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Reactive astrocyte-derived exosomes enhance intracranial lymphatic drainage in mice after intracranial hemorrhage

Fluids and Barriers of the CNS volume 22, Article number: 37 (2025) Cite this article

Abstract

Background

After intracranial hemorrhage (ICH), the formation of primary hematoma foci leads to the development of secondary brain injury factors such as perihematomal edema (PHE) and accumulation of toxic metabolites, which severely affect the survival and prognosis of patients. The intracerebral lymphatic system, proposed by Jeffrey J. Iliff et al., plays an important role in central nervous system (CNS) fluid homeostasis and waste removal, while reactive astrocyte-derived exosomes have shown therapeutic potential in CNS disorders. Our study focuses on the effects of hemin-treated reactive astrocyte-derived exosomes on the functional integrity of the glymphatic system (GLS) after ICH and their potential mechanism of action in repairing brain injury.

Methods

Hemin, an iron-rich porphyrin compound, was used to construct the in vitro model of ICH. Primary astrocytes were treated with complete medium supplemented with different concentrations of hemin to obtain exosomes secreted by them, and mice with ICH induced by the collagenase method were intervened by intranasal administration. Solute clearance efficiency was assessed by intracranial injection of cerebrospinal fluid tracers and fluorescent magnetic beads. Immunofluorescence analysis of Aquaporin 4 (AQP4) polarization and astrocyte proliferation. Magnetic Resonance Imaging was used to visualize and quantify the volume of hematoma foci and PHE, and Western Blot was used to analyze the accumulation of toxic metabolites, while neuronal apoptosis was detected by a combination of TUNEL assay apoptosis detection kit and Nissl staining, and their functional status was analyzed. Gait analysis software was used to detect functional recovery of the affected limb in mice.

Results

Exosomes from hemin treated astrocytes facilitated the recovery of AQP4 polarization and attenuated astrocyte proliferation around hematoma foci in mice with ICH, thereby promoting the recovery of the GLS. Meanwhile, exosomes from hemin treated astrocytes reduced PHE and toxic protein accumulation, decreased apoptosis of cortical neurons on the affected side, and facilitated recovery of motor function of the affected limb, and these effects were blocked by TGN020, an AQP4-specific inhibitor.

Conclusions

Exosomes from hemin treated astrocytes attenuated secondary brain injury and neurological deficits in mice with ICH by promoting the repair of GLS injury.

Background

ICH is a relatively common type of stroke, accounting for approximately 15% of all strokes [1], with a mortality rate of up to 50% within 30 days [2]. In patients with primary ICH, microangiopathy due to hypertension is a major risk factor and occurs in the deeper portions of the brain parenchyma that are supplied by small blood vessels, including the chiasmatic nuclei, caudate nucleus, thalamus, and brainstem [3]. In patients with ICH, microvascular disease due to hypertension is a major risk factor. Prolonged high blood pressure causes fast-flowing blood to strike the walls of these vessels, which run vertically from the middle cerebral artery into the brain parenchyma. The damaged walls eventually rupture and bleed, and blood flows into the brain parenchyma, causing a variety of primary and secondary brain injuries. Primary brain injury is primarily caused by early hematoma foci with occupying effects and can develop within hours of the hemorrhage. Secondary brain injury is often caused by inflammatory cell recruitment, accumulation of toxic metabolites, etc. secondary to the primary brain injury, resulting in hematoma expansion, PHE formation, and other damage [4, 5].

Long before Jeffrey J. Iliff’s research team first introduced the concept of GLS in 2012, it was thought that the CNS lacked parenchymal lymphatic vasculature to support humoral homeostasis [6,7,8,9,10]. The absence of lymphatic vessels in the brain presents a conceptual challenge to understanding how fluid homeostasis and waste removal can be achieved within the confines of the CNS [11]. The GLS can be roughly divided into three parts: the arterial end, the CSF-interstitial fluid exchange zone, and the venous end. First, cerebrospinal fluid is generated from the choroid plexus and flows from the subarachnoid space to the brain parenchyma through the periarterial space, known as the Virchow-Robin lumen [12]. Then, cerebrospinal fluid exits the periarterial space and mixes and exchanges with the interstitial fluid, carrying with it substances such as metabolic waste produced by the cells of the brain parenchyma. Finally, it leaves the brain through the perivenous space and enters the lymphatic network, such as the meningeal lymphatics and the deep cervical lymph nodes [13]. The large amount of AQP4 expressed in the polarized distribution of perivascular astrocytes is thought to play a critical role in the flow of cerebrospinal fluid from the arterial end into the interstitium [14]. In animal models, AQP4 knockout mice showed significant inhibition of both cerebrospinal fluid tracer drainage and β-amyloid clearance compared to wild-type mice [15]. In patients with ICH, damage to the GLS caused by reactive proliferation of astrocytes around hematoma foci and depolarizing overexpression of AQP4 contributes to disease onset and progression by inhibiting clearance of toxic metabolic waste from the brain and promoting PHE formation [16,17,18,19].

Extracellular Vesicles are spherical structures formed by a phospholipid bilayer wrapped around a content, with sizes ranging from 30 nm to 1000 nm. The sizes between 30 nm and 150 nm are known as exosomes, which contain a variety of biologically active molecules [20]. Astrocytes release large amounts of exosomes when resting, and their vesicles contain a variety of substances such as proteins, lipids, cytokines, RNA, DNA, and other substances [21, 22]. Because of hemoglobin degradation products (such as hemin) and cytokine release from recruited inflammatory cells around the hematoma foci, astrocytes are reactive and produce a combination of effects after ICH [20, 23]. Similarly, reactive astrocytes have been observed to secrete a variety of exosomes in different disease models, containing different contents and playing different roles in exerting damaging or restorative effects on the CNS [20, 24]. Under the action of pro-inflammatory factors such as TNFα and IL1β, astrocytes reactively proliferate [25]. Reactive oxygen species, complement, and other components by astrocyte-derived exosomes have been shown to induce neuronal apoptosis and cognitive dysfunction in hippocampal regions in an animal model of Traumatic brain injury mice. However, exosomes from reactive astrocytes produced by pretreatment with berberine, oxygen-glucose deprivation, or a small amount of Aβ produced neuroprotective effects in animal models of cerebral ischemia, ICH, and Parkinson’s syndrome, respectively that were closely related to their secretion of exosomes rich in heat shock proteins, apolipoprotein D, nuclear factor E2-related factors, and other substances [26,27,28,29,30].

The relevant role of astrocyte-derived exosomes in disease progression after ICH has been less well studied, and it remains to be investigated what combined effect reactive astrocyte-derived exosomes has on GLS function in the context of ICH. Therefore, in this experiment, we used a low concentration of hemin to construct an in vitro ICH model to reactive astrocytes and obtain the exosomes produced by them, and used the collagenase method to construct an in vivo mouse ICH model for in-depth study [23, 31, 32].

Methods

Animals

All animals used in this study were obtained from Beijing Vital River Laboratory Animal Technology Co., Ltd (license number SCXK (Jing) 2021 0006). Male C57BL/6 mice (22–25 g), 8–10 weeks old, were housed under pathogen-free conditions with free access to food and water in a temperature-controlled environment simulating day and night. All animal experiments in this study were approved by the Animal Care and Use Committee of the Tianjin Medical University General Hospital, China (APPROVAL NUMBER IRB2022-DWFL-074) and were performed according to ARRIVE guidelines. Mice were randomly assigned to different groups and subjected to the respective experimental treatments (sTable. 1).

ICH modeling

In this study, ICH was induced by the collagenase method [33]. Mice were anesthetized (5% induction, 2% maintenance) with isoflurane (792632, Sigma-Aldrich), and the skull was exposed and fixed on a stereotactic frame after skin preparation and disinfection. A hole was made on the right side of the fontanel at 2.0 mm and 0.5 mm anteriorly, and a microinjection needle (ga33/10 mm, Hamilton) was used to aspirate an appropriate amount of collagenase VII (0.1 U/ul, C2399, Sigma-Aldrich), which was then placed in a microinjection pump (Shenzhen Ruiwode Life Technology Co, China) and vertically injected into the hole at a depth of 3.5 mm. 0.5ul of collagenase was injected at a rate of 0.2ul/min, and the needle was withdrawn 5 min after the injection was complete. After the skin was again sterilized and the wound sutured, the mice were resuscitated by placing them on a heating pad. Mice other than the sham group all underwent the above manipulations, and 0.5 ul of PBS was injected into the brain parenchyma of mice in the sham group using the microinjection needle as a substitute for collagenase VII. We observed the degree of contralateral limb dysfunction (hemiparesis) of the mice and performed the mNSS score to determine whether the ICH modeling was successful after 24 h.

Administration program

Exosomes were administered nasally at 1 h, 1 day, and 2 days after ICH. 10ul of PBS solution containing 108 exosomes was slowly dripped into the right nostril of mice for natural inhalation [34]. All other mice that did not receive exosome treatment were given 10ul of PBS nasally in the same way as a substitute. TGN020 (HY-W008574, MedChem Express), a specific AQP4 inhibitor, was used to block GLS, and the solvent was configured according to the instructions (5% DMSO, 40% PEG300, 5% TWEEN-80, 50% saline) to dissolve TGN020 sequentially. TGN020 was injected intraperitoneally (200ul, 100 mg/kg) or the solvent required to dissolve TGN020 30 min before each exosomes administration [35].

Tracer injection

The procedure was performed as previously described [36, 37].

Cisterna magna injection: Three days after ICH, mice were anesthetized and fixed in the stereotaxic apparatus to prepare the skin on the back of the neck, and the cisterna magna, which had an inverted triangular shape, was fully exposed. The microinjection needle was fixed in a microinjector, and 10 ul of cerebrospinal fluid tracer (25 mg/ml, RITC-Dextran, R9379, Sigma-Aldrich) was injected perpendicular to the cisterna magna at a depth of 1 mm at a rate of 1 ul/min. The needle was allowed to circulate in the cisterna magna for 50 min before mice were euthanized, perfused with cold PBS and brain tissue removed. Brain tissues were fixed in 4% paraformaldehyde for 24 h and then sectioned at 100 μm thickness and examined under a fluorescence microscope. Meningeal lymphatic vessel (MLV) and deep cervical limbic node (DCLN) drainage experiments were performed by bead (F8813, Invitrogen) injection. A microinjection needle injected 5 ul of beads into the cisterna magna at a rate of 1 ul/min. 50 min later, the right DCLN of the mice was harvested under a microscope, and the skull was harvested after the mice were euthanized and perfused with cold PBS. Lymph nodes and skulls were fixed in 4% paraformaldehyde for 24 h. The meninges were dissected under a microscope to await immunofluorescence staining. Lymph nodes were embedded in OCT and sectioned at 10 μm for immunofluorescence staining analysis.

Brain parenchyma injection: as described in the method of preparing the ICH model. Three days after ICH, 0.5 ul of cerebrospinal fluid tracer was injected at a rate of 0.1 ul/min after the microinjection needle was inserted 3.5 mm into the injection site in the model preparation. The needle was left in place for 5 min, then withdrawn and the cycle continued for 50 min. Brain tissue collection and processing were the same as for the cisterna magna injection.

Modified neurological severity score

The Modified Neurological Severity Score (mNSS) measures neurological deficits at 1D, 3D, 5D, 7D, and 14D after ICH in the areas of motor, sensory, reflex, and balance. A total of 18 points were scored in all aspects, with higher scores associated with more severe neurological deficits [38].

Gait analysis

Fourteen days after ICH, mice were placed on a gait analysis track, and a series of data generated by mice walking the 21 cm long by 3.6 cm wide section of the track were recorded without external stimuli using Cat Walk XT software [39]. Data from mice that completed the entire length of the track within 4 s were included in the recordings, and data from approximately 20% of mice that consistently failed to cooperate with the experiment were not included in the statistical analysis. Motor dysfunction in mice recovering from ICH is assessed using the maximum pressure generated by contact between the affected limb and the track as the mice walk, the maximum contact area, and the single-support chronophase.

Magnetic resonance imaging and brain water content determination

MRI was performed on mice 1 and 3 days after ICH. Mice were anesthetized with isoflurane and then scanned with coronal T2-weighted imaging using a 9.4 T MRI scanner (Bruker BioSpec 94/30) to visualize their hematoma foci with PHE. Hematoma volume calculation formula: maximum area of coronal hematoma foci × number of layers of hematoma foci × 0.5. Mice were euthanized after MRI, and their affected cerebral hemispheres were immediately removed and their wet weights were measured using a balance. They were then dried in an oven at 100 °C for 24 h to obtain the dry weight. The formula used to calculate brain water content was: brain water content (%) = (wet weight - dry weight)/wet weight × 100% [40].

In vitro cell assays

A 1 mM stock solution was prepared by sequential dissolution of hemin (HY-19424, MedChem Express) using 0.2% volume DMSO (D8371, Solarbio), 2% volume ammonia (G1823, Solarbio) and 97.8% volume complete medium (CM-M157, Pricella). The complete medium was used to dilute the stock solution to different concentrations to be used. The same generation of 3*106 primary astrocytes (CP-M157, Pricella) were inoculated into 10 cm dishes, cultured for 24 h, then replaced with medium containing different concentrations of hemin and cultured for 48 h before being used for cellular protein extraction.

Cell viability assay

According to the instructions of the CCK-8 Cell Proliferation and Cytotoxicity Assay Kit (CA1210, Solarbio), 100 μL of cell suspension containing 3*104 primary astrocytes were added to 96-well plates and incubated for 24 h. After 24 h, the medium was changed to complete medium with hemin at different concentrations. After 48 h of incubation, the medium was changed to 100 ul of complete medium without hemin, 10 μL of CCK-8 solution was added to each well and the cells were incubated in the incubator for 2 h and the absorbance at 450 nm was measured using an enzyme marker. The cell viability was calculated as follows Cell viability (%) = [OD(hemin)-OD(cell-free)]/[ OD(no hemin)-OD(cell-free)]×100% [41].

Extraction and identification of exosomes

When the density of primary astrocytes reached 70%, the exosome-free complete medium (obtained by ultracentrifugation) was replaced and the supernatant of the medium was collected for exosomes extraction after 1 day of culture. The extraction method was performed by ultracentrifugation, with the temperature controlled at 4 degrees Celsius throughout the procedure [42]. Centrifugation at 300 g for 10 min was used to remove live cells, and the supernatant was collected and centrifuged at 2000 g for 10 min to remove dead cells. The supernatant was centrifuged at 10,000 g for 30 min and the centrifuged supernatant was filtered through a 0.22 μm filter to remove debris and particles. The filtered supernatant was ultracentrifuged at 110,000 g for 70 min and the final precipitate was exosomes. The exosomes were dispensed and stored at -80 °C in a refrigerator. Transmission electron microscopy (TEM) and Nano tracer analysis (NTA) were used to analyze the morphology and size of the exosomes. Western blot was used to identify exosomes surface markers [43].

Intracranial localization of exosomes

Follow the instructions of the PKH26 Red Cell Membrane Staining Kit (D0030, Solarbio). Mix 100ul (containing 109 exosomes) of PBS solution with 900ul of Dilution C in the same centrifuge tube and add 1 ml of Dilution C with 4ul of PKH26 dye in another centrifuge tube. Quickly mix and blow the liquid in both centrifuge tubes and incubate for 4 min at room temperature in the dark. The staining was stopped by adding 8 mL of fetal bovine serum and then ultracentrifuged at 110,000 g for 70 min to obtain PKH26-labeled exosomes. According to the above administration method, brain tissue was extracted and fixed in formaldehyde 1 h after the last administration, dehydrated, and cut into 30 μm thick frozen sections for immunofluorescence staining.

Immunofluorescence

Mice were euthanized 3 days after ICH and brain tissues were fixed in 4% paraformaldehyde after in vivo infusion of cold PBS. After 24 h, the tissues were dehydrated through a gradient of 15%, and 30% sucrose, and after adequate dehydration, the tissues were placed in molds and embedded in optimal cutting temperature compound (OCT, 4583, SAKURA). Frozen sections were cut at 10 μm coronal using a cryosectioning machine, and OCT was washed off the sections with PBS on a shaker at room temperature. A circle was drawn around the tissue using an immunohistochemistry pen, and the tissue was incubated with a membrane-rupting sealer (PBS containing 5% BSA, 3% Triton, and 5% Tween-20) for 1 h at room temperature. Primary antibodies were incubated for 12 h at 4 °C, and secondary antibodies incubated for 1 h at room temperature. The following primary antibodies were used: GFAP mouse mAb (1:500, 3670, Cell Signaling Technology), AQP4 rabbit mAb (1:500, 59678, Cell Signaling Technology), CD31 goat pAb (1:500, AF3628, R&D Systems), NeuN rabbit mAb (1:500, ab177487, Abcam), LYVE1 rabbit pAb (1:500, ab14917, Abcam). Fluorescent secondary antibodies were purchased from Thermo Fisher Scientific.

Western blot

Protein extraction, electrophoresis, transfer, incubation, and exposure were performed according to the experimental methods described previously [44]. The consumables used in the experiments are described below. Color PAGE Gel Rapid Preparation Kit (PG112 and PG113, Yamei) and Immobilon® -P PVDF membrane (IPVH00010, Merck) were used. The primary antibodies used include: GFAP rabbit mAb(1:5000, ab68428, Abcam)、Alix rabbit mAb (1:1000, 92880, Cell Signaling Technology)、CD9 rabbit mAb (1: 1000, 13174, Cell Signaling Technology)、HSP70 rabbit mAb(1:2000, A21180, ABclonal)、TSG101/VPS23 rabbit pAb(1:1000, A2216, ABclonal)、Tau rabbit mAb(1: 5000, A23490, ABclonal)、Phospho-Tau-T205 rabbit pAb (1:1000, AP0168, ABclonal), Phospho-Tau-S404 rabbit pAb (1:1000, AP0170, ABclonal), Phospho-Tau-S396 rabbit mAb(1: 1000, AP1028, Abclonal)、Bax rabbit mAb(1:1000, A19684, Abclonal)、Bcl-2 mouse mAb(1:500, A20777, Abclonal)、Occludin rabbit mAb (1:1000, A24601, Abclonal)、ZO-1 mouse mAb(1:1000, 33-9100, Thermo Fisher Scientific)、β-Actin rabbit mAb(1:20000, AC026, Abclonal). The grayscale values of each band were analyzed using PS.

Detection of neuronal apoptosis

After Neun labeling was completed in the immunofluorescence steps, apoptotic cell labeling was performed using the TUNEL Assay Apoptosis Detection Kit (abs50033, absin). According to the kit instructions, 50 ul of Proteinase K (20 μg/mL) was added to the Neun-labeled frozen sections and incubated for 20 min under light protection, washed well with PBS, and incubated for 2 h under light protection with 50 ul of TUNEL working solution. Sections were sealed and observed under a microscope.

Nissl staining

Mice were euthanized 3 days after ICH and brain tissues were fixed in 4% paraformaldehyde after in vivo infusion of cold PBS. After 24 h, 3-μm paraffin sections were prepared by ethanol gradient dehydration, paraffin embedding, and sectioning. Sections were deparaffinized in xylene and ethanol and hydrated for Nissl staining. According to the instructions of the Nissl staining kit (G1036, Servicebio), the sections were immersed in Niehl’s staining solution for 5 min and then washed with water until the water on the slides was colorless. The sections were dried with excess water, sealed with neutral gum, and observed under a bright field microscope.

Statistical analysis

The sample size of the present study was consistent with that of previous studies. Results were expressed as mean standard deviation (SD). All data sets were tested for Shapiro-Wilk normality. Unpaired two-tailed Student’s t-test was used to analyze statistical differences between the two groups. Comparisons between multiple groups were made using one- or two-way analysis of variance (ANOVA) followed by Tukey’s post hoc test. p < 0.05 was considered statistically different. GraphPad Prism software (version 9.0) was used for all data statistics and image generation.

Results

Results 1. H-Exo promotes AQP4 polarization and attenuates astrocyte proliferation around hematoma foci in acute-phase ICH mice

Morphology of primary astrocytes under bright field microscopy (Fig. 1, A). GFAP was used as a marker of astrocyte activation and cell expression of GFAP under different culture conditions was quantified by Western Blot. The results showed a statistically significant increase in GFAP expression at hemin concentrations of 50 μm and 100 μm (Fig. 1, B-C). The CCK-8 showed that astrocyte viability began to decrease when the hemin concentration reached 200 μm (Fig. 1, D). Therefore, cells were cultured with complete medium containing 50 μm hemin concentration for the in vitro construction of the ICH model. Primary astrocytes were reactive and their viability was not inhibited under that culture conditions.

Fig. 1
figure 1

Activation of primary astrocytes with hemin and characterization of astrocyte-derived exosomes. (A) Morphology of primary astrocytes under bright field microscopy. (B, C) Western Blot quantification of GFAP relative expression after different concentrations of hemin treatment of primary astrocytes. (D) CCK-8 detection of the viability of primary astrocytes under different concentrations of hemin treatment. (E) The size distribution of exosomes secreted by astrocytes was detected by NTA, and the graph shows the average value of three experiments after 500-fold dilution of the stock solution. (F) Morphological features of exosomes under TEM. (G) Western Blot analysis of the expression of exosomes markers Alix, CD9, HSP70, and TSG101. (H) Distribution of exosomes around hematoma foci in mice after exosomes administration by nasal feeding after ICH. All data are presented as mean ± SD (n = 4 per group). p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

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The exosomes were extracted and identified by ultracentrifugation, and the particle size of the extracts was mainly distributed in the range of 30–150 nm by NTA, with the peak at 109 nm (Fig. 1, E). The morphology of the extracts was observed by TEM, and the double-membrane round vesicles could be seen under the microscope (Fig. 1, F). The extracts were qualitatively analyzed by Western Blot for Alix, CD9, HSP70, and TSG101 showed positive results (Fig. 1, G). The identification results showed that the extracts were consistent with exosomes characteristics. Exosomes secreted by astrocytes without hemin addition were defined as astrocyte-derived exosomes in the resting state (R-Exo), and exosomes secreted by astrocytes with hemin addition were defined as astrocyte-derived exosomes in the reactive state by hemin treatment (H-Exo). Administered by the nasal method, PKH26-labeled exosomes were seen densely distributed around the hematoma foci by immunofluorescence (Fig. 1, H).

Previous studies have shown that loss of AQP4 polarization is closely associated with reactive proliferation of astrocytes. Immunofluorescence showed that astrocytes proliferated around the hematoma foci after ICH under the combined effect of primary and secondary injury (Fig. 2, A-B), and their AQP4 polarization anchored to the perivascular endfeet disappeared (Fig. 2, C-D). Compared with the ICH group, astrocyte proliferation was inhibited in the H-Exo group and accompanied to some extent by repolarization of AQP4, whereas the R-Exo group showed no significant differences.

Fig. 2
figure 2

H-Exo promotes AQP4 polarization and reduces astrocyte proliferation in the acute phase of ICH. (A) Immunofluorescence staining of AQP4 with GFAP around hematoma foci. (B) Quantitative analysis of the percentage area of the GFAP-positive region in (A). (C) Immunofluorescence staining of AQP4 with CD31 around hematoma foci. (D) Quantitative analysis of the percentage area of double-positive areas of AQP4 versus CD31 in (B) to reflect AQP4 polarization. All data are presented as mean ± SD (n = 6 per group). p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

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Results 2. H-Exo improves intracranial lymphatic drainage in mice in the acute phase of ICH

Previous reports have shown that the GLS function is dependent on the paravascular polarization distribution of AQP4. Therefore, we evaluated intracerebral lymphatic drainage by intracranial injection of tracers. To evaluate the solute clearance effect of the GLS, we injected the tracer RITC-Dextran into the basal ganglia and left a representative coronal section of brain tissue after a total of 1 h of circulation. The results showed that the amount of tracer residue in the brains of mice in the H-Exo group was greatly reduced, while the tracer clearance in the ICH and R-Exo groups was difficult (Fig. 3, A-B). The influx of tracer into the brain parenchyma injected via the cisterna magna was reduced after ICH, but the results showed that the tracer influx in the H-Exo group was significantly greater than that in the ICH and R-Exo groups, and the tracer accumulated more in the skull base (Fig. 3, C-D). We further injected beads into the cisterna magna and observed the efficiency of magnetic bead influx into the MLV & DCLN. More beads converged on the dorsal MLV and DCLN in the H-Exo group compared with the ICH and R-Exo groups. The results further suggested that H-Exo facilitated the drainage of intracerebral solutes to the extracranial area while enhancing the function of the GLS (Fig. 3, E-H).

Fig. 3
figure 3

H-Exo improves intracranial lymphatic drainage in the acute phase of ICH. (A) Representative coronal fluorescence images after basal ganglia injection of RITC-Dextran. (B) Quantitative analysis of the percentage area of RITC-Dextran-positive area in the basal ganglia injected in (A). (C) Representative coronal fluorescence images after RITC-Dextran injection in the cisterna magna. (D) Quantitative analysis of the percentage area of RITC-Dextran-positive area in the cisterna magna injected in (C). (E) Immunofluorescence images of beads injected into the cisterna magna converging on the dorsal MLV. (F) Quantitative analysis of the percentage of positive area of beads within the transverse sinus in (E). (G) Immunofluorescence images of cisterna magna injected beads draining into the largest diameter section of the DCLN. (H) Quantitative analysis of the percentage area of the bead-positive region in (G). All data are presented as mean ± SD (n = 6 per group). p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

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Results 3. H-Exo reduces phosphorylated tau deposition and PHE volume in the affected cerebral cortex of mice in the acute phase of ICH

Having validated that H-Exo treatment favors GLS to promote exogenous tracer drainage and clearance, we further investigated its role in adverse events during ICH pathology. The expression of tau protein in the affected cerebral cortex was examined by Western Blot to estimate the clearance capacity of toxic solutes. The results showed that tau protein phosphorylation increased after ICH without significant changes in total tau levels compared to the sham group. However, the level of cortical phosphorylated tau deposition was reduced in the H-Exo group compared with the ICH and R-Exo groups (Fig. 4, A-E).

Meanwhile, after ICH, PHE occurs due to blood-brain barrier disruption, reactive oxygen species (ROS) release, and inflammatory cell aggregation, the development of PHE directly affects the early mortality and late prognosis of ICH patients. It has been reported that in mice models, PHE increases rapidly at 24 h after hemorrhage and peaks at 72 h. Intracranial hematoma and PHE were observed in mice at 1 and 3 days after ICH using T2-weighted magnetic resonance imaging (Fig. 4, F). Analysis of the results showed that there was no significant difference in hematoma volume between the groups (Fig. 4, G). H-Exo treatment failed to prevent further progression of PHE over time, but the brain water content of the affected hemisphere in the H-Exo group of mice was significantly reduced compared with the ICH and R-Exo groups at the same time point (Fig. 4, H). At the same time, the level of tight junction proteins in the brain tissue of mice was relatively increased after H-Exo treatment compared with that in the ICH and R-Exo groups (Fig. 4, I-K). This partly reflects that the physical integrity of the blood-brain barrier was better restored in H-Exo treated mice.

Fig. 4
figure 4

H-Exo reduces intracranial accumulation of toxic metabolites and attenuates cerebral edema. (A-E) Western Blot quantification of phosphorylated tau protein expression in the hemorrhagic side hemisphere. (F) MRI T2-weighted images showing intracranial hematoma foci and PHE in mice. (G) Quantitative analysis of hematoma foci volume in (F). (H) Quantitative analysis of water content of the cerebral hemisphere on the hemorrhagic side. (I-K) Quantitative analysis of BBB-related protein expression in the hemorrhage side cerebral hemisphere by Western Blot. All data are presented as mean ± SD (n = 6 per group). p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

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Results 4. Functional improvement of GLS facilitates functional recovery of the affected limb during ICH recovery in mice

Neurological deficits in mice after ICH were assessed using the mNSS, and the higher the score, the more severe the neurological deficits. Mice showed significant motor and sensory deficits on the first day after ICH and gradually recovered over time (Fig. 5, A).

Fig. 5
figure 5

H-Exo promotes functional recovery of the affected limb in ICH-recovery mice. (A) mNss scores of mice at 1, 3, 5, 7, and 14 days after ICH. (B) Representative images of crawling footprints of mice during ICH recovery. (C) Quantitative analysis of the single-support phase of the affected limb as a percentage of the walking cycle in mice during ICH recovery. (D-G) Quantitative analysis of maximum force and maximum contact area of the affected anterior and posterior limbs after contact with the treadmill in mice recovering from ICH. All data are presented as mean ± SD (n = 8 per group). p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

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To further investigate the recovery of motor function after ICH, we used a gait analysis system to obtain the walking footprints and a series of related data at 14 days after ICH (Fig. 5, B). The uni-support temporal phase, which refers to the time required for the studied paw to completely leave the ground from the initial contact with the ground, should be 30–40% of the gait cycle, and a decrease in this percentage suggests that utilizing obstacles in the use of the limb on the studied side. Analysis of the uni-support temporal phase, maximum support force, and maximum contact area of the left (affected) limb of the mice at 14 days after ICH suggested that the decrease in the use of the affected limb, the weakening of muscle strength, and the sensory deficits of the mice in the H-Exo treated group were restored to a greater extent and were statistically significant when compared to those in the ICH group and the R-Exo group (Fig. 5, C-G).

Application of TGN020, an AQP4-specific blocker, successfully reversed the ability of H-Exo to promote solute efflux into the brain parenchyma during the acute phase of ICH by increasing GLS (Fig. 6, A-B). It was also found that the role of H-Exo in promoting tracer influx into the brain parenchyma was inhibited in the context of ICH (Fig. 6, C-D).

Fig. 6
figure 6

TGN020 reverses outcome in ICH-recovery mice by inhibiting GLS function. (A) Representative coronal fluorescence images after basal ganglia injection of RITC-Dextran. (B) Quantitative analysis of the percentage area of RITC-Dextran-positive area in the basal ganglia injected in (A). (C) Representative coronal fluorescence images after RITC-Dextran injection in the cisterna magna. (D) Quantitative analysis of the percentage area of RITC-Dextran-positive area in the cisterna magna injected in (C). (E) mNss scores of mice at 1, 3, 5, 7, and 14 days after ICH. (F) Representative images of crawling footprints of mice during ICH recovery. (G) Quantitative analysis of the percentage of walking cycles during the single-support phase of the affected limb in mice during ICH recovery. (H-K) Quantitative analysis of maximum force and maximum contact area of the affected anterior and posterior limbs after contact with the treadmill in mice during ICH recovery. All data are presented as mean ± SD (n = 6–8 per group). p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

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To explore the effect of blocking GLS in the acute phase of ICH on the long-term outcome of H-Exo therapy, we designed the following experiment. The subgroups were named as follows: placebo group (ICH + placebo), control group (ICH + TGN020), treatment group (H-Exo + placebo), and inhibitor group (H-Exo + TGN020). We recorded the mNSS scores of these four groups of mice for 14 days after ICH (Fig. 6, E). On day 14 after ICH, we performed gait analysis and obtained a series of data on these four groups of mice (Fig. 6, F-K). The results of the treatment and inhibitor groups showed that the administration of TGN020 blocked the beneficial effect of H-Exo on the recovery of neurological function in ICH mice. The placebo and control groups indicated that the application of TGN020 alone had no significant beneficial or detrimental effects on neurological deficits after ICH in mice. The results of the control and inhibitor groups showed that the application of TGN020 followed by the administration of H-Exo had no effect on the behavioral outcome of ICH mice. The results of the intergroup control suggested that H-Exo could effectively improve neurological impairment after ICH, and the integrity of GLS function played an important role in this process.

Results 5. H-Exo reduces neuronal apoptosis in the cerebral cortex during the acute phase of ICH by enhancing the function of GLS

The impaired neurological recovery caused by ICH is mainly attributed to the inability to effectively regenerate neurons and axons after destruction caused by primary and secondary brain injury. Early survival of neurons and conduction bundles in ICH is critical for later recovery of limb function, and Western blot quantified apoptosis and phosphorylated tau protein tangles in the mouse cerebral cortex 3 days after ICH (Fig. 7, A). Consistent with previous studies, abnormal phosphorylation of tau proteins in the affected cortex was reduced after administration of H-Exo, but further studies showed that this effect could be blocked by TGN020, and administration of TGN020 alone did not show a statistical difference compared to the placebo group (Fig. 7, B-E). It was also found that apoptosis in the cerebral cortex was significantly reduced in the treatment group compared to the control and inhibitor groups (Fig. 7, F-H). The percentage of neurons in apoptotic cells was analyzed using TUNEL and Neun immunofluorescence co-localization (Fig. 7, I). The control group showed a large number of apoptotic neurons in the cortex after ICH and administration of H-Exo was effective in increasing neuronal survival, but this effect disappeared with GLS blockade (Fig. 7, J). Nissl body are distributed in the cytosol and dendrites of neurons and are often regarded as a sign of the functional state of neurons, which can be reduced or even disappear when neurons are damaged. In Nissl staining, the shrunken and necrotic nucleus of cells are dark blue, the granular Nissl body distributed around the neuronal cytosol are blue, and the nucleus of normal cells located in the center of the cell is light blue (Fig. 7, K). Quantitative analysis of the number of Nissl body in the hemorrhagic side of the cerebral cortex in the high magnification field of view showed that neuronal survival was improved after the application of H-Exo, together with a better preservation of their functional status (Fig. 7, L).

Fig. 7
figure 7

GLS functional status affects neuronal survival in the acute phase of ICH. (A-H) Western blot quantification of phosphorylated tau protein with BAX and BCL expression in the affected cerebral hemisphere of ICH acute phase mice. (I) Immunofluorescence staining of Tunel and Neun in the cerebral cortex around the hematoma foci. (J) Quantification of the percentage of Neun-positive areas in the Tunel and Neun double-positive areas in (I) to detect the percentage of neuronal apoptosis. (K) Nissl staining of the cerebral cortex around the hematoma foci. (L) Quantitative analysis of Nissl body expression in the cerebral cortex under high magnification field of view in (K). All data are presented as mean ± SD (n = 6 per group). p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

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Discussion

This study describes the critical role of reactive astrocyte-derived exosomes in intracranial solute clearance after ICH. We obtained exosomes produced by astrocytes through hemin treatment. Administered nasally to mice in the acute phase of ICH, the exosomes were found to enhance intracranial lymphatic drainage in mice in the acute phase of ICH and promote functional limb recovery in the recovery phase of ICH. In addition, intraperitoneal injection of the AQP4 blocker TGN020 was used to explore the potential mechanisms of action of H-Exo. The results showed that the effects of H-Exo in promoting clearance of intracranial toxic solutes and reducing neuronal apoptosis were reversed after application of TGN020. The functionality of the H-Exo is dependent on the integrity of the GLS functionality.

Previous studies have shown that nasally administered exosomes allow their components to rapidly pass through the olfactory nerve pathway, bypass the blood-brain barrier, and act directly on the CNS for efficient and rapid treatment [45]. Experimentally, PKH26-labeled exosomes were observed to be widely distributed around the hematoma foci, and some were taken up by astrocytes and neurons [46].

Astrocytes play important roles in the CNS, including neurotrophic support, synapse-associated repair, and maintenance of the blood-brain barrier [47]. When ICH occurs, astrocytes are induced to enter a reactive state (reactive astrocytes). In some disease models reactive astrocytes can mediate aberrant synthesis and release of neurotransmitters, synaptic disruption, and loss of homeostasis of other homeostatic functions, and their release of reactive oxygen species, complement, and other components contained in exosomes can lead to events such as neuronal apoptosis and inflammatory cell recruitment [21]. However, in other diseases, reactive astrocytes have more protective functions, and their released exosomes containing substances such as Hsp70, Apo D, and Nrf2 have been observed to be neuroprotective in various disease models, including stroke [30]. The role of reactive astrocytes in the progression of the same disease is complex and variable [48], and the outcome of their role in different diseases varies or is even completely opposite [19]. In this experiment, we explored the results of the role of exosomes produced by reactive astrocytes in ICH after hemin treatment. For the first time, we demonstrated that H-Exo significantly improved GLS dysfunction, alleviated neurological deficits, and reduced neuronal apoptosis in ICH model mice.

In our experiments, we found that the efficiency of RITC-Dextran influx from cerebrospinal fluid into the brain parenchyma and its eventual clearance was significantly enhanced after H-Exo treatment. Intracerebral solutes partially flowed into the MLV after cerebrospinal fluid-interstitial fluid exchange and finally entered the DCLN to reach the peripheral lymphatic system [49]. After injection of fluorescent magnetic beads via cisterna magna and adequate circulation, more fluorescent magnetic beads were accumulated in the dorsal MLV and DCLN in the H-Exo group. A series of results showed that H-Exo significantly improved intracranial lymphatic drainage in mice after ICH. In pathological conditions such as Alzheimer’s disease, traumatic brain injury and intracranial hemorrhage, beta-amyloid and phosphorylated tau proteins accumulate pathologically [50] which cause and exacerbate neurodegenerative lesions [51]. Enhanced GLS function has been shown to significantly reduce pathological tau protein accumulation in models of Alzheimer’s disease and traumatic brain injury [52, 53], and ameliorated cognitive and motor dysfunction in diseased mice. This is consistent with our experimental findings and conclusions.

Different pathophysiological mechanisms dominate the development of PHE in the different phases of ICH, which are usually divided into vasogenic and cytotoxic edema [5]. Human ICH studies typically show rapid PHE accumulation within 1–3 days after ICH, and the absolute or relative PHE growth rate at 3 days is considered an independent predictor of death and poor functional prognosis [54]. After the blood-brain barrier is damaged, solutions and solutes from the blood enter the brain interstitium with impunity, and factors such as changes in osmotic pressure and inflammatory cell recruitment allow PHE to occur and develop [55]. Numerous studies have described the attenuation and regression of PHE from the perspective of repairing blood-brain barrier damage [56, 57]. AQP4 is the most abundant water channel protein in the brain, and its main functions are to maintain brain water homeostasis and clear neurotransmitters, which are thought to be related to the formation and regression of brain edema in disease states [58]. As part of the neurovascular unit, AQP4 is mainly polarized in the endfeet of perivascular astrocytes, and either overexpression and depolarization of AQP4 or its deletion is detrimental to the performance of its physiological functions [59, 60]. The GLS is anatomically tightly connected to the blood-brain barrier, and together they regulate the transport and exchange of body fluids and substances throughout the brain [61]. Our study suggests that H-Exo promotes the regression of PHE in the acute phase of ICH from the perspective of enhanced cerebrospinal fluid-interstitial fluid exchange. H-Exo attenuated the reactive proliferation of astrocytes around the hematoma foci and promoted the repolarization of AQP4. Application of H-Exo significantly improved the fluid exchange and solute clearance function of the GLS, resulting in a significant reduction in brain water content during the acute phase of ICH, although its application did not reduce hematoma volume.

To further demonstrate that H-Exo functions by enhancing GLS, we injected mice intraperitoneally with TGN020, an AQP4-specific blocker, to inhibit GLS [62]. To rule out interference from the solvent used to dissolve TGN020 and the side effects of TGN020, we set up four experimental groups for the between-group control. Analysis of the results in the placebo and control groups showed that the side effects of TGN020 did not significantly interfere with the experiments [63], while the control and inhibitor groups showed that H-Exo did not effectively exert its beneficial effects after administration of TGN020. Therefore, the results of the control between the treatment group and the inhibitor group suggest that the effects of H-Exo in promoting the recovery of neurological deficits, reducing the deposition of toxic proteins, inhibiting neuronal apoptosis, and maintaining the functional state of neurons are dependent on the integrity of GLS function and that the restoration of GLS function is a potential pathway for the amelioration of neurological deficits in mice with ICH by H-Exo.

It has been reported that reactive astrocyte-derived exosomes contain apolipoprotein D, which facilitates neuronal survival and maintains their functional integrity in pathological situations [64]. In addition, astrocytes are reactive to release more HSP70-containing exosomes, which mediate neuronal resistance to physical trauma or toxicity [65]. This reflects the shortcomings of the present experiments in that we considered the exosomes and its contents as a whole to explore its functional mechanism of action and did not specifically investigate the role of the single component contained in H-Exo. Understanding the interaction between reactive astrocyte-derived exosomes and GLS requires further investigation.

Conclusions

Our study explores for the first time the results of reactive astrocyte-derived exosomes in mice with ICH from the perspective of regulating intracranial lymphatic drainage and the possible mechanisms of action. Reactive astrocyte-derived exosomes obtained after hemin treatment was found to promote AQP4 repolarization and enhance intracranial lymphatic drainage. By blocking AQP4, the neuroprotective effect of H-Exo was shown to be dependent on the functional integrity of the GLS. Together with published studies, this expands the potential avenues for reactive astrocyte-derived exosomes to exert neuroprotective effects in the context of ICH.

Data availability

No datasets were generated or analysed during the current study.

Abbreviations

AQP4:

Aquaporin 4

CNS:

Central nervous system

DCLN:

Deep cervical limbic node

GLS:

Glymphatic system

H-Exo:

Astrocyte-derived exosomes in the reactive state by hemin treatment

ICH:

Intracranial hemorrhage

MLV:

Meningeal lymphatic vessel

MNSS:

Modified Neurological Severity Score

NTA:

Nano tracer analysis

PHE:

Perihematomal edema

R-Exo:

Astrocyte-derived exosomes in the resting state

TEM:

Transmission electron microscopy

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Acknowledgements

Not applicable.

Funding

This work was supported by the Tianjin Health Commission Science and Technology Key Discipline Project (via Grant No. TJWJ2024XK001 to Zengguang Wang)and Tianjin Education Commission Research Program Project (via Grant No. 2024KJ164 to Yuheng Liu).

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Author notes

  1. Kexin Li and Yuheng Liu contributed equally to this work.

Authors and Affiliations

  1. Department of Neurosurgery, Tianjin Medical University General Hospital, Tianjin, China

    Kexin Li, Yuheng Liu, Junjie Gong, Jing Li, Mingyu Zhao, Chengyou Hong, Yuchi Zhang, Mengyao He, Zhenye Zhu, Zhijuan Chen & Zengguang Wang

  2. Key Laboratory of Post-Neuroinjury Neuro-Repair and Regeneration in Central Nervous System, Tianjin Neurological Institute, Ministry of Education and Tianjin, Tianjin, China

    Kexin Li, Yuheng Liu, Junjie Gong, Jing Li, Mingyu Zhao, Chengyou Hong, Yuchi Zhang, Mengyao He & Zhenye Zhu

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  9. Zhenye Zhu
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  10. Zhijuan Chen
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  11. Zengguang Wang
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Contributions

ZW, YL and KL designed the study. YL and MZ developed the experimental methodology. KL, CH and ZZ carried out the experiments. ZC, YZ and MH carried out the data summary and analysis. JL and JG provided technical assistance with the software. KL drafted the manuscript. ZW and YL read and approved the final version of the manuscript. All authors read and approved the final manuscript.

Corresponding authors

Correspondence to Zhijuan Chen or Zengguang Wang.

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Ethics approval and consent to participate

All animal experiments in this study were approved by the Animal Care and Use Committee of the Tianjin Medical University General Hospital, China (APPROVAL NUMBER IRB2022-DWFL-074) and were performed according to ARRIVE guidelines.

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Not applicable.

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The authors declare no competing interests.

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Li, K., Liu, Y., Gong, J. et al. Reactive astrocyte-derived exosomes enhance intracranial lymphatic drainage in mice after intracranial hemorrhage. Fluids Barriers CNS 22, 37 (2025). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s12987-025-00651-y

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  • DOI: https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s12987-025-00651-y

Keywords

Fluids and Barriers of the CNS

ISSN: 2045-8118