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. 2025 Jul 11;169(7):e70146. doi: 10.1111/jnc.70146

Astrocytes Contribute to Motor Neuron Degeneration in ALS via the TRAIL‐DR5 Signaling Pathway

Kangqin Yang 1, Yang Liu 1, Wenhua Deng 1, Zhenxiang Gong 1, Lifang Huang 1, Zehui Li 1, Min Zhang 2,3,✉
PMCID: PMC12246772  PMID: 40641248

ABSTRACT

Amyotrophic lateral sclerosis (ALS) is a neurodegenerative disorder characterized by the degeneration of both upper and lower motor neurons. The mechanisms underlying the selective degeneration of motor neurons in ALS remain poorly understood, underscoring the need for further investigation into the factors driving this process. In this study, we utilized ALS mouse models and an in vitro NSC34 motor neuron cell line expressing the SOD1G93A mutation to identify a novel pathogenic mechanism wherein astrocyte‐secreted Tumor necrosis factor‐related apoptosis‐inducing ligand (TRAIL) binds to Death Receptor 5 (DR5)on motor neurons, leading to caspase‐8 activation and subsequent neuronal death. Blocking DR5 with neutralizing antibodies significantly attenuated TRAIL‐induced motor neuron death. These findings provide the first evidence that TRAIL may serve as a potential therapeutic target in ALS, offering new insights into the mechanisms of motor neuron degeneration in this disease.

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Keywords: amyotrophic lateral sclerosis, apoptosis, astrocytes, DR5, motor neurons, TRAIL

Schematic diagram of astrocytes inducing motor neuron damage through the TRAIL‐DR5 pathway. In ALS, astrocytes induce motor neuron death through the secretion of TRAIL. TRAIL binds to death receptor 5 (DR5) on motor neurons, leading to the upregulation of genes associated with oxidative stress, apoptosis, pyroptosis, and necroptosis. This molecular cascade ultimately facilitates motor neuron death.

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Abbreviations

ALS

amyotrophic lateral sclerosis

CCK8

Cell Counting Kit‐8

CNS

central nervous system

DMEM

Dulbecco's modified Eagle medium

DR5

death receptor 5

EAE

experimental autoimmune encephalomyelitis

FBS

fetal bovine serum

IF

immunofluorescence

LDH

lactate dehydrogenase

NC

negative control

PCD

programmed cell death

qRT‐PCR

quantitative real‐time polymerase chain reaction

RRID

Research Resource Identifier

SOD1

Cu, Zn‐superoxide dismutase‐1

Tg

transgenic mice

TRAIL

tumor necrosis factor‐related apoptosis‐inducing ligand

WB

western blot

WT

wild‐type

1. Introduction

Amyotrophic lateral sclerosis (ALS) is a progressive neurodegenerative disorder characterized by the simultaneous degeneration of both upper and lower motor neurons. The loss of motor neurons leads to progressive muscle atrophy, weakness, and paralysis, with death typically occurring 3–5 years after disease onset due to respiratory failure, dysphagia, and pulmonary infections (Kiernan et al. 2011; Mejzini et al. 2019). Current treatment options are limited and largely ineffective, which may be attributed to the unclear etiology and pathogenesis of ALS (Aschenbrenner 2023; Brooks et al. 2022; Fels et al. 2022; Jaiswal 2019; Miller et al. 2020; Paganoni et al. 2020). Our understanding of the underlying mechanisms has been primarily derived from studies on transgenic mice overexpressing a toxic mutant variant of SOD1 (Cu, Zn‐superoxide dismutase‐1), responsible for ALS in a subset of familial cases. Mutant SOD1 induces neurodegeneration in motor neurons through both cell‐autonomous processes and non‐cell‐autonomous mechanisms mediated by microglia and astrocytes. Numerous studies have demonstrated that reactive astrocytes contribute to ALS pathology by reducing trophic support to neurons, influencing microglial activation, and secreting neurotoxic substances (Guttenplan et al. 2020; Haidet‐Phillips et al. 2011; Marchetto et al. 2008; Taha et al. 2022; Yang et al. 2024). However, the specific mechanisms by which astrocytes mediate selective motor neuron degeneration remain incompletely understood—particularly their involvement in triggering motor neuron death through programmed cell death pathways (PCD).

PCD, including apoptotic and non‐apoptotic pathways, has been implicated in the pathogenesis of ALS (Thal et al. 2024). Apoptosis, particularly caspase‐dependent apoptotic pathways, is well‐studied and shown to be activated in degenerating motor neurons (Wang et al. 2022). Recent research has also highlighted the potential involvement of non‐apoptotic PCD forms like ferroptosis and pyroptosis in ALS (Thal et al. 2024; Wang et al. 2022; Neel et al. 2022; Van Schoor et al. 2022). However, the contribution of astrocyte‐derived signals in these processes remains elusive, underscoring the necessity for further investigation.

Tumor necrosis factor‐related apoptosis‐inducing ligand (TRAIL), also known as Apo2L/TNFSF10, is a new member of the tumor necrosis factor superfamily that plays a critical role in regulating apoptosis, immune responses, and inflammation (Burgaletto et al. 2020; Gao et al. 2020). It exists in both membrane‐bound protein (mTRAIL) and soluble forms (sTRAIL). It is expressed on the surface or released by various immune effector cells, including natural killer cells, T cells, dendritic cells, macrophages, monocytes, and neutrophils. In pathological conditions such as ischemia, trauma, and neurodegeneration, TRAIL expression has been observed in the central nervous system (CNS), particularly in astrocytes, microglia, and neurons (Huang, Erdmann, Peng, et al. 2005). TRAIL binds to its death receptor DR5, initiating apoptotic and non‐apoptotic signaling pathways (Lv et al. 2024; Guerrache and Micheau 2024). Due to its selective cytotoxicity toward tumor cells while sparing normal cells, TRAIL has emerged as a promising candidate for cancer therapy (Maji et al. 2024). However, as research into TRAIL and its pathways advances, its involvement in CNS diseases has gathered increasing attention. Abnormal TRAIL protein expression has been detected in the blood and cerebrospinal fluid of patients with central nervous system disorders such as multiple sclerosis, Alzheimer's disease, and ischemic stroke, correlating well with clinical symptoms (Wandinger et al. 2003; Kouchaki et al. 2022; Wu et al. 2015; Uberti et al. 2004; Kang et al. 2015; Pan et al. 2015). These findings suggest that TRAIL may be involved in the pathophysiological processes of neurological diseases. Blocking the TRAIL‐DR5 pathway in animal models of these diseases has significantly improved clinical outcomes (Di Benedetto et al. 2022; Burgaletto et al. 2021; Cantarella et al. 2015; Chyuan et al. 2018; Cui et al. 2010). Additionally, abnormal TRAIL expression has been observed in the blood and cerebrospinal fluid of ALS patients, negatively correlating with survival time (Iłzecka 2008; Olesen et al. 2020). However, the precise mechanism by which TRAIL contributes to ALS remains unclear. In this study, we employed transgenic mice and cellular models to investigate the role of the TRAIL‐DR5 pathway in the pathogenesis of ALS.

2. Methods and Materials

2.1. Mice

Thirty female SOD1G93A transgenic mice (B6‐Cg‐Tg [SOD1*G93A] 1Gur/J; stock number 004435) were used as experimental subjects. Mice were divided into three groups based on disease progression: pre‐symptomatic (10 weeks old, n = 10), during which mice exhibited no detectable motor deficits and appeared healthy; symptomatic (14 weeks old, n = 10), characterized by the development of progressive motor symptoms such as reduced performance in the rotarod test, decreased grip strength, or hindlimb tremors; and end‐stage (17 weeks old, n = 10), marked by severe motor impairment, an inability to right themselves, or significant weight loss, and euthanasia conducted in accordance with ethical guidelines (Magnus et al. 2008; Xu et al. 2021). A priori sample size calculation was not performed; instead, the sample size was determined based on previous similar studies (Wang et al. 2022; Magnus et al. 2008; Cheng et al. 2023). Age‐ and sex‐matched littermates served as WT controls. The transgenic mice were originally sourced from Jackson Laboratory (USA) and bred at Huachuang Xinnuo Co. Ltd., Jiangsu, China. SOD1G93A male mice were crossed with WT female mice to generate heterozygous offspring, which were consistently maintained on a C57BL/6 background. All mice were housed in an SPF facility at the Experimental Animal Center of Tongji Medical College, Huazhong University of Science and Technology, under controlled conditions with unrestricted access to food and water. Mice were housed in groups of no more than 5 per cage with a 12‐h light/dark cycle. This study was approved by the Institutional Animal Care and Use Committee, Huazhong University of Science and Technology (IACUC Number: 4311). To minimize animal suffering, especially when hind limb weakness was pronounced, food and water were placed at the bottom of the cage, as the mice had difficulty reaching the food and water placed on the cage cover. Additionally, the bedding in the cages was kept dry and clean. Mice were regularly monitored for changes in body weight and neurological function twice a week as previously described (Garofalo et al. 2020; Hatzipetros et al. 2015). If the mice showed a weight loss of more than 20% or experienced difficulty in turning over, euthanasia was performed following humane guidelines (Wang et al. 2022). Specifically, euthanasia was carried out by intraperitoneal injection of sodium pentobarbital at a dose of 150 mg/kg, which induces rapid and painless death through central nervous system depression. No pre‐determined exclusion criteria were set. During the course of the experiment, two mice in the end‐stage group died unexpectedly due to disease progression before tissue collection and were therefore excluded from the final analysis. No animals were replaced. Due to the inherent nature of the genetic model, complete blinding was not feasible. However, to minimize bias, we used automated data acquisition systems, blinded histopathological analysis, and predefined statistical thresholds.

2.2. Spinal Cord Quantitative Real‐Time PCR (qRT‐PCR)

Symptomatic SOD1G93A mice (120 days) were anesthetized with sodium pentobarbital at a concentration of 50 mg/kg via intraperitoneal (i.p.) injection, and their hearts were perfused with pre‐chilled PBS. The lumbar spinal cords were dissected, snap‐frozen in isopentane, and stored at −80°C. Total RNA extraction was performed using TRIzol reagent (Invitrogen, cat. no. 15596026), followed by cDNA synthesized using the Takara PrimeScript RT Reagent Kit (Takara, cat. no. RR036A). qRT‐PCR was performed on a Roche LightCycler 480 system (384‐well format) using SYBR Green PCR Master Mix (Yeasen, cat. no. 11201ES08), with actin as the internal control. Gene expression levels were quantified using the 2−ΔΔCT method, and the primer sequences are provided in Table 1.

TABLE 1.

Primer sequences used in the qPCR experiments of this study.

Primer Forward Reverse
Actin GGCTGTATTCCCCTCCATCG CCAGTTGGTAACAATGCCATGT
TRAIL ATGGTGATTTGCATAGTGCTCC GCAAGCAGGGTCTGTTCAAGA
hSOD1 GGTGGGCCAAAGGATGAAGAG CCACAAGCCAAACGACTTCC
caspase3 CTGACTGGAAAGCCGAAACTC CGACCCGTCCTTTGAATTTCT
caspase6 GGAAGTGTTCGATCCAGCCG GGAGGGTCAGGTGCCAAAAG
caspase7 GGACCGAGTGCCCACTTATC TCGCTTTGTCGAAGTTCTTGTT
caspase8 TGCTTGGACTACATCCCACAC GTTGCAGTCTAGGAAGTTGACC
caspase9 TCCTGGTACATCGAGACCTTG AAGTCCCTTTCGCAGAAACAG
Bcl2 ATGCCTTTGTGGAACTATATGGC GGTATGCACCCAGAGTGATGC
bax TGAAGACAGGGGCCTTTTTG AATTCGCCGGAGACACTCG
caspase1 ACAAGGCACGGGACCTATG TCCCAGTCAGTCCTGGAAATG
caspase11 AGCGTTGGGTTTTTGTAGATGC CCTTGTGAACTCTTCAGGGGA
NLRP3 ATTACCCGCCCGAGAAAGG TCGCAGCAAAGATCCACACAG
gsdmd CCATCGGCCTTTGAGAAAGTG ACACATGAATAACGGGGTTTCC
IL1β GAAATGCCACCTTTTGACAGTG TGGATGCTCTCATCAGGACAG
IL18 GACTCTTGCGTCAACTTCAAGG CAGGCTGTCTTTTGTCAACGA
MLKL AATTGTACTCTGGGAAATTGCCA TCTCCAAGATTCCGTCCACAG
RIPK1 GAAGACAGACCTAGACAGCGG CCAGTAGCTTCACCACTCGAC
RIPK3 TCTGTCAAGTTATGGCCTACTGG GGAACACGACTCCGAACCC
CHOP CTCGCTCTCCAGATTCCAGTC CTTCATGCGTTGCTTCCCA
iNOS GTTCTCAGCCCAACAATACAAGA GTGGACGGGTCGATGTCAC
GPX1 AGTCCACCGTGTATGCCTTCT GAGACGCGACATTCTCAATGA
Nrf2 TCTTGGAGTAAGTCGAGAAGTGT GTTGAAACTGAGCGAAAAAGGC
COX2 TTCAACACACTCTATCACTGGC AGAAGCGTTTGCGGTACTCAT
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2.3. Immunofluorescent Staining (IF)

SOD1G93A mice were anesthetized with sodium pentobarbital (i.p.,50 mg/kg). Subsequently, transcardiac perfusion was performed using pre‐chilled PBS and 4% paraformaldehyde to fix the spinal cord. The lumbar spinal cord was dissected and fixed overnight in 4% paraformaldehyde at 4°C. Dehydration was achieved using sucrose solutions (20% for 1 day, 30% for 1 day). The tissue was then embedded in OCT (Sakura, cat. no. 4583), frozen, and sectioned into 15 μm coronal slices using a Leica microtome. Sections were stored at −80°C until further use. Before staining, sections were thawed at 37°C for 15 min, washed with PBS, and permeabilized with 0.3% Triton X‐100 for 15 min. Non‐specific binding was minimized by incubating the sections with a blocking solution at room temperature for 15 min. Primary antibodies against TRAIL (1:100, RRID: AB_10855951, Bioss), DR5 (1:100, RRID: AB_10859874, Bioss), GFAP (1:400, RRID: AB_561049, CST), Iba‐1 (1:200, RRID: AB_839504, Abcam), Olig2 (1:200, RRID: AB_2157554, R&D), NeuN (1:500, RRID: AB_11205592, Millipore), and ChAT (1:400, RRID: AB_2079751, Millipore) were applied and incubated overnight at 4°C. Following PBS washes, the sections were incubated with fluorophore‐conjugated secondary antibodies (Donkey anti‐mouse Alexa Fluor 594, 1:200; Cat# 34112ES60, Yeasen Biotech; Donkey anti‐rabbit Alexa Fluor 488, 1:200; RRID: AB_2909605, Yeasen Biotech; Donkey anti‐goat Alexa Fluor 488, 1:200; Cat# 34306ES60, Yeasen Biotech; Donkey anti‐guinea pig Alexa Fluor 488, 1:200; Cat# 34506ES60, Yeasen Biotech) in the dark at room temperature for 1 h. Finally, sections were counterstained with DAPI and mounted for observation. Confocal microscopy was used to examine and scan the sections, focusing on the anterior horn region of the spinal cord at 20× magnification. For quantification, the number of TRAIL‐ or DR5‐positive cells was counted throughout the entire anterior horn of the lumbar spinal cord in each section. At least 3 sections per mouse and 3 mice per group were analyzed. Imaging processing was performed using ImageJ software, an open‐source image analysis software (version 1.53t, NIH, Bethesda, MD, USA; https://imagej.nih.gov/ij/).

2.4. Cell Culture and Transfection

In this experiment, the NSC34 cell line (kindly provided by Professor Cashman at the Djavad Mowafaghian Centre for Brain Health, University of British Columbia, Vancouver, Canada), commonly utilized in ALS research, was employed (Cashman et al. 1992; Guo et al. 2024; Shvil et al. 2018). The NSC34 cell line is not listed as a commonly misidentified cell line by the International Cell Line Authentication Committee (ICLAC; http://iclac.org/databases/cross‐contaminations/) and was not further authenticated upon receipt. We used NSC34 cells within a passage range of 5–15 after thawing. The culture medium comprised 10% fetal bovine serum (FBS), 90% Dulbecco's modified Eagle medium (DMEM), and 1% penicillin–streptomycin (Cashman et al. 1992; Guo et al. 2024). All cell culture reagents were sourced from Gibco. Cells were maintained in a humidified incubator at 37°C with 5% CO2. Upon reaching approximately 80% confluence, cells were passaged at 1:2–1:4, contingent on their growth rate. Transfection was carried out using the plasmid pCDNA3.1‐CMV‐H_SOD1(p. G93A)‐T2A‐ZsGreen1‐P2A‐Puro (encoding the human mutant SOD1 protein) and the empty plasmid pcDNA3.1‐CMV‐T2A‐ZsGreen1‐P2A‐Puro (used as a negative control, NC), both obtained from Jima Biotechnology. The transfection was performed using Lipofectamine 2000 reagent (Thermo Fisher, cat. no. 11668019) according to the manufacturer's protocol (Huo et al. 2024; Richardson et al. 2013). After 6–8 h, the medium was exchanged for culture medium supplemented with 10% FBS, and the cells were further cultured. The transfected cells constituted an in vitro ALS model, widely recognized for investigating ALS mechanisms. Notably, for all in vitro experiments in this study, each condition was performed in three or more separate wells treated at the same time (technical replicates). Key experiments were independently repeated using different cell passages and on different days to confirm reproducibility.

2.5. TRAIL and Anti‐DR5 Treatment

To assess the effect of TRAIL, NSC34 cells transfected with the human Sod1 G93A gene for 48 h were divided into two groups. The control group did not receive TRAIL treatment. In contrast, the experimental group was exposed to soluble TRAIL (MCE, cat. no. HY‐P7089A) dissolved in a complete medium at concentrations of 1, 5, and 25 ng/mL for incubation periods of 6, 12, or 24 h. To evaluate the impact of the DR5 antagonist, the model cells were allocated into two groups. One group was treated with 25 ng/mL TRAIL for 24 h, whereas the other group was pre‐incubated with 1 μg/mL anti‐DR5 neutralizing antibody (R and D Systems, RRID: AB_2205069) for 4 h before co‐incubation with TRAIL and the antibody for an additional 24 h (Brunetti et al. 2013; Liang et al. 2017; Wu et al. 2008).

2.6. Cell qRT‐PCR

After transfecting and treating the NSC34 cells, total RNA was extracted using a TRIzol reagent. The subsequent procedures for cDNA synthesis and qRT‐PCR followed the protocol described for the spinal cord.

2.7. Western Blot

Cells were lysed using RIPA buffer (BOSTER, cat. no. AR0102) supplemented with PMSF (BOSTER, cat. no. AR1192) and phosphatase inhibitors (MCE, cat. no. HY‐L081). Protein concentrations were measured using the BCA Protein Assay Kit (BOSTER, cat. no. AR0146). After quantification, proteins were mixed with 5× loading buffer and denatured by boiling at 100°C for 10 min. Samples not analyzed immediately were stored at −80°C. Equal amounts of protein (20 μg) were separated by 12% SDS‐PAGE (Vazyme, cat. no. E304‐01) and transferred to a 0.22 μm nitrocellulose membrane (BOSTER, cat. no. AR0135‐02) at 220 mA for 60 min on ice. The membrane was then incubated with a quick blocking buffer containing BSA (Yamei, cat. no. PS108P) at room temperature for 20 min to minimize nonspecific binding. Subsequently, the membrane was incubated overnight at 4°C with primary antibodies: rabbit anti‐SOD1 (Abcam, RRID: AB_2193891, 1:3000), rabbit anti‐DR5 (Abclone, RRID: AB 2862536, 1:1000), and rabbit anti‐β‐actin (Abclone, RRID: AB_2768234, 1:10000). Following a 30‐min wash with TBST buffer, the membrane was incubated with HRP‐conjugated secondary antibodies at room temperature for 1 h, including HRP Conjugated AffiniPure Goat Anti‐rabbit IgG (H+L) (BOSTER, cat. no. BA1054, 1:5000). Protein signals were detected using ECL chemiluminescent reagents (Abbikine, cat. no. BMU102) and visualized with a CCD camera (BLT, GelView 6000pro). The intensity of the target bands t was quantified using ImageJ software.

2.8. Flow Cytometry

After transfection and treatment, cell apoptosis was assessed using the Annexin V‐APC/PI Apoptosis Detection Kit (Vazyme, cat. no. A214‐01/02) following the manufacturer's instructions. Detached and trypsinized adherent cells were harvested and resuspended in 1× Binding Buffer. The cells were then stained with Annexin V‐APC and PI d, followed by a 10‐min incubation at room temperature in the dark. Post‐incubation, the samples were diluted with 1× Binding Buffer and analyzed by flow cytometry using a NeonSYS flow cytometer. Both a negative control (untreated cells) and a positive control (cells exposed to 56°C heat for 10 min) were included to validate reagent efficacy and result specificity. A blank control (unstained cells) was also incorporated to account for autofluorescence.

The APC channel (660 nm) was used to detect Annexin V signals, while the PE channel (585 nm) was employed to capture PI signals. Cells were categorized into four groups based on their staining patterns: Annexin V−/PI−, indicating cells that are neither apoptotic nor necrotic; Annexin V+/PI−, identifying early apoptotic cells characterized by phosphatidylserine externalization without membrane permeability; Annexin V+/PI+, indicating late apoptotic or necrotic cells, which suggests advanced apoptosis or necrosis with compromised membrane integrity; and Annexin V−/PI+, representing necrotic cells that have lost membrane integrity without phosphatidylserine exposure.

2.9. Cell Viability Assay (CCK‐8)

A Cell Counting Kit‐8 (CCK‐8, MCE, cat. no. HY‐K0301) assay was performed to assess cell viability and proliferation according to the manufacturer's instructions. Following treatment (e.g., transfection, TRAIL, or anti‐DR5 intervention), 10 μL of CCK8 reagent was added into each well. During incubation, viable cells metabolized the tetrazolium salt in CCK‐8 into an orange formazan product. Absorbance was measured at 450 nm using a microplate reader (Bio‐Rad). A blank control consisting of media and CCK‐8 without cells was included to account for background interference.

2.10. Lactate Dehydrogenase (LDH) Release Assay

The LDH release assay, performed using the LDH Cytotoxicity Assay Kit (MCE, cat. no. HY‐K1090), was employed to evaluate cell damage and cytotoxicity. Lactate dehydrogenase (LDH) is an intracellular enzyme released into the culture medium upon cell membrane disruption. By quantifying the LDH levels in the medium, we can indirectly assess the extent of cell damage and cytotoxicity. Following the kit's protocol, 50 μL of the cell culture supernatant from each well of the 96‐well plate was removed, and 50 μL of the LDH detection reagent was added to each well. The plate was incubated at room temperature in the dark for 30 min. After the incubation, 50 μL of stop solution was added to each well. Absorbance was measured at 490 nm using a microplate reader. Four control groups were established as per the kit's instructions: a blank control (medium only, no cells) to account for background absorbance; a high blank control (medium plus 10 μL of Lysis Solution) to correct for non‐specific absorbance in the high control; a low control (cells in medium, no lysis treatment) to measure spontaneous LDH release from untreated cells; and a high control (cells in medium, with 10 μL of Lysis Solution) to determine the maximum possible LDH release from lysed cells.

2.11. Statistical Analysis

Data are presented as mean ± SEM. Normality was not assessed prior to analysis due to the small sample size. No formal test for outliers was conducted. For pairwise comparisons, a two‐tailed Student's t‐test was employed; for multiple groups comparisons, one‐way ANOVA with Tukey's post hoc test was applied. Statistical significance was set at p < 0.05. Statistical analysis was conducted using GraphPad Prism 9.5, and visualizations were created using Adobe Illustrator.

3. Results

3.1. Increased Expression of TRAIL and DR5 in Astrocytes of the Lumbar Spinal Cord in SOD1G93A Mice

To investigate the involvement of the TRAIL‐DR5 pathway in ALS, we first assessed the temporal dynamics of TRAIL expression levels in SOD1G93A mice. Longitudinal analyses revealed a progressive increase in TRAIL protein levels from the pre‐symptomatic to advanced disease stages. Quantitative measurements at multiple time points (pre‐symptomatic, onset, and advanced stages) confirmed this gradual elevation. In contrast, TRAIL expression was almost undetectable in the anterior horn of the spinal cord in WT mice (Figure 1A,B). Notably, while TRAIL expression was absent in the ventral horn gray matter of WT mice, immunofluorescence analysis revealed that TRAIL was present at baseline levels in the white matter of both WT and SOD1G93A mice (Figure S1). Subsequently, we performed immunofluorescent double‐labeling experiments to investigate the distribution of TRAIL across various cell types throughout all disease phases. The results demonstrated that in the anterior horn of the lumbar spinal cord in SOD1G93A mice, TRAIL expression was predominantly localized in astrocytes, with minimal presence in oligodendrocytes, microglia, and neurons (Figure 1C,D). The phenotypic features of SOD1G93A mice were validated, as shown in Figure S2.

FIGURE 1.

FIGURE 1

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Increased Expression of TRAIL in Astrocytes of the Lumbar Spinal Cord in SOD1G93A mice. (A) Representative immunofluorescence images of TRAIL in the lumbar spinal cord anterior horn of WT and SOD1G93A mice at 10, 14, and 17 weeks of age (scale bar: 100 μm). Red color denotes TRAIL. (B) Statistical graph of the average fluorescence intensity of TRAIL in the lumbar spinal cord anterior horn of WT and SOD1G93A mice at 10, 14, and 17 weeks of age, n = 3 mice per group, mean ± SEM, one‐way ANOVA. (C) Representative immunofluorescence images of TRAIL co‐localized with GFAP (label astrocytes), Iba‐1 (label microglia), Olig2 (label oligodendrocytes), and NeuN (neurons) in the lumbar spinal cord anterior horn of SOD1G93A mice, with magnified images (scale bar for representative image: 100 μm; scale bar for magnified image: 50 μm). Red color denotes TRAIL. Green color denotes GFAP, Iba‐1, Olig2, and NeuN. (D) A bar chart showing the percentage of double‐positive cells. *p < 0.05.

Given the progressive increase in TRAIL expression, we next explored the distribution of its receptor, DR5, at the late stage when motor neuron loss is most pronounced. Unlike TRAIL, DR5 was expressed in astrocytes, microglia, oligodendrocytes, and neurons (including motor neurons) within the anterior horn of both WT and SOD1G93A mice (Figure 2A). Notably, in SOD1G93A mice, there was a significant increase in the number of DR5‐expressing astrocytes and microglia (Figure 2B,C). Conversely, DR5 expression was reduced in oligodendrocytes and neurons, including motor neurons (Figure 2D–F). Significantly, DR5 fluorescence intensity was elevated in motor neurons exhibiting morphological abnormalities rather than those morphologically intact (Figure 2G). To confirm the specificity of ChAT immunolabeling for motor neurons in the spinal cord, we performed additional immunostaining. As shown in Figure S3, ChAT‐positive cells were exclusively localized to the ventral horn, with no punctate staining detected in the dorsal horn. These findings further substantiate the reliability of ChAT as a specific marker for motor neurons.

FIGURE 2.

FIGURE 2

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Increased Expression of DR5 in the Lumbar Spinal Cord in SOD1G93A mice. (A) Representative immunofluorescence images of DR5 co‐localized with GFAP (label astrocytes), Iba‐1 (label microglia), Olig2 (label oligodendrocytes), NeuN (neurons), and ChAT (label Cholinergic neurons, mainly motor neurons) in the lumbar spinal cord anterior horn of WT and SOD1G93A mice (scale bar for representative image: 100 μm). The red color denotes DR5. The green color denotes GFAP, Iba‐1, Olig2, and NeuN. The gray color denotes ChAT. (B) Statistical graph of GFAP+DR5+/DR5+ cells in the lumbar spinal cord anterior horn of WT and SOD1G93A mice, n = 3 mice per group, mean ± SEM, unpaired t‐test. (C) Statistical graph of Iba‐1+DR5+/DR5+ cells in the lumbar spinal cord anterior horn of WT and SOD1G93A mice, n = 3 mice per group, mean ± SEM, unpaired t‐test. (D) Statistical graph of Olig2+DR5+/DR5+ cells in the lumbar spinal cord anterior horn of WT and SOD1G93A mice, n = 3 mice per group, mean ± SEM, unpaired t‐test. (E) Statistical graph of NeuN+DR5+/DR5+ cells in the lumbar spinal cord anterior horn of WT and SOD1G93A mice, n = 3 mice per group, mean ± SEM, unpaired t‐test. (F) Statistical graph of ChAT+DR5+/DR5+ cells in the lumbar spinal cord anterior horn of WT and SOD1G93A mice, n = 3 mice per group, mean ± SEM, unpaired t‐test. (G) Quantification of DR5 fluorescence intensity in morphologically intact (com) and morphologically compromised (incom) motor neurons in the anterior horn of the spinal cord in SOD1G93A mice. White arrows indicate double‐positive cells shown in the corresponding fluorescence images, and blue arrows indicate morphologically intact motor neurons in the anterior horn of SOD1G93A mice. ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001.

3.2. TRAIL Induces Neurotoxic Effects on NSC34‐G93A Cells

To further investigate the role of TRAIL, which is highly expressed in astrocytes, in ALS, we simulated the ALS pathological process by transfecting the mutant plasmid hSOD1‐G93A into the NSC34 motor neuron cell line (Figure 3A). First, we confirmed the transfection efficiency using PCR and Western blot (WB) analysis. The results indicated that, compared to cells transfected with the control plasmid NC, NSC34 cells transfected with hSOD1‐G93A exhibited a significant increase in both mRNA and protein expression levels of hSOD1 after 48 h (Figure 3B,C). In addition, the transfection efficiency was further validated by flow cytometry analysis (Figure S4). Subsequently, to evaluate whether the functional changes in NSC34 cells transfected with hSOD1 aligned with previous findings, we assessed cell viability and cytotoxicity using the CCK8 and LDH release assay, respectively. Compared to the NC group, cells transfected with hSOD1‐G93A exhibited significantly reduced viability and increased LDH release (Figure 3D,E), consistent with previously published data.

FIGURE 3.

FIGURE 3

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TRAIL treatment is detrimental to NSC34‐G93A cells. (A) Flowchart of the cell experiment. (B) hSOD1 mRNA expression levels in NSC34 cells transfected with hSOD1‐G93A plasmid or NC control plasmid for 48 h, n = 3 wells of cells per group. (C) Representative western blot image of hSOD1 protein expression in NSC34 cells transfected with hSOD1‐G93A plasmid or NC control plasmid for 48 h, n = 3 wells of cells per group. (D) CCK‐8 assay results of NSC34 cells transfected with hSOD1‐G93A plasmid or NC control plasmid for 48 h, n = 10 wells of cells per group, mean ± SEM, unpaired t‐test. (E) LDH release rate in NSC34 cells transfected with hSOD1‐G93A plasmid or NC control plasmid for 48 h, n = 8 wells of cells per group, mean ± SEM, unpaired t‐test. (F) Flowchart of the cell experiment. (G) Cell viability measured by the CCK‐8 assay in NSC34 cells transfected with hSOD1‐G93A plasmid for 48 h and treated with TRAIL at different concentrations (1, 5, and 25 ng/mL) and for various durations (6, 12, 24 h), n = 7–8 wells of cells per group, mean ± SEM, two‐way ANOVA. (H) LDH release rate measured in NSC34 cells transfected with hSOD1‐G93A plasmid for 48 h and treated with TRAIL at different concentrations (1, 5, and 25 ng/mL) and for various durations (6, 12, and 24 h), n = 5–6 wells of cells per group, mean ± SEM, two‐way ANOVA. (I) Representative flow cytometry images showing apoptotic and viable cells after NSC34 cells transfected with hSOD1‐G93A plasmid for 48 h were treated with TRAIL at 0 or 25 ng/mL for 24 h. (J) Statistical analysis of apoptotic cell percentages between the experimental and control groups, n = 8 wells of cells per group, mean ± SEM, unpaired t‐test. (K) Statistical analysis of early apoptotic cell percentages between the experimental and control groups, n = 8 wells of cells per group, mean ± SEM, unpaired t‐test. (L) Statistical analysis of late apoptotic cell percentages between the experimental and control groups, n = 8 wells of cells per group, mean ± SEM, unpaired t‐test. (M) Heatmap showing the expression levels of genes related to apoptosis, pyroptosis, necroptosis, and oxidative stress, as detected by qPCR. ns, not significant; *p < 0.05, **p < 0.01.

We subsequently investigated the impact of TRAIL on the survival of NSC34 motor neuron cells transfected with hSOD1‐G93A (Figure 3F). Our results demonstrated that escalating TRAIL concentrations led to a significant reduction in cell viability and a marked increase in LDH release, with the most pronounced cytotoxicity observed at a TRAIL concentration of 25 ng/mL (Figure 3G,H). Moreover, we verified that 25 ng/mL TRAIL exerted minimal effects on non‐transfected NSC34 cells, thereby underscoring the specificity of its cytotoxicity toward SOD1‐transfected cells (Figure S5). Prolonged exposure to TRAIL further decreased cell viability and significantly increased LDH release, with the most severe toxicity manifesting after 24 h of treatment (Figure 3G,H). These findings confirmed 24‐h exposure to 25 ng/mL TRAIL elicited the most substantial neurotoxic effect. To further validate this observation, we evaluated cell survival and apoptosis using flow cytometry. The data revealed that 24‐h treatment with 25 ng/mL TRAIL significantly elevated the apoptosis rate compared to untreated controls, particularly in early apoptosis, while late apoptosis remained relatively unchanged (Figure 3I–L). Our experiments indicate that TRAIL induces neurotoxic effects in an in vitro ALS cell model.

To elucidate the mechanisms by which TRAIL induces neurotoxicity in this in vitro ALS model, we analyzed the expression levels of genes associated with pathways potentially activated by TRAIL, as reported in the literature (Lv et al. 2024). Our results revealed that, compared to the control group, TRAIL‐treated cells exhibited significant upregulation of mRNA levels for apoptosis‐related genes, including Bcl2 and caspases 3,6,7,8. Genes linked to pyroptosis, such as Caspases 1, 11 and IL‐18, and those related to necroptosis, such as Mlkl, were also markedly elevated. Notably, the oxidative stress‐related gene Chop was significantly upregulated (Figure 3M). As further shown in Figure S6, TRAIL treatment failed to significantly modulate the expression of genes associated with apoptosis, necroptosis, pyroptosis, or oxidative stress‐related genes in NSC34 cells. This suggests that the neurotoxic effects of TRAIL are predominantly observed in the context of SOD1‐G93A overexpression. To determine whether SOD1‐G93A overexpression alone could activate similar pathways independently of TRAIL, we performed a complementary gene expression analysis. As presented in Figure S7, the expression level of genes linked to apoptosis, necroptosis, pyroptosis, and oxidative stress was evaluated 72 h post‐transfection in NSC34 cells. Among all the examined genes, only inducible nitric oxide synthase (iNOS) exhibited significant upregulation in the SOD1‐G93A group, indicative of a mild oxidative stress response, while no significant changes were detected for other genes. Collectively, these findings indicate that TRAIL exerts neurotoxic effects on NSC34‐G93A cells by upregulating genes involved in apoptosis, pyroptosis, necroptosis, and oxidative stress.

3.3. DR5 Neutralizing Antibody Significantly Inhibits TRAIL‐Mediated Motor Neuron Damage

To explore the role of DR5, the TRAIL effector receptor, in mediating the downstream effects of TRAIL, we first examined DR5 expression in NSC34 cells using Western blot analysis. The results showed that DR5 is expressed in NSC34 cells and its expression markedly increased following TRAIL treatment (Figure 4B,C), thereby enhancing the binding affinity between TRAIL and DR5. To further investigate the role of DR5 in TRAIL‐induced neurotoxicity, we pretreated the cell model with a DR5‐neutralizing antibody before TRAIL addition. After 24 h of co‐incubation with TRAIL and the neutralizing antibody, we assessed cell viability using CCK8, LDH release assays, and flow cytometry. The findings demonstrated that, compared to the control group, the DR5 neutralization group exhibited significantly higher cell viability, reduced LDH release, and decreased apoptosis, particularly early apoptosis (Figure 4E–J).

FIGURE 4.

FIGURE 4

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DR5 neutralizing antibody inhibits TRAIL‐induced cytotoxicity. (A, D) Flowchart of the cell experiment. (B) Representative western blot images of DR5 protein expression in different groups (NSC34‐NC, NSC34‐G93A, and NSC34‐G93A + TRAIL). (C) Statistical analysis of the ratio of DR5 protein expression relative to β‐actin protein expression in different groups, n = 3 wells of cells per group, mean ± SEM, one‐way ANOVA. (E) CCK‐8 assay results comparing the antagonist and control groups. Cell viability is expressed as a percentage relative to the untreated NSC34‐G93A control group, which was set as 100%. n = 8 wells of cells per group, mean ± SEM, unpaired t‐test. (F) LDH release rate comparison between the antagonist and control groups. LDH release is expressed as a percentage relative to the untreated NSC34‐G93A control group, which was set as 100%. n = 8 wells of cells per group, mean ± SEM, unpaired t‐test. (G) Representative flow cytometry images of the antagonist group and the control group. (H–J) Statistical analysis of flow cytometry results comparing the antagonist and control groups, n = 6 wells of cells per group, mean ± SEM, unpaired t‐test. (K) Heatmap showing the expression levels of genes related to oxidative stress, apoptosis, pyroptosis, and necroptosis as detected by qPCR. ns, not significant; *p < 0.05, **p < 0.01, ***p < 0.001.

In addition, fluorescence imaging of GFP‐positive cells showed improved cell morphology following DR5 antibody treatment (Figure S9). The neutralizing efficiency of the DR5 antibody was further validated, as shown in Figure S8. To further confirm the role of oxidative stress in vivo, we investigated Chop mRNA expression in the lumbar spinal cord of WT and SOD1‐G93A mice at various disease stages. Quantitative PCR analysis revealed that Chop expression was consistently elevated in SOD1‐G93A mice compared to WT controls at 10, 14, and 17 weeks of age, with the peak expression level observed at 17 weeks (Figure S10). These findings highlight the activation of oxidative stress pathways in both the ALS mouse model and the in vitro cell model.

Next, we performed qPCR to investigate alterations in gene expression following TRAIL treatment. The results demonstrated that the expression of TRAIL‐induced genes, including Bcl2, Caspase 6, 8, Caspase1, IL‐18, Gsdmd, Mlkl, and Chop, was significantly downregulated in the DR5 neutralization group. Moreover, the modest upregulation of IL‐1β, Nlrp3, Ripk1, Ripk3, Cox2, Nrf2, and Gpx1 observed in response to TRAIL was also significantly diminished in this group (Figure 4K). These data provide compelling evidence that DR5 is essential for mediating the neurotoxic effects of TRAIL. Neutralizing DR5 with a specific antibody mitigates the toxic effects induced by TRAIL.

4. Discussion

In this study, we observed a significant elevation in TRAIL protein expression in the lumbar spinal cord of SOD1G93A mice as the disease progressed. Specifically, TRAIL levels progressively increased from the pre‐symptomatic stage through the symptomatic to the end stage, with the highest expression observed in the latter stages. Importantly, TRAIL was predominantly expressed by astrocytes, while detectable expression in neurons, microglia, or oligodendrocytes was minimal or absent across all stages of disease progression. In vitro experiments revealed that TRAIL mediates the death of motor neurons transfected with the mutant plasmid hSOD1‐G93A and upregulates genes associated with oxidative stress, apoptosis, pyroptosis, and necroptosis (Figure 5). Furthermore, a DR5‐neutralizing antibody inhibited TRAIL‐induced downstream effects and alleviated motor neuron damage.

FIGURE 5.

FIGURE 5

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Schematic diagram of astrocytes inducing motor neuron damage through the TRAIL‐DR5 pathway. In ALS, astrocytes induce motor neuron death through the secretion of TRAIL. TRAIL binds to death receptor 5 (DR5) on motor neurons, leading to the upregulation of genes associated with oxidative stress, apoptosis, pyroptosis, and necroptosis. This molecular cascade ultimately facilitates motor neuron death. (by figdraw).

Our study utilized both SOD1‐G93A mouse and cell models to investigate the role of TRAIL in ALS. Although SOD1 mutations account for only a subset of familial ALS cases, these models were extensively utilized owing to their well‐documented neurodegenerative progression. Nevertheless, given the clinical and pathological heterogeneity of ALS, the insights gained from SOD1‐mutant models might not fully capture the disease mechanisms underlying sporadic ALS. In our study, we observed a progressive increase in TRAIL protein expression in the spinal cords of SOD1‐G93A mice as the disease advanced, with astrocytes identified as the predominant source. This observation raises the question of whether a similar upregulation of TRAIL occurs in sporadic ALS, where distinct pathological mechanisms such as TDP‐43 aggregation predominate. Future studies using patient‐derived motor neurons or alternative ALS models are necessary to determine whether TRAIL‐mediated neurotoxicity represents a common pathogenic feature of ALS or is specific to SOD1 mutations.

Previous studies have demonstrated that TRAIL is absent in the normal brain. However, both TRAIL and DR5 mRNA and protein expression levels are significantly elevated in the brain tissues of animal models of diseases such as Alzheimer's disease, multiple sclerosis, and ischemic stroke (Burgaletto et al. 2020; Gao et al. 2020), consistent with our findings in the ALS animal model. Notably, in our study, we found a progressive increase in TRAIL expression in the lumbar spinal cord of SOD1‐G93A mice as the disease progressed, with astrocytes identified as the predominant source. Interestingly, we also detected TRAIL expression in the white matter of healthy WT mice, a result consistent with the findings of Sanmarco et al., who demonstrated that astrocytes located in the meninges and white matter of the healthy CNS express TRAIL (Sanmarco et al. 2021).

Despite the elevated spinal cord expression observed in our model, two clinical studies have reported reduced TRAIL protein levels in both the plasma and cerebrospinal fluid (CSF) of ALS patients compared to healthy controls (Iłzecka 2008; Olesen et al. 2020). This apparent contradiction may be attributed to several factors. First, TRAIL exists in two biologically active forms—membrane‐bound TRAIL (mTRAIL) and soluble TRAIL (sTRAIL). Elevated mTRAIL in spinal cord tissue might not proportionally translate into increased sTRAIL in CSF or plasma, especially since the latter form is generated through cleavage by metalloproteases. Additionally, differences in the cellular sources of TRAIL, such as peripheral immune cells versus CNS‐resident astrocytes, could also contribute to these discrepancies.

TRAIL exerts its effects as a ligand by binding to its homologous receptors. In mice, three TRAIL receptors have been identified: DR5, DcR1, and DcR2, which exhibit varying affinities for TRAIL under normal physiological conditions (Gao et al. 2020; Hoffmann et al. 2009). Among these, only the DR5 receptor contains a death domain (DD) in its cytoplasmic region, transmitting apoptosis signals and exhibiting agonistic activity. The other two receptors, DcR1 and DcR2, lack a complete death domain and do not possess agonistic activity; however, they can bind TRAIL competitively, modulating receptor function (Gao et al. 2020; Haase et al. 2008). Interestingly, our in vitro experiments revealed that DR5 protein levels were significantly elevated following TRAIL supplementation in cell models. Previous studies have reported that chop is an upstream regulator of DR5 (Jeon et al. 2018; Kim et al. 2019; Song et al. 2023; Yue and Sun 2019). Our experiments demonstrated that TRAIL upregulated the Chop gene in the cell model. Therefore, we hypothesized that TRAIL further upregulates DR5 expression by promoting endoplasmic reticulum oxidative stress and CHOP production, thus creating positive feedback and amplifying the effect of the TRAIL‐DR5 pathway in ALS cell model. Oxidative stress is a well‐established pathogenic factor in ALS (Barber and Shaw 2010; Filézac De L'etang et al. 2015), linking ER stress and the TRAIL‐DR5 pathway in ALS. Edaravone, an antioxidant, has been reported to reduce neuronal death and decrease in TRAIL levels in mice with ischemic stroke (Li et al. 2018). Based on these findings, it’ is reasonable to infer that Edaravone's protective effects in ALS are associated with its anti‐oxidative properties and a concomitant reduction in TRAIL levels in the spinal cord of SOD1G93A mice (our unpublished data). It is worth noting that gene expression analysis was performed 72 h post‐transfection, while functional assays revealed cell injury phenotypes at 48 h. This timing was selected to align with the time point of TRAIL intervention (48 h post‐transfection followed by a 24‐h treatment period). The subtle changes in gene expression may reflect the transient and dynamic characteristics of stress responses during the progression of ALS‐like pathology in this in vitro model.

In contrast to the robust DR5 upregulation observed in vitro upon TRAIL treatment, DR5 expression in the spinal cords of SOD1‐G93A mice displayed a more restricted pattern. While overall DR5 levels were elevated compared to those in WT mice, DR5 expression within motor neurons was primarily detected in morphologically abnormal or degenerating neurons. In contrast, morphologically intact neurons exhibited minimal or no DR5 expression. This observation suggests that DR5 may be selectively upregulated in neurons undergoing degeneration. Additionally, the reduction in DR5 expression in motor neurons could be attributed to either the overall decrease in motor neuron numbers or alterations in transcriptional regulation mechanisms. However, further investigation is necessary to elucidate the precise mechanisms underlying these changes.

The apparent discrepancy between in vivo and in vitro DR5 expression profiles can be attributed to differences in microenvironmental context. In vivo, DR5 upregulation is likely tightly regulated and modulated by complex cellular interactions involving glial cells, cytokines, and local inflammation responses. In contrast, in vitro conditions lack these regulatory components, potentially allowing TRAIL to directly induce DR5 expression in a feed‐forward manner. This phenomenon has been documented in other systems and reinforces the concept that TRAIL signaling not only initiates but also amplifies cell death pathways through DR5 induction.

Previous studies have shown that in animal models of Alzheimer's disease (AD) and ischemic stroke, TRAIL is predominantly expressed by activated astrocytes and microglia (Cantarella et al. 2015; Cui et al. 2010). In contrast, DR5 is selectively expressed in vulnerable neurons, leading to the induction of neuronal death by TRAIL. In SOD1G93A mice, we observed a notable phenomenon: DR5 expression was virtually undetectable in morphologically intact motor neurons but was evident in a few morphologically compromised motor neurons.

Another intriguing finding in our study was the high expression of DR5 in the astrocytes and microglia of SOD1G93A mice. Previous studies have demonstrated that astrocytes and microglia in ALS undergo activation and proliferation rather than apoptosis (Humphrey et al. 2023; Van Harten et al. 2021). Consistent with these findings, we observed increased proliferation of astrocytes and microglia in SOD1G93A mice (Figures 2A). Therefore, the impact of the TRAIL‐DR5 pathway on astrocytes and microglia in ALS may differ from its pro‐apoptotic effect on motor neurons. While the TRAIL‐DR5 pathway is traditionally recognized for inducing apoptosis, emerging evidence suggests it can also promote survival by activating ERK1/2 downstream signaling in epithelial cells (Secchiero et al. 2003). Prior studies have shown that TRAIL does not induce apoptosis in astrocytes, regardless of pretreatment with a complete cytokine mix (CCM) (Song et al. 2006; Huang, Erdmann, Zhao, et al. 2005). In addition, microglia exhibit some resistance to TRAIL‐induced apoptosis, although this resistance can be reversed by viral infections or immune activation (Huang, Erdmann, Zhao, et al. 2005; Matysiak et al. 2002). Based on these observations and relevant literature, we hypothesize that the TRAIL‐DR5 pathway may drive the proliferation of astrocytes and microglia in ALS, which has been implicated in motor neuron damage. However, further experimental validation is required to confirm these hypotheses.

These findings and inferences, which appear to present contradictory phenomena—where the TRAIL‐DR5 pathway simultaneously promotes both survival and apoptosis—can be explained by the differential susceptibility of various cell types to TRAIL. As reported in the literature, oligodendrocytes and neurons exhibit greater sensitivity to TRAIL‐induced apoptosis, while astrocytes and microglia resist this effect (Song et al. 2006; Huang, Erdmann, Zhao, et al. 2005; Matysiak et al. 2002). The anti‐apoptotic effects observed in astrocytes and microglia may be attributed to their high expression of decoy receptors, which effectively mitigate the pro‐apoptotic actions typically induced by TRAIL.

Compared to previous studies on TRAIL/DR5 in ALS, our research is the first to investigate the effects and mechanisms of the TRAIL/DR5 pathway in ALS using both animal and cell models. Jiang et al. (2007) reported that DR5 expression is upregulated in subpopulations of residual motor neurons and correlates with pathological markers such as phosphorylated neurofilament heavy chain (pNF‐H) accumulation and the number of surviving motor neurons, suggesting its involvement in later stages of motor neuron degeneration. In contrast, our study using the SOD1G93A mouse model observed that DR5 was absent in morphologically intact motor neurons but present in a few degenerating motor neurons. Despite being derived from different ALS contexts, these findings collectively suggest that DR5 expression may be closely associated with motor neuron vulnerability and degeneration.

Our study demonstrates that blocking the TRAIL‐DR5 pathway in vitro using DR5‐neutralizing antibodies significantly reduces motor neuron loss induced by TRAIL and substantially decreases the transcription of genes associated with cell death. This finding not only offers a novel potential therapeutic approach for ALS but also provides a robust theoretical foundation for the clinical application of DR5‐neutralizing antibodies in ALS treatment. Previous research has shown that intraperitoneal injection of TRAIL‐neutralizing antibodies alleviates cognitive impairments in 3xTg‐AD mice and mitigates inflammatory responses (Cantarella et al. 2015). However, intraperitoneal administration of TRAIL or DR5‐neutralizing antibodies exacerbates clinical symptoms in Experimental Autoimmune Encephalomyelitis (EAE, an animal model commonly used to study the pathogenesis of multiple sclerosis) mice, promoting inflammation (Cretney et al. 2005). In contrast, intraventricular or intrathecal injection of TRAIL or DR5‐neutralizing antibodies significantly improves clinical symptoms and suppresses inflammation in EAE mice (Aktas et al. 2005), suggesting that TRAIL plays a role in immune regulation in multiple sclerosis. Therefore, we aim to investigate whether blocking the TRAIL‐DR5 pathway in SOD1G93A mice affects motor abilities and survival and whether peripheral or central blockade yields comparable outcomes in these mice.

However, this study has several limitations. First, to confirm whether TRAIL secreted by astrocytes binds to DR5 on motor neurons, it would have been ideal to culture and analyze primary astrocytes from SOD1G93A mice for elevated TRAIL secretion in the cell supernatant. Unfortunately, this experiment was not conducted due to technical constraints. Second, although we have made efforts to validate the specificity of the DR5‐neutralizing antibody through literature review and concentration‐dependent response analysis, we recognize that off‐target effects cannot be entirely excluded. Antibodies, particularly when applied in complex biological systems, may exhibit cross‐reactivity or non‐specific binding, which could potentially influence the observed outcomes. Future studies employing genetic approaches such as DR5 knockout or knockdown models, or employing alternative neutralizing strategies, will be essential for further confirming the specificity of DR5‐mediated effects in SOD1‐mutant motor neurons. Third, while this study demonstrates the neurotoxic effects of TRAIL in an ALS in vitro model, these findings need to be validated through in vivo experiments and, ideally, in human tissue samples. However, due to the current unavailability of postmortem human spinal cord specimens, such validation could not be performed in this study and remains an important goal for future research. Finally, a methodological limitation exists in the use of LDH and CCK8 assays, as these methods evaluate the entire cell population rather than specifically distinguishing between transfected and non‐transfected cells. Although our supplementary data indicate that 25 ng/mL TRAIL has minimal effects on non‐transfected NSC34 cells regarding viability and gene expression, we cannot entirely rule out the possibility of non‐specific toxicity. Future studies should incorporate fluorescence‐activated cell sorting (FACS) to separate transfected from non‐transfected cells, followed by viability assessments such as LDH release and CCK8 assays, to further clarify the specific effects of TRAIL on SOD1‐mutant motor neurons.

5. Conclusion and Outlook

This study is the first to demonstrate that TRAIL, predominantly secreted by astrocytes in an ALS animal model, exerts neurotoxic effects on motor neurons via the DR5 receptor. Additionally, DR5‐neutralizing antibodies were shown to effectively mitigate TRAIL‐induced motor neuron damage. These findings not only elucidate a novel mechanism underlying motor neuron degeneration but also highlight TRAIL/DR5 signaling as a potential therapeutic target for ALS.

Nevertheless, the precise mechanisms through which TRAIL induces motor neuron injury remain to be fully elucidated. Future studies should concentrate on validating these findings in vivo and exploring the cellular and molecular interactions between astrocytes and motor neurons within this pathway. We anticipate that our work will stimulate the development of novel therapeutic strategies for ALS.

Author Contributions

Kangqin Yang: conceptualization, data curation, formal analysis, visualization, writing – original draft, methodology, investigation, project administration, writing – review and editing, software. Yang Liu: conceptualization, data curation, formal analysis, writing – review and editing, supervision. Wenhua Deng: methodology. Zhenxiang Gong: supervision. Lifang Huang: methodology. Zehui Li: methodology. Min Zhang: conceptualization, writing – review and editing, supervision, resources, funding acquisition, validation.

Consent

Informed consent was achieved for all subjects, and the experiments were approved by the local ethics committee.

Conflicts of Interest

The authors declare no conflicts of interest.

Peer Review

The peer review history for this article is available at https://www.webofscience.com/api/gateway/wos/peer‐review/10.1111/jnc.70146.

Supporting information

Data S1.

JNC-169-0-s001.zip (17.9MB, zip)

Acknowledgements

We would like to thank Professor Neil R. Cashman at the Djavad Mowafaghian Centre for Brain Health, University of British Columbia, Vancouver, BC, V6T 1Z3, Canada, for granting us permission to use the NSC34 cell line. We are grateful to Professor Yaling Liu from The Second Hospital of Hebei Medical University (China) for her instrumental role in facilitating the transfer of this cell line. We also extend our sincere appreciation to the members of the Neurology Laboratory at Tongji Hospital, Tongji Medical College, Huazhong University of Science and Technology, for their invaluable support in experimental design and technical assistance, especially Dr. Xiaowei Pang, Dr. Yunfan You, and Dr. Hang Zhang.

Yang, K. , Liu Y., Deng W., et al. 2025. “Astrocytes Contribute to Motor Neuron Degeneration in ALS via the TRAIL‐DR5 Signaling Pathway.” Journal of Neurochemistry 169, no. 7: e70146. 10.1111/jnc.70146.

Funding: This study was supported by the National Natural Science Foundation of China (Grant 82271478) and the Research and Innovation Team Project for Scientific Breakthroughs at Shanxi Bethune Hospital (Grant 2024AOXIANG05).

Data Availability Statement

The datasets generated and analyzed during the current study are available from the corresponding author upon reasonable request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Data S1.

JNC-169-0-s001.zip (17.9MB, zip)

Data Availability Statement

The datasets generated and analyzed during the current study are available from the corresponding author upon reasonable request.

Articles from Journal of Neurochemistry are provided here courtesy of Wiley

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