigate the CSF circulation and cerebral lymphatic drainage in a rat model of chronic hydrocephalus induced by kaolin injection, we performed time-dependent evaluations of intraparenchymal injection of tracers as well as intraventricular injection of Evans blue. The study systemically evaluated the dysfunction of CSF circulation and lymphatic drainage in the brain from various perspectives, including the glymphatic system, transependymal CSF flow, subarachnoid CSF flow, meningeal lymphatic drainage, and peripheral lymphatic drainage to deep cervical lymph nodes. Results The results indicated delayed CSF circulation, including glymphatic system, and cerebral lymphatic drainage in the kaolin-induced chronic hydrocephalus model. Based on these findings, our research indicated that dysfunction of CSF circulation, as observed in conditions such as chronic hydrocephalus or NPH, may act as an initiating or exacerbating factor in neurodegenerative diseases. Conclusions This can lead to the accumulation of metabolic waste, as seen in Alzheimer's disease. Our research can help identify risk factors and provide insight into the underlying pathophysiology of neurodegenerative diseases, which may lead to the development of novel therapeutic strategies. Cerebrospinal fluid circulation hydrocephalus glymphatic meningeal lymphatic kaolin animal model Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Figure 8 Introduction The human brain is surrounded by cerebrospinal fluid (CSF), which is primarily produced by the choroid plexus and circulates within the intraventricular and subarachnoid spaces, eventually being absorbed through the arachnoid granulations into the dural venous sinuses [ 1 ]. The CSF plays an important role in protecting the brain from physical trauma. However, the CSF plays an even more crucial role in interacting with the brain parenchyma, helping to maintain homeostasis by removing metabolic waste generated by brain activity [ 1 ]. Recent research has highlighted the perivascular space and meningeal lymphatic channels as pathways for metabolic waste clearance, providing a new perspective on the role of CSF circulation in the brain [ 2 – 4 ]. In clinical practice, dysfunction of CSF circulation can manifest as communicating or non-communicating hydrocephalus depending on the presence of intraventricular obstruction [ 5 ]. Normal pressure hydrocephalus (NPH) is a type of communicating and chronic hydrocephalus characterized by increased CSF in the ventricles, often due to previous subarachnoid hemorrhage, infection, or head trauma, that chronically disrupts CSF circulation [ 1 , 6 ]. However, it has been observed that many NPH patients do not have these pre-existing medical conditions, suggesting that NPH may be a manifestation of age-related or neurodegenerative dysfunction in CSF circulation [ 7 ]. Impaired glymphatic clearance has been documented in NPH patients using MRI studies after intrathecal contrast agent injection [ 3 , 8 ] or diffusion tensor imaging [ 9 ]. Furthermore, detailed identification of meningeal lymphatic drainage channels has been achieved in both rodent models [ 4 , 10 ] and humans [ 11 , 12 ]. The circulation of CSF seems to be tightly regulated by the complex interplay between intraparenchymal glymphatic clearance activity and meningeal lymphatic drainage. In this study, our aim was to assess CSF circulatory dysfunction and cerebral lymphatic drainage from various perspectives, including the glymphatic system, transependymal CSF flow, subarachnoid CSF flow, meningeal lymphatic drainage, and peripheral lymphatic drainage to deep cervical lymph nodes. To achieve this, we used an animal model of chronic hydrocephalus induced by kaolin injection, a widely accepted method for replicating chronic hydrocephalus [ 13 – 17 ] or NPH [ 17 ] in rats. It was hypothesized that obstructing the intra-cisternal space through kaolin injection into the cisterna magna would impair CSF circulation. To validate the relevance of the model to chronic hydrocephalus, we performed measurements of ventricular enlargement and behavioral tests such as beam walking test. Intraparenchymal glymphatic clearance was evaluated by analyzing the dispersion of intraparenchymally injected CSF tracers. In addition, unbound Evans blue (EB), injected intraventricularly, was tracked at various sites to assess transependymal flow, subarachnoid flow, meningeal lymphatic drainage, and peripheral lymphatic drainage to deep cervical lymph nodes, providing insight into CSF circulation pathways and cerebral lymphatic drainage. Our investigation aimed to understand how clinical conditions such as chronic hydrocephalus or NPH, which impair CSF circulation, glymphatic clearance, and cerebral lymphatic drainage, may contribute to the progression of neurodegenerative diseases by impairing the mechanism responsible for removing metabolic waste. Material and Methods Animal preparation Male Wistar rats (10 week old, 300–320 g, Orient Bio) were used in all experiments after a two-week acclimatization period in the Konkuk University vivarium before the start of the experiment. Rats were housed under standard laboratory conditions (22 ± 1°C temperature, 50 ± 10% humidity, 12 h alternate light/dark cycle) with ad libitum access to food and water. Animal experimental procedures were performed in accordance with the ethical approval of the Institutional Animal and Use Committee of Konkuk University and ARRIVE guidelines ( https://www.nc3rs.org.uk/arrive-guidelines ). Rats were induced with 3% isoflurane and then anesthetized intraperitoneally with Zoletil 50 (30 mg/kg) and Rompun (10 mg/kg) before being secured in a stereotaxic frame. All stereotaxic injection experiments (kaolin with intra-cisterna magna injection, icm; CSF tracer with intraparenchymal injection; EB with intracerebroventricular injection, icv) had common preparation procedures, including anesthesia, head fixation in the stereotaxic frame, and reflow prevention by leaving the injection cannula in place for an additional 3 min, differing only in the specifications of the Hamilton syringe connected to the syringe pump (Harvard) and the needle size. Rats with kaolin-induced hydrocephalus were assigned to the CH group. Meanwhile rats with sham surgery were classified as the sham group. Induction of kaolin-induced hydrocephalus A midline incision was made to expose the atlantooccipital membrane overlying the cisterna magna. A 60 µL of sterile suspension of kaolin (100 mg/mL in 0.9% saline, Sigma-Aldrich) was injected into the cisterna magna at a rate of 2.0 µL/s via a 28-gauge cannula connected to polyethylene tubing, attached to a 100 µL Hamilton syringe with a 22-gauge needle. Based on documented findings of normalized CSF dynamics [ 18 , 19 ] and cerebral blood flow [ 16 ], changes in cerebral metabolism [ 16 , 19 ], cessation of ventricular enlargement [ 16 , 20 , 21 ], cognitive decline [ 22 , 23 ] and behavioral changes [ 24 ], our study established an 8-week post-kaolin infusion period for the development of a chronic hydrocephalus model in adult rats. All experimental evaluations were performed 8 weeks after kaolin induction of hydrocephalus. Ventricular volume Ventricular volumes were measured from multiple brain slices stained with cresyl violet acetate (Sigma-Aldrich) according to the methods described in our previous studies [ 25 , 26 ]. Briefly, ventricular volumes, including the lateral and third ventricles, were calculated from the anterior horn of the lateral ventricle to the cerebral aqueduct using seven representative coronal slices at 2-mm intervals from bregma: AP + 1.92 mm to -4.92mm. Ventricular volume was calculated by multiplying the slice interval by the ventricular area obtained using the National Institutes of Health (NIH) Image J program. Beam walking test The beam walking test was performed as previously described [ 27 – 29 ], with minor modifications. Rats were assessed for their ability to traverse two parallel plastic beams, each 2.0 cm in diameter and 95 cm in length, positioned horizontally 50 cm above the ground. Prior to the kaolin icm (before 1 week), the rats participated in an adaptation training session to freely traverse the beams, eliminating any signs of fear or hesitation, and subsequent tests were conducted on consecutive days 1, 3, 7, 14, 28, 42, and 56 after the kaolin injection. After being placed on a platform (30 × 30 cm), we recorded the latency time and the number of footsteps (for each forelimb and hindlimb) for up to 60 s as the rats traversed from the moment their entire body entered the beam until they reached the opposite platform. Intraparenchymal injection of cerebrospinal fluid tracers We performed intraparenchymal CSF tracer injection experiments as described in our previous study [ 26 ]. Briefly, a 3 µL tracer solution containing Texas - red conjugated dextran (TR-d3, MW 3 kDa, Thermo Fisher Scientific) and fluorescein isothiocyanate (FITC) conjugated dextran (FITC-d40, MW 40 kDa, Sigma-Aldrich), both at 0.5% concentration in a 1:1 ratio, was dissolved in artificial CSF (Tocris Bioscience). In the co-infusion experiment, the pre-prepared 3 µL tracer solution was administered into the brain parenchyma at a rate of 0.5 µL/min at bregma-related coordinates (AP + 2.0 mm, ML + 2.6 mm, DV -5.4 mm) using a 30-gauge injection cannula connected to polyethylene tubing, attached to a 10 µL Hamilton syringe with a 26s-gauge needle. Six hours after injection, rats were transcardially perfused with 0.01 M phosphate-buffered saline, pH 7.4 (1x PBS) and fixed with 4% paraformaldehyde (PFA) for fluorescence imaging analysis. Intracerebroventricular injection of EB EB (DB 53, MW 0.96 kDA, Sigma-Aldrich) was suspended in 0.9% and injected into the ventricle saline (10 µL at 1.0 µL/min, 20 mg/mL) at coordinates relative to bregma (AP -0.9 mm, ML + 1.4 mm, and DV -3.8 mm) using a 30-gauge injection cannula connected to polyethylene tubing, attached to a 25 µL Hamilton syringe with a 22-gauge needle. For imaging and quantitative analysis at 3 or 24 h after icv EB injection, rats were perfused with PBS, omitting the fixative solution, to facilitate neck incision for deep cervical lymph nodes harvesting, which was also used to obtain coronal brain slices and whole brain tissue to ensure experimental consistency. Meanwhile, EB-stained meninges were dissected without transcardiac perfusion of both PBS and PFA to preserve the integrity of meningeal lymphatic vessels. Tissue preparation and immunohistochemistry After isoflurane anesthesia and transcardiac perfusion, rat brains were immediately removed, post-fixed overnight in the same fixative, sequentially cryoprotected in 30% sucrose solution, and were embedded in an optimal cutting temperature compound (Sakura Finetek). Brains were serially sectioned into 40 µm coronal slices using a CM1520 cryostat (Leica) for immunohistochemistry and tracer studies. To obtain whole mounts of rat dural meninges, we adapted methods from previously reported mouse experiments [ 30 – 34 ], with minor modifications. Briefly, at 3 or 24 h after icv EB injection, anesthetized rats were decapitated above the shoulders, and the skin and muscle were removed from the outer skull. Using a bone cutter (Roboz), a precise dissection was made from the foramen magnum to the lateral end of the occipital bone, extending transversely in the longitudinal plane across the squamous and frontal bones to the junction of the frontal and nasal bones, taking care to minimize damage to the brain tissue. In order to obtain preserved meninges firmly attached to the adjacent skull, the skull bone was levered anteriorly outwards and meninges were gently separated from the bone using an elevator (Roboz). The separated EB-stained meninges, which were gently washed with PBS for 1 min to remove blood while retaining EB, were then used for immunohistochemistry and quantitative analysis of EB distribution. To obtain deep cervical lymph nodes, rats perfused only with PBS underwent a midline incision (5 mm above the clavicle) to expose the sternocleidomastoid muscle, which was then retracted with forceps, followed by post-fixation, cryoprotection, and embedding procedures (similar to brain sections) for quantitative analyses of EB distribution. After sample preparation, immunofluorescence staining was performed to assess various pathological outcomes. Brain sections (either intact or stained with EB) were washed (1x PBS/0.3% Triton X-100) and then incubated in blocking serum (1x PBS/10% normal donkey serum/0.3% Triton X-100) for 1 h at room temperature (RT), while the meninges started the incubation process. Double-label immunofluorescence was performed overnight at 4°C in a solution (1x PBS/0.15% normal donkey serum/0.3% Triton X-100) containing the following primary antibodies: mouse anti-glial fibrillary acidic protein (GFAP, 1:1000, BD Bioscience), rabbit anti-ionized calcium binding adapter molecule-1 (Iba-1, 1:1000, Wako), rabbit anti-collagen IV (COLIV, 1:100, Abcam), mouse anti-aquaporin 4 (AQP4, 1:100, Abcam), mouse anti-collagen IV (COLIV, 1:100, Sigma-Aldrich), rabbit anti-lymphatic vessel endothelial hyaluronan receptor 1 (LYVE1, 1:100, Abcam). The sections were then washed (1x PBS/0.15% Triton X-100) and incubated for 3 h at RT in a secondary antibody solution as follows: anti-mouse Alexa Fluor 488 and 568; anti-rabbit Alexa Fluor 488 and 568; all from donkey (1:200, Invitrogen). Some of the coronal sections were counterstained with TO-PRO-3 (1:1,000, Invitrogen) for 30 min at RT. Stained sections were mounted on slides, dried for 30 min, and coverslipped with ProLong® Gold antifade reagent (Invitrogen), excluding structurally uneven meninges. Aquaporin 4 polarization Astrocytic AQP4 polarization was assessed according to our previous study [ 25 ]. Briefly, a low stringency threshold defined the total area of AQP4 immunoreactivity, whereas a high stringency threshold identified the vascular AQP4 colocalized with COLIV. Image J (NIH) was used for area calculations after black and white thresholding. The ratio of low stringency area to high stringency area was defined as AQP4 polarization. Image analysis and quantification Bright field photographs were taken with a digital camera using a standard exposure time for all brains. Immunofluorescence images were obtained using a Carl Zeiss LSM 900 confocal laser scanning microscope at different magnifications (10× air, NA 0.45; 20× air, NA 0.80; 40× water, NA 1.2; 63× oil, NA 1.4) with a resolution of 1024 × 1024 pixels, acquired by multichannel scanning in either a single or tiled frame. Capillary filling of EB into the microvessels in the corpus callosum was analyzed both by z-stack three-dimensional reconstruction (image slices taken at 22 µm intervals consisting of 12 z-sections), with upper and lower limits set based on positively visualized vascular morphology for COLIV and AQP4, and by fluorescence co-localization in the XY and orthogonal XZ, YZ projections of the image stacks. Image analysis and quantification was performed using Image J (NIH) and Zen blue image analysis wizard (Carl Zeiss). Statistics Behavioral data were analyzed using two-way repeated measures analysis of variance (ANOVA) and unpaired t-tests. Two-way ANOVA was used to assess the interaction between group and time, followed by a post-hoc Tukey’s honest comparison to analyze differences between groups over time. Time or group dependent differences were analyzed using unpaired t-tests, where applicable. Pearson’s correlation coefficient (r) was used for correlation analyses between ventricular volume and other variables. All data are expressed as the mean ± SEM. A value of p < 0.05 was considered to be statistically significant. Data analysis was performed with SPSS software version 27.0. Results Characteristics of rats with chronic hydrocephalus Eight weeks after icm injection of kaolin, the extracted brains showed a nearly uniform distribution of kaolin deposits throughout the ventral regions, extending from the optic nerves to the medulla and covering the interpeduncular fossa, pons, and cerebellum as indicated by the black lining (Fig. 1 A) and kaolin-contacted inflammation as indicated by black arrowheads in the CH group (Fig. 1 B). Marked hydrocephalus was observed, characterized by symmetrical enlargement of the lateral ventricles, cerebral aqueduct, and the fourth ventricle (Fig. 1 B) with a quantitatively significant increase in total ventricular volume in the CH group (p < 0.001, Fig. 1 C). Modified beam walking test showed a significant difference in patterns (F (7,273) = 15.359, p < 0.001, latency time in Fig. 1 D; F (7,273) = 5.664, p < 0.001, the number of footsteps in Fig. 1 E). At the various time points, there was a significant increase in latency time (p < 0.001, post-op day 1; p < 0.01, post-op day 3, 56; p < 0.05, post-op day 7, 28; Fig. 1 D) and the number of footsteps (p < 0.001, post-op day 1; p < 0.01, post-op day 3; p < 0.05, post-op day 14, 56; Fig. 1 E) in the CH group compared to the sham group. A more frequent use of the forelimbs than the hindlimbs was observed in the CH group compared to the sham group on post-op day 1 (p < 0.01, Fig. 1 F). The gait impairment in the CH group was pronounced within the first 7 days post-operation, with the impairment persisting for the entire 8-week period, suggesting a chronic motor function deficit in rats with kaolin-induced hydrocephalus. Neuroinflammation associated with kaolin-induced chronic hydrocephalus Meningeal fibrosis, characterized by chemical inflammatory arachnoiditis in the basal cistern due to direct contact with kaolin (Fig. 2 A) and indirect inflammation in the corpus callosum influenced by ventricular enlargement (Fig. 2 B) are notable features in the CH group. Quantitative analysis revealed a significant increase in reactive astro- and microgliosis within the CH group, measured by area (Fig. 2 C and 2 M in GFAP; Fig. 2 E and 2 O in Iba1), fluorescence intensity (Fig. 2 D and 2 N in GFAP; Fig. 2 F and 2 P in Iba1), and the ratio of the number of Iba1-positive cells to total number of ToPr3-positive cells (Fig. 2 G and 2 Q). Furthermore, within the CH group, a positive correlation was observed between ventricular volume and GFAP signal measured by area (r = 0.653, p < 0.05, Fig. 2 H) and fluorescence intensity (r = 0.605, p < 0.05, Fig. 2 I), as well as Iba1 signal measured by area in the basal cistern (r = 0.669, p < 0.05, Fig. 2 J). In addition, the Iba1 to To-pro3 positive cell ratio in the basal cistern (r = 0.877, p < 0.001, Fig. 2 L) and corpus callosum (r = 0.437, p < 0.05, Fig. 2 V) correlated with ventricular volume, suggesting a positive relationship between ventricular dilatation and the severity of neuroinflammation. AQP4 depolarization in the white matter in rats with chronic hydrocephalus Changes in AQP4 expression were evident in the corpus callosum of the CH group (Fig. 3 A). In addition to the increase in parenchymal AQP4 expression (p < 0.001, Fig. 3 B), the distribution pattern of AQP4 also changed, indicating redistribution into the brain parenchyma from the location on perivascular astrocyte endfeet in close contact with COLIV-positive endothelial vessels (Fig. 3 C). Perivascular AQP4 localization, indicative of the AQP4 polarization ratio, was significantly reduced in the CH group compared to the sham group (p < 0.001, Fig. 3 D). This loss of AQP4 polarization also correlated well with ventricular dilatation (r = -0.586, p < 0.05, Fig. 3 E). Delayed dispersion of intraparenchymally injected tracers in rats with chronic hydrocephalus Intraparenchymally injected tracers of different molecular weights exhibited delayed dispersion patterns at 6 h post-injection in the CH group compared to the sham group (Fig. 4 A). The black star in Fig. 4 A indicates the intraparenchymal tracer injection site. Tracer dispersion into the brain parenchyma was impaired in the CH group in multiple brain slices in both cases of TR-d3 (p < 0.05 at 4.20 mm and 1.08 mm, p < 0.01 at 3.24 mm, assessed by area in Fig. 4 B; p < 0.05 at 4.20 mm, p < 0.01 at 3.24 mm, assessed by fluorescence intensity in Fig. 4 C) and FITC-d40 (p < 0.05 at 3.24 mm, assessed by area in Fig. 4 D; p < 0.01 at 3.24 mm, assessed by fluorescence intensity in Fig. 4 E). The lighter TR-d3 tracer exhibited greater dispersion both anteriorly and posteriorly from the injection site than the heavier FITC-d40 tracer. Transependymal and subarachnoid dispersion of icv-injected EB in rats with chronic hydrocephalus Transependymal dispersion of icv-injected EB was visibly present at 3 hours post-injection and disappeared by 24 hours in the sham group, whereas it was scarce in the CH group (Fig. 5 A). The red star in Fig. 5 A indicates the icv EB injection site. Five sections around the injection site were selected to assess the pattern of dispersion depending on time or model. In multiple sections, transependymal dispersion was significantly impeded in the CH group compared to the sham group throughout the 24 hours when assessed by the EB-stained area (p < 0.05 at 0.00 mm, p < 0.001 at -0.96 mm, CH3h vs. Sham3h; p < 0.01 at 0.00 mm, p < 0.05 at -0.96 mm, CH24h vs. Sham24h; Fig. 5 B) and fluorescence intensity (p < 0.001 at 0.96, 0.00, and − 0.96 mm, p < 0.05 at -1.92mm, CH3h vs. Sham3h; p < 0.001 at 0.96, 0.00, and − 0.96 mm, p < 0.05 at -2.92 mm, CH24h vs. Sham24h; Fig. 5 C). While the sham group showed a time-dependent washout pattern with a peak at 3 hours post-injection and a subsequent decrease at 24 hours (p < 0.05 at -0.96 mm, Sham3h vs. Sham24h, assessed by area in Fig. 5 B; p < 0.05 at 0.00 mm, p < 0.01 at -0.96mm, p < 0.001 at 0.96, -1.92 and − 2.92 mm, Sham3h vs. Sham24h, assessed by fluorescence intensity in Fig. 5 C), the CH group showed minimal initial dispersion into the transependymal region and a persistent pattern. Subarachnoid dispersion of icv-injected EB was also hindered in the CH group throughout the 24 h compared to the sham group (Fig. 5 D). EB staining was quantified by assessing views from the four different angles including caudal, ventral, lateral, and dorsal angels. Subarachnoid dispersion was significantly impeded in the CH group compared to the sham group when assessed by the EB-stained area (p < 0.001 at ventral angle, CH3h vs. Sham3h in Fig. 5 E) and fluorescence intensity (p < 0.001 at ventral, lateral, and caudal angles, CH3h vs. Sham3h in Fig. 5 F). The time-dependent washout pattern was observed in the sham group (p < 0.05 at caudal, p < 0.001 at ventral, lateral, and dorsal angles, Sham3h vs. Sham 24h, assessed by fluorescence intensity in Fig. 5 F), whereas it was not evident in the CH group except at ventral angle (p < 0.05, assessed by fluorescence intensity in Fig. 5 F). Transependymal capillary filling of icv-injected EB in rats with chronic hydrocephalus Examination by confocal microscopy revealed apparent transependymal capillary filling of the icv-injected EB in the corpus callosum of the sham group, whereas it was impaired in the CH group (orthogonal views in Fig. 6 A and z-stack 3D in Fig. 6 B). Capillary density measured by COLIV-positive blood vessels within the unit region (%) showed a significant decrease in the CH group compared to the sham group (p < 0.001, Fig. 6 C). In the sham group, the capillary filling of the EB, which was evident at 3 hours post-injection, disappeared at 24 hours. In contrast, transependymal capillary filling of EB in the CH group was minimal throughout the 24 hours. Quantitative evaluation using the ratio of EB-filled capillary length to the total capillary length showed peaks at 3 hours, indicating rapid capillary filling in the sham group compared to the CH group (p < 0.001, CH3h vs. Sham3h, Fig. 6 D), and declines at 24 hours, indicating subsequent capillary emptying (p < 0.05, Sham3h vs. Sham24h, Fig. 6 D). In contrast, the CH group showed peaks at 24 hours instead of at 3 hours, indicating delayed capillary filling. Meningeal lymphatic drainage in rats with chronic hydrocephalus Immunohistochemistry using whole-mount meninges revealed the colocalization of scattered lymphatic endothelial signals (LYVE1) alongside meningeal vascular signals (COLIV), as shown in Fig. 7 A. Cross-sectional analysis of multi-immunofluorescence signals showed red peaks (LYVE1) surrounding green peaks (COLIV) in Fig. 7 B. After 3 hours of icv-injected EB in the sham group, distinct blue peaks (EB) were observed between the two red peaks, as shown by the black dotted lines indicating the lumen of meningeal lymphatic vessels in Fig. 7 B, suggesting effective EB influx into the meningeal lymphatic vessels. This distinct pattern of blue peaks disappears at post-24 hours of icv-injected EB in the sham group, indicating subsequent efficient drainage of EB via meningeal lymphatic vessels. In contrast to the sham group, the CH group showed no clear influx of EB into the meningeal lymphatic vessels and subsequent drainage. Quantitative analysis using EB signals on the meninges revealed a consistent impairment of meningeal lymphatic drainage in the CH group throughout the 24 hours compared to the sham group. This impairment is evident in the CH group when assessing CH3h vs. Sham3h, as indicated by the EB-stained area (p < 0.01, Fig. 7 C) or fluorescence intensity (p < 0.01, Fig. 7 D). In contrast to the normal meningeal lymphatic drainage pattern in the sham group, which peaked at 3 hours and then disappeared at 24 hours (p < 0.05, Sham3h vs. Sham24h, Fig. 7 C), the CH group showed minimal influx into the meningeal lymphatic vessels and a stagnant pattern. Peripheral lymphatic drainage in rats with chronic hydrocephalus Time-dependent peripheral lymphatic drainage of icv-injected EB into the deep cervical lymph nodes is shown in both groups. Comparing the two groups at different time points, EB signals were evident at 3 hours post-injection and peaked at 24 hours in the sham group, while a relatively weaker signal was observed in the CH group (Fig. 8 A). Peripheral lymphatic drainage of EB was consistently impaired in the CH group compared to the sham group throughout the 24 hours (p < 0.01, CH3h vs. Sham3h; p < 0.001, CH24h vs. Sham24 h, assessed by area in Fig. 8 B; p < 0.01, CH3h vs. Sham3h; p < 0.001, CH24 h vs. Sham24 h, assessed by fluorescence intensity in Fig. 8 C). Although both groups showed a time-dependent increase in peripheral lymphatic drainage of EB to the deep cervical lymph nodes, this was more pronounced in the sham group (p < 0.01, Sham3h vs. Sham24h, assessed by area in Fig. 8 B or fluorescence intensity in Fig. 8 C). Discussion The experimental findings using the kaolin-induced hydrocephalus rat model demonstrate how chronic hydrocephalus can disrupt the dynamics of CSF circulation associated with the glymphatic system and cerebral lymphatic drainage. It is hypothesized that the injection of kaolin into the rat cisterna magna, which limits CSF absorption through the subarachnoid space, could serve as a reliable model for studying clinical conditions such as chronic hydrocephalus or NPH, a communicating type of hydrocephalus. Although we acknowledge that the kaolin-induced hydrocephalus model, in which the CSF absorption pathway is artificially obstructed by injected kaolin, may not perfectly mimic the pathophysioloy and clinical features of chronic hydrocephalus or NPH, we have modified our experimental settings to resemble chronic hydrocephalus or NPH observed in humans. The aim was to create a chronic and mild condition similar to NPH without directly obstructing the intraventricular circulation of CSF. To achieve this, we used a minimal amount of kaolin and prolonged the duration of the hydrocephalus. Injection of kaolin into the cisterna magna is known to induce inflammation, which leads to meningeal fibrosis, specifically chemical arachnoiditis [ 35 – 38 ]. The majority of the kaolin deposits were distributed throughout the basal subarachnoid space, evenly on both sides of the ventral brainstem, including the interpeduncular fossa to the medulla and pons. No significant presence of kaolin was observed within the intraventricular system, particularly blocking the outlet of the fourth ventricle. The icv-injected EB diffused throughout the entire subarachnoid space through the outlet of the fourth ventricle. This diffusion pattern of EB indicates that CSF flow through the intraventricular and subarachnoid spaces was not mechanically obstructed in the kaolin-induced chronic hydrocephalus rat model. When discussing hydrocephalus, it is important to differentiate between acute and chronic conditions based on duration. The kaolin-induced hydrocephalus model is a well-known animal model for chronic hydrocephalus as it develops and progresses hydrocephalus over several weeks to months [ 21 , 35 , 39 – 41 ]. Injected kaolin in the basal subarachnoid space can cause acute hydrocephalic symptoms within two weeks, leading to an increase in resistance to CSF outflow and subsequent elevation of intracranial pressure [ 35 , 39 ]. As hydrocephalus progresses from the acute phase to the chronic phase over a longer period of four to six weeks or more, the resistance to CSF outflow and intracranial pressure undergo changes [ 35 , 39 ]. Animal studies using MRI have shown ventricular enlargement, with a peak at 6 weeks, followed by a subsequent decline at 10 weeks [ 21 ]. The progressive increase in ventricular size up to 8 weeks reaches a new steady state characterized by the restoration of normal intracranial pressure and an increase in resistance to CSF outflow [ 42 ]. Moreover, the normalization of cerebral blood flow after 8 weeks indicates that the post-operative 8-week time point we chose for evaluation is optimal for investigating chronic hydrocephalus, such as NPH [ 40 ]. Our model also identified gait impairments, which are clinical manifestations observed in human NPH [ 6 ]. Although the kaolin-induced hydrocephalus rat model has inherent limitations, it has potential for investigating the impact of chronic hydrocephalus on the glymphatic system and cerebral lymphatic drainage in neurodegenerative diseases, particularly in relation to the accumulation of metabolic waste. The kaolin-induced hydrocephalus rat model’s pathophysiology is mainly focused on neuroinflammatory changes and alterations of AQP4, the brain’s most abundant water channel that regulates water homeostasis [ 43 ]. Neuroinflammation with gliosis in the basal forebrain and corpus callosum may be indirectly triggered by increased pressure in the subarachnoid or intraventricular space, in addition to the reactive gliosis due to direct contact with kaolin. The degree of gliosis is strongly correlated with enlarged ventricular volume, which may indirectly support the possibility of pressure-related gliosis. A significant depolarization of AQP4 was observed, resulting in the translocation of the normal perivascular pattern of AQP4 to the parenchymal pattern with an increased total AQP4 signal in the CH group. The evidence indicates that neuroinflammation and depolarization of AQP4 play a significant role in the pathophysiology of impaired CSF circulation in chronic hydrocephalus. Previous studies have reported impaired glymphatic system associated with the depolarization of AQP4 [ 25 , 26 ]. In this study, we investigated the glymphatic efflux of intraparenchymally injected tracers with two different molecular weights. As anticipated, the molecular weight-dependent intraparenchymal dispersion of the tracers at 6 hours post-injection was hindered in the CH group, indicating a compromised glymphatic system in the chronic hydrocephalus. Previous studies have evaluated the classical CSF circulation pathway, which involves production in the choroid plexus, intraventricular circulation, and absorption into the dural venous sinuses through the arachnoid villi [ 7 , 44 ]. To assess this pathway, we measured the subarachnoid dispersion of icv-injected EB. Analysis of the subarachnoid brain surface area covered by EB in the sham group revealed rapid subarachnoid dispersion at 3 hours post-injection, which was subsequently eliminated by 24 hours post-injection. In contrast, the CH group exhibited delayed and stagnated dispersion into the subarachnoid space, indicating a disturbance of the classical CSF circulation pathway in the chronic hydrocephalus. Furthermore, we investigated an alternative transependymal pathway for CSF circulation. This pathway was discovered in a study that utilized precise MRI mapping of CSF flow with a tracer injected into the ventricle of healthy rats [ 45 ]. The investigation identified previously unrecognized parenchymal perivascular space connections that spread across various brain regions, facilitating the direct transport of CSF from the ventricles to the subarachnoid space [ 45 ]. In studies of hydrocephalus in humans, the absorption of CSF by periventricular tissues serves as a compensatory mechanism for increased intracranial pressure and sheds light on an alternative pathway for CSF circulation [ 46 – 48 ]. Consistent with these studies, our analysis of coronal brain slices stained by EB revealed rapid periventricular diffusion of icv-injected EB at 3 hours post-injection and subsequent elimination at 24 hours post-injection in the sham group, indicating active transependymal CSF flow in the normal condition. Conversely, minimal periventricular diffusion of EB was observed in the CH group. Based on the confocal microscopic image of the corpus callosum in the sham group, the capillaries were completely filled with EB three hours after injection and showed clear drainage within 24 hours after injection. The microvascular density was found to be deceased in the CH group compared to the sham group, which is consistent with previous studies reporting a decrease in microvascular density in chronic hydrocephalus [ 49 – 51 ]. In the CH group, there was a decrease in capillary density and no visible diffusion of EB into the capillaries even after 24 hours. These findings suggest a disturbance of the alternative transependymal CSF circulation pathway in chronic hydrocephalus, as evidenced by compromised periventricular diffusion and capillary drainage of EB. In the animal study, the final step of CSF efflux into the venous sinuses through the arachnoid villi was demonstrated using serial high resolution MRI tracking of injected tracer into the lateral ventricles [ 45 ]. Time-dependent EB drainage through the venous sinus was observed in the sham group, supporting the classical CSF circulation pathway via venous sinus drainage. Analysis of EB staining on the meninges showed strong staining around the venous sinuses and the middle meningeal artery. Recent research has also revealed the presence of lymphatic vessels in the meninges responsible for draining CSF, interstitial fluid, macromolecules, and immune cells to the cervical lymph nodes [ 52 , 53 ]. In a previous study using novel lymphatic reporter rats, it was demonstrated that meningeal lymphatic vessels were located alongside the middle meningeal artery, superior sagittal sinus, and transverse sinuses [ 54 ]. In our study, we used a whole-mount dissection of the dura mater and found that LYVE1-positive meningeal lymphatic vessels appeared as discontinuous dotted lines scattered along the COLIV-positive meningeal arteries. Cross-sectional analysis of multi-immunofluorescence signals revealed intraluminal filling and subsequent drainage of EB through the meningeal lymphatic vessels as a CSF efflux pathway. In the CH group compared to the sham group, the EB signal did not appear within the meningeal lymphatic vessels, although the structure of the meningeal lymphatic vessels seemed intact. This suggests a diminished functional CSF efflux through the meningeal lymphatic vessels in chronic hydrocephalus. As expected, peripheral lymphatic drainage to the deep cervical lymph nodes via the meningeal lymphatic vessels or other possible routes was also delayed in the CH group. Conclusions Our study evaluated several crucial steps in the CSF circulation pathway over time, including the glymphatic system, transependymal CSF flow, subarachnoid CSF flow, efflux through the venous sinus and meningeal lymphatic vessels, as well as peripheral lymph nodes drainage. Recognizing the role of the glymphatic system and its connection with cerebral lymphatic drainage, which functions as a clearance system for amyloid, tau, and other brain metabolic waste [ 3 , 55 ], clinical conditions such as chronic hydrocephalus or NPH may pose a significant risk for the progression of various neurodegenerative diseases, including Alzheimer’s disease. Therefore, it will be critical to conduct extensive research on the circulation of CSF and cerebral lymphatic drainage to identify risk factors and understand the pathophysiology of neurodegenerative diseases, which may lead to novel therapeutic strategies. Abbreviations CSF cerebrospinal fluid NPH normal pressure hydrocephalus EB Evans blue NIH National Institutes of Health FITC fluorescein isothiocyanate PFA paraformaldehyde RT room temperature GFAP glial fibrillary acidic protein Iba-1 ionized calcium binding adapter molecule-1 COLIV collagen IV LYVE1 lymphatic vessel endothelial hyaluronan receptor 1 ANOVA analysis of variance Declarations Ethics approval and consent to participate All animal experimental procedures were in accordance with the approved ethical standards of the Institutional Animal and Use Committee of Konkuk University. Consent for publication Not applicable Availability of data and material The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request. Competing interests The authors declare that they have no competing interests. Funding This work was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education (2020R1F1A1076085). Authors' contributions DB, BC, and HK participated in the conception and design of the study. DB and HK drafted the manuscript and participated in general management. DB, KK, and DC helped acquisition and analysis of data. CS and JL helped to draft the manuscript. All authors reviewed and approved the final manuscript. 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Reeves BC, Karimy JK, Kundishora AJ, Mestre H, Cerci HM, Matouk C, Alper SL, Lundgaard I, Nedergaard M, Kahle KT: Glymphatic System Impairment in Alzheimer's Disease and Idiopathic Normal Pressure Hydrocephalus. Trends Mol Med 2020, 26(3):285-295. Additional Declarations No competing interests reported. Cite Share Download PDF Status: Posted Version 1 posted You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. Our growing team is made up of researchers and industry professionals working together to solve the most critical problems facing scientific publishing. Also discoverable on Platform About Our Team In Review Editorial Policies Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-3989278","acceptedTermsAndConditions":true,"allowDirectSubmit":true,"archivedVersions":[],"articleType":"Research Article","associatedPublications":[],"authors":[{"id":275247049,"identity":"651d8669-2891-4cce-a5c3-c2567f0b40b0","order_by":0,"name":"Dong Bin Back","email":"","orcid":"","institution":"Konkuk University School of Medicine","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"Dong","middleName":"Bin","lastName":"Back","suffix":""},{"id":275247050,"identity":"47bb25a8-7b9b-47e0-b921-afb5f73aa08a","order_by":1,"name":"Bo-Ryoung Choi","email":"","orcid":"","institution":"Konkuk University School of Medicine","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"Bo-Ryoung","middleName":"","lastName":"Choi","suffix":""},{"id":275247051,"identity":"e75f1cd9-bdb1-433e-aa30-ec7bdb4c8b01","order_by":2,"name":"Kyoung Ja Kwon","email":"","orcid":"","institution":"Konkuk University School of Medicine","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"Kyoung","middleName":"Ja","lastName":"Kwon","suffix":""},{"id":275247052,"identity":"78e6c9d2-7941-45cc-a591-fe848d43af5c","order_by":3,"name":"Dong-Hee Choi","email":"","orcid":"","institution":"Konkuk University School of Medicine","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"Dong-Hee","middleName":"","lastName":"Choi","suffix":""},{"id":275247053,"identity":"5e174b2e-c6a5-4b48-9f34-6c14a8edebf6","order_by":4,"name":"Chan Young Shin","email":"","orcid":"","institution":"Konkuk University School of Medicine","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"Chan","middleName":"Young","lastName":"Shin","suffix":""},{"id":275247054,"identity":"1170c1af-d0b1-4883-a97b-c26e2b7a3759","order_by":5,"name":"Jongmin Lee","email":"","orcid":"","institution":"Konkuk University School of Medicine","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"Jongmin","middleName":"","lastName":"Lee","suffix":""},{"id":275247055,"identity":"4be74d08-4ad8-4a7f-b6eb-3901e94c2e18","order_by":6,"name":"Hahn Young Kim","email":"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAAAvklEQVRIiWNgGAWjYBACAwY2hgMMDMxyIM6BB6RoMQZrSSBWCxAwJzaAKKK0mEukJR66mWOdPj/s8EOgLXZyug0EtFjOSDtwOHdbeu7G22kGQC3JxmYHCDnsRnoDUMvh3I2zE0BaDiRuI1ZLuuHs9A/EagE77HCCvHQOsbaceZYA8ovhBumcggMJBsT45Xia8efcbdby8rPTN3/4UGEnR1ALg0ACVC9YpQEh5SDADzVUvoEY1aNgFIyCUTAiAQC+sE1ywDFtlwAAAABJRU5ErkJggg==","orcid":"","institution":"Konkuk University Medical Center and Konkuk University School of Medicine","correspondingAuthor":true,"submittingAuthor":false,"prefix":"","firstName":"Hahn","middleName":"Young","lastName":"Kim","suffix":""}],"badges":[],"createdAt":"2024-02-25 23:44:37","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-3989278/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-3989278/v1","draftVersion":[],"editorialEvents":[],"editorialNote":"","failedWorkflow":false,"files":[{"id":51801037,"identity":"5e55a3ba-2a83-4b94-a868-eb4d8186873b","added_by":"auto","created_at":"2024-02-29 09:10:16","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":920488,"visible":true,"origin":"","legend":"\u003cp\u003eInduction of chronic hydrocephalus via intra-cisterna magna kaolin injection and behavioral test results.\u003c/p\u003e\n\u003cp\u003e(A) Representative brain regions showing kaolin deposition indicated by the black lining in the CH group. (B) Representative cresyl violet stain depicting ventricular enlargement with kaolin-contacted inflammation indicated by black arrowheads in the CH group. (C) Quantification of ventricular volume, n = 7-10. (D) Latency time and (E) footsteps, and (F) forelimb-use asymmetry in the beam walking test, n = 3-23 at various time points. Scale bar = 2 mm; *p \u0026lt; 0.05, **p \u0026lt; 0.01, ***p \u0026lt; 0.001; †††p \u0026lt; 0.001; CH, chronic hydrocephalus; LV, lateral ventricle; Aq, cerebral aqueduct; 4V, fourth ventricle.\u003c/p\u003e","description":"","filename":"floatimage1.png","url":"https://assets-eu.researchsquare.com/files/rs-3989278/v1/f963985c76c32d2d30486098.png"},{"id":51801038,"identity":"623bfe35-5031-4208-b2bb-ea2e3673c61a","added_by":"auto","created_at":"2024-02-29 09:10:16","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":517073,"visible":true,"origin":"","legend":"\u003cp\u003eNeuroinflammation in kaolin-induced chronic hydrocephalus.\u003c/p\u003e\n\u003cp\u003eNeuroinflammation in the basal cistern (A) or corpus callosum (B) measured by GFAP and Iba1, quantified by covered area (C and M in GFAP, E and O in Iba1), by fluorescence intensity (D and N in GFAP, F and P in Iba1), or Iba1/TP3 ratio (G and Q) and its correlation with ventricular volume (H-L and R-V). n = 11-13; scale bar = 500 µm (100 µm in the inset); *p \u0026lt; 0.05, **p \u0026lt; 0.01, ***p \u0026lt; 0.001; CH, chronic hydrocephalus; MFI, mean fluorescence intensity; au, arbitrary unit; TP3, TO-PRO-3.\u003c/p\u003e","description":"","filename":"floatimage3.png","url":"https://assets-eu.researchsquare.com/files/rs-3989278/v1/888c272d3ed9f292cb207ddb.png"},{"id":51801040,"identity":"5fef3d0d-7fd5-4a6e-8117-a8681c496823","added_by":"auto","created_at":"2024-02-29 09:10:17","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":1016486,"visible":true,"origin":"","legend":"\u003cp\u003eAlterations in AQP4 expression and AQP4 depolarization.\u003c/p\u003e\n\u003cp\u003e(A) Representative images of AQP4 expression in the corpus callosum, depicting depolarization of AQP4 in the CH group. (B) Quantification of AQP4 expression. (C) Schematic diagram illustrating the calculation of AQP4 polarization using low and high stringency. (D) AQP4 polarization index. (E) Correlation between AQP4 polarization index and ventricular volume. n = 12-14; scale bar = 500 µm (100 µm in the inset); *p \u0026lt; 0.05, ***p \u0026lt; 0.001; CH, chronic hydrocephalus; MFI, mean fluorescence intensity; au, arbitrary unit.\u003c/p\u003e","description":"","filename":"floatimage4.png","url":"https://assets-eu.researchsquare.com/files/rs-3989278/v1/47b2d83cdbe755b507d4db90.png"},{"id":51801043,"identity":"b3d23073-2d33-4864-89c9-b3ad3c40e643","added_by":"auto","created_at":"2024-02-29 09:10:18","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":905446,"visible":true,"origin":"","legend":"\u003cp\u003eDispersion of intraparenchymally injected tracers.\u003c/p\u003e\n\u003cp\u003e(A) Representative images illustrating intraparenchymal dispersion of CSF tracers at 6 h post-injection, quantified by covered area (B and D) or by fluorescence intensity (C and E). The black star in (A) indicates the intraparenchymal tracer injection site. n = 4-5; scale bar = 2 mm; *p \u0026lt; 0.05, **p \u0026lt; 0.01; CH, chronic hydrocephalus; TR-d3, Texas-red conjugated dextran; FITC-d49, fluorescein isothiocyanate conjugated dextran; MFI, mean fluorescence intensity; au, arbitrary unit.\u003c/p\u003e","description":"","filename":"floatimage6.png","url":"https://assets-eu.researchsquare.com/files/rs-3989278/v1/1246a5edce2f5302162728b1.png"},{"id":51801042,"identity":"05019e59-35fb-4c48-8fd0-a41a00f00ec5","added_by":"auto","created_at":"2024-02-29 09:10:18","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":1234256,"visible":true,"origin":"","legend":"\u003cp\u003eTransependymal and subarachnoid dispersion of icv injected EB.\u003c/p\u003e\n\u003cp\u003e(A) Representative images illustrating time-dependent transependymal dispersion of EB, quantified by covered area (B) or by fluorescence intensity (C), The red star in (A) indicates the icv EB injection site. n = 6-7. (D) Representative images illustrating time-dependent subarachnoid dispersion of EB, quantified by covered area (E) or by fluorescence intensity (F), n = 6-8. Scale bar = 2 mm; *p \u0026lt; 0.05, ***p \u0026lt; 0.001 (CH3h vs. Sham3h); †p \u0026lt; 0.05, ††p \u0026lt; 0.01, †††p \u0026lt; 0.001 (Sham3h vs. Sham24h); #p \u0026lt; 0.05 (CH3h vs. CH24h); §p \u0026lt; 0.05, §§p \u0026lt; 0.01, §§§p \u0026lt; 0.001 (Sham24h vs. CH24h); icv, intracerebroventricular injection; EB, Evans blue; CH, chronic hydrocephalus; MFI, mean fluorescence intensity; MGI, mean gray intensity; au, arbitrary unit.\u003c/p\u003e","description":"","filename":"floatimage8.png","url":"https://assets-eu.researchsquare.com/files/rs-3989278/v1/faa26c671762d84755971986.png"},{"id":51801036,"identity":"13257a34-b536-4544-bb56-e9076ddc1183","added_by":"auto","created_at":"2024-02-29 09:10:16","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":3205284,"visible":true,"origin":"","legend":"\u003cp\u003eTransependymal capillary filling of icv injected EB.\u003c/p\u003e\n\u003cp\u003eRepresentative images illustrating time-dependent capillary filling of EB in the corpus callosum imaged by confocal immunofluorescence with orthogonal view (A) and three-dimensional reconstruction from z-stacks (B). Single capillary vessels focused with a black arrowhead in (B) are magnified as insets. (C) Quantification of capillary densities, ***p \u0026lt; 0.001. (D) Time-dependent profiles of capillary filling, ***p \u0026lt; 0.001 (CH3h vs. Sham3h); †p \u0026lt; 0.05 (Sham3h vs. Sham24h). n = 14; Scale bar = 100 µm (50 µm in the inset); CH, chronic hydrocephalus; icv, intracerebroventricular injection; EB, Evans blue.\u003c/p\u003e","description":"","filename":"floatimage10.png","url":"https://assets-eu.researchsquare.com/files/rs-3989278/v1/0a38d95bce2318e0beec0053.png"},{"id":51801041,"identity":"557adadc-edef-4dec-81bd-58597cff6c7f","added_by":"auto","created_at":"2024-02-29 09:10:18","extension":"png","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":3085424,"visible":true,"origin":"","legend":"\u003cp\u003eMeningeal lymphatic drainage of icv injected EB.\u003c/p\u003e\n\u003cp\u003e(A) Representative images illustrating time-dependent EB staining of whole-mount meninges, co-stained with COLIV (meningeal vasculature) and LYVE1 (meningeal lymphatics). White boxes are focused on the middle meningeal arteries and further magnified with red boxes (i-xii). (B) Cross-section analysis of the meningeal lymphatic vessels with multi-signal intensities of COLIV/LYVE1/EB in the red boxes (i-xii). The lumen of meningeal lymphatic vessels is indicated by black dotted lines in (B). Meningeal EB staining quantified by covered area (C) or mean fluorescence intensity (D). **p \u0026lt; 0.01 (CH3h vs. Sham3h); †p \u0026lt; 0.05 (Sham3h vs. Sham24h). n = 11-13; scale bar = 2 mm (100 µm in the red boxes); CH, chronic hydrocephalus; icv, intracerebroventricular injection; EB, Evans blue; SSS, superior sagittal sinus; TS, transverse sinus; MMA, middle meningeal artery; MFI, mean fluorescence intensity; au, arbitrary unit.\u003c/p\u003e","description":"","filename":"floatimage12.png","url":"https://assets-eu.researchsquare.com/files/rs-3989278/v1/bef51b3e27f46f0befb71776.png"},{"id":51801045,"identity":"c49fe6f3-fb3e-40e3-b9d6-705696751321","added_by":"auto","created_at":"2024-02-29 09:10:19","extension":"png","order_by":8,"title":"Figure 8","display":"","copyAsset":false,"role":"figure","size":486921,"visible":true,"origin":"","legend":"\u003cp\u003ePeripheral lymphatic drainage of icv injected EB into the deep cervical lymph nodes.\u003c/p\u003e\n\u003cp\u003e(A) Representative images illustrating time-dependent EB-stained deep cervical lymph nodes (white arrowheads) and magnified with fluorescence images, quantified by covered area (B) or mean fluorescence intensity (C). n = 7-9; scale bar = 1 cm (100 µm in the inset); **p \u0026lt; 0.01 (CH3h vs. Sham3h); ††p \u0026lt; 0.01 (Sham3h vs. Sham24h); §§§p \u0026lt; 0.001 (Sham24h vs. CH24h); CH, chronic hydrocephalus; icv, intracerebroventricular injection; EB, Evans blue; MFI, mean fluorescence intensity; au, arbitrary unit.\u003c/p\u003e","description":"","filename":"floatimage14.png","url":"https://assets-eu.researchsquare.com/files/rs-3989278/v1/cfd7d1f7e654ee397f56ec40.png"},{"id":51829360,"identity":"bb532445-ec6a-4ad5-85d3-9944a7f675eb","added_by":"auto","created_at":"2024-02-29 17:48:12","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":7877550,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-3989278/v1/97d9992a-9a7f-42b4-a1ea-fe7efa2148e4.pdf"}],"financialInterests":"No competing interests reported.","formattedTitle":"Impaired Cerebrospinal Fluid Circulation and Cerebral Lymphatic Drainage in a Rat Model of Chronic Hydrocephalus","fulltext":[{"header":"Introduction","content":"\u003cp\u003eThe human brain is surrounded by cerebrospinal fluid (CSF), which is primarily produced by the choroid plexus and circulates within the intraventricular and subarachnoid spaces, eventually being absorbed through the arachnoid granulations into the dural venous sinuses [\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e]. The CSF plays an important role in protecting the brain from physical trauma. However, the CSF plays an even more crucial role in interacting with the brain parenchyma, helping to maintain homeostasis by removing metabolic waste generated by brain activity [\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e]. Recent research has highlighted the perivascular space and meningeal lymphatic channels as pathways for metabolic waste clearance, providing a new perspective on the role of CSF circulation in the brain [\u003cspan additionalcitationids=\"CR3\" citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eIn clinical practice, dysfunction of CSF circulation can manifest as communicating or non-communicating hydrocephalus depending on the presence of intraventricular obstruction [\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e]. Normal pressure hydrocephalus (NPH) is a type of communicating and chronic hydrocephalus characterized by increased CSF in the ventricles, often due to previous subarachnoid hemorrhage, infection, or head trauma, that chronically disrupts CSF circulation [\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e, \u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e]. However, it has been observed that many NPH patients do not have these pre-existing medical conditions, suggesting that NPH may be a manifestation of age-related or neurodegenerative dysfunction in CSF circulation [\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e]. Impaired glymphatic clearance has been documented in NPH patients using MRI studies after intrathecal contrast agent injection [\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e, \u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e] or diffusion tensor imaging [\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e]. Furthermore, detailed identification of meningeal lymphatic drainage channels has been achieved in both rodent models [\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e, \u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e] and humans [\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e, \u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e]. The circulation of CSF seems to be tightly regulated by the complex interplay between intraparenchymal glymphatic clearance activity and meningeal lymphatic drainage.\u003c/p\u003e \u003cp\u003eIn this study, our aim was to assess CSF circulatory dysfunction and cerebral lymphatic drainage from various perspectives, including the glymphatic system, transependymal CSF flow, subarachnoid CSF flow, meningeal lymphatic drainage, and peripheral lymphatic drainage to deep cervical lymph nodes. To achieve this, we used an animal model of chronic hydrocephalus induced by kaolin injection, a widely accepted method for replicating chronic hydrocephalus [\u003cspan additionalcitationids=\"CR14 CR15 CR16\" citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e] or NPH [\u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e] in rats. It was hypothesized that obstructing the intra-cisternal space through kaolin injection into the cisterna magna would impair CSF circulation.\u003c/p\u003e \u003cp\u003eTo validate the relevance of the model to chronic hydrocephalus, we performed measurements of ventricular enlargement and behavioral tests such as beam walking test. Intraparenchymal glymphatic clearance was evaluated by analyzing the dispersion of intraparenchymally injected CSF tracers. In addition, unbound Evans blue (EB), injected intraventricularly, was tracked at various sites to assess transependymal flow, subarachnoid flow, meningeal lymphatic drainage, and peripheral lymphatic drainage to deep cervical lymph nodes, providing insight into CSF circulation pathways and cerebral lymphatic drainage.\u003c/p\u003e \u003cp\u003eOur investigation aimed to understand how clinical conditions such as chronic hydrocephalus or NPH, which impair CSF circulation, glymphatic clearance, and cerebral lymphatic drainage, may contribute to the progression of neurodegenerative diseases by impairing the mechanism responsible for removing metabolic waste.\u003c/p\u003e"},{"header":"Material and Methods","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003eAnimal preparation\u003c/h2\u003e \u003cp\u003eMale Wistar rats (10 week old, 300\u0026ndash;320 g, Orient Bio) were used in all experiments after a two-week acclimatization period in the Konkuk University vivarium before the start of the experiment. Rats were housed under standard laboratory conditions (22\u0026thinsp;\u0026plusmn;\u0026thinsp;1\u0026deg;C temperature, 50\u0026thinsp;\u0026plusmn;\u0026thinsp;10% humidity, 12 h alternate light/dark cycle) with \u003cem\u003ead libitum\u003c/em\u003e access to food and water. Animal experimental procedures were performed in accordance with the ethical approval of the Institutional Animal and Use Committee of Konkuk University and ARRIVE guidelines (\u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003ehttps://www.nc3rs.org.uk/arrive-guidelines\u003c/span\u003e\u003cspan address=\"https://www.nc3rs.org.uk/arrive-guidelines\" targettype=\"URL\" class=\"RefTarget\"\u003e\u003c/span\u003e\u003c/span\u003e). Rats were induced with 3% isoflurane and then anesthetized intraperitoneally with Zoletil 50 (30 mg/kg) and Rompun (10 mg/kg) before being secured in a stereotaxic frame. All stereotaxic injection experiments (kaolin with intra-cisterna magna injection, icm; CSF tracer with intraparenchymal injection; EB with intracerebroventricular injection, icv) had common preparation procedures, including anesthesia, head fixation in the stereotaxic frame, and reflow prevention by leaving the injection cannula in place for an additional 3 min, differing only in the specifications of the Hamilton syringe connected to the syringe pump (Harvard) and the needle size. Rats with kaolin-induced hydrocephalus were assigned to the CH group. Meanwhile rats with sham surgery were classified as the sham group.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec4\" class=\"Section2\"\u003e \u003ch2\u003eInduction of kaolin-induced hydrocephalus\u003c/h2\u003e \u003cp\u003eA midline incision was made to expose the atlantooccipital membrane overlying the cisterna magna. A 60 \u0026micro;L of sterile suspension of kaolin (100 mg/mL in 0.9% saline, Sigma-Aldrich) was injected into the cisterna magna at a rate of 2.0 \u0026micro;L/s via a 28-gauge cannula connected to polyethylene tubing, attached to a 100 \u0026micro;L Hamilton syringe with a 22-gauge needle. Based on documented findings of normalized CSF dynamics [\u003cspan citationid=\"CR18\" class=\"CitationRef\"\u003e18\u003c/span\u003e, \u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e] and cerebral blood flow [\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e], changes in cerebral metabolism [\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e, \u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e], cessation of ventricular enlargement [\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e, \u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e, \u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e], cognitive decline [\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e, \u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e] and behavioral changes [\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e], our study established an 8-week post-kaolin infusion period for the development of a chronic hydrocephalus model in adult rats. All experimental evaluations were performed 8 weeks after kaolin induction of hydrocephalus.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec5\" class=\"Section2\"\u003e \u003ch2\u003eVentricular volume\u003c/h2\u003e \u003cp\u003eVentricular volumes were measured from multiple brain slices stained with cresyl violet acetate (Sigma-Aldrich) according to the methods described in our previous studies [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e, \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e]. Briefly, ventricular volumes, including the lateral and third ventricles, were calculated from the anterior horn of the lateral ventricle to the cerebral aqueduct using seven representative coronal slices at 2-mm intervals from bregma: AP\u0026thinsp;+\u0026thinsp;1.92 mm to -4.92mm. Ventricular volume was calculated by multiplying the slice interval by the ventricular area obtained using the National Institutes of Health (NIH) Image J program.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec6\" class=\"Section2\"\u003e \u003ch2\u003eBeam walking test\u003c/h2\u003e \u003cp\u003eThe beam walking test was performed as previously described [\u003cspan additionalcitationids=\"CR28\" citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e], with minor modifications. Rats were assessed for their ability to traverse two parallel plastic beams, each 2.0 cm in diameter and 95 cm in length, positioned horizontally 50 cm above the ground. Prior to the kaolin icm (before 1 week), the rats participated in an adaptation training session to freely traverse the beams, eliminating any signs of fear or hesitation, and subsequent tests were conducted on consecutive days 1, 3, 7, 14, 28, 42, and 56 after the kaolin injection. After being placed on a platform (30 \u0026times; 30 cm), we recorded the latency time and the number of footsteps (for each forelimb and hindlimb) for up to 60 s as the rats traversed from the moment their entire body entered the beam until they reached the opposite platform.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec7\" class=\"Section2\"\u003e \u003ch2\u003eIntraparenchymal injection of cerebrospinal fluid tracers\u003c/h2\u003e \u003cp\u003eWe performed intraparenchymal CSF tracer injection experiments as described in our previous study [\u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e]. Briefly, a 3 \u0026micro;L tracer solution containing Texas\u003cem\u003e-\u003c/em\u003ered conjugated dextran (TR-d3, MW 3 kDa, Thermo Fisher Scientific) and fluorescein isothiocyanate (FITC) conjugated dextran (FITC-d40, MW 40 kDa, Sigma-Aldrich), both at 0.5% concentration in a 1:1 ratio, was dissolved in artificial CSF (Tocris Bioscience). In the co-infusion experiment, the pre-prepared 3 \u0026micro;L tracer solution was administered into the brain parenchyma at a rate of 0.5 \u0026micro;L/min at bregma-related coordinates (AP\u0026thinsp;+\u0026thinsp;2.0 mm, ML\u0026thinsp;+\u0026thinsp;2.6 mm, DV -5.4 mm) using a 30-gauge injection cannula connected to polyethylene tubing, attached to a 10 \u0026micro;L Hamilton syringe with a 26s-gauge needle. Six hours after injection, rats were transcardially perfused with 0.01 M phosphate-buffered saline, pH 7.4 (1x PBS) and fixed with 4% paraformaldehyde (PFA) for fluorescence imaging analysis.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003ch2\u003eIntracerebroventricular injection of EB\u003c/h2\u003e \u003cp\u003eEB (DB 53, MW 0.96 kDA, Sigma-Aldrich) was suspended in 0.9% and injected into the ventricle saline (10 \u0026micro;L at 1.0 \u0026micro;L/min, 20 mg/mL) at coordinates relative to bregma (AP -0.9 mm, ML\u0026thinsp;+\u0026thinsp;1.4 mm, and DV -3.8 mm) using a 30-gauge injection cannula connected to polyethylene tubing, attached to a 25 \u0026micro;L Hamilton syringe with a 22-gauge needle. For imaging and quantitative analysis at 3 or 24 h after icv EB injection, rats were perfused with PBS, omitting the fixative solution, to facilitate neck incision for deep cervical lymph nodes harvesting, which was also used to obtain coronal brain slices and whole brain tissue to ensure experimental consistency. Meanwhile, EB-stained meninges were dissected without transcardiac perfusion of both PBS and PFA to preserve the integrity of meningeal lymphatic vessels.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec9\" class=\"Section2\"\u003e \u003ch2\u003eTissue preparation and immunohistochemistry\u003c/h2\u003e \u003cp\u003eAfter isoflurane anesthesia and transcardiac perfusion, rat brains were immediately removed, post-fixed overnight in the same fixative, sequentially cryoprotected in 30% sucrose solution, and were embedded in an optimal cutting temperature compound (Sakura Finetek). Brains were serially sectioned into 40 \u0026micro;m coronal slices using a CM1520 cryostat (Leica) for immunohistochemistry and tracer studies.\u003c/p\u003e \u003cp\u003eTo obtain whole mounts of rat dural meninges, we adapted methods from previously reported mouse experiments [\u003cspan additionalcitationids=\"CR31 CR32 CR33\" citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e], with minor modifications. Briefly, at 3 or 24 h after icv EB injection, anesthetized rats were decapitated above the shoulders, and the skin and muscle were removed from the outer skull. Using a bone cutter (Roboz), a precise dissection was made from the foramen magnum to the lateral end of the occipital bone, extending transversely in the longitudinal plane across the squamous and frontal bones to the junction of the frontal and nasal bones, taking care to minimize damage to the brain tissue. In order to obtain preserved meninges firmly attached to the adjacent skull, the skull bone was levered anteriorly outwards and meninges were gently separated from the bone using an elevator (Roboz). The separated EB-stained meninges, which were gently washed with PBS for 1 min to remove blood while retaining EB, were then used for immunohistochemistry and quantitative analysis of EB distribution.\u003c/p\u003e \u003cp\u003eTo obtain deep cervical lymph nodes, rats perfused only with PBS underwent a midline incision (5 mm above the clavicle) to expose the sternocleidomastoid muscle, which was then retracted with forceps, followed by post-fixation, cryoprotection, and embedding procedures (similar to brain sections) for quantitative analyses of EB distribution.\u003c/p\u003e \u003cp\u003eAfter sample preparation, immunofluorescence staining was performed to assess various pathological outcomes. Brain sections (either intact or stained with EB) were washed (1x PBS/0.3% Triton X-100) and then incubated in blocking serum (1x PBS/10% normal donkey serum/0.3% Triton X-100) for 1 h at room temperature (RT), while the meninges started the incubation process. Double-label immunofluorescence was performed overnight at 4\u0026deg;C in a solution (1x PBS/0.15% normal donkey serum/0.3% Triton X-100) containing the following primary antibodies: mouse anti-glial fibrillary acidic protein (GFAP, 1:1000, BD Bioscience), rabbit anti-ionized calcium binding adapter molecule-1 (Iba-1, 1:1000, Wako), rabbit anti-collagen IV (COLIV, 1:100, Abcam), mouse anti-aquaporin 4 (AQP4, 1:100, Abcam), mouse anti-collagen IV (COLIV, 1:100, Sigma-Aldrich), rabbit anti-lymphatic vessel endothelial hyaluronan receptor 1 (LYVE1, 1:100, Abcam). The sections were then washed (1x PBS/0.15% Triton X-100) and incubated for 3 h at RT in a secondary antibody solution as follows: anti-mouse Alexa Fluor 488 and 568; anti-rabbit Alexa Fluor 488 and 568; all from donkey (1:200, Invitrogen). Some of the coronal sections were counterstained with TO-PRO-3 (1:1,000, Invitrogen) for 30 min at RT. Stained sections were mounted on slides, dried for 30 min, and coverslipped with ProLong\u0026reg; Gold antifade reagent (Invitrogen), excluding structurally uneven meninges.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec10\" class=\"Section2\"\u003e \u003ch2\u003eAquaporin 4 polarization\u003c/h2\u003e \u003cp\u003eAstrocytic AQP4 polarization was assessed according to our previous study [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e]. Briefly, a low stringency threshold defined the total area of AQP4 immunoreactivity, whereas a high stringency threshold identified the vascular AQP4 colocalized with COLIV. Image J (NIH) was used for area calculations after black and white thresholding. The ratio of low stringency area to high stringency area was defined as AQP4 polarization.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec11\" class=\"Section2\"\u003e \u003ch2\u003eImage analysis and quantification\u003c/h2\u003e \u003cp\u003eBright field photographs were taken with a digital camera using a standard exposure time for all brains. Immunofluorescence images were obtained using a Carl Zeiss LSM 900 confocal laser scanning microscope at different magnifications (10\u0026times; air, NA 0.45; 20\u0026times; air, NA 0.80; 40\u0026times; water, NA 1.2; 63\u0026times; oil, NA 1.4) with a resolution of 1024 \u0026times; 1024 pixels, acquired by multichannel scanning in either a single or tiled frame. Capillary filling of EB into the microvessels in the corpus callosum was analyzed both by z-stack three-dimensional reconstruction (image slices taken at 22 \u0026micro;m intervals consisting of 12 z-sections), with upper and lower limits set based on positively visualized vascular morphology for COLIV and AQP4, and by fluorescence co-localization in the XY and orthogonal XZ, YZ projections of the image stacks. Image analysis and quantification was performed using Image J (NIH) and Zen blue image analysis wizard (Carl Zeiss).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec12\" class=\"Section2\"\u003e \u003ch2\u003eStatistics\u003c/h2\u003e \u003cp\u003eBehavioral data were analyzed using two-way repeated measures analysis of variance (ANOVA) and unpaired t-tests. Two-way ANOVA was used to assess the interaction between group and time, followed by a post-hoc Tukey\u0026rsquo;s honest comparison to analyze differences between groups over time. Time or group dependent differences were analyzed using unpaired t-tests, where applicable. Pearson\u0026rsquo;s correlation coefficient (r) was used for correlation analyses between ventricular volume and other variables. All data are expressed as the mean\u0026thinsp;\u0026plusmn;\u0026thinsp;SEM. A value of p\u0026thinsp;\u0026lt;\u0026thinsp;0.05 was considered to be statistically significant. Data analysis was performed with SPSS software version 27.0.\u003c/p\u003e \u003c/div\u003e"},{"header":"Results","content":"\u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003ch2\u003eCharacteristics of rats with chronic hydrocephalus\u003c/h2\u003e \u003cp\u003eEight weeks after icm injection of kaolin, the extracted brains showed a nearly uniform distribution of kaolin deposits throughout the ventral regions, extending from the optic nerves to the medulla and covering the interpeduncular fossa, pons, and cerebellum as indicated by the black lining (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eA) and kaolin-contacted inflammation as indicated by black arrowheads in the CH group (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eB). Marked hydrocephalus was observed, characterized by symmetrical enlargement of the lateral ventricles, cerebral aqueduct, and the fourth ventricle (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eB) with a quantitatively significant increase in total ventricular volume in the CH group (p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eC).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eModified beam walking test showed a significant difference in patterns (F\u003csub\u003e(7,273)\u003c/sub\u003e\u0026thinsp;=\u0026thinsp;15.359, p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, latency time in Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eD; F\u003csub\u003e(7,273)\u003c/sub\u003e\u0026thinsp;=\u0026thinsp;5.664, p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, the number of footsteps in Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eE). At the various time points, there was a significant increase in latency time (p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, post-op day 1; p\u0026thinsp;\u0026lt;\u0026thinsp;0.01, post-op day 3, 56; p\u0026thinsp;\u0026lt;\u0026thinsp;0.05, post-op day 7, 28; Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eD) and the number of footsteps (p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, post-op day 1; p\u0026thinsp;\u0026lt;\u0026thinsp;0.01, post-op day 3; p\u0026thinsp;\u0026lt;\u0026thinsp;0.05, post-op day 14, 56; Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eE) in the CH group compared to the sham group. A more frequent use of the forelimbs than the hindlimbs was observed in the CH group compared to the sham group on post-op day 1 (p\u0026thinsp;\u0026lt;\u0026thinsp;0.01, Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eF). The gait impairment in the CH group was pronounced within the first 7 days post-operation, with the impairment persisting for the entire 8-week period, suggesting a chronic motor function deficit in rats with kaolin-induced hydrocephalus.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec15\" class=\"Section2\"\u003e \u003ch2\u003eNeuroinflammation associated with kaolin-induced chronic hydrocephalus\u003c/h2\u003e \u003cp\u003eMeningeal fibrosis, characterized by chemical inflammatory arachnoiditis in the basal cistern due to direct contact with kaolin (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA) and indirect inflammation in the corpus callosum influenced by ventricular enlargement (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eB) are notable features in the CH group. Quantitative analysis revealed a significant increase in reactive astro- and microgliosis within the CH group, measured by area (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eC and \u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eM in GFAP; Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eE and \u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eO in Iba1), fluorescence intensity (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eD and \u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eN in GFAP; Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eF and \u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eP in Iba1), and the ratio of the number of Iba1-positive cells to total number of ToPr3-positive cells (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eG and \u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eQ). Furthermore, within the CH group, a positive correlation was observed between ventricular volume and GFAP signal measured by area (r\u0026thinsp;=\u0026thinsp;0.653, p\u0026thinsp;\u0026lt;\u0026thinsp;0.05, Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eH) and fluorescence intensity (r\u0026thinsp;=\u0026thinsp;0.605, p\u0026thinsp;\u0026lt;\u0026thinsp;0.05, Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eI), as well as Iba1 signal measured by area in the basal cistern (r\u0026thinsp;=\u0026thinsp;0.669, p\u0026thinsp;\u0026lt;\u0026thinsp;0.05, Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eJ). In addition, the Iba1 to To-pro3 positive cell ratio in the basal cistern (r\u0026thinsp;=\u0026thinsp;0.877, p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eL) and corpus callosum (r\u0026thinsp;=\u0026thinsp;0.437, p\u0026thinsp;\u0026lt;\u0026thinsp;0.05, Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eV) correlated with ventricular volume, suggesting a positive relationship between ventricular dilatation and the severity of neuroinflammation.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec16\" class=\"Section2\"\u003e \u003ch2\u003eAQP4 depolarization in the white matter in rats with chronic hydrocephalus\u003c/h2\u003e \u003cp\u003eChanges in AQP4 expression were evident in the corpus callosum of the CH group (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA). In addition to the increase in parenchymal AQP4 expression (p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB), the distribution pattern of AQP4 also changed, indicating redistribution into the brain parenchyma from the location on perivascular astrocyte endfeet in close contact with COLIV-positive endothelial vessels (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eC). Perivascular AQP4 localization, indicative of the AQP4 polarization ratio, was significantly reduced in the CH group compared to the sham group (p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eD). This loss of AQP4 polarization also correlated well with ventricular dilatation (r = -0.586, p\u0026thinsp;\u0026lt;\u0026thinsp;0.05, Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eE).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec17\" class=\"Section2\"\u003e \u003ch2\u003eDelayed dispersion of intraparenchymally injected tracers in rats with chronic hydrocephalus\u003c/h2\u003e \u003cp\u003eIntraparenchymally injected tracers of different molecular weights exhibited delayed dispersion patterns at 6 h post-injection in the CH group compared to the sham group (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eA). The black star in Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eA indicates the intraparenchymal tracer injection site. Tracer dispersion into the brain parenchyma was impaired in the CH group in multiple brain slices in both cases of TR-d3 (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05 at 4.20 mm and 1.08 mm, p\u0026thinsp;\u0026lt;\u0026thinsp;0.01 at 3.24 mm, assessed by area in Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eB; p\u0026thinsp;\u0026lt;\u0026thinsp;0.05 at 4.20 mm, p\u0026thinsp;\u0026lt;\u0026thinsp;0.01 at 3.24 mm, assessed by fluorescence intensity in Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eC) and FITC-d40 (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05 at 3.24 mm, assessed by area in Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eD; p\u0026thinsp;\u0026lt;\u0026thinsp;0.01 at 3.24 mm, assessed by fluorescence intensity in Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eE). The lighter TR-d3 tracer exhibited greater dispersion both anteriorly and posteriorly from the injection site than the heavier FITC-d40 tracer.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec18\" class=\"Section2\"\u003e \u003ch2\u003eTransependymal and subarachnoid dispersion of icv-injected EB in rats with chronic hydrocephalus\u003c/h2\u003e \u003cp\u003eTransependymal dispersion of icv-injected EB was visibly present at 3 hours post-injection and disappeared by 24 hours in the sham group, whereas it was scarce in the CH group (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eA). The red star in Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eA indicates the icv EB injection site. Five sections around the injection site were selected to assess the pattern of dispersion depending on time or model. In multiple sections, transependymal dispersion was significantly impeded in the CH group compared to the sham group throughout the 24 hours when assessed by the EB-stained area (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05 at 0.00 mm, p\u0026thinsp;\u0026lt;\u0026thinsp;0.001 at -0.96 mm, CH3h vs. Sham3h; p\u0026thinsp;\u0026lt;\u0026thinsp;0.01 at 0.00 mm, p\u0026thinsp;\u0026lt;\u0026thinsp;0.05 at -0.96 mm, CH24h vs. Sham24h; Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eB) and fluorescence intensity (p\u0026thinsp;\u0026lt;\u0026thinsp;0.001 at 0.96, 0.00, and \u0026minus;\u0026thinsp;0.96 mm, p\u0026thinsp;\u0026lt;\u0026thinsp;0.05 at -1.92mm, CH3h vs. Sham3h; p\u0026thinsp;\u0026lt;\u0026thinsp;0.001 at 0.96, 0.00, and \u0026minus;\u0026thinsp;0.96 mm, p\u0026thinsp;\u0026lt;\u0026thinsp;0.05 at -2.92 mm, CH24h vs. Sham24h; Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eC). While the sham group showed a time-dependent washout pattern with a peak at 3 hours post-injection and a subsequent decrease at 24 hours (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05 at -0.96 mm, Sham3h vs. Sham24h, assessed by area in Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eB; p\u0026thinsp;\u0026lt;\u0026thinsp;0.05 at 0.00 mm, p\u0026thinsp;\u0026lt;\u0026thinsp;0.01 at -0.96mm, p\u0026thinsp;\u0026lt;\u0026thinsp;0.001 at 0.96, -1.92 and \u0026minus;\u0026thinsp;2.92 mm, Sham3h vs. Sham24h, assessed by fluorescence intensity in Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eC), the CH group showed minimal initial dispersion into the transependymal region and a persistent pattern.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eSubarachnoid dispersion of icv-injected EB was also hindered in the CH group throughout the 24 h compared to the sham group (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eD). EB staining was quantified by assessing views from the four different angles including caudal, ventral, lateral, and dorsal angels. Subarachnoid dispersion was significantly impeded in the CH group compared to the sham group when assessed by the EB-stained area (p\u0026thinsp;\u0026lt;\u0026thinsp;0.001 at ventral angle, CH3h vs. Sham3h in Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eE) and fluorescence intensity (p\u0026thinsp;\u0026lt;\u0026thinsp;0.001 at ventral, lateral, and caudal angles, CH3h vs. Sham3h in Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eF). The time-dependent washout pattern was observed in the sham group (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05 at caudal, p\u0026thinsp;\u0026lt;\u0026thinsp;0.001 at ventral, lateral, and dorsal angles, Sham3h vs. Sham 24h, assessed by fluorescence intensity in Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eF), whereas it was not evident in the CH group except at ventral angle (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05, assessed by fluorescence intensity in Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eF).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec19\" class=\"Section2\"\u003e \u003ch2\u003eTransependymal capillary filling of icv-injected EB in rats with chronic hydrocephalus\u003c/h2\u003e \u003cp\u003eExamination by confocal microscopy revealed apparent transependymal capillary filling of the icv-injected EB in the corpus callosum of the sham group, whereas it was impaired in the CH group (orthogonal views in Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eA and z-stack 3D in Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eB). Capillary density measured by COLIV-positive blood vessels within the unit region (%) showed a significant decrease in the CH group compared to the sham group (p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eC).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eIn the sham group, the capillary filling of the EB, which was evident at 3 hours post-injection, disappeared at 24 hours. In contrast, transependymal capillary filling of EB in the CH group was minimal throughout the 24 hours. Quantitative evaluation using the ratio of EB-filled capillary length to the total capillary length showed peaks at 3 hours, indicating rapid capillary filling in the sham group compared to the CH group (p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, CH3h vs. Sham3h, Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eD), and declines at 24 hours, indicating subsequent capillary emptying (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05, Sham3h vs. Sham24h, Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eD). In contrast, the CH group showed peaks at 24 hours instead of at 3 hours, indicating delayed capillary filling.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec20\" class=\"Section2\"\u003e \u003ch2\u003eMeningeal lymphatic drainage in rats with chronic hydrocephalus\u003c/h2\u003e \u003cp\u003eImmunohistochemistry using whole-mount meninges revealed the colocalization of scattered lymphatic endothelial signals (LYVE1) alongside meningeal vascular signals (COLIV), as shown in Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eA. Cross-sectional analysis of multi-immunofluorescence signals showed red peaks (LYVE1) surrounding green peaks (COLIV) in Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eB. After 3 hours of icv-injected EB in the sham group, distinct blue peaks (EB) were observed between the two red peaks, as shown by the black dotted lines indicating the lumen of meningeal lymphatic vessels in Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eB, suggesting effective EB influx into the meningeal lymphatic vessels. This distinct pattern of blue peaks disappears at post-24 hours of icv-injected EB in the sham group, indicating subsequent efficient drainage of EB via meningeal lymphatic vessels. In contrast to the sham group, the CH group showed no clear influx of EB into the meningeal lymphatic vessels and subsequent drainage.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eQuantitative analysis using EB signals on the meninges revealed a consistent impairment of meningeal lymphatic drainage in the CH group throughout the 24 hours compared to the sham group. This impairment is evident in the CH group when assessing CH3h vs. Sham3h, as indicated by the EB-stained area (p\u0026thinsp;\u0026lt;\u0026thinsp;0.01, Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eC) or fluorescence intensity (p\u0026thinsp;\u0026lt;\u0026thinsp;0.01, Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eD). In contrast to the normal meningeal lymphatic drainage pattern in the sham group, which peaked at 3 hours and then disappeared at 24 hours (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05, Sham3h vs. Sham24h, Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eC), the CH group showed minimal influx into the meningeal lymphatic vessels and a stagnant pattern.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec21\" class=\"Section2\"\u003e \u003ch2\u003ePeripheral lymphatic drainage in rats with chronic hydrocephalus\u003c/h2\u003e \u003cp\u003eTime-dependent peripheral lymphatic drainage of icv-injected EB into the deep cervical lymph nodes is shown in both groups. Comparing the two groups at different time points, EB signals were evident at 3 hours post-injection and peaked at 24 hours in the sham group, while a relatively weaker signal was observed in the CH group (Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eA). Peripheral lymphatic drainage of EB was consistently impaired in the CH group compared to the sham group throughout the 24 hours (p\u0026thinsp;\u0026lt;\u0026thinsp;0.01, CH3h vs. Sham3h; p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, CH24h vs. Sham24 h, assessed by area in Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eB; p\u0026thinsp;\u0026lt;\u0026thinsp;0.01, CH3h vs. Sham3h; p\u0026thinsp;\u0026lt;\u0026thinsp;0.001, CH24 h vs. Sham24 h, assessed by fluorescence intensity in Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eC). Although both groups showed a time-dependent increase in peripheral lymphatic drainage of EB to the deep cervical lymph nodes, this was more pronounced in the sham group (p\u0026thinsp;\u0026lt;\u0026thinsp;0.01, Sham3h vs. Sham24h, assessed by area in Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eB or fluorescence intensity in Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003eC).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e"},{"header":"Discussion","content":"\u003cp\u003eThe experimental findings using the kaolin-induced hydrocephalus rat model demonstrate how chronic hydrocephalus can disrupt the dynamics of CSF circulation associated with the glymphatic system and cerebral lymphatic drainage. It is hypothesized that the injection of kaolin into the rat cisterna magna, which limits CSF absorption through the subarachnoid space, could serve as a reliable model for studying clinical conditions such as chronic hydrocephalus or NPH, a communicating type of hydrocephalus. Although we acknowledge that the kaolin-induced hydrocephalus model, in which the CSF absorption pathway is artificially obstructed by injected kaolin, may not perfectly mimic the pathophysioloy and clinical features of chronic hydrocephalus or NPH, we have modified our experimental settings to resemble chronic hydrocephalus or NPH observed in humans. The aim was to create a chronic and mild condition similar to NPH without directly obstructing the intraventricular circulation of CSF. To achieve this, we used a minimal amount of kaolin and prolonged the duration of the hydrocephalus.\u003c/p\u003e \u003cp\u003eInjection of kaolin into the cisterna magna is known to induce inflammation, which leads to meningeal fibrosis, specifically chemical arachnoiditis [\u003cspan additionalcitationids=\"CR36 CR37\" citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e]. The majority of the kaolin deposits were distributed throughout the basal subarachnoid space, evenly on both sides of the ventral brainstem, including the interpeduncular fossa to the medulla and pons. No significant presence of kaolin was observed within the intraventricular system, particularly blocking the outlet of the fourth ventricle. The icv-injected EB diffused throughout the entire subarachnoid space through the outlet of the fourth ventricle. This diffusion pattern of EB indicates that CSF flow through the intraventricular and subarachnoid spaces was not mechanically obstructed in the kaolin-induced chronic hydrocephalus rat model.\u003c/p\u003e \u003cp\u003eWhen discussing hydrocephalus, it is important to differentiate between acute and chronic conditions based on duration. The kaolin-induced hydrocephalus model is a well-known animal model for chronic hydrocephalus as it develops and progresses hydrocephalus over several weeks to months [\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e, \u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e, \u003cspan additionalcitationids=\"CR40\" citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e41\u003c/span\u003e]. Injected kaolin in the basal subarachnoid space can cause acute hydrocephalic symptoms within two weeks, leading to an increase in resistance to CSF outflow and subsequent elevation of intracranial pressure [\u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e, \u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e]. As hydrocephalus progresses from the acute phase to the chronic phase over a longer period of four to six weeks or more, the resistance to CSF outflow and intracranial pressure undergo changes [\u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e, \u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e]. Animal studies using MRI have shown ventricular enlargement, with a peak at 6 weeks, followed by a subsequent decline at 10 weeks [\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e]. The progressive increase in ventricular size up to 8 weeks reaches a new steady state characterized by the restoration of normal intracranial pressure and an increase in resistance to CSF outflow [\u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e]. Moreover, the normalization of cerebral blood flow after 8 weeks indicates that the post-operative 8-week time point we chose for evaluation is optimal for investigating chronic hydrocephalus, such as NPH [\u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e40\u003c/span\u003e]. Our model also identified gait impairments, which are clinical manifestations observed in human NPH [\u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e]. Although the kaolin-induced hydrocephalus rat model has inherent limitations, it has potential for investigating the impact of chronic hydrocephalus on the glymphatic system and cerebral lymphatic drainage in neurodegenerative diseases, particularly in relation to the accumulation of metabolic waste.\u003c/p\u003e \u003cp\u003eThe kaolin-induced hydrocephalus rat model\u0026rsquo;s pathophysiology is mainly focused on neuroinflammatory changes and alterations of AQP4, the brain\u0026rsquo;s most abundant water channel that regulates water homeostasis [\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e]. Neuroinflammation with gliosis in the basal forebrain and corpus callosum may be indirectly triggered by increased pressure in the subarachnoid or intraventricular space, in addition to the reactive gliosis due to direct contact with kaolin. The degree of gliosis is strongly correlated with enlarged ventricular volume, which may indirectly support the possibility of pressure-related gliosis. A significant depolarization of AQP4 was observed, resulting in the translocation of the normal perivascular pattern of AQP4 to the parenchymal pattern with an increased total AQP4 signal in the CH group. The evidence indicates that neuroinflammation and depolarization of AQP4 play a significant role in the pathophysiology of impaired CSF circulation in chronic hydrocephalus.\u003c/p\u003e \u003cp\u003ePrevious studies have reported impaired glymphatic system associated with the depolarization of AQP4 [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e, \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e]. In this study, we investigated the glymphatic efflux of intraparenchymally injected tracers with two different molecular weights. As anticipated, the molecular weight-dependent intraparenchymal dispersion of the tracers at 6 hours post-injection was hindered in the CH group, indicating a compromised glymphatic system in the chronic hydrocephalus.\u003c/p\u003e \u003cp\u003ePrevious studies have evaluated the classical CSF circulation pathway, which involves production in the choroid plexus, intraventricular circulation, and absorption into the dural venous sinuses through the arachnoid villi [\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e, \u003cspan citationid=\"CR44\" class=\"CitationRef\"\u003e44\u003c/span\u003e]. To assess this pathway, we measured the subarachnoid dispersion of icv-injected EB. Analysis of the subarachnoid brain surface area covered by EB in the sham group revealed rapid subarachnoid dispersion at 3 hours post-injection, which was subsequently eliminated by 24 hours post-injection. In contrast, the CH group exhibited delayed and stagnated dispersion into the subarachnoid space, indicating a disturbance of the classical CSF circulation pathway in the chronic hydrocephalus.\u003c/p\u003e \u003cp\u003eFurthermore, we investigated an alternative transependymal pathway for CSF circulation. This pathway was discovered in a study that utilized precise MRI mapping of CSF flow with a tracer injected into the ventricle of healthy rats [\u003cspan citationid=\"CR45\" class=\"CitationRef\"\u003e45\u003c/span\u003e]. The investigation identified previously unrecognized parenchymal perivascular space connections that spread across various brain regions, facilitating the direct transport of CSF from the ventricles to the subarachnoid space [\u003cspan citationid=\"CR45\" class=\"CitationRef\"\u003e45\u003c/span\u003e]. In studies of hydrocephalus in humans, the absorption of CSF by periventricular tissues serves as a compensatory mechanism for increased intracranial pressure and sheds light on an alternative pathway for CSF circulation [\u003cspan additionalcitationids=\"CR47\" citationid=\"CR46\" class=\"CitationRef\"\u003e46\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR48\" class=\"CitationRef\"\u003e48\u003c/span\u003e]. Consistent with these studies, our analysis of coronal brain slices stained by EB revealed rapid periventricular diffusion of icv-injected EB at 3 hours post-injection and subsequent elimination at 24 hours post-injection in the sham group, indicating active transependymal CSF flow in the normal condition. Conversely, minimal periventricular diffusion of EB was observed in the CH group. Based on the confocal microscopic image of the corpus callosum in the sham group, the capillaries were completely filled with EB three hours after injection and showed clear drainage within 24 hours after injection. The microvascular density was found to be deceased in the CH group compared to the sham group, which is consistent with previous studies reporting a decrease in microvascular density in chronic hydrocephalus [\u003cspan additionalcitationids=\"CR50\" citationid=\"CR49\" class=\"CitationRef\"\u003e49\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e51\u003c/span\u003e]. In the CH group, there was a decrease in capillary density and no visible diffusion of EB into the capillaries even after 24 hours. These findings suggest a disturbance of the alternative transependymal CSF circulation pathway in chronic hydrocephalus, as evidenced by compromised periventricular diffusion and capillary drainage of EB.\u003c/p\u003e \u003cp\u003eIn the animal study, the final step of CSF efflux into the venous sinuses through the arachnoid villi was demonstrated using serial high resolution MRI tracking of injected tracer into the lateral ventricles [\u003cspan citationid=\"CR45\" class=\"CitationRef\"\u003e45\u003c/span\u003e]. Time-dependent EB drainage through the venous sinus was observed in the sham group, supporting the classical CSF circulation pathway via venous sinus drainage. Analysis of EB staining on the meninges showed strong staining around the venous sinuses and the middle meningeal artery.\u003c/p\u003e \u003cp\u003eRecent research has also revealed the presence of lymphatic vessels in the meninges responsible for draining CSF, interstitial fluid, macromolecules, and immune cells to the cervical lymph nodes [\u003cspan citationid=\"CR52\" class=\"CitationRef\"\u003e52\u003c/span\u003e, \u003cspan citationid=\"CR53\" class=\"CitationRef\"\u003e53\u003c/span\u003e]. In a previous study using novel lymphatic reporter rats, it was demonstrated that meningeal lymphatic vessels were located alongside the middle meningeal artery, superior sagittal sinus, and transverse sinuses [\u003cspan citationid=\"CR54\" class=\"CitationRef\"\u003e54\u003c/span\u003e]. In our study, we used a whole-mount dissection of the dura mater and found that LYVE1-positive meningeal lymphatic vessels appeared as discontinuous dotted lines scattered along the COLIV-positive meningeal arteries. Cross-sectional analysis of multi-immunofluorescence signals revealed intraluminal filling and subsequent drainage of EB through the meningeal lymphatic vessels as a CSF efflux pathway. In the CH group compared to the sham group, the EB signal did not appear within the meningeal lymphatic vessels, although the structure of the meningeal lymphatic vessels seemed intact. This suggests a diminished functional CSF efflux through the meningeal lymphatic vessels in chronic hydrocephalus. As expected, peripheral lymphatic drainage to the deep cervical lymph nodes via the meningeal lymphatic vessels or other possible routes was also delayed in the CH group.\u003c/p\u003e"},{"header":"Conclusions","content":"\u003cp\u003eOur study evaluated several crucial steps in the CSF circulation pathway over time, including the glymphatic system, transependymal CSF flow, subarachnoid CSF flow, efflux through the venous sinus and meningeal lymphatic vessels, as well as peripheral lymph nodes drainage. Recognizing the role of the glymphatic system and its connection with cerebral lymphatic drainage, which functions as a clearance system for amyloid, tau, and other brain metabolic waste [\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e, \u003cspan citationid=\"CR55\" class=\"CitationRef\"\u003e55\u003c/span\u003e], clinical conditions such as chronic hydrocephalus or NPH may pose a significant risk for the progression of various neurodegenerative diseases, including Alzheimer\u0026rsquo;s disease. Therefore, it will be critical to conduct extensive research on the circulation of CSF and cerebral lymphatic drainage to identify risk factors and understand the pathophysiology of neurodegenerative diseases, which may lead to novel therapeutic strategies.\u003c/p\u003e"},{"header":"Abbreviations","content":"\u003cp\u003eCSF\u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp;\u0026nbsp;cerebrospinal fluid\u003c/p\u003e\n\u003cp\u003eNPH\u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp;\u0026nbsp;normal pressure hydrocephalus\u003c/p\u003e\n\u003cp\u003eEB\u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp;\u0026nbsp;Evans blue\u003c/p\u003e\n\u003cp\u003eNIH\u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp;National Institutes of Health\u003c/p\u003e\n\u003cp\u003eFITC\u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp;fluorescein isothiocyanate\u003c/p\u003e\n\u003cp\u003ePFA\u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp;paraformaldehyde\u003c/p\u003e\n\u003cp\u003eRT\u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp;room temperature\u003c/p\u003e\n\u003cp\u003eGFAP\u0026nbsp; \u0026nbsp; \u0026nbsp;\u0026nbsp;glial fibrillary acidic protein\u003c/p\u003e\n\u003cp\u003eIba-1\u0026nbsp; \u0026nbsp; \u0026nbsp; \u0026nbsp;ionized calcium binding adapter molecule-1\u003c/p\u003e\n\u003cp\u003eCOLIV\u0026nbsp; \u0026nbsp;collagen IV\u003c/p\u003e\n\u003cp\u003eLYVE1\u0026nbsp; \u0026nbsp;lymphatic vessel endothelial hyaluronan receptor 1\u003c/p\u003e\n\u003cp\u003eANOVA\u0026nbsp;analysis of variance\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cbr\u003e\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eEthics approval and consent to participate\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll animal experimental procedures were in accordance with the approved ethical standards of the Institutional Animal and Use Committee of Konkuk University.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eConsent for publication\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eNot applicable\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAvailability of data and material\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eCompeting interests\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare that they have no competing interests.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunding\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis work was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education (2020R1F1A1076085).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthors\u0026apos; contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eDB, BC, and HK participated in the conception and design of the study. DB and HK drafted the manuscript and participated in general management. DB, KK, and DC helped acquisition and analysis of data. CS and JL helped to draft the manuscript. All authors reviewed and approved the final manuscript.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eAcknowledgements\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eNot applicable\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n\u003cli\u003eKarimy JK, Reeves BC, Damisah E, Duy PQ, Antwi P, David W, Wang K, Schiff SJ, Limbrick DD, Jr., Alper SL\u003cem\u003e et al\u003c/em\u003e: Inflammation in acquired hydrocephalus: pathogenic mechanisms and therapeutic targets. \u003cem\u003eNat Rev Neurol \u003c/em\u003e2020, 16(5):285-296.\u003c/li\u003e\n\u003cli\u003eTarasoff-Conway JM, Carare RO, Osorio RS, Glodzik L, Butler T, Fieremans E, Axel L, Rusinek H, Nicholson C, Zlokovic BV\u003cem\u003e et al\u003c/em\u003e: Clearance systems in the brain-implications for Alzheimer disease. \u003cem\u003eNat Rev Neurol \u003c/em\u003e2015, 11(8):457-470.\u003c/li\u003e\n\u003cli\u003eRasmussen MK, Mestre H, Nedergaard M: The 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Alterations in brain metabolism, CNS morphology and CSF dynamics in adult rats with kaolin-induced hydrocephalus. \u003cem\u003eBrain research \u003c/em\u003e2002, 927(1):35-41.\u003c/li\u003e\n\u003cli\u003eLi J, McAllister II JP, Shen Y, Wagshul ME, Miller JM, Egnor MR, Johnston MG, Haacke EM, Walker ML: Communicating hydrocephalus in adult rats with kaolin obstruction of the basal cisterns or the cortical subarachnoid space. \u003cem\u003eExperimental neurology \u003c/em\u003e2008, 211(2):351-361.\u003c/li\u003e\n\u003cli\u003eNagra G, Li J, McAllister J, Miller J, Wagshul M, Johnston M: Impaired lymphatic cerebrospinal fluid absorption in a rat model of kaolin-induced communicating hydrocephalus. \u003cem\u003eAmerican Journal of Physiology-Regulatory, Integrative and Comparative Physiology \u003c/em\u003e2008, 294(5):R1752-R1759.\u003c/li\u003e\n\u003cli\u003eNagra G, Wagshul ME, Rashid S, Li J, McAllister JP, Johnston M: Elevated CSF outflow resistance associated with impaired 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Schlemm\u0026rsquo;s canal reporter rat. \u003cem\u003eScientific reports \u003c/em\u003e2017, 7(1):5577.\u003c/li\u003e\n\u003cli\u003eReeves BC, Karimy JK, Kundishora AJ, Mestre H, Cerci HM, Matouk C, Alper SL, Lundgaard I, Nedergaard M, Kahle KT: Glymphatic System Impairment in Alzheimer\u0026apos;s Disease and Idiopathic Normal Pressure Hydrocephalus. \u003cem\u003eTrends Mol Med \u003c/em\u003e2020, 26(3):285-295.\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":true,"highlight":"","institution":"","isAcceptedByJournal":false,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true},"keywords":"Cerebrospinal fluid circulation, hydrocephalus, glymphatic, meningeal lymphatic, kaolin, animal model","lastPublishedDoi":"10.21203/rs.3.rs-3989278/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-3989278/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003ch2\u003eBackground\u003c/h2\u003e \u003cp\u003eThe cerebrospinal fluid (CSF) not only protects the brain but also maintains homeostasis by removing metabolic waste produced by brain activity. This study hypothesizes that chronic CSF circulatory dysfunction, such as chronic hydrocephalus or normal pressure hydrocephalus (NPH), may be a critical condition in neurodegenerative diseases associated with metabolic waste accumulation.\u003c/p\u003e\u003ch2\u003eMethods\u003c/h2\u003e \u003cp\u003eTo investigate the CSF circulation and cerebral lymphatic drainage in a rat model of chronic hydrocephalus induced by kaolin injection, we performed time-dependent evaluations of intraparenchymal injection of tracers as well as intraventricular injection of Evans blue. The study systemically evaluated the dysfunction of CSF circulation and lymphatic drainage in the brain from various perspectives, including the glymphatic system, transependymal CSF flow, subarachnoid CSF flow, meningeal lymphatic drainage, and peripheral lymphatic drainage to deep cervical lymph nodes.\u003c/p\u003e\u003ch2\u003eResults\u003c/h2\u003e \u003cp\u003eThe results indicated delayed CSF circulation, including glymphatic system, and cerebral lymphatic drainage in the kaolin-induced chronic hydrocephalus model. Based on these findings, our research indicated that dysfunction of CSF circulation, as observed in conditions such as chronic hydrocephalus or NPH, may act as an initiating or exacerbating factor in neurodegenerative diseases.\u003c/p\u003e\u003ch2\u003eConclusions\u003c/h2\u003e \u003cp\u003eThis can lead to the accumulation of metabolic waste, as seen in Alzheimer's disease. Our research can help identify risk factors and provide insight into the underlying pathophysiology of neurodegenerative diseases, which may lead to the development of novel therapeutic strategies.\u003c/p\u003e","manuscriptTitle":"Impaired Cerebrospinal Fluid Circulation and Cerebral Lymphatic Drainage in a Rat Model of Chronic Hydrocephalus","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2024-02-29 09:10:07","doi":"10.21203/rs.3.rs-3989278/v1","editorialEvents":[{"type":"communityComments","content":0}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"15c65ff1-b51e-4590-97ea-20c5b377efb1","owner":[],"postedDate":"February 29th, 2024","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"posted","subjectAreas":[],"tags":[],"updatedAt":"2024-03-18T15:27:30+00:00","versionOfRecord":[],"versionCreatedAt":"2024-02-29 09:10:07","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-3989278","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-3989278","identity":"rs-3989278","version":["v1"]},"buildId":"omnImTCwR2MFx8CMYfrG7","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}