Analytical-grade artemisinin, CORT, and Fluoxetine were obtained from Meilunbio (Dalian, China). Dulbecco’s modified Eagle’s medium (DMEM), fetal bovine serum (FBS), and 0.25% trypsin were from GIBCO (USA). DMSO and penicillin/streptomycin were sourced from Sigma-Aldrich (USA). Poly-D-lysine, MTT, Lipofectamine 3000, and BCA Protein Assay Kit were from Thermo Fisher Scientific (USA). Annexin V: FITC Apoptosis Detection Kit was from BD Biosciences (USA). SDS-PAGE gels (4%–20%) were purchased from Genscript Biology (China). Primary antibodies for GAPDH, phospho-AKT1, phospho-ERK1/2, phospho-CREB, phospho-TrkB, phospho-GSK3β, NeuN, and HRP-conjugated secondary antibodies were from CST (USA). BDNF antibody was from Abcam (UK), while GFAP, NRF2, and HO1 antibodies were from Signalway Antibody (USA). MEK inhibitor PD98059 and PI3K inhibitor LY294002 were from Calbiochem (USA). Lactate dehydrogenase (LDH) cytotoxicity, mitochondrial membrane potential (MMP, △ψm; JC-1) and reactive oxygen species (ROS) detection kits were from Beyotime Biotechnology (China).

PC12 cells were provided by Dr. Gordon Guroff (NIH, USA) and cultured in DMEM with 10% FBS and penicillin/streptomycin at 37 °C in 5% CO2. Cells were sub-cultured 2–3 times per week at a 1:5 split. Primary neurons were obtained from one-day-old C57BL/6 mice as per Zhao and colleagues (Zhao et al., 2019b). Artemisinin and CORT were dissolved in DMSO (≤0.1% final concentration). Experimental groups included: (1) Control: serum-free medium, (2) CORT group: 200 μM CORT, and (3) CORT + Artemisinin: 200 μM CORT with 3.125–100 μM artemisinin for 48 h.

Cell viability was assessed using the MTT assay. Cells (6–8 × 10³/well) were treated with artemisinin and CORT for 48 h, followed by incubation with 0.5 mg/mL MTT for 3–4 h. Formazan crystals were dissolved in 150 μL DMSO, and absorbance was measured at 490 nm using an Infinite M200 PRO microplate reader (Tecan, Switzerland). Cell viability was expressed as a percentage of control.

LDH release was measured using a commercial kit (Beyotime). Cells (6–8 × 10³/well) were treated as described, and fluorescence intensity was measured at 560/590 nm using an Infinite M200 PRO microplate reader. LDH release was normalized to the control group.

ROS levels were detected using DCFH-DA (Beyotime). Cells were incubated with 10 μM DCFH-DA in DMEM at 37 °C for 30 min, washed, and fluorescence was observed using an EVOS FL Imaging System. ROS levels were semi-quantified using ImageJ.

MMP (∆ψm) was assessed using the JC-1 kit (Beyotime). Cells (1 × 10⁴ cells/cm²) were treated for 48 h, stained with JC-1 (10 μg/mL) at 37 °C for 20 min, washed, and fluorescence was analyzed using a fluorescence microscope. The red/green fluorescence ratio was normalized to the control group.

Apoptosis was measured using an Annexin V-FITC/PI apoptosis detection kit (BD Biosciences, San Diego, CA, USA) according to the manufacturer’s instructions. Cells were treated for 48 h, then 1 × 10⁵ cells were collected and stained with Annexin V-FITC and PI. Based on the assay principle, intact cells were double-negative for Annexin V and PI; early apoptotic cells were identified as Annexin V-positive/PI-negative, while late apoptotic or necrotic cells were positive for both Annexin V and PI. To quantify the apoptotic cells, CellQuest™ Pro Software (BD Biosciences, San Diego, CA) was utilized, and the apoptosis rate was determined as the percentage of cells positive for Annexin V.

The gene editing was carried out as previously described (Ran et al., 2013). Rat AKT1 was targeted using sgRNA sequences rAKT1-gRNA-F1: CAC​CGA​GGT​GCC​ATC​ATT​CTT​GAG​G and rAKT1-gRNA-R1: AAA​CCC​TCA​AGA​ATG​ATG​GCA​CCT​C. The sgRNA was cloned into PX459 V2.0 and transfected into PC12 cells using lipofectamine 3000. Cells were screened with puromycin, and AKT1 expression was verified by Western blot.

Protein lysates were prepared from the mouse hippocampus or PC12 cells using RIPA buffer with protease and phosphatase inhibitors. Protein concentrations were determined using the BCA assay. Samples (20 μg) were separated via 4%–20% SDS-PAGE, transferred to PVDF membranes, blocked with 5% BSA, and incubated overnight with primary antibodies (1:500–2000) at 4 °C. Membranes were incubated with HRP-conjugated secondary antibodies (1:4000) for 2 h, and bands were detected using ECL. Protein expression was normalized to GAPDH. The intensity of the bands was semi-quantified by using ImageJ software.

Male C57BL/6 mice (n = 48, 6–8 weeks, 18–21 g) were housed under standard conditions (25 °C, 12 h light/dark cycle) with food and water ad libitum. All procedures followed the University of Macau Animal Ethics Committee guidelines. Mice were divided into six groups: Control, CUMS, artemisinin (0.3, 1 and 3 mg/kg), and fluoxetine (10 mg/kg) which is a selective serotonin reuptake inhibitor (SSRI) and well-established antidepressant used as a positive control in this study (Dulawa et al., 2004; Levy et al., 2019). Drugs were administered via intraperitoneal injection 30 min after stress daily for 4 weeks. Mice were sacrificed post-behavioral tests.

The CUMS procedures were performed according to previous reports (Xu et al., 2020; Zhao Y.-N. et al., 2020; Zhu et al., 2020), with some modifications. CUMS was applied for 5 weeks with random exposure to stressors, with each stressors applied 3–4 times per week: (1) overnight flash illumination, (2) 8-h ultrasonic noise, (3) 12-h wet bedding, (4) 12-h water deprivation, (5) 24-h food deprivation, (6) 4-h cage tilt, (7) 1-min tail nip, (8) 5-min cold swim (4 °C), (9) 2-h physical restraint, (10) 8-h exposure to a pungent odor, (11) overnight light exposure. After 5 weeks of CUMS exposure, mice were subjected to different behavioral tests such as sucrose preference test (SPT), Tail suspension test (TST) and forced swim test (FST).

SPT: The test was carried out based on previously established methods, with a few alterations (Liu et al., 2015). Briefly, mice were trained to consume 1% sucrose solution, followed by 12 h water deprivation. Sucrose preference was calculated as (sucrose intake/total liquid intake) × 100%. TST: This test was performed based on the previous description (Steru et al., 1985), with slight modification. Briefly, mice were suspended by the tail (60 cm above ground) for 5 min. Immobility duration was recorded. FST: The test was conducted according to the method of Porsolt, with some modifications (Porsolt et al., 1977). Briefly, mice were placed in water-filled cylinders (25 °C ± 2 °C) for 5 min. Immobility was recorded during the last 4 min.

Mice were perfused with PBS, and brains were fixed in paraformaldehyde. Sections (5 μm) were dewaxed, rehydrated, blocked, and incubated with primary antibodies (1:200) overnight at 4 °C. For DAB staining, sections were treated with secondary antibodies, followed by color development. For immunofluorescence, sections were incubated with Alexa Fluor 488-conjugated secondary antibodies (1:500) and counterstained with DAPI. Images were acquired using an EVOS FL Imaging System or Carl Zeiss Axio Observer.

In vitro experiments were performed in at least three independent replicates. In vivo data are based on a sample size of 8 animals per group. The data analysis was conducted by GraphPad Prism 8.0 statistical software (GraphPad software, Inc., SanDiego, CA, USA) and presented as mean ± SD. Statistical significance was determined using unpaired t-test for comparisons between two groups, and one-way ANOVA followed by post hoc Tukey’s test for multiple group comparisons. Statistical significance was defined as p < 0.05, P < 0.01 or P < 0.001.

We first examined whether artemisinin could protect PC12 cells from CORT-induced cell toxicity. Figure 1A displays the chemical structure of artemisinin. The results of MTT assay showed that the viability of PC12 cells exposed to 200 μM of CORT for 48 h was significant decreased by 30% (Figure 1B), thus this concentration was chosen for further study. We further co-treated PC12 cells with 200 μM of CORT and different concentrations of artemisinin. The MTT results showed that artemisinin dose-dependently attenuated the loss of cell viability caused by CORT. Specifically, 25 μM of artemisinin restored cell viability to over 95% of that in the control group (Figure 1C). Furthermore, the effect of artemisinin on LDH release in CORT-treated PC12 cells was examined. The results indicated that treating PC12 cells with 200 μM corticosterone for 48 h induced a significant increase in LDH leakage compared to the control group (P < 0.01). Conversely, the addition of artemisinin significantly reduced corticosterone-induced LDH release, as shown in Figure 1D.

As shown in Figure 2A, exposure of PC12 cells to 200 μM CORT for 48 h resulted in a significant increase in green fluorescent signals, indicating that CORT induced oxidative stress. In contrast, co-treatment with artemisinin (25 μM) for 48 h resulted in a reduction of green fluorescent signals. As shown in Figure 2C, quantitative analysis demonstrates that the intracellular ROS level in CORT-treated PC12 cells significantly increased to 179.5% compared to the control value (100%, P < 0.001). However, co-treatment with 25 μM artemisinin significantly reduced that value to 137% (P < 0.01).

Mitochondria generate membrane potential through the activity of enzymes in the electron transport chain. During apoptosis, the collapse of MMP coincides with the opening of mitochondrial permeability transition pores, leading to the release of cytochrome C into the cytoplasm and triggering downstream events in the apoptotic cascade. To study whether CORT-induced apoptosis is associated with the loss of MMP, we performed JC-1 assay and found that MMP was indeed significantly decreased in PC12 cells after 48 h of treatment with 200 μM CORT, whereas artemisinin reversed this effect (Figures 2B,D). These findings suggest that artemisinin may have a positive impact on mitochondrial function.

Annexin V-FITC is a fluorescent probe that binds to phosphatidylserine in the presence of calcium. As shown in Figure 3, treatment with 200 μM CORT for 48 h significantly increased the percentage of Annexin V+/PI + cells, indicating a substantial increase in the apoptosis rate of PC12 cells following CORT-induced damage. However, treatment with 25 μM artemisinin significantly reversed this effect, suggesting that artemisinin can mitigate corticosterone-induced apoptosis in PC12 cells.

The AKT and ERK signaling pathways have been shown to play crucial roles in promoting cell survival and combating apoptosis. To determine whether artemisinin exerts its anti-apoptotic effects by modulating the AKT and ERK pathways, we treated PC12 cells with varying concentrations of artemisinin for different durations and then extracted proteins to analyze the expression levels of related proteins. We found that artemisinin upregulated the expression of phosphorylated AKT and ERK proteins in a time-dependent manner (Figure 4). Additionally, artemisinin also increased the expression of BDNF and phosphorylated GSK in a time-dependent manner. To further confirm whether the PI3K/AKT and ERK signaling pathways mediate the neuroprotective effect of artemisinin, PC12 cells were pre-treated with the specific PI3K inhibitor LY294002 (25 μM) and the MEK inhibitor PD98059 (25 μM) for 30 min. Subsequently, the cells were exposed to 200 μM CORT in the presence or absence of artemisinin, and cell viability was assessed using the MTT assay. The results (Figure 4B) showed that both inhibitors significantly blocked the cytoprotective effect of artemisinin. Moreover, knocking down AKT1 (Figure 4C) in PC12 cells also declined the protective effect of artemisinin against CORT (Figure 4D). These results further confirm the involvement of the AKT and ERK pathways in the neuroprotective action of artemisinin.

To investigate if the neuroprotective effect of artemisinin against CORT-induced toxicity is not only limit to PC12 cells line, the neuroprotective effect of artemisinin on primary hippocampal neurons was also examined. The viability of primary cultured neurons was significantly reduced in a dose-dependent manner following treatment with CORT concentrations of 50, 100, and 200 μM (Figure 5A). As shown in Figure 5B, artemisinin (25 μM) was also able to protect hippocampal neurons from the deleterious effects of CORT.

In this study, we evaluated the antidepressant-like activity of artemisinin in the CUMS mice model and used fluoxetine as a reference positive drug. As shown in Figures 6A,B, the immobility duration significantly increased in the CUMS-induced depressive mice in the tail suspension and the forced swimming test. While treatment with artemisinin or fluoxetine can all significantly decreased the immobility duration in both tail suspension and forced swimming tests compared to the CUMS group, and there was no significant difference in the immobility duration between the artemisinin group and the fluoxetine group in the two tests. Similarly, we also assessed depression-like behavior by sucrose preference test in CUMS mice. As shown in Figure. 6C, 5-week CUMS exposure significantly decreased the percentage of sucrose consumption in the CUMS mice as compared to control group; 1 mg/kg dose artemisinin administration significantly and dose-dependently reversed the decrease in sucrose preference in the stressed mice. However, the doses of 0.3 mg/kg and 3 mg/kg artemisinin did not show significant difference in sucrose consumption after 4 weeks of treatment.

To examine whether Artemisinin confers anti-depression effect via AKT/GSK/NRF2/HO1 and BDNF/TrkB/ERK/CREB pathway, we performed Western blot analysis to check the levels of phosphorylation of those proteins (Figure 7A). The results indicated that artemisinin treatment increased the level of AKT, GSK, ERK and CREB phosphorylation, as well as stimulated the expression of NRF2, HO1 and BDNF in brain extracts of CUMS mice (Figure 7B). Quantification results of the protein bands from the western blots are presented in Figures 7C–J.

CUMS induces pathological alterations in hippocampal astrocytes and neurogenesis, both hallmarks of depression. To evaluate artemisinin’s cellular effects in this context, we analyzed astrocyte activity and neuronal maturation in the CA1 region, with fluoxetine serving as a positive control. Immunohistochemical staining for GFAP, a marker of astrocyte activation, revealed a significant increase in GFAP + cells in CUMS mice compared to controls, reflecting stress-induced astrocytic hyperactivity. Artemisinin treatment markedly reduced GFAP levels, similar to the effect of fluoxetine, suggesting its potential to normalize astrocyte activity in the depressed hippocampus (Figure 8A). Concurrently, we assessed neurogenesis by quantifying NeuN + mature neurons, which are critical for functional hippocampal circuitry. CUMS exposure caused a severe loss of NeuN + neurons, consistent with impaired neurogenesis in depression. Artemisinin administration significantly restored NeuN⁺ cell numbers, indicating its capacity to counteract stress-induced neuronal deficits (Figure 8B). These results collectively highlight artemisinin’s dual role in ameliorating depression-associated glial dysregulation and neurogenesis impairment.