The chemicals used were hemin (Sigma, St. Louis, MO, United States) and afatinib (AdooQ Bioscience, Irvine, CA, United States). Hemin was dissolved in ammonia water and pH value was corrected to pH 7.4. Afatinib was dissolved in dimethyl sulfoxide (DMSO, Sigma) and diluted with DMEM or Neurobasal (NB, Thermo Fisher Scientific, Waltham, MA, United States) medium.

Twenty-three pregnant female Sprague-Dawley (SD) rats were supplied by BioLASCO Taiwan Co., Ltd. (Yilan, Taiwan). All animals (one rat/individually ventilated cage) were housed in an air-conditioned room (22 ± 2°C) on a 12 hr-light/dark cycle (07:00–19:00h light) and had free access to food and water. Pregnant female Sprague-Dawley (SD) rats of 17-day gestation were sacrificed by an overdose of Zoletil^® (Virbac, Taiwan) to minimize pain or discomfort. Embryonic day 17 fetal rat brains obtained from pregnant female SD rats were used to prepare primary cultured cortical neurons. The use of animals and all experiments conducted were under approved protocols from the Institutional Animal Care and Use Committee (IACUC) of Taipei Veterans General Hospital, Taipei, Taiwan. The approval number is IACUC2022-235. All experiments were performed in accordance with relevant guidelines and regulations.

Cerebral cortices of fetal rats were isolated and dissociated mechanically. The dissociated cells were suspended in the Basal Medium Eagle (BME, Thermo Fisher Scientific) medium containing 20% fetal bovine serum, and were seeded onto 35-mm culture dishes (IWAKI, Tokyo, Japan) with a density of 5 × 10⁶ cells per dish. Afterwards, cells were maintained with serum-free Neurobasal medium supplemented with B27 (Thermo Fisher Scientific) in the incubator with 5% CO₂ at 37°C. Four experimental treatments included vehicle (as control), hemin (30 μM), hemin (30 μM) plus afatinib (10 nM) and afatinib (10 nM).

In brief, primary cultured cortical neurons were seeded on a 24-well plate. Concentration-dependent effects (10–60 μM) of hemin were performed for 16 and 24 h. The effect of afatinib was investigated 16 h after simultaneous addition of afatinib (10 nM) and hemin (30 μM). Cytotoxicity was determined by a Lactate Dehydrogenase (LDH) assay. The LDH released in the culture medium was assessed by adding β-nicotinamide adenine dinucleotide and sodium pyruvate (Sigma). LDH activity was determined by measuring the absorbance at 340 nm for 6 min using an enzyme-linked immunosorbent assay (ELISA) reader (TECAN Sunrise, Männedorf, Switzerland). The LDH activity of cells treated with 0.1% Triton X-100 was used as control set to 100%.

At the end of 16-h treatments, the cells were collected, washed with phosphate buffered saline (PBS), and lysed in a radioimmunoprecipitation assay (RIPA, Cell Signaling Tech., Beverly, MA, United States) lysis buffer containing 20 mM Tris HCl, 150 mM NaCl, 1% (v/v) NP-40, 1% (w/v) sodium deoxycholate, 1 mM ethylenediaminetetraacetates (EDTA), 0.1% (w/v) sodium dodecyl sulfate polyacrylamide (SDS) and 0.01% (w/v) sodium azide (pH 7.5) for 20 min on ice. Lysates were then centrifuged at 13,800xg for 10 min, and the protein concentrations of supernatants were determined by Pierce BCA Protein Assay Kit (Thermo Fisher Scientific). Protein samples (30 μg) were run on 12%–13.5% SDS-polyacrylamide gel electrophoresis and then transferred onto a polyvinylidene difluoride (PVDF, Bio-Rad, Hercules, CA, United States) at 100 V for 120 min. Blots were probed with primary antibodies including antibodies against p-EGFR/total-EGFR, p-AKT/total-AKT (Cell Signaling Tech.), Tumor necrosis factor (TNF)-α, Cyclooxygenase 2 (COX2), Heme oxygenase-1 (HO-1) (StressGen, Victoria, CA, United States), Glutathione hydroperoxidase 4 (GPX4) and Receptor-interacting serine/threonine-protein kinase 3 (RIP3) (Cell Signaling Tech.) overnight at 4°C. After incubation of primary antibodies, the membrane was washed and incubated with a secondary antibody for 1 h at room temperature. The secondary antibodies were horseradish peroxidase-conjugated secondary IgG (Chemicon, Temecula, CA, United States). The immunoreaction was visualized using Amersham Enhanced Chemiluminescence (Amersham Pharmacia Biotech, Piscataway, NJ, United States). After this measurement, the bound primary and secondary antibodies were stripped by incubating the membrane in stripping buffer (100 mM 2-mercaptoethanol, 2% SDS) at 50°C for 5 min. The membrane was reprobed with a primary antibody against β-actin (Millipore, Billerica, MA, United States).

At the end of 1-hr treatments, the cells were fixed with 4% paraformaldehyde (Merck, Boston, MA, United States). Cells were then washed with 0.1 M PBS, incubated with 0.3% Triton X-100 (Sigma) and 1% goat serum (GS; Jackson ImmunoResearch, West Grove, PA, United States), and blocked with 3% GS for 60 min. Next, cells were processed for immunostaining using mouse monoclonal antibody specific for rat p-EGFR and microtubule-associated protein 2 (MAP-2, Millipore) in 1% GS-PBS at 4°C for 24 h. The cells were then incubated in fluorescein conjugated-IgG (FITC) (Jackson ImmunoResearch) and Texas Red dye-conjugated IgG fraction monoclonal mouse anti-biotin (Jackson ImmunoResearch) for 1 h at room temperature, mounted in glycerol (Merck). Controls consisted of omission of primary antibodies. The sections were visualized by a fluorescence confocal microscope (Olympus FluoView, Norfolk, VA, United States).

At the end of 24-h treatments, the culture medium was collected to measure NO production. The culture medium was mixed with an equal volume of the Griess reagent (1% sulfanilamide, 0.1% N-(1-Naphthyl)ethylenediamine in 2.5% H₃PO₄) and incubated for 15 min at room temperature in the dark. Nitrite concentration was determined by measuring the absorbance at 550 nm using an ELISA plate reader (TECAN Sunrise, Männedorf, Schweiz).

All data are expressed as the mean ± standard error of the mean (S.E.M.). The results were analyzed by one-way analysis of variance (one-way ANOVA) followed by the least significance difference (LSD) test as post-hoc method. The significance level was set at p < 0.05.

To mimic the neurodegeneration in ICH, a hemin-induced neurotoxicity model was established in primary cultured cortical neurons. The LDH assay showed that incubation of hemin (10–60 μM) for 16 and 24 h increased neuronal death in time- and concentration-dependent manners (Figure 1A). The IC₅₀ of hemin in primary cortical neurons was about 30 μM after 24-hr incubation. Co-incubation with afatinib (10 nM) for 16 h significantly attenuated hemin-induced cell death in primary cultured cortical neurons (Figure 1B), indicating that afatinib is capable of inhibiting hemin-induced neuronal death.

The involvement of EGFR signaling in the hemin-induced neurotoxicity was investigated by measuring p-EGFR levels in primary cultured cortical neurons. We found that 1-h incubation of hemin (10–60 μM) concentration-dependently increased EGFR phosphorylation (Figure 2A). EGFR phosphorylation was evident when cells were treated with 30 μM hemin and maintained elevated with 60 μM hemin (Figure 2A). Similarly, incubation of hemin for 1 h significantly elevated p-AKT levels in a concentration-dependent manner (Figure 2B). Co-incubation of afatinib (10 nM) significantly attenuated hemin-induced EGFR phosphorylation (Figure 2C) and AKT phosphorylation (Figure 2D), indicating hemin indeed activated EGFR-AKT signaling in primary cultured cortical neurons.

Furthermore, we investigated the effect of afatinib on hemin-induced morphological changes in primary cultured cortical neurons. Compared with the vehicle-treated cells, immunofluorescent staining data showed that hemin concentration-dependently (10–60 μM) increased p-EGFR immunoreactivity and damaged neurite outgrowth (Figure 3A). Incubation of hemin (10 μM) for 8 h did not cause significant changes in the neurite outgrowth. However, higher concentrations of hemin (30–60 μM) induced strong p-EGFR immunoreactivity. At the same time, hemin caused focal bead-like swellings, neuritic beading and discontinuities of neurites in primary cultured cortical neurons (Figure 3A). Co-incubation with afatinib (10 nM) attenuated hemin-induced elevation in p-EGFR immunoreactivity and impairment in neurite outgrowth (Figure 3B), suggesting that the EGFR signaling pathway is responsible for the hemin-induced damage to neurite outgrowth in primary cultured cortical neurons.

To further confirm the involvement of EGFR-AKT signaling in ICH, afatinib was employed to block hemin-induced activation of EGFR-TK signaling and related neurotoxicity. First, we established hemin-induced neuroinflammation in primary cultured cortical neurons. Western blot assay showed that a 16-hr incubation of hemin concentration-dependently (10–60 μM) increased TNF-α (Figure 4A) and COX2 protein levels (Figure 4B) in primary cultured cortical neurons. Co-incubation with afatinib (10 nM) prevented hemin (30 μM)-induced elevations in TNF-α (Figure 4C) and COX2 (Figure 4D) as well as NO production in the culture medium of treated neurons (Figure 4E), suggesting that afatinib is capable of reducing hemin-induced neuroinflammation. At the same time, we investigated the effect of afatinib on HO-1 expression (an enzyme catalyzing hemin). Western blot assay showed that a 16-h incubation of hemin (10–60 μM) increased HO-1 levels (Figure 5A). Co-incubation with afatinib (10 nM) prevented hemin (30 μM)-induced elevation in HO-1 (Figure 5B), suggesting that afatinib is capable of reducing hemin-induced HO-1 expression.

The cell death mechanisms underlying hemin-induced neurotoxicity were investigated by measuring GPX4 (a biomarker of ferroptosis) and receptor-interacting protein 3 (RIP3, a biomarker of necroptosis). Western blot assay demonstrated that hemin concentration-dependently (10–60 μM) reduced GPX4 (Figure 6A) and increased RIP3 (Figure 6B). Co-incubation with afatinib (10 nM) inhibited the hemin (30 μM)-induced reduction in GPX4 (Figure 6C) and elevation in RIP3 (Figure 6D). These data indicate that afatinib is capable of reducing hemin-induced ferroptosis and necroptosis.