Differential Protective Effects of Edaravone in Cerebellar and Hippocampal Ischemic Injury Models

Abstract

Ischemic stroke is a leading cause of mortality and disability, with cerebellar strokes posing severe complications such as herniation and brainstem compression. Edaravone, a radical scavenger known for reducing oxidative stress, has shown neuroprotective effects in cerebral strokes, but its impact on cerebellar strokes remains unclear. This study investigates Edaravone’s protective properties in organotypic slice cultures of rat cerebellum and hippocampus, employing an oxygen-glucose deprivation (OGD) model to simulate ischemic stroke. The hippocampus served as comparative structure due to its high hypoxia sensitivity. Our results confirmed effective hypoxic induction with increases in HIF-1α and HIF-2α expression. Edaravone significantly reduced lactate dehydrogenase (LDH) levels, indicating diminished cellular damage, with cerebellar tissues showing greater vulnerability. Additionally, Edaravone reduced reactive oxygen species (ROS) in both tissues, though its efficacy may be limited by higher oxidative stress in cerebellar cultures. Seahorse XF analysis revealed that Edaravone preserved mitochondrial respiration and tissue integrity in cerebellar and hippocampal slice cultures. However, Edaravone was more effective in preserving mitochondrial respiration in hippocampal slices, suggesting that OGD-induced damage is more severe in cerebellar tissue. In conclusion, Edaravone demonstrates significant cell protective effects in both cerebellar and hippocampal tissues under OGD conditions, preserving tissue integrity and enhancing mitochondrial function in a tissue-dependent manner. These findings suggest Edaravone as a promising therapeutic candidate for cerebellar stroke. Further in vivo studies are required to assess its full clinical potential.

Supplementary Information

The online version contains supplementary material available at 10.1007/s12311-025-01804-3.

Introduction

Stroke is one of the leading causes of death worldwide and the primary cause of acquired and permanent disability in adults. According to the World Health Organization (WHO), a stroke occurs every two seconds worldwide, and someone dies from a stroke every six seconds, surpassing the combined mortality rates of malaria, tuberculosis, and HIV/AIDS [1]. Survivors often face significant limitations in their ability to live independently, burdening healthcare systems with extensive medical and nursing needs. Ischemic strokes, caused by intravascular thrombosis or embolism blocking a cerebral vessel, account for over 80% of strokes [2]. The clinical presentation and symptoms of a stroke largely depend on the affected brain region. Cerebellar ischemic stroke involving one or more of the three main cerebellar arteries (posterior inferior cerebellar artery (PICA), anterior inferior cerebellar artery (AICA) and/or superior cerebellar artery (SCA)) [3], are relatively rare, accounting for 2–3% of strokes, but they cause disproportionately high morbidity and mortality [4, 5]. The distinct symptoms of cerebellar strokes, such as dizziness, nausea or vomiting are often mistaken for less severe conditions, complicating diagnosis [3]. Treatment options for cerebellar strokes generally mirror those for cerebral strokes, including lysis therapy and mechanical thrombectomy. However, cerebellar strokes pose a higher risk of complications such as herniation and brain stem compression, requiring specialized therapeutic approaches. Cerebral oedema resulting from strokes can lead to ventricular system compression, requiring additional therapeutic measures [4–6]. The pathophysiology of hypoxic neuronal brain damage in ischemic strokes involves several signaling pathways leading to necrosis and apoptosis, including energy deficiency, loss of ion homeostasis, acidosis, increased intracellular calcium levels, excitotoxicity, free oxygen radicals, complement activation, cytokine-mediated mechanisms, and immune system activation. These complex mechanisms can be categorized into three basic mechanisms: direct neuronal loss through apoptotic signaling pathways [7], production of reactive oxygen species (ROS) causing oxidative stress, and ischemia-induced inflammation of neuronal tissue [8]. Massive cell damage occurs not only during acute ischemia but also during reperfusion, exacerbated by ROS and neuroinflammatory processes [9]. Expanding current therapies to target these pathophysiological processes could significantly advance stroke treatment. Edaravone (3-methyl-1-phenyl-pyrazolin-5-one) represents a promising option for innovative stroke therapy. Approved by the US Food and Drug Administration (FDA) in 2017 for treatment of amyotrophic lateral sclerosis, Edaravone acts as a radical scavenger, mitigating oxidative stress [10]. Its favorable acid dissociation constant (pKa) allows it to exist in both lipophilic and hydrophilic forms, facilitating passage through cell membranes and the blood-brain barrier [11]. Edaravone significantly reduces oxidative stress during a stroke, lowering cell apoptosis and tissue damage. Further, it inhibits vascular endothelial growth factor (VGEF) expression in astrocytes, protecting against brain oedema [12]. It also increases the expression of BDNF and Bcl-2, which have anti-apoptotic effects, and reduces caspase-3 activity, crucial for apoptosis. Edaravone likely lowers post-ischemic leukotriene levels, reducing oedema, with 5-LOX activation playing a key role in regulating arachidonic acid metabolism [13]. Although Edaravone has shown promise in human trials for cerebral strokes, its potential in cerebellar strokes remains underexplored, presenting an opportunity to advance stroke treatment [14–17]. This research aims to investigate Edaravone’s protective effects in cerebellar ischemic stroke using a hypoxia model with organotypic slice cultures. The hypoxia-sensitive hippocampus will also be used as a comparative structure [18] to further clarify its treatment efficacy.

Materials and Methods

Animals

All procedures were conducted under established standards of the German federal state of North Rhine Westphalia, in accordance with the European Communities Council Directive 2010/63/EU on the protection of animals used for scientific purposes. According to current German and European legislation, the removal of organs or cells from vertebrates for scientific purposes is not considered an animal experiment if the animals have not been subject to surgical interventions or invasive treatments prior to sacrifice. Thus, the euthanasia of rats for the removal of brain tissue does not require approval or permission from local or federal authorities.

Wistar rats of both sexes from our in-house breeding stock were used and housed under a 12:12 light-dark cycle with food and water provided ad libitum. For organotypic slice culture preparation, rats aged postnatal days 7–10 (P7–10) were utilized. In accordance with established ethical standards and relevant German and European legislation, the animals were decapitated without prior euthanasia.

Organotypic Cell Cultures and Oxygen-Glucose Deprivation (OGD)

For this study, organotypic slice cultures of the cerebellum and hippocampus were prepared from seven Wistar rats following the method described by Wolters and Reuther et al. [19]. In some preparations, an additional animal was used to ensure consistent slice quality. Brain tissue was sectioned using a McLain Tissue Chopper to obtain cerebellar slices (275 μm thickness) and hippocampal slices (350 μm thickness), which were subsequently cultivated on inserts (Millicell; REF: PICM0RG50) in six-well plates containing 1 ml of culture medium per well. The culture medium consisted of 25% heat-inactivated horse serum, 25% Hanks’ Balanced Salt Solution (Sigma; REF: H8264), 2.6% glucose (Merck; REF: 8342), 1% Nerve Growth Factor, 1% penicillin, 1% glutamine (Gibco; REF: 35050-061), 0.2% phenol red (Sigma; REF: P0290) in Basal Medium Eagle (Sigma; REF: B1522). Cultures were maintained in an incubator at 5% CO₂ and 37 °C for seven days. To ensure robust and unbiased experimental conditions, slices were randomly pooled and distributed across wells. Importantly, each well contained either four cerebellar or five hippocampal slices, with slices in a single well always originating from different rats to prevent intra-animal redundancy. This distribution strategy ensured that slices from different animals were evenly represented across all experimental groups. Only slices with a preserved architecture were included, enhancing the reproducibility and reliability of the experimental results. A minimum of three wells per condition was prepared for each experimental group, and each experiment was independently repeated at least three times with new preparations, accounting for biological variability across animals and technical variability across experimental runs. After seven days in culture, oxygen-glucose deprivation (OGD) was induced using two specific media formulations. The OGD medium lacked glucose and contained 25% Hanks’ Balanced Salu Solution (Sigma; REF: H8264], 1% Nerve Growth Factor, 1% penicillin, 1% glutamine (gibco; REF: 35050-061), 0.2% phenol red (Sigma; REF: P0290) in Dulbecco’s Modified Eagle’s Medium (Sigma; REF: D6046). A control medium with identical composition but with glucose supplementation (6.5 mg/ml) was used for normoxic conditions. Both media were pre-gassed for at least one hour before use. The OGD procedure was conducted in a hypoxic chamber (Hypoxic Chamber Polymer O₂ Control Glove Box; Coy Lab, Grass Lake, MI, USA) under controlled conditions of 37 °C, 5% CO₂ and 0.4% O₂. Following the designated OGD duration, the inserts were transferred back to normoxic conditions with fresh glucose-containing medium to initiate reoxygenation. Experimental time points and protocols were conducted as specified in the respective test procedures.

Real Time Quantitative PCR

To evaluate the impact of OGD on the expression of hypoxia-inducible factors (HIF), qPCR was performed. Total RNA was extracted from organotypic slice cultures of the cerebellum and hippocampus after 1, 4, and 8 h of OGD, as well as from normoxic control conditions. Each well representing one timepoint included 4 cerebellar and 5 hippocampal slice cultures. The isolated RNA was further processed to analyse the mRNA levels of HIF-1α and HIF-2α using the RNA isolation kit from Machery-Nagel (REF: 740955.50). The manufacturer’s protocol was followed with slight modifications, particularly an adjusted elution volume of 40 µl to achieve a higher RNA concentration. cDNA was synthesized using oligo(dT) primers and the GoScript Reverse Transcription Mix (Promega: A2790). qPCR was performed using the GoTaq qPCR Master Mix (Promega: REF A6002). Primer sequences were as follows: Hif1a: forward 5’-AAT GTA CCC TAA CAA GCC GGG–3’, reverse 5’-GTT TCT TGT AGC CAC ACT GCG-3’; Hif2a: forward 5’- AGT GGT CTG TGG GCA ATC AG-3’, reverse 5’-AAC ATG GAG ACA TGA GGC GG-3; Actb: forward 5’-CTA AGG CCA ACC GTG AAA AG-3’, reverse 5’-AAC ACA GCC TGG ATG GCT AC-3’. The differences in the expression levels were calculated using the 2^-ΔΔCt method with endogenous normalization to Actin.

Western Blotting

To prevent the degradation of HIF-1α and HIF-2α proteins during the transition to normoxic conditions, protein isolation was performed directly within the hypoxia chamber. Organotypic slice cultures were carefully removed from the cultivation membranes by pipetting with PBS containing a protease and metalloprotease inhibitor cocktail (ThermoFisher; Halt Protease Inhibitor Cocktail (100X); REF: 87786). The detached samples were transferred to Eppendorf tubes, and excess PBS was removed. The samples were then snap-frozen in liquid nitrogen. Each sample consisted of either 5 hippocampal or 4 cerebellar organotypic slice cultures.

Frozen slices from the rat cerebellum and hippocampus were homogenized in 100 µL of lysis buffer with a protease/phosphatase inhibitor cocktail (Cell Signaling Technology, REF: 5872 S). The lysates were incubated on ice for 2 h in a rotating incubator, followed by centrifugation at 13,400 × g for 20 min at 4 °C. Supernatants were stored at -80 °C until further use. For SDS-PAGE, 30 µg of total protein were incubated in Laemmli sample buffer (0.1 M Tris, pH 6.8, 4% SDS, 10% β-mercapto-ethanol, 0.025% bromophenol blue, 20% glycerol, 5% ddH2O) for 5 min at 95 °C. Proteins were separated by SDS-PAGE and transferred onto a PVDF membrane using a Trans-Blot Turbo Transfer System (Bio-Rad Laboratories). Membranes were blocked with 5% skim milk in TBS-T for 1 h at room temperature. Primary antibodies against HIF-1α (Cayman Chemical, REF: CAY-10006421), HIF-2α (Novus Biologicals, REF: NB100-122), and Actin (Sigma-Aldrich, REF: A2103) were diluted in blocking buffer and incubated overnight at 4 °C. Secondary antibodies—goat anti-mouse (Sigma-Aldrich, REF: A2304) and goat anti-rabbit (Sigma-Aldrich; REF: A0545)—were diluted in blocking buffer and incubated for 1 h at room temperature. Protein signals were developed using SuperSignal West Femto Maximum Sensitivity Substrate (Thermo Fisher Scientific; REF: 34096) and detected with the Fusion FX System (Vilber). Band intensities were measured using ImageJ2 (version 2.9.0, National Institutes of Health, Bethesda), with normalization to actin blots as a loading control. Full, uncropped Western Blots for HIF-1α, HIF-2α, and the corresponding Actin bands are shown in Supplementary Fig. 1.

LDH Assay

The organotypic slice cultures were prepared as previously described, and before the oxygen-glucose deprivation (OGD) procedure, the samples were divided into two groups: an OGD group and a control group. Each group contained four inserts—two inserts with 4 cerebellar slice cultures and two with 5 hippocampal slice cultures each. Before transferring the OGD group inserts to the hypoxia chamber, 2 µl of media were collected from all wells to determine the background signal. The samples were stored in 48 µl of storage buffer at -80 °C. The storage buffer consisted of Tris-HCL (200mM, pH 7.3) (Fisher BioReagents; REF: BP153-1), 10% glycerol (J.T.Baker) & 1% BSA (ROTH; REF: 8076.2).

The control group remained under normoxic conditions in the previously described control medium. The OGD group was placed in the hypoxia chamber for 4 h using the established OGD medium. During this period, samples were taken after 2 h and 4 h under hypoxic conditions. After 4 h, the six-well plates were removed from the hypoxia chamber, and 26 µl of glucose (250 mg/ml) were added to the wells to simulate reoxygenation. Additionally, 1 µl of Edaravone (10 mM) (Aldrich; REF: M70800-5G) was added to the intervention parts of both the control and OGD groups, resulting in a final concentration of 10 µM Edaravone. This setup created an intervention group in both the OGD and control conditions, while the remaining OGD and control groups did not receive Edaravone. As illustrated in Fig. 1 further samples were taken 30 min, 1 h, 4 h, and 24 h after reoxygenation, and stored in 48 µl of storage buffer at -80 °C. The samples were then analyzed using the LDH-Glo Cytotoxicity Assay (Promega; REF: J2380), according to the manufacturer’s instructions. In the assay we used the recommended volumes of 50 µl LDH detection reagent and 50 µl sample with a 60 min incubation time.

Fig. 1.

Experimental timeline for an Oxygen-Glucose Deprivation (OGD) study. The sequence starts with the transfer of inserts into the OGD medium and the initiation of OGD, followed by sample collection at different time points (0, 2, and 4 h). After the termination of OGD at 4 h, glucose and Edaravone are added to the respective media. Additional samples are collected at various time points post-OGD (0.5, 1, 4, and 24 h). The experiment concludes with the final sample collection at 24 h post-OGD

Raw fluorescence values of each experimental group were normalized relative to the control group, with the mean values of the control group defined as 100%. This normalization allows a comparable representation of the results across different conditions.

ROS Assay

The organotypic slice cultures were subjected to OGD as previously described. After 4 h of OGD, the samples were reoxygenated by placing them back in the normoxic incubator and adding 26 µl of glucose (250 mg/ml) to the culture medium. At the same time, the intervention groups received 1 µl of Edaravone (10 mM) (Aldrich; REF: M70800-5G). Following reoxygenation, the samples were incubated for 4 h under these conditions, after which they were snap-frozen in liquid nitrogen. The samples were treated with 200 µl PBS and 1 µl of 1% Triton X-100, followed by sonication on ice. Insoluble particles were removed after centrifugation at 10,000 × g for 5 min. Subsequently, the samples were analyzed using the ROS assay OxiSelect In Vitro ROS/RNS Assay Kit (Green Fluorescence; REF: STA-347-T) according to the manufacturer’s instructions with a 45 min incubation time. The raw fluorescence values of each experimental group were normalized relative to the control group, with the mean values of the control group defined as 100%. This normalization allows for a comparable representation of the results across different conditions.

Seahorse Metabolic Flux Analysis of Organotypic Slice Cultures

To assess mitochondrial respiration, oxygen consumption rate (OCR) was measured in cerebellar and hippocampal organotypic slice cultures using the Seahorse XFe24 Analyzer (Agilent Technologies, USA). After the OGD procedure, slices were subjected to a 4-hour reperfusion phase in Seahorse XF DMEM medium (Agilent; REF: 103575-100) containing 36 mM glucose and 1x GlutaMAX (Thermo Fisher Scientific; REF: 35050061), with or without 10 µM Edaravone. Following the reperfusion period, culture inserts were quickly washed in Assay Medium (Seahorse XF DMEM medium, 5 mM glucose, 1x GlutaMAX) prior to collecting 1 mm tissue punches using a biopsy punch. Tissue punches were transferred to Seahorse XF24 Islet Capture Microplates (Agilent; REF: 101122-100) in 500 µL Assay Medium and secured using specialized mesh inserts to ensure consistent positioning within the wells during measurement. Following an 1 h equilibration period in a cell culture incubator (21% O₂, 0% CO₂, 37 °C), OCR was measured in response to sequential injections of metabolic modulators, including glucose (36 mM), pyruvate (10 mM), rotenone/antimycin A (Rot/AA; inhibitors of mitochondrial complexes I and III; 2 µM each), and 2-deoxy-D-glucose (2-DG; a glycolysis inhibitor; 100 mM). Measurements (3x baseline, 5x after each injection) consisted of the following phases: Mix 2 min, Wait 2 min, Measure 5 min. Each condition was measured three times using tissue punches derived from three different animals to ensure biological replicates and statistical reliability. Raw OCR values were normalized to DNA content of the respective tissue punch to account for potential differences in tissue integrity and cell number. Basal respiration was calculated by subtracting the last OCR value after Rot/AA injection from the last baseline OCR level. Glucose-induced respiration was calculated by subtracting the difference between the last baseline OCR value and the highest glucose-induced OCR value and pyruvate-induced OCR was calculated by subtracting the last glucose-induced OCR from the highest pyruvate-induced OCR value.

Statistical Analysis

The data were analyzed statistically in accordance with the methodology previously described [20]. Data analysis was conducted using GraphPad Prism 9 software (GraphPad Software, USA). The results are presented as mean values from at least three independent experiments, along with the standard error of the mean (SEM). Kolmogorov-Smirnov normality test was used to confirm normal distribution. To assess statistical significance between two groups, a two-tailed Student’s t-test was applied, while comparisons across multiple groups were performed using a one-way analysis of variance (ANOVA). Pairwise comparisons were conducted using Tukey’s post-hoc test. A p-value of less than 0.05 was considered indicative of statistical significance.

Results

OGD Effectively Induces Expression of Hypoxia Inducible Factors in Cerebellar and Hippocampal Organotypic Slice Cultures

To confirm that the OGD procedure effectively induces hypoxia and oxygen deprivation in the tissue, the expression levels of HIF-1α and HIF-2α were assessed at both the mRNA level, using qPCR, and the protein level, using Western blot analysis. mRNA expression was measured after 1, 4, and 8 h of OGD to determine the optimal duration of OGD that induces a consistent hypoxic response in the organotypic slice cultures for subsequent experiments.

qPCR analysis of the cerebellar organotypic slice cultures revealed a significant increase in HIF-1α and HIF-2α mRNA levels in the samples subjected to OGD. This upregulation of HIF expression was detectable as early as 1 h of OGD and persisted through 4 and 8 h. However, a decrease in Hif1a and Hif2a expression levels was observed after 8 h (Fig. 2A). In hippocampal slice cultures, qPCR similarly showed a significant increase in Hif1a and Hif2a levels following OGD. Unlike the cerebellar samples, both Hif1a and Hif2a expression continued to increase from 1 to 4 h of OGD in the hippocampus. Hif1a levels continued to rise after 8 h of OGD, whereas HIF2A expression decreased between 4 and 8 h (Fig. 2B). Since significant mRNA upregulation was observed at the 4 h time point in both cerebellum and hippocampus, this duration was selected for subsequent experiments.

Fig. 2.

HIF-1α and HIF-2α expression levels showed a tissue-specific upregulation following OGD in cerebellar and hippocampal organotypic slice cultures. (A) Hif1a and Hif2a expression levels normalized to Actb in cerebellar samples after 1, 4, and 8 h of OGD. Significant upregulation of Hif1a and Hif2a was observed at 1 h and 4 h of OGD. (B) Hif1a and Hif2a expression levels normalized to Actb in hippocampal samples after 1, 4, and 8 h of OGD. Hif1a showed significant upregulation at 4 h and 8 h of OGD, while HIF2A was significantly upregulated only at 4 h of OGD. (C) Representative Western blot images for HIF-1α and HIF-2α in cerebellar slice cultures after 4 h of OGD. Semiquantitative analyses indicated a trend toward increased HIF-1α protein levels and a significant upregulation of HIF-2α. Actin was used as loading control. (D) Representative Western blot images of HIF-1α and HIF-1α in hippocampal slice cultures after 4 h of OGD. Semiquantitative analyses showed a significant increase in HIF-1α protein levels, with no significant changes in HIF-2α protein expression. Actin was again used as a loading control. Data are expressed as mean ± SEM; *p < 0.05, **p < 0.01, ****p < 0.0001 (A, B One-way ANOVA with Tukey’s multiple comparisons test; C, D two-tailed Student’s t-test). (A, B) n = 40 cerebellar and 35 hippocampal slice cultures per group, five independent preparations; (C, D) n = 16 cerebellar and 20 hippocampal slice cultures per group, four independent preparations

Western blot analysis confirmed the findings from qPCR. In the cerebellar slice cultures, HIF-1α protein levels increased after OGD, though this increase did not reach statistical significance, while HIF-2α protein expression was significantly elevated after 4 h of OGD (Fig. 2C). In hippocampal slice cultures, Western blots showed a significant increase in HIF-1α protein levels in the OGD-treated samples. In contrast, no significant changes were observed in HIF-2α protein expression in the hippocampus (Fig. 2D).

Edaravone Displays Protective Properties in both Cerebellar and Hippocampal Organotypic Slice Cultures after OGD

To investigate whether Edaravone exhibits protective properties on organotypic slice cultures of the cerebellum and hippocampus following OGD-reperfusion conditions, the drug was added to the culture medium after a 4 h OGD period. The protective effect was assessed using an LDH assay, allowing for a quantitative comparison of cell damage between the treatment and control groups.

In cerebellar slice cultures, samples subjected to OGD exhibited significantly higher LDH levels in the medium compared to non-OGD controls (Fig. 3A). Cultures treated with Edaravone demonstrated a reduction in LDH levels compared to untreated OGD cultures, indicating a protective effect of the drug over the duration of the experiment (Fig. 3A). Four h after reoxygenation, the OGD group showed a marked increase in LDH levels relative to the control, but a substantial reduction was observed in Edaravone-treated cultures compared to the untreated OGD group, further supporting the protective effect of Edaravone (Fig. 3B).

Fig. 3.

Edaravone significantly reduced LDH levels and cell damage in organotypic slice cultures of the cerebellum and hippocampus following OGD. (A) Time course of LDH levels in cerebellar slices at different time points, with the X-axis representing time (h) and the Y-axis representing absolute luminescence values, indicating LDH levels. Over time, LDH levels increased in both the OGD and control groups, with Edaravone-treated samples consistently showing lower LDH levels compared to untreated samples. (B) Bar chart showing the relative LDH levels in cerebellar slices 4 h after reoxygenation (ReOx). Since hypoxia-induced cell damage was clearly evident at this point, the 4-hour post-reoxygenation time point was selected for analysis. Absolute LDH values were normalized to the control group and presented as percentages. Cerebellar samples in the untreated OGD group exhibited significantly elevated LDH levels compared to controls. A significant reduction in LDH levels was observed following Edaravone treatment, indicating a marked decrease in cell damage in cerebellar cultures. (C) Time course of LDH levels in hippocampal slices, showing increased LDH levels in the OGD groups compared to controls. (D) Bar chart showing the relative LDH levels in hippocampal slices 4 h after reoxygenation. Since hypoxia-induced cell damage was clearly evident at this point, the 4-hour post-reoxygenation time point was selected for analysis. Absolute LDH values were normalized to the control group and presented as percentages. In hippocampal samples, untreated OGD groups showed significantly elevated LDH levels compared to controls. Edaravone treatment resulted in a significant reduction in LDH levels, highlighting a substantial decrease in cell damage also in hippocampal cultures. Data are expressed as mean ± SEM; *p < 0.05, **p < 0.01 (One-way ANOVA with Tukey’s multiple comparisons test). (A, B) n = 20 slice cultures per group, five independent preparations; (C, D) n = 25 slice cultures per group, five independent preparations. ReOx in (A) and (C) indicates the time point of reoxygenation and glucose supplementation, and for the Edaravone group, the addition of Edaravone as well

Similarly, hippocampal slice cultures exposed to OGD showed higher LDH concentrations over time compared to control groups. Edaravone treatment of OGD-treated slices led to significantly reduced LDH levels compared to untreated OGD cultures (Fig. 3C). This trend continued 4 h after reoxygenation, with Edaravone-treated hippocampal slices showing markedly lower LDH levels than untreated OGD cultures (Fig. 3D). No significant reduction in LDH levels was observed in either cerebellar or hippocampal slices not exposed to OGD following Edaravone treatment.

Comparing the extent of LDH level increase between the cerebellum and hippocampus, OGD-exposed cerebellar slices exhibited a 390% higher mean LDH level compared to non-OGD controls (Fig. 3B), while hippocampal slices showed a smaller increase of 108% (Fig. 3D). In terms of LDH reduction following Edaravone treatment, cerebellar cultures displayed a 45.9% decrease in mean LDH levels, whereas hippocampal cultures demonstrated a greater reduction of 54.9%.

Edaravone Effectively Reduces ROS Levels in Cerebellar and Hippocampal Organotypic Slice Cultures After OGD

To investigate the potential protective mechanism of Edaravone as a radical scavenger, ROS levels were analysed in the various treatment groups using a ROS assay. For comparison, ROS levels in the control groups were normalized to 100% for both cerebellar and hippocampal organotypic slice cultures. In the cerebellar cultures, a significant increase in ROS levels was observed in the OGD group compared to the control group (Fig. 4A). However, in the Edaravone-treated OGD group, ROS levels were significantly reduced compared to the untreated OGD group (Fig. 4A). A similar pattern was observed in hippocampal slice cultures (Fig. 4B). The OGD group exhibited a marked increase in ROS levels, while Edaravone treatment significantly reduced ROS levels compared to the untreated OGD group (Fig. 4B).

Fig. 4.

Edaravone significantly reduces ROS levels in both cerebellar and hippocampal organotypic slice cultures following OGD. (A) ROS levels in cerebellar slices, and (B) ROS levels in hippocampal slices, are shown for the control and OGD groups, with and without Edaravone treatment. ROS levels were normalized to the control group and set to 100%. As with the LDH assay, samples were collected 4 h post-reoxygenation, coinciding with 4 h after Edaravone application. In both tissues, OGD groups exhibited a significant increase in ROS levels compared to controls. The application of Edaravone in OGD groups led to a significant reduction in ROS levels in both cerebellum and hippocampus. Fluorescence values for ROS measurements were obtained in triplicates. Data are presented as mean ± SEM; *p < 0.05, **p < 0.01, ****p < 0.0001 (One-way ANOVA with Tukey’s multiple comparisons test). (A, B) n = 48 cerebellar and 60 hippocampal slice cultures per group, four independent preparations

When comparing the magnitude of ROS level increase between the cerebellum and hippocampus in OGD-exposed samples without Edaravone, ROS levels in cerebellar slices were 39.1% higher than in non-OGD controls, while the increase in hippocampal slices was 25.8%. Edaravone treatment resulted in a 22.0% reduction in ROS levels in cerebellar slices, whereas the reduction in hippocampal slices was 11.2%.

Tissue-Dependent Metabolic Response to Edaravone Treatment After OGD

To further elucidate the protective mechanisms of Edaravone, mitochondrial respiration was assessed using the Seahorse XFe24 Analyzer in cerebellar and hippocampal slice cultures subjected to OGD conditions, followed by reoxygenation with and without Edaravone treatment.

Following OGD, a marked decline in raw OCR was observed in both cerebellar and hippocampal slices (Fig. 5A, B). Edaravone treatment fully restored raw OCR levels in both tissues (Fig. 5A, B). To account for differences in cell density, OCR values were normalized to DNA content, reflecting mitochondrial respiration per cell rather than total tissue oxygen consumption. This normalization revealed distinct recovery patterns between the two tissues and helped differentiate whether Edaravone’s effects resulted from preserved cellular integrity or direct mitochondrial enhancement. In this DNA-normalized dataset, OCR in the OGD-group of cerebellar slice cultures (Fig. 5C) was reduced in the OGD group compared to normoxic controls, although not significant. Edaravone treatment during reperfusion did not restore OCR, as OGD + Edaravone cultures exhibited respiration levels similar to the untreated OGD group. This suggests that the observed reduction in raw OCR in cerebellar slices was primarily due to cell loss rather than direct mitochondrial dysfunction. In contrast, hippocampal slice cultures (Fig. 5D) showed a different response to Edaravone treatment. Following OGD, OCR was reduced compared to normoxic controls. In the DNA-normalized OCR levels, Edaravone-induced improvement of respiration was still apparent. This tissue-dependent metabolic response is further supported by the calculation of basal respiration, glucose-induced maximal respiration, and pyruvate-induced maximal respiration in OGD and OGD + Edaravone-treated groups. For this purpose, OCR levels of both groups were normalized to the respective control group to calculate relative change after treatment. Basal respiration analysis revealed distinct tissue-specific responses to Edaravone. In cerebellar slices (Fig. 5E), calculations based on raw OCR data showed that OGD led to a decrease in basal respiration compared to control data, though this reduction was not statistically significant compared to Edaravone-treated OGD slices. Still, Edaravone treatment restored basal respiration to control levels, consistent with its cell-protective effects. In the DNA-normalized dataset, basal respiration remained at approximately 100% across all experimental conditions, reinforcing that Edaravone’s primary effect in the cerebellum was the preservation of tissue integrity rather than direct mitochondrial enhancement. In contrast, hippocampal slices (Fig. 5F) responded differently to Edaravone treatment. In this tissue, Edaravone fully restored basal respiration of OGD slices and even increased it beyond control levels, resulting in significantly higher basal respiration compared to OGD slices without Edaravone treatment. This suggests that in the hippocampus, Edaravone supports mitochondrial function beyond cell protection, potentially through enhanced metabolic efficiency or mitochondrial biogenesis. Substrate-specific further underscored these tissue-dependent effects. In cerebellar slices (Fig. 5E), glucose-induced OCR was slightly reduced in both OGD groups, though the difference did not reach statistical significance, and pyruvate-induced maximal respiration remained unchanged across all groups. In hippocampal slices (Fig. 5F), Edaravone treatment significantly increased glucose-driven respiration following glucose injection in the OGD + Edaravone group compared to untreated OGD slice cultures. Pyruvate-induced respiration exhibited a trend toward higher values in the OGD + Edaravone condition, though this effect did not reach statistical significance. These findings suggest a tissue-specific response to Edaravone, with its primary effect in the cerebellum being the preservation of tissue integrity, whereas in the hippocampus, it appears to support mitochondrial respiration after ischemic stress.

Fig. 5.

Tissue-Dependent Mitochondrial Response to Edaravone Treatment After OGD. (A, B) Raw oxygen consumption rate (OCR) data for cerebellar (A) and hippocampal (B) organotypic slice cultures under normoxic conditions (blue line), after OGD (red line), and with (dashed lines) or without (solid lines) Edaravone treatment. Thin dashed lines indicate the standard error of the mean (SEM). Time points of glucose, pyruvate, rotenone/antimycin A (Rot/AA), and 2-deoxy-D-glucose (2-DG) injections are indicated. (C, D) DNA-normalized OCR data for cerebellar (C) and hippocampal (D) cultures. In cerebellar slices, OGD led to a slight reduction in normalized OCR compared to normoxic controls. Edaravone treatment did not restore OCR, as OGD + Edaravone cultures exhibited similar respiration levels to the untreated OGD group. In hippocampal slices, OGD resulted in a decrease in OCR compared to normoxic controls, whereas Edaravone treatment restored OCR to control levels. (E) Basal and substrate-driven respiration in cerebellar slice cultures. Raw OCR (left) and normalized OCR (right) levels were expressed relative to their respective control groups. Edaravone treatment did not significantly improve respiration across all conditions. (F) Basal and substrate-driven respiration in hippocampal slice cultures. Raw OCR (left) and normalized OCR (right) levels were expressed relative to their respective control groups. Hippocampal slice cultures exhibited a significant increase in basal- and glucose-driven OCR following Edaravone treatment. Pyruvate-induced OCR showed a non-significant upward trend with Edaravone treatment. The 100% control values are indicated by a dashed line. Data are presented as mean ± SEM, *p < 0.05, **p < 0.01, **p < 0.001, ****p < 0.0001 (two-tailed Student’s t-test); n = 3 slice cultures per group

Discussion

Ischemic stroke is a leading cause of disability and death worldwide, with the cerebellum being particularly vulnerable due to its critical role in motor coordination and balance [5]. Neuronal damage in cerebellar stroke arises from oxygen-glucose deprivation (OGD), which triggers complex pathological mechanisms such as ionic dysregulation, altered metabolism, oxidative stress, and inflammation [21, 22]. Purkinje cells, essential for cerebellar function, are highly susceptible to hypoxic injury, leading to severe and often irreversible consequences for stroke survivors [5, 23]. Current stroke therapies, such as thrombolysis or thrombectomy, primarily focus on restoring blood flow and preventing recurrence [24], with few options that directly target the molecular cascades underlying neuronal injury. Thus, developing cell protective strategies that mitigate cell death could significantly improve patient outcomes.

In this study, we investigated the potential cell protective effects of Edaravone, a radical scavenger known for its efficacy in mitigating oxidative stress, particularly in the context of stroke [10, 25, 26]. Using an OGD model in organotypic slice cultures from rat cerebellum and hippocampus, we simulated ischemic stroke conditions and evaluated Edaravone’s ability to reduce cellular damage [19]. The hippocampus, a region highly sensitive to hypoxia [27], was used as a comparative structure in this study, as prior studies have demonstrated Edaravone’s neuroprotective effects specifically in hippocampal tissue [28–30]. This provided a robust benchmark to assess the strength of Edaravone’s protective effects in the cerebellum.

Our findings confirm that OGD effectively induces hypoxia, as evidenced by significant upregulation of HIF-1α and HIF-2α mRNA in both cerebellar and hippocampal tissues [31]. These markers are crucial indicators of hypoxic response [32]. Interestingly, while both tissues exhibited elevated HIF mRNA levels, the protein analysis revealed tissue-specific differences. In the cerebellum, HIF-2α protein levels were significantly increased, whereas HIF-1α was significantly upregulated in the hippocampus. These discrepancies could be attributed to the cellular composition of the tissues. HIF-1α is predominantly expressed in neurons, whereas HIF-2α is primarily found in non-neuronal cells such as endothelial cells and astrocytes [33]. Especially endothelial cells play a central role in the expression of HIF-2α under hypoxic conditions [34]. Given the cerebellum’s relatively higher density of endothelial cells, the stronger HIF-2α response likely reflects the endothelial contribution to hypoxic adaptation in this tissue [35]. The lack of significant HIF-1α upregulation in the cerebellum at the 4-hour time point might be explained by differences in mechanisms of proteostasis or the metabolic activity of cerebellar neurons compared to hippocampal neurons. This difference could be attributed to the fact that the hippocampus is a brain region that shows a particularly strong accumulation of HIF-1α under hypoxia [36]. Additionally, the higher LDH levels observed in cerebellar cultures after OGD suggest more pronounced cellular damage compared to the hippocampus, which could significantly influence the observed differences in HIF expression. The selective loss of cerebellar granule cells, as demonstrated in our previous work [19], may have led to a reduced contribution of HIF-1α-producing neuronal populations, whereas the remaining non-neuronal cells, such as astrocytes and endothelial cells, could continue to upregulate HIF-2α. The greater extent of cell damage in the cerebellum could thus contribute to the differential expression patterns observed. While cerebellar neurons might degrade HIF-1α protein more rapidly during reoxygenation, limiting its accumulation, the cerebellum’s hypoxic response might be dominated by endothelial pathways, reducing the relative contribution of neuronal HIF-1α. These tissue-specific dynamics highlight the complexity of the cerebellum’s response to hypoxia and suggest that its vulnerability might not solely depend on neuronal factors. Further studies focusing on cell-type-specific dynamics of HIF expression and stability are needed to clarify these mechanisms and their implications for therapeutic interventions like Edaravone.

The selection of the 4 h OGD time point for Edaravone treatment was based on our observation that hypoxia-induced cell responses were well-established at this stage. Although earlier cellular responses were detectable after 1 h, longer OGD durations likely provide a more clinically relevant scenario, as stroke patients typically present after a delay [37, 38]. This approach aligns with therapeutic windows for lysis or thrombectomy, making Edaravone a potential adjunct treatment during revascularization and reoxygenation [24]. Future studies should explore the timing and efficacy of Edaravone administration, including pre-hypoxia applications, particularly in high-risk stroke models or surgical scenarios where ischemic events are anticipated. Considering its potent antioxidant properties and established safety profile, Edaravone has the potential to serve as a prophylactic agent, preconditioning neuronal tissue against ischemic injury, mitigating oxidative damage, and reducing the impact of anticipated hypoxic conditions [39].

While previous studies have already indicated the neuroprotective effect of Edaravone in hypoxic cell damage [40], in our study Edaravone also demonstrated significant protective effects in both cerebellar and hippocampal tissues, as indicated by reduced LDH levels in OGD-treated samples. LDH release, a marker of cellular damage, is triggered by membrane permeability loss during hypoxia [41]. Edaravone significantly reduced LDH levels in both tissues, indicating comparable cellular protection in cerebellar and hippocampal tissues, a finding consistent with previous studies on hippocampal protection [28, 30]. However, we observed that the cerebellum experienced a higher percentage increase in LDH levels compared to the hippocampus under OGD conditions. This could suggest greater vulnerability to hypoxia in the cerebellum, though the hippocampus is known to be highly sensitive to hypoxic damage [42]. While the cerebellum showed more pronounced cellular damage, the hippocampus may suffer more from functional impairments due to its involvement in cognitive functions such as memory and learning [43]. This heightened susceptibility is attributed to the vulnerability of hippocampal pyramidal cells and fast-spiking interneurons, which are essential for hippocampal function [44, 45]. Their loss leads to significant functional deficits [43]. Since our assay focused on cellular damage (LDH release), rather than functional deficits, we may not fully capture the hippocampal vulnerability. Future studies incorporating functional assessments would provide a clearer picture of Edaravone’s protective effects.

In addition to its effects on cellular damage, Edaravone significantly reduced ROS levels in both cerebellar and hippocampal slice cultures, emphasizing its role as an effective radical scavenger [10]. OGD conditions induced an increase in ROS production, which persisted through the reoxygenation phase, further exacerbating cell damage. Edaravone’s ability to mitigate this oxidative burst highlights its therapeutic potential in limiting reperfusion injury, a major contributor to neuronal death following stroke [9, 46]. Notably, cerebellar cultures showed higher ROS levels than hippocampal cultures after 4 h OGD, which might reflect the cerebellum’s distinct cellular composition and metabolic demands. Previous studies have shown that the cerebellum, along with the brain stem, exhibits the highest basal rate of ROS production [47]. The relatively high density of astrocytes in the hippocampus, which are known for their antioxidative metabolism and increased glutathione synthesis, may contribute to lower ROS levels under hypoxic conditions [48]. In contrast, Purkinje cells, with their extensive dendritic arborization and elevated metabolic activity, may further amplify ROS production in the cerebellum [49]. These factors could explain the cerebellum’s heightened oxidative response and its relatively higher ROS levels during hypoxia. Our findings indicate that Edaravone’s radical-scavenging effects were more pronounced in the cerebellum, potentially due to the higher initial ROS levels, which provide a greater window for antioxidative intervention. However, excessive ROS levels might also overwhelm Edaravone’s scavenging capacity, partially contributing to less pronounced reduction in LDH levels observed in the cerebellum. It is important to note that elevated ROS levels do not necessarily correlate with greater functional damage. Recent studies have shown that following OGD in hippocampal slice cultures, Edaravone treatment did not attenuate the decline in neuronal transmission [50]. Since our study focused on quantitative cell survival rather than functional assessment, this represents a limitation of our approach and should be further investigated in future studies. The cited study also highlights the importance of superoxide radicals and suggests that Edaravone, as a non-specific radical scavenger, may be less effective compared to other agents specifically targeting super oxides. This aspect requires further exploration to determine the most effective antioxidant strategy.

Mitochondrial dysfunction plays a central role in ischemic neuronal damage, as OGD disrupts oxidative phosphorylation and ATP production [51]. Our findings reveal tissue-specific differences in mitochondrial response to OGD and Edaravone treatment. In cerebellar tissue, Edaravone primarily preserved tissue integrity during reperfusion, as indicated by higher OCR levels after normalization, suggesting that the observed reductions in raw OCR values after OGD were primarily due to cell loss rather than mitochondrial dysfunction. The higher LDH release in cerebellar tissues under OGD conditions reflects greater cellular damage, consistent with the heightened vulnerability of metabolically active Purkinje cells to hypoxic injury [52], which might explain why Edaravone did not influence mitochondrial respiration in cerebellar tissue. In contrast, hippocampal cultures exhibited a distinct mitochondrial response. Both, raw and DNA-normalized OCR data showed a significant enhancement of mitochondrial function in the OGD + Edaravone group which resulted in increased basal respiration and glucose-induced maximal respiration. This suggests that Edaravone not only maintains cellular integrity but also actively enhances mitochondrial activity in this tissue. This may involve mechanisms such as increased metabolic efficiency or mitochondrial biogenesis, potentially mediated by transcription factors like PGC-1α, which regulate mitochondrial adaptation to oxidative stress [53]. The improved mitochondrial function in the hippocampus is further supported by reduced LDH release and lower ROS levels, suggesting a more robust recovery of metabolic and antioxidative capacity. Our findings further support a tissue-specific role of Edaravone in stroke recovery. In cerebellar tissue, the lack of significant changes in DNA-normalized OCR across all conditions suggests that Edaravone’s primary function is cell preservation rather than direct mitochondrial enhancement. The recovery of basal OCR in raw data, despite unchanged DNA-normalized OCR, reinforces this notion, indicating that Edaravone prevents cell loss but does not necessarily improve mitochondrial respiration per cell. In contrast, hippocampal slices exhibited a clear enhancement of mitochondrial function, particularly in glucose-induced OCR, suggesting Edaravone promotes metabolic efficiency post-ischemia. The fact that pyruvate-driven respiration remained unaffected suggests that Edaravone’s effect is primarily linked to glucose metabolism, which represents the primary source of energy for the brain as it fuels the TCA cycle to enhance oxidative phosphorylation. These findings highlight the importance of region-specific metabolic responses in ischemic brain injury and recovery. Edaravone’s effect in the cerebellum appears more structural, whereas in the hippocampus, it enhances cellular metabolism and mitochondrial function, which could have implications for functional recovery and neuroplasticity. These insights emphasize the need for tailored therapeutic approaches depending on the affected brain region, particularly in stroke therapy where metabolic and cellular vulnerabilities differ. The interplay between ROS scavenging and mitochondrial function is particularly evident in the hippocampus. The marked improvement in mitochondrial respiration in hippocampal slices treated with Edaravone suggests that ROS reduction facilitated mitochondrial recovery and may have activated pathways supporting mitochondrial repair or biogenesis. In contrast, the cerebellum’s less efficient mitochondrial recovery, as indicated by basal respiration as well as glucose-induced and pyruvate-induced maximal respiration, could reflect a diminished capacity for adaptive responses under hypoxic and/or OGD stress, potentially due to its distinct cellular composition and metabolic profile [49].

These findings suggest potential tissue-specific roles for Edaravone in stroke therapy. In the cerebellum, Edaravone may primarily help mitigate cell loss and reduce post-stroke atrophy, which could contribute to preserving motor coordination. In the hippocampus, the observed enhancement of mitochondrial function raises the possibility that Edaravone supports cognitive recovery and neuroplasticity. This highlights the broader potential of Edaravone’s neuroprotective effects, which might extend beyond ROS scavenging to include a role in preventing mitochondrial dysfunction. The observed regional disparities further emphasize the importance of tailoring therapeutic strategies to the specific vulnerabilities and recovery capacities of different brain regions. During reperfusion, where oxidative stress and mitochondrial impairment contribute significantly to secondary injury, Edaravone’s ability to mitigate ROS and support mitochondrial recovery highlights its potential as a region-specific therapeutic agent.

Furthermore Edaravone’s neuroprotective effects go beyond ROS scavenging, potentially involving the prevention of brain oedema and modulation of inflammatory pathways [54, 55]. These additional mechanisms could be particularly relevant in cerebellar stroke, where oedema and brainstem compression pose significant risks [56]. While this study focused on cellular damage, future in vivo studies should investigate Edaravone’s broader therapeutic potential, particularly its role in preventing oedema and managing inflammation.

Edaravone might also have potential applications beyond ischemic stroke. For instance, during myocardial infarction or cardiac pump failure, reduced cardiac output leads to cerebral hypoxia, which our OGD model simulates [57]. Edaravone could offer neuroprotection in these contexts, particularly during resuscitation when reperfusion injury may exacerbate neuronal damage. Additionally, its application in perinatal and neonatal asphyxia, where therapeutic hypothermia remains the only treatment for reducing long-term neurological damage, could represent a critical area for future investigation [58].

In summary, our findings indicate that Edaravone provides significant cell protection in both cerebellar and hippocampal tissues under OGD conditions, primarily through its antioxidant properties. These results suggest that Edaravone holds promise as a therapeutic agent for cerebellar stroke, particularly in combination with reperfusion strategies. Future studies should explore Edaravone’s full therapeutic potential in vivo, including its effects on functional recovery, oedema prevention, and broader clinical applications in hypoxic conditions beyond stroke.

Conclusion

This study highlights the cell protective potential of Edaravone in reducing cellular damage and oxidative stress in cerebellar and hippocampal organotypic slice cultures under OGD-reoxygenation conditions. Our findings reveal tissue-specific responses to ischemic injury and antioxidant treatment, with distinct differences in ROS production, LDH release, and mitochondrial function between the cerebellum and hippocampus. Edaravone’s effectiveness in mitigating these effects highlights its therapeutic value for addressing oxidative stress and ischemic damage. The observed reductions in ROS and LDH levels offer mechanistic insights into its protective action, supporting its application as a post-ischemic intervention. Importantly, our results demonstrate that Edaravone primarily preserves tissue integrity in cerebellar cultures by preventing cell loss rather than directly enhancing mitochondrial function. In contrast, hippocampal cultures exhibited a significant improvement in mitochondrial activity after Edaravone treatment, suggesting additional metabolic benefits such as enhanced mitochondrial efficiency or biogenesis. These findings indicate that Edaravone’s protective effects are not uniform but are dependent on the tissue type, highlighting the need for targeted therapeutic approaches.

In conclusion, Edaravone emerges as a promising therapeutic candidate for cerebellar stroke and other hypoxic brain injuries. Future in vivo studies should investigate its broader therapeutic potential, particularly in conjunction with reperfusion strategies, to prevent neuronal death, preserve tissue integrity, reduce oedema, and enhance long-term neurological recovery.

Electronic Supplementary Material

Below is the link to the electronic supplementary material.

Author Contributions

VM, CT, and TW contributed to the study conception and design. Material preparation, data collection and analysis were performed by JD, YH, SS, and VM. The first draft of the manuscript was written by JD and VM and YH, SS, TW, and CT commented on previous versions of the manuscript. All authors read and approved the final manuscript.

Funding

Open Access funding enabled and organized by Projekt DEAL.

The author(s) received no specific funding for this work.

Data Availability

All data supporting the findings of this study are included within the manuscript.

Declarations

Ethics Approval

All procedures were conducted under established standards of the German federal state of North Rhine Westphalia, in accordance with the European Communities Council Directive 2010/63/EU on the protection of animals used for scientific purposes. According to current German and European legislation, the removal of organs or cells from vertebrates for scientific purposes is not considered an animal experiment if the animals have not been subject to surgical interventions or invasive treatments prior to sacrifice. Thus, the euthanasia of rats for the removal of brain tissue does not require approval or permission from local or federal authorities.

Human Ethics and Consent to Participate

Not applicable, as no human participants were involved in this research.

Competing Interests

The authors declare no competing interests.