Microglial HVCN1 Deficiency Improves Movement and Survival of SOD1G93A ALS Mice by Enhancing Microglial Migration and Neuroprotection
Fan Wang, Ke‐Yu Zhang, Lang‐Jian Zhu, Wei‐Jue Li, Yang Wu, Xiang Gao, Xiao‐Ru Ma, Xiu‐Hua Yin, Jian‐Bin Wu, Xiao‐Kang Ye, Zhao‐Jun Dong, Di‐Xian Wang, Zhe Zhou, Shao‐Dong Wang, Lei Han, Zhi‐Nong Jiang, Jing‐Wei Zhao
TL;DR
Deleting HVCN1 in microglia improves motor function and survival in ALS mice by enhancing microglial migration and neuroprotection.
Contribution
Identifies HVCN1 as a novel, druggable target for ALS therapy through microglial modulation.
Findings
HVCN1 deletion in microglia improves motor neuron survival and neuromuscular junction integrity in ALS mice.
HVCN1 deficiency suppresses Akt signaling, enhancing microglial migration and neurotrophic functions.
HVCN1 ablation leads to better motor function and survival in ALS mice compared to current clinical drugs.
Abstract
Amyotrophic lateral sclerosis (ALS) is an incurable motor neuron disease characterized by progressive loss of motor neurons. Current clinically available drugs targeting neurons show minor survival extension and no motor improvement in ALS patients. This shifts the focus of ALS research toward non‐neuronal cells, particularly microglia, a critical driver of ALS pathogenesis. Highly druggable ion channels are key regulators of microglia function. Here, Hydrogen voltage gated channel 1 (HVCN1) was screened out as the most highly expressed ion channel in microglia, and was upregulated in microglia of SOD1G93A mice and patients. Deletion of HVCN1 in microglia increased motor neuron survival, rescued the innervated neuromuscular junctions in the muscle, reduced glial activation and decreased the level of both misfolded protein and myelin debris in the ALS mice. Importantly, these…
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FIGURE 7- —National Key R&D Program of China10.13039/501100012166
- —National Natural Science Foundation of China10.13039/501100001809
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Taxonomy
TopicsAmyotrophic Lateral Sclerosis Research · Neuroinflammation and Neurodegeneration Mechanisms · Hydrogen's biological and therapeutic effects
Introduction
1
Amyotrophic lateral sclerosis (ALS) is the most prevalent form of motor neuron diseases [1]. The primary pathological feature of ALS is the loss of motor neurons in the motor cortex, brainstem, and anterior horn of the spinal cord. This leads to progressive atrophy of innervated muscles and ultimately results in the loss of muscle control, speech ability, and respiratory function [2]. ALS progresses rapidly, with most patients succumbing within 3–5 years after symptom onset [3, 4]. The pathogenesis of ALS remains incompletely understood, though glutamate‐mediated excitotoxicity and oxidative stress in neurons have been considered major pathogenesis mechanisms underlying ALS [5]. To date, directly targeting these two pathological mechanisms has led to two clinical drugs: riluzole and edaravone. However, both drugs offered only modest survival benefits without meaningful improvement in functional outcomes [6, 7]. The other drug, tofersen, targeting mRNA of superoxide dismutase 1 (SOD1), shows potential effects only in a subset of SOD1‐ALS patients [8, 9, 10]. These disappointing results suggest that therapeutic strategies focusing exclusively on neuronal targets may be insufficient. Consequently, recent research has shifted toward investigating the role of non‐neuronal cells, particularly microglia.
Microglia, the primary innate immune cells in the central nervous system (CNS), maintain homeostasis through surveillance, phagocytosis and inflammation [11, 12]. In both patients and animal models of ALS, microglia exhibit widespread hyperactivation in the CNS [13, 14, 15]. Activated microglia primarily exert neuroprotective effects during early stages of ALS but shift to predominantly neurotoxic roles in late disease phases [11, 16]. Given the critical roles of ion homeostasis and membrane potential changes in maintaining homeostatic microglia function, targeting ion channels in microglia has demonstrated therapeutic benefits in neuronal diseases such as Alzheimer's disease, Parkinson's disease and ischemia [17, 18, 19]. However, research on targeting microglial ion channels in ALS remains limited, with only P2X4, P2X7 and KCa3.1 knockouts shown to improve motor function in ALS mouse models thus far [20, 21, 22]. Therefore, there is an urgent need to explore new microglial ion channel targets for better intervention strategies for ALS.
In this study, we used a transcriptomic database to analyze ion channel genes and screened out hydrogen voltage gated channel 1 (HVCN1) as the most abundantly and specifically expressed ion channel gene in microglia, with its expression upregulated in microglia from both patients and mouse models of ALS. Using the classic and widely adopted *SOD1^G93A^
- ALS mouse model, which phenocopies motor neuron degeneration and muscle atrophy symptoms in ALS patients [23], we further validated the increased expression of HVCN1 in ALS‐associated microglia. By crossing Cx3cr1‐creERT mice with *HVCN1^flox/flox^
- mice to specifically knockout HVCN1 in microglia, our results demonstrated that HVCN1 deletion in microglia improved motor function and extended survival of the ALS mice. Neurons co‐cultured with HVCN1‐deficient primary microglia of the ALS mice exhibited a significant increase in both their survival and branching complexity compared to controls. Using bone marrow‐derived macrophages (BMDM) as a practically accessible cellular model based on their functional and molecular similarities to microglia, both in vivo and in vitro results revealed that HVCN1 deficiency enhanced microglial migration and phagocytosis, and their increased migration was mediated through the PI3K/Akt signaling pathway. Collectively, our results suggest that upregulated HVCN1 expression in ALS microglia impairs their chemotactic and neurotrophic capacities likely via the PI3K/Akt pathway, and thereby extends lifespan and improves motor function. This highlights HVCN1's potential as a novel therapeutic target for ALS intervention.
Results
2
Upregulation of HVCN1 in Microglia of ALS
2.1
To systematically identify ion channels specifically and highly expressed in microglia, we first retrieved all known ion channel genes from the International Union of Basic and Clinical Pharmacology database (IUPHAR‐DB, www.guidetopharmacology.org). Using transcriptomic data from acutely isolated mouse brain cells (GSE52564) [24], we found 182 ion channel‐encoding genes with detectable expression in brain cells (Supporting Table S1). Among these, 24 genes showed microglia‐specific expression patterns (Figure 1A). Strikingly, Hvcn1 exhibited the highest expression level in microglia (Figure 1B). The microglial specificity of Hvcn1 expression was further validated by quantitative real‐time PCR (qPCR) in primary microglial cultures of mouse (Figure 1C).
*Upregulation of Hvcn1 in microglia of ALS mice and patients. (A,B) Expression profiles of ion channel genes in cells acutely isolated from mouse brains based on RNA‐seq data (GSE52564, n = 2 independent samples per group). OPC, oligodendrocyte precursor cells; NFO, newly formed oligodendrocyte; MO, mature oligodendrocyte. (C) Hvcn1 mRNA level in primary cultured microglia, oligodendrocyte, astrocyte and neuron (n = 3 independent experiments). (D) Quantitative real‐time PCR (qPCR) analysis of Hvcn1 mRNA in the lumbar spinal cord of the SOD1G93A ALS mice (n = 3 mice per group). (E) Immunofluorescence staining of HVCN1 and Iba1 in the lumbar spinal cord of the ALS mice. Arrows, HVCN1+Iba1+ cells. (F) Quantification of HVCN1 positive microglia in the lumbar spinal cord of the ALS mice (n = 4 mice per group). (G) The mRNA level of HVCN1 in transcriptomic datasets from brains and spinal cords of ALS patients carrying SOD1 or FUS mutation, or assumed to have TDP‐43 pathology (GSE153960). Sample sizes for each region are as follows: lumbar spinal cord (Ctrl: 44, ALS: 147), cervical spinal cord (Ctrl: 41, ALS: 161), cerebellum (Ctrl: 38, ALS: 173), occipital cortex (Ctrl: 12, ALS: 51), temporal cortex (Ctrl: 26, ALS: 28), and motor cortex (Ctrl: 19, ALS: 83). (H) The mRNA level of HVCN1 in transcriptomic datasets from the lumbar spinal cord of the ALS mice (GSE18597, GSE220705). n = 3 mice per group. (I) The RNA level of HVCN1 in microglia acutely isolated from the spinal cord tissues of the ALS mice (GSE252050). n = 8 independent samples per group. FPKM, fragments per kilobase of transcript per million mapped fragments. TPM, transcripts per million. Data are presented as mean ± S.E.M. Significance was assessed by unpaired two‐tailed Student's t‐test (D, F, G, H and I), one‐way ANOVA with Dunnett's multiple comparisons test (C, H). *, p < 0.05; **, p < 0.01; **, p < 0.001.
HVCN1, so far the only one known proton‐selective ion channel in the mammalian nervous system, is predominantly expressed in microglia in the CNS [25, 26, 27]. This tetra‐spanning channel selectively conducts hydrogen ions, maintaining intracellular and organellar pH homeostasis under physiological conditions. Expression of HVCN1 in microglia is upregulated in multiple neural injury and neurodegenerative diseases [26]. Systemic knockout of HVCN1 exhibited beneficial results in neural diseases, suggesting its therapeutic potential [25, 28, 29, 30]. However, the function and mechanism of HVCN1 in ALS remains unexplored.
To investigate the expression level of HVCN1 in ALS, we performed qPCR and immunofluorescence staining on the lumbar spinal cord tissues from the ALS mice. The results showed significant upregulation of HVCN1 in the lumbar spinal cord tissues at RNA level (Figure 1D). Immunofluorescence confirmed that HVCN1 was specifically expressed in microglia at protein level, and microglial HVCN1 expression was significantly upregulated in the lumbar spinal cord of the ALS mice (Figure 1E,F). Importantly, transcriptome analysis of multiple ALS‐related datasets revealed consistent HVCN1 upregulation across different ALS models, and the HVCN1 upregulation was detected in both tissues (Figure 1G–I), including the brains and spinal cords of ALS patients carrying SOD1 or FUS mutation, or assumed to exhibit TDP‐43 pathology (GSE153960) [31, 32, 33], the spinal cords of *SOD1^G93A^
- mice (GSE18597, GSE220705) [34, 35], and cells‐acutely isolated microglia from *SOD1^G93A^
- mice (GSE252050) [36]. These results collectively indicate that HVCN1 is predominantly and specifically expressed in microglia, and is further upregulated in microglia in ALS, both in mouse models and in patients. These results suggest a potential role of microglial HVCN1 in ALS pathogenesis.
Microglial HVCN1 Deficiency Rescues Motor Neuron Degeneration and Ameliorates ALS Pathology in Mice
2.2
Given that HVCN1 is upregulated in microglia of ALS, we wonder whether microglia‐specific HVCN1 ablation modulates ALS pathology. We first constructed microglia‐specific Hvcn1 knockout ALS mouse model, *Cx3cr1‐creERT, Hvcn1^flox/flox^; SOD1^G93A^
- (subsequently named *Hvcn1^ΔMG^; SOD1^G93A^
- for short) mice and *Hvcn1^flox/flox^; SOD1^G93A^
- control mice. At age of 8 weeks, mice were injected with tamoxifen to induce HVCN1 knockout, a time point when neuronal development has completed but prior to the onset of disease symptoms. The mice were then sacrificed for pathological analysis at age of 20 weeks, when ALS symptoms have become evident (Figure 2A). In the lumbar spinal cord of the ALS mice, microglia‐specific Hvcn1 knockout resulted in an approximately 50‐fold downregulation of Hvcn1 mRNA (Figure 2B). Immunofluorescence data further showed that the density of HVCN1^+^ microglia and the HVCN1 signal within these cells were nearly undetectable following the knockout (Figure 2C–G). Furthermore, the HVCN1 deletion was validated by Western blot analysis of acutely isolated microglia from the brain and spinal cord of the ALS mice (Supporting Figure S1A–F). Collectively, our results validate the successful ablation of HVCN1 in microglia in the ALS mouse model.
*Conditional knockout of Hvcn1 in microglia ameliorates motor neuron loss and ALS pathology in the ALS mice. (A) Experimental design for microglial HVCN1 knockout in the ALS mice. (B) Quantification of Hvcn1 mRNA level in the lumbar spinal cord of the ALS mice following microglial HVCN1 knockout (n = 3 mice per group). (C—E) Quantification of HVCN1+ microglia density, mean fluorescence intensity (MFI) of HVCN1 in TMEM119+HVCN1+ microglia and their percentage against Iba1+ cells in the lumbar spinal cord of the ALS mice. (n = 4 mice per group). (F,G) Representative immunofluorescence images for HVCN1 co‐stained with TMEM119 and Iba1, GFAP and SOX10 in the lumbar spinal cord of the ALS mice. Arrows, HVCN1+TMEM119+Iba1+ cells. (H,I) Immunofluorescence images and their statistics show reduced microglial density in the lumbar spinal cord of the ALS mice following microglial HVCN1 knockout (n = 4 mice per group). (J,K) Immunofluorescence images and their statistics showed decreased astrocyte density in the lumbar spinal cord of the ALS mice following microglial HVCN1 knockout (n = 4 mice per group). (L—N) Immunofluorescence staining and its quantification showed reduced dMBP+ myelin debris in the lumbar spinal cord of the ALS mice following microglial HVCN1 knockout (n = 4 mice per group). (O—Q) Immunofluorescence staining and its quantification indicate diminished deposition of misfolded SOD1 protein (B8H10+) in the lumbar spinal cord of the ALS mice following microglial HVCN1 knockout (n = 4 mice per group). (R,S) Immunofluorescence staining and its statistics show more ChAT+ motor neuron in the lumbar spinal cord of ALS the mice following microglial HVCN1 knockout (n = 4 mice per group). (T) Quantification of Chat mRNA level in the lumbar spinal cord of the ALS mice following microglial HVCN1 knockout (n = 3 mice per group). (U) Experimental layout to detect ALS‐associated muscle pathology. (V) Microglial HVCN1 knockout increased the mass of the tibialis anterior muscle in the ALS mice (n = 4 mice per group). (W,X) Representative immunofluorescence images and corresponding quantification show that microglial HVCN1 knockout significantly increased the percentage of innervated neuromuscular junctions in the tibialis anterior muscle of the ALS mice (n = 4 mice per group). Arrow, innervated neuromuscular. Data are presented as mean ± S.E.M. Significance was assessed by unpaired two‐tailed Student's t‐test (B ‐E, I, K, M, N, P, Q, S, T and V), two‐way ANOVA with Sidak's multiple comparisons test (X). *, p < 0.05; **, p < 0.01; **, p < 0.001.
To further investigate the impact of microglial HVCN1 knockout on disease pathology in the ALS mice, immunofluorescence revealed reduction in microglial density (Figure 2H,I), astroglial density (Figure 2J,K), and accumulation of both degenerated myelin (Figure 2L–N) and misfolded SOD1 protein (Figure 2O–Q) in the lumbar spinal cord of the ALS mice following HVCN1 knockout. Moreover, the number of surviving motor neuron was increased in the anterior horn of the lumbar spinal cord of *Hvcn1^ΔMG^; SOD1^G93A^
- mice (Figure 2R–T), whose axons directly innervate the corresponding skeleton muscles of the hindlimbs. We speculate that more surviving motor neurons could indicate better motor control ability. As expected, the tibialis anterior muscle was heavier in the knockout mice (Figure 2U,V). Moreover, immunofluorescence revealed that microglial HVCN1 deletion caused an increase in the innervation of the neuromuscular junction (NMJ) in this muscle of the ALS mice (Figure 2W,X). Altogether, these results indicate that microglial specific Hvcn1 deletion enhances muscle control and motor neuron survival, attenuates microgliosis and astrogliosis, and reduces the accumulation of myelin debris and misfolded protein in the ALS mice.
Loss of Microglial HVCN1 Prolongs Survival and Ameliorates Symptoms of the ALS Mice
2.3
To further investigate whether the effects of microglial HVCN1 deletion can translate into functional outcomes, we induced HVCN1 knockout by injecting tamoxifen intraperitoneally daily for four consecutive days in 8‐week‐old *Hvcn1^ΔMG^; SOD1^G93A^
- mice and *Hvcn1^flox/flox^; SOD1^G93A^
- control mice (Figure 3A, Supporting Figure S2A). Behavioral tests were conducted every 4 weeks to monitor disease progression in the ALS mice. Results showed that conditional knockout of microglial HVCN1 significantly extended the survival of the ALS mice (Figure 3B, Supporting Figure S2B,C). The median survival increased from 198.5 days in *Hvcn1^flox/flox^; SOD1^G93A^
- control mice to 212.0 days in the *Hvcn1^ΔMG^; SOD1^G93A^
- mice, roughly equivalent to 7% extension of their lifespan. During the disease progression phase, *Hvcn1^ΔMG^; SOD1^G93A^
- mice exhibited significantly lower neurological function scores compared to controls (Figure 3C, Supporting Figure S2D,E), indicating their improved neurological symptoms. The rotarod test revealed a significant improvement in motor coordination in the ALS mice caused by microglial Hvcn1 deletion (Figure 3D), while the screen test revealed that *Hvcn1^ΔMG^; SOD1^G93A^
- mice exhibited markedly longer muscle endurance than control mice (Figure 3E). No significant difference in body weight was detected between *Hvcn1^ΔMG^; SOD1^G93A^
- mice and control mice (Figure 3F). The slow initial disease course and the minor mass of the affected muscles together could explain the stable total body weight of the ALS mice. Thus, while a sustained maximal 10% weight loss (∼2 g) to achieve humane endpoint for ALS mice, the considerable inherent variation in mouse body weight (up to 8 g) can thereby mask the specific effect of HVCN1 on muscle wasting. Collectively, these findings demonstrate that microglia‐specific HVCN1 knockout improves motor function and prolongs lifespan of the ALS mice, indicating that microglial HVCN1 is a potential translational target for ALS therapy.
*Microglial HVCN1 knockout alleviates motor function and prolongs survival of male SOD1G93A mice. (A) Experimental layout of microglial HVCN1 deletion in the ALS mice. (B) The Kaplan–Meier survival curve shows that microglial HVCN1 deletion significantly prolongs the survival of the male ALS mice (n = 18 mice in Hvcn1flox/flox; SOD1G93A group, n = 17 mice in Hvcn1ΔMG; SOD1G93A group). (C) Neurological score assay shows that microglial HVCN1 deletion reduces neurological symptoms in male SOD1G93A mice (n = 10 mice per group). (D) Rotarod test shows that microglial HVCN1 deletion improves motor coordination in male SOD1G93A mice (n = 10 mice per group). (E) Screen test shows increased muscle endurance of male SOD1G93A mice following microglial HVCN1 deletion (n = 10 mice per group). (F) Body weight curve of male SOD1G93A mice following microglial HVCN1 deletion (n = 10 mice per group). Data are presented as mean ± S.E.M. Significance was assessed by Kaplan‐Meier survival curves with log‐rank test (B), two‐way ANOVA with Sidak's multiple comparisons test (C—F). *, p < 0.05; **, p < 0.01; **, p < 0.001.
Microglial HVCN1 Deletion Enhances Their Own Migration
2.4
Given that microglial HVCN1 deletion exhibits therapeutic efficacy in the ALS mice, particularly by enhancing motor neuron survival, we wonder whether such beneficial effect is achieved through the surrounding microglia. 4‐month‐old *Hvcn1^ΔMG^; SOD1^G93A^
- mice and *Hvcn1^flox/flox^; SOD1^G93A^
- control mice received daily tamoxifen injections for four consecutive days to induce HVCN1 knockout, by which time the disease symptoms had already started. 3 days post induction, mice were perfused, and the lumbosacral enlargement of the spinal cord tissues were analyzed (Figure 4A). Using 2D immunofluorescence images, we quantified microglial soma (Iba1^+^DAPI^+^) within concentric 10 µm rings extending from the border of each motor neuron cell body (ChAT^+^DAPI^+^). The results showed a significant increase in microglial proximal distribution in the lumbar spinal cords toward motor neurons of *Hvcn1^ΔMG^; SOD1^G93A^
- mice compared to *Hvcn1^flox/flox^; SOD1^G93A^
- control mice (Figure 4B,C). These data suggest that, at least partially, HVCN1 in microglia rescues motor neuron through surrounding more microglia which have migrated likely from the surroundings into the anterior horn.
*Microglial HVCN1 deletion promotes microglia migration. (A) Experimental layout of microglial HVCN1 knockout in the ALS mice. (B,C) Representative images and their quantification show that microglial HVCN1 knockout enhanced microglial migration in the ALS mice (n = 5 mice per group). Arrows, microglia nearby motor neurons. (D) Experimental design for RNA sequencing in bone marrow derived macrophages (BMDMs) from the ALS mice following HVCN1 knockout. (E) Principal component analysis plot of RNA sequencing in BMDMs from the ALS mice following HVCN1 knockout (n = 3 independent samples per group). (F) Volcano plot shows differentially expressed genes (|log2FC|>1 and p_adjust < 0.05 analyzed by DESeq2) in BMDMs from the ALS mice following HVCN1 knockout (n = 3 independent samples per group). The representative migration related genes were labeled. Red dot, upregulated genes; Blue dot, downregulated genes; Gray dot, non‐significant changed genes. (G) Top 12 significantly enriched GO pathways based on ranks of adjusted p value in BMDMs from the ALS mice following HVCN1 knockout (n = 3 independent samples per group). (H) Gene network analysis of migration‐related pathways (n = 3 independent samples per group). (I) Heatmap of migration‐related gene expression patterns based on transcriptomic sequencing of BMDMs from the ALS mice following HVCN1 depletion (n = 3 independent samples per group). (J) Quantification of migration‐related gene expression in BMDMs from the ALS mice following HVCN1 knockout (n = 3 independent samples per group). (K,L) Representative transwell assay images and quantitative analysis show enhanced migration in BMDMs from the ALS mice following HVCN1 knockout (n = 4 independent samples per group). (M) Experimental layout for evaluating microglia migration in vivo by an early myelin injury assay. (N,O) Representative immunofluorescence images and quantitative analyses show significantly increased density of TMEM119⁺Iba1⁺ microglia in the peripheral lesion area of the brain in microglial HVCN1‐knockout ALS mice at 4 h post‐myelin injury (n = 4 mice per group). C, central lesion area; P, peripheral lesion area. P) Representative immunofluorescence images display the absence of BrdU⁺ proliferating microglia in the peripheral lesion area of the brain in microglial HVCN1‐knockout ALS mice at 4 h post‐myelin injury. Data are presented as mean ± S.E.M. Significance was assessed by unpaired two‐tailed Student's t‐test (L), two‐way ANOVA with Sidak's multiple comparisons test (C, J, O). *, p < 0.05; **, p < 0.01; **, p < 0.001.
To further explore whether HVCN1 deletion could affect migration of microglia in the ALS mice, bone marrow‐derived macrophages (BMDMs) from *Hvcn1^ΔMG^; SOD1^G93A^
- mice and *Hvcn1^flox/flox^; SOD1^G93A^
- control mice during the disease progression stage were cultured. BMDMs serve as a practically accessible in vitro model for microglia based on their similarity in morphology, gene expression and functions, particularly in injury and disease contexts. HVCN1 knockout was induced by treatment with 4 mg/ml tamoxifen during culture. BMDMs of *Hvcn1^ΔMG^; SOD1^G93A^
- mice and *Hvcn1^flox/flox^; SOD1^G93A^
- control mice were collected for transcriptomic sequencing (Figure 4D). Principal component analysis (PCA) plot revealed significant divergence between HVCN1‐knockout cells and control cells (Figure 4E). Volcano plot of differentially expressed genes (DEGs) demonstrated that most cell migration‐related genes were significant upregulated (Figure 4F). Gene Ontology enrichment analysis of upregulated DEGs highlighted that 8 out of the top 12 significant enriched pathways were associated with cell migration (Figure 4G). Further network analysis identified 5 genes as core genes within these 8 migration‐related pathways, included Col1a1, Itga11, Snai2, Ccn4 and Smoc2 (Figure 4H). Heatmap showed that, 29 migration related genes, identified through literature mining and PathCards analysis (www.pathcards.genecards.org), including Col1a1, Fn1, and Rhob, were significantly altered following HVCN1 knockout (Figure 4I). These results suggest a potential critical role of HVCN1 in regulating cell migration capacity. Subsequent qPCR data confirmed significant upregulation of migration‐related genes (Col1a1, Cxcl12, Rhob, etc.) in BMDMs from *Hvcn1^ΔMG^; SOD1^G93A^
- mice compare to *Hvcn1^flox/flox^; SOD1^G93A^
- control mice (Figure 4J).
To directly assess whether HVCN1 deletion affects cell migration in vitro, transwell assays further revealed enhanced chemotactic capacity of HVCN1‐knockout BMDMs compared to control cells (Figure 4K,L), confirming that loss of HVCN1 in BMDMs indeed enhances cell migration. Considering the similarity in gene expression and function between BMDMs and microglia, these data made us speculate that HVCN1 deletion in microglia rescuing motor neurons was achieved likely by redistributing more microglia to the vicinity of motor neuron, and these microglia likely have migrated from the surrounding areas.
More microglia surrounding motor neurons in the anterior horn could be also due to proliferation of microglia in situ. To clarify whether HVCN1 in microglia affects proliferation of microglia, gene set enrichment analysis and qPCR assay revealed significant downregulation of proliferation‐related pathways and gene expression signatures in Hvcn1‐deficient ALS BMDMs (Supporting Figure S3A–C). Immunofluorescence staining of the spinal cords from the ALS mice further showed decreased density and the proportion of proliferative microglia following Hvcn1 ablation (Supporting Figure S3D–F). Taking into accounts that HVCN1 deletion enhances cell migration, these data indicate that HVCN1 deletion suppressed microglia proliferation. Thus, the increased presence of microglia surrounding motor neurons is highly likely due to microglia migration.
To assess the effects of HVCN1 deletion on microglial migration in vivo, we stereotactically injected myelin debris into the brains of *Hvcn1^ΔMG^; SOD1^G93A^
- mice and *Hvcn1^flox/flox^; SOD1^G93A^
- control mice. Following the injection, 5‐bromo‐2'‐deoxyuridine (BrdU) was administered intraperitoneally to concurrently label proliferating cells. (Figure 4M, Supporting Figure S4A). 4 h post‐injection, immunofluorescence staining revealed a significantly higher Iba1^+^TMEM119^+^ microglia density in the peripheral lesion areas of *Hvcn1^ΔMG^; SOD1^G93A^
- mice compared to controls, whereas nearly no Iba1^+^TMEM119^−^ macrophage or Iba1^+^TMEM119^+^ BrdU^+^ proliferating microglia was detected (Figure 4N–P). 3 days post‐injection, immunofluorescence staining revealed significantly higher Iba1^+^TMEM119^+^ microglial density, higher Iba1^+^TMEM119^−^ macrophage density, lower Ki67^+^ proliferative microglia/macrophage percentage, and lower BrdU^+^ proliferative microglia percentage in the lesion areas of *Hvcn1^ΔMG^; SOD1^G93A^
- mice compared to controls (Supporting Figure S4B–H). Similarly, employing the lysolecithin induced demyelination model, we observed a significant increase in Iba1^+^ microglia/macrophage density alongside a decrease in the proportion of Ki67^+^ proliferative microglia/macrophage in the lesion areas of *Hvcn1^ΔMG^; SOD1^G93A^
- mice compared to controls (Supporting Figure S4I–O). These results indicate that HVCN1 deficiency enhances microglial chemotaxis in vivo. Collectively, our in vitro and in vivo results indicate that microglial HVCN1 knockout enhances microglia chemotaxis and migration.
HVCN1 Deficiency Enhances Neurotrophic Capacity of Microglia
2.5
To investigate the effects of microglial HVCN1 deletion on neuronal survival, we employed a co‐culture system using microglia from adult ALS mice CNS and primary hippocampal neurons from E18 wild type mice. As previously described (Supporting Figure S1A), tamoxifen was administrated to 16‐week old *Hvcn1^ΔMG^; SOD1^G93A^
- mice and *Hvcn1^flox/flox^; SOD1^G93A^
- control mice to induce microglial HVCN1 knockout. Microglia were subsequently isolated from both the brain and the spinal cord via fluorescence‐activated cell sorting and cultured. These microglia were then reseeded into 1 µm Transwell chambers and placed above the primary neuronal cultures (Figure 5A). Immunofluorescence results exhibited a significant increase in the number of surviving neurons co‐cultured with microglia from *Hvcn1^ΔMG^; SOD1^G93A^
- mice compared to *Hvcn1^flox/flox^; SOD1^G93A^
- control mice (Figure 5B,C). Further morphological analysis indicated that the surviving neurons in the *Hvcn1^ΔMG^; SOD1^G93A^
- group also exhibited more branches than those in the control group (Figure 5D), implying their healthier status. Consistent with the immunofluorescence data, conditioned culture medium was collected from the microglial cultures and was added to primary hippocampal neurons. Subsequent CCK‐8 assays confirmed that neuronal viability was significantly enhanced when treated with the conditioned medium from *Hvcn1^ΔMG^; SOD1^G93A^
- microglia in comparison with the controls (Figure 5E). We further validated these findings using a BMDM‐neuron transwell co‐culture system. Following tamoxifen‐induced HVCN1 knockout in BMDMs (Supporting Figure S5A), immunofluorescence quantification showed that neurons co‐cultured with *Hvcn1^ΔMG^; SOD1^G93A^
- BMDMs exhibited significantly more neuronal numbers and more neurite branches compared to the controls (Supporting Figure S5B–D). Consistently, CCK‐8 assay revealed significantly higher neuronal viability when cultured with conditioned medium from *Hvcn1^ΔMG^; SOD1^G93A^
- BMDMs than the controls (Supporting Figure S5E). These data demonstrated the beneficial effects of both microglia and BMDMs on neurons following HVCN1 knockout. Collectively, these data indicate that HVCN1 deletion in microglia augments their neurotrophic support in the ALS mice.
*HVCN1 knockout in microglia enhances neurotrophic capacity of microglia. (A) Schematic layout of the co‐culture assays employing primary adult microglia and neurons, including the transwell system and conditioned medium application. (B—D) Representative immunofluorescence images and quantitative analyses show that primary hippocampal neurons exhibited enhanced neuronal survival and denser neurite branching when co‐cultured with microglia from Hvcn1ΔMG; SOD1G93A mice than from Hvcn1flox/flox; SOD1G93A mice (n = 4 independent samples per group). (E) CCK‐8 assay reveals enhanced viability of neurons when co‐cultured with microglia from Hvcn1ΔMG; SOD1G93A mice than from Hvcn1flox/flox; SOD1G93A mice (n = 4 independent samples per group). (F) Gene set enrichment analysis shows upregulated TGF‐β pathway activity in BMDMs following HVCN1 knockout(n = 3 independent samples per group). (G) Upregulated Tgfb2 mRNA level in BMDMs from the ALS mice following HVCN1 knockout (n = 3 independent samples per group). (H) Upregulated Tgfb2 mRNA level in the lumbar spinal cord of the ALS mice following microglial HVCN1 knockout (n = 3 mice per group). (I) Quantification the mRNA level of Nos2, Ym1 and Vsig4 in BMDMs from the ALS mice following HVCN1 knockout (n = 3 independent samples per group). (J,K) Representative flow cytometry plots and their quantitative results show reduced proportion of CD86+ BMDMs from the ALS mice following HVCN1 knockout (n = 3 independent samples per group). (L,M) Representative flow cytometry images and statistical analyses show increased proportion of CD206+ BMDMs from the ALS mice following HVCN1 knockout (n = 3 independent samples per group). (N—P) Immunofluorescence staining and statistical analyses reveal decreased density and proportion of iNOS⁺Iba1⁺ microglia in the lumbar spinal cord of the ALS mice following HVCN1 knockout (n = 4 mice per group). Arrows, co‐localization of iNOS and Iba1. (Q—S) Representative immunofluorescence images and their quantification show increased density and proportion of TGF‐β⁺Iba1⁺ microglia in the lumbar spinal cord of the ALS mice following HVCN1 knockout (n=4 mice per group). Arrows, co‐localization of TGF‐β and Iba1. (T U) Representative immunofluorescence images and their quantification show that the mean fluorescence intensity of TMEM119 was significantly increased in TMEM119⁺Iba1⁺ microglia in the lumbar spinal cord of the ALS mice following HVCN1 knockout (n = 4 mice per group). Data are presented as mean ± S.E.M. Significance was assessed by unpaired two‐tailed Student's t‐test (C, E, K, M, O, P, R, S, U), two‐way ANOVA with Sidak's multiple comparisons test (D, G—I). *, p < 0.05; **, p < 0.01; **, p < 0.001.
Sustained microglial activation is a recognized feature for ALS that drives disease progression [11, 37]. Importantly, the restoration of homeostatic microglia has been demonstrated to attenuate neuronal loss and ameliorate disease progression in ALS mouse models [38, 39]. We further explored the effects of HVCN1 knockout on microglial activation in the ALS mice. Gene set enrichment analysis (GSEA) of transcriptomic data from BMDMs of HVCN1‐knockout ALS mice showed a significant upregulation of TGF‐β signaling activity (Figure 5F). qPCR results further revealed elevated expression of Bmp1 and Tgfb2 in both spinal cords and BMDMs of *Hvcn1^ΔMG^; SOD1^G93A^
- mice compared to *Hvcn1^flox/flox^; SOD1^G93A^
- control mice (Figure 5G,H). Given the established role of TGF‐β signaling in maintaining microglial homeostasis and promoting neuroprotection by suppressing their excessive activation in neurodegenerative disease models [39, 40, 41], these data suggest that HVCN1 knockout possibly alters the activation of BMDMs in the ALS mice. To test this possibility, qPCR results revealed reduced mRNA levels of the Nos2 but increased expression of Ym1 and Vsig4 in HVCN1 deficient BMDMs (Figure 5I). Flow cytometry analysis also showed that HVCN1 knockout significantly decreased the proportion of CD86 positive BMDMs, and in contrast, it increased the proportion of CD206 positive cells (Figure 5J–M). In vivo immunofluorescence staining showed that HVCN1 knockout in microglia significantly reduced the number and proportion of iNOS positive microglia (Figure 5N–P), on the contrary, the number and proportion of TGF‐β positive microglia (Figure 5Q–S), and the mean fluorescence intensity of TMEM119 in microglia were significantly increased in the lumbar spinal cord of the ALS mice following HVCN1 knockout (Figure 5T,U). These in vitro and in vivo data indicate that HVCN1 deficiency in microglia promotes a shift of microglia toward a homeostatic or more anti‐inflammatory state.
Oxidative stress and cellular debris accumulation contribute to the neural microenvironment deterioration and neuronal death in neurodegenerative diseases [42]. In both the lumbar spinal cord tissue and BMDMs of the ALS mice with HVCN1 knockout, qPCR analysis showed a coordinated downregulation of pro‐oxidant genes and upregulation of antioxidant genes. Furthermore, flow cytometry confirmed that ROS production was significantly reduced in BMDMs from the Hvcn1 deficiency ALS mice compared to the controls (Supporting Figure S6A–J). Gene set variation analysis and transcriptomic heatmap profiling exhibited modest upregulation of phagocytosis‐related pathways and genes following Hvcn1 ablation. Flow cytometry and immunofluorescence staining also confirmed that HVCN1 deficiency enhanced microglial phagocytosis in BMDMs from the ALS mice (Supporting Figure S7A–I). These results collectively indicate that microglial HVCN1 knockout both suppresses their pro‐inflammatory response and enhances their neuroprotective function in the ALS mice.
HVCN1 Deletion Reduces Akt Pathway Activity
2.6
To dissect the mechanisms underlying that HVCN1 knockout affects microglial migration and activation, we performed KEGG enrichment analysis on transcriptomic data from BMDMs of HVCN1‐knockout ALS mice and the control mice. The results showed that PI3K‐Akt pathway was among the top 5 significant enrichment pathways (Figure 6A, Supporting Table S2), suggesting that HVCN1 knockout may affect PI3K‐Akt activity in microglia of the ALS mice. To detect the protein level of Akt, BMDMs were cultured from *Hvcn1^ΔMG^; SOD1^G93A^
- mice and the control mice. Tamoxifen was administrated throughout the culture of BMDMs to induce HVCN1 deletion. Further Western blot experiments showed that phosphorylated Akt (p‐Akt) levels in BMDMs of the ALS mice was significantly decreased by HVCN1 knockout, while the total Akt level remained unchanged (Figure 6B–E). We observed that HVCN1 protein was not completely eliminated in the HVCN1‑knockout group. This may be attributed to the heterogeneous nature of the BMDM population, a notion supported by previous reports indicating the presence of a minor subpopulation of CX3CR1^low^ monocytes in the bone marrow [43, 44, 45]. These results show that HVCN1 knockout decreases Akt pathway activity in BMDMs of the ALS mice. It is plausible that Akt signaling is also suppressed in microglia by HVCN1 knockout, given that microglia share high functional and gene expression similarity with BMDMs.
*Decreased activity of Akt pathway in BMDMs from the ALS mice caused by HVCN1 knockout. (A) Top 5 significantly enriched KEGG pathways based on ranks of p_adjust value in BMDMs following HVCN1 knockout (n = 3 independent samples per group). (B—E) Representative immunoblot images and their quantification show HVCN1 knockout reduced p‐Akt level in BMDMs (n = 3 independent samples per group). Data are presented as mean ± S.E.M. Significance was assessed by unpaired two‐tailed Student's t‐test (C—E) *, p < 0.01.
Akt Activation Reverses the Enhanced Migration and Decreases Pro‐inflammatory Activation of Microglia Caused by HVCN1 Deletion
2.7
Now that loss of HVCN1 in microglia reduces Akt pathway activity, to further investigate whether HVCN1 regulates microglial migration in ALS through the Akt pathway, the Akt‐specific agonist SC79 was used to enhance Akt activity in BMDMs from *Hvcn1^ΔMG^; SOD1^G93A^
- mice. The qPCR results showed that SC79 treatment counteracted the upregulation of migration‐related genes such as Col1a1, Cxcl12, Rhob in BMDMs caused by HVCN1 knockout (Figure 7A). Transwell assays further demonstrated that SC79 abolished the enhanced migration of HVCN1‐knockout BMDMs (Figure 7B,C). To explore whether HVCN1 modulates the pro‐inflammatory activation state of microglia in the ALS mice via the Akt pathway, flow cytometry revealed that SC79 treatment restored the HVCN1 knockout‐induced decrease in the proportion of CD86 positive cells (Figure 7D,E), and eliminated the increase in CD206 positive cell proportion (Figure 7F,G) in BMDMs from the ALS mice. The qPCR results showed that activation of Akt by SC79 counteracted the expression of Nos2 from downregulation to a sharp upregulation, while reversed the expression of Ym1 and Vsig4 from upregulation to significant downregulation in BMDMs of HVCN1 knockout ALS mice (Figure 7H). Finally, the qPCR results indicated that SC79 reversed the upregulation of Bmp1 and Tgfb2 expression to significant downregulation in BMDMs from the ALS mice caused by HVCN1 knockout (Figure 7I). Collectively, these data indicate that HVCN1 regulates both migration and pro‐inflammatory activation of BMDMs in the ALS mice, and such effects are mediated likely via Akt pathway. These results suggest that HVCN1 modulates microglial migration and their pro‐inflammatory activation likely via the Akt pathway.
*Pharmacological activation of Akt counteracts the increased microglia migration and decreased pro‐inflammatory response caused by HVCN1 deletion. (A) Akt activation by SC79 downregulates migration‐related gene expression in BMDMs from the HVCN1‐knockout ALS mice (n = 3 independent samples per group). (B,C) Representative transwell assay images and their quantification show SC79 reduced migration ability of BMDMs from the HVCN1‐knockout ALS mice (n = 4 independent samples per group). (D,E) Representative flow cytometry images and statistical analyses show SC79 increased the proportion of CD86+ BMDMs from the HVCN1‐knockout ALS mice (n = 4 independent samples per group). (F,G) Representative flow cytometry images and quantitative analysis show decreased proportion of CD206+ BMDMs from the HVCN1‐knockout ALS mice (n = 4 independent samples per group). (H) SC79 upregulates the mRNA level of Nos2 and downregulated the mRNA level of Ym1, Vsig4 in BMDMs from the HVCN1‐knockout ALS mice (n = 3 independent samples per group). (I) SC79 reduces Tgfb2 mRNA level in BMDMs from the HVCN1‐knockout ALS mice (n = 3 independent samples per group). Data are presented as mean ± S.E.M. Significance was assessed by one‐way ANOVA with Tukey's multiple comparisons test (A, C, E, G—I) *, p < 0.05; **, p < 0.01; **, p < 0.001.
Discussion
3
ALS is an incurable and fatal motor neuron disease. So far, there are only three clinically available drugs, riluzole, edaravone and tofersen, which directly target glutamate‐mediated excitotoxicity, oxidative stress in neurons and SOD1 mRNA, respectively [46]. Unfortunately, toferson shows potential effects only confined to a subset of SOD1‐ALS patients [8, 9, 10], while riluzole and edaravone exhibit limited efficacy in extending life span without significant improvement in functional outcomes [5, 6, 7]. Microglia, the resident innate immune cells in the CNS, play crucial roles in ALS pathogenesis [11, 37]. Ion channels, the second largest class of drug targets after GPCRs, shape microglia function in physiological and pathological conditions [17, 18]. Therefore, exploring novel druggable ion channels in microglia of ALS is a promising strategy to devise new disease‐modifying therapies for ALS.
Here, using transcriptomic data from acutely isolated mouse brain cells, we screened out, in microglia, HVCN1 exhibits the highest expression among all known ion channel genes. Subsequent analysis further revealed upregulated HVCN1 expression in microglia of not only ALS mouse models but also ALS patients. This established the association between microglial HVCN1 and ALS. To further explore whether HVCN1 in microglia affects pathogenesis of ALS, we specifically knocked out HVCN1 in microglia of the *SOD1^G93A^
- ALS mouse. Our results revealed that microglial HVCN1 deletion increased motor neuron survival, ameliorated gliosis, and reduced toxic myelin and misfolded protein level in the spinal cord of the ALS mice. These pathological improvements were ultimately translated into enhancing motor function and extending survival of the ALS mice. Compared to the minor survival extension and lack of significant functional outcomes observed with the current clinical drugs, our findings highlight that targeting HVCN1 in microglia, identified in this work, has great potential to achieve significant improvement of motor function and survival for ALS. Understandably, further preclinical studies are needed to fully elucidate the effects of HVCN1 in microglia in ALS. To date, drug discovery efforts targeting ion channels in ALS have primarily focused on ion channels in neurons [47]. However, the highly similar ion channel expression patterns across different neuronal subtypes, combined with the fact that motor neurons, which are severely affected in ALS, constitute only a small fraction of all neurons, may contribute to the limited efficacy and undesirable side effects of these drugs. Our study shifts the focus from neuronal ion channels to microglia‐specific ion channels, pointing a new direction for ALS drug development.
Our work has established the principle of concept that inhibiting microglial HVCN1 is a promising therapeutic target for ALS. Based on this, the exploration of potent and selective inhibitors of HVCN1 is promising. Currently, several candidate compounds have been identified that show inhibitory effects on HVCN1 in cellular models [13, 27, 48]. However, their specificity and efficacy require further validation. Among these candidates, YHV98‐4 has been shown to inhibit HVCN1 in dorsal root ganglion neurons in mice without affecting hERG, KCNQ2, BK, and Nav1.7 channels [29]. More comprehensive screenings are needed to characterize the selectivity of YHV98‐4 against a broader range of ion channels and receptors. The blood‐brain barrier (BBB) poses a significant challenge for drug delivery to the CNS, and has been a major obstacle for ALS drug discovery. Therefore, more systematic screening efforts are necessary to identify HVCN1 inhibitors that can effectively penetrate BBB and exert their therapeutic effects in the CNS.
This work demonstrates that targeting microglial HVCN1 protects neurons in ALS. Together with previous findings that targeting microglial genes can ameliorate ALS pathology [38, 49], we propose that directly targeting microglial HVCN1 is a promising and effective strategy for promoting neuronal survival in ALS. Since ALS pathology develops much earlier than appearance of the symptoms. To explore the roles of microglial HVCN1 on pathologies of ALS, we induced microglial HVCN1 knockout in mice at age of 8 weeks, when neuronal development has nearly completed but overt gliosis or motor neuronal loss has not occurred yet, aiming to maximize efficacy while reduce the developmental side effects. In clinical practice, however, treatment usually begins after symptom onset. This might explain why frequent failures has been hard to avoid. Further investigation is needed to explore HVCN1 as a viable target for future ALS drug discovery.
The technical challenge of directly targeting microglia with genetic tools like CRISPR has hindered the translational advancement of microglia‐focused therapies in ALS. This has evoked interest in more druggable targets, such as ion channels and receptors in microglia. Meanwhile, a recent study showed that indirectly influencing microglia via galectin‐1 alongside AAV9‐SOD1 shRNA not only reduced neuroinflammation but also led to therapeutic outcomes in a ALS mouse model, underscoring the role of microenvironment [50]. These findings pave the way for future strategies that combining direct microglial gene targeting with indirect microenvironmental modulation (e.g., tofersen or galectins) may lead to more effective therapies in ALS.
Given that the survival, maintenance and proliferation of microglia are critically dependent on CSF1R, several compounds have been developed to selectively ablate microglia, providing powerful tools for microglial manipulation [51, 52]. Clinical trials exploring the depletion of microglia in ALS are currently in progress [53, 54]. Future studies should systematically evaluate the effects of both microglial depletion and microglial replacement therapies in ALS models. Such investigations are crucial for evaluating whether microglial targeting represents a viable therapeutic strategy for ALS patients.
Our results reveal that microglial HVCN1 deletion strengthens the neurotrophic functions of microglia, characterized by reducing pro‐inflammation genes, and elevated expression of TGF‐β and TMEM119. It is reported that microglia are activated to an anti‐inflammatory state during early stages of ALS, exerting neuroprotective effects through secreting neurotrophic factors. As the disease progresses, microglia gradually shift to a pro‐inflammatory state in the late stages, accelerating disease progression by releasing pro‐inflammatory cytokines [11, 16]. Notably, in late‐stages of ALS, microglial inflammatory state become dysregulated, often exhibiting simultaneous upregulation of both pro‐ and anti‐inflammatory factors, or concurrent activation and suppression signals within the same cytokine pathway [55, 56]. In line with this complexity, a recent study reported co‐upregulation of CD109, a TGF‐β suppressor, and TGF‐β itself in microglia of the spinal cord in the ALS mice [57]. Together, these findings underscore the therapeutic potential of restoring microglial homeostasis rather than merely inhibiting pro‑inflammatory microglia in ALS.
Mechanistically, our transcriptomic analysis reveals that the PI3K/Akt pathway exhibits the most pronounced inhibition in activity in HVCN1‐deficient microglia. Furthermore, Akt activation eliminated the enhanced cell migration caused by HVCN1 deficiency. Previous studies have reported that HVCN1 knockout affects multiple downstream signaling pathways, including PI3K/Akt, Erk, AMPK, and calcium signaling [29, 58, 59, 60, 61, 62]. These results highlight the critical role of the Akt pathway in mediating the function of HVCN1. It is reported that disruption of Akt signaling contributes to the pathogenesis of neurodegenerative diseases [63]. The Akt kinase family comprises three isoforms (Akt1, Akt2 and Akt3). So far, their distinct roles in specific CNS cell types in ALS require further elucidation [64]. Notably, Akt exerts pleiotropic effects on processes such as proliferation, survival, and metabolism through central downstream mediators including FOXO, GSK3 and mTOR [65, 66]. Therefore, it seems hard to selective target Akt pathway to modulate HVCN1 for therapeutic purpose.
In addition, our screening of ion channels across different cell types in the CNS provides valuable resources for future drug discovery efforts. The dataset identifies cell type‐specific ion channel expression patterns and potential therapeutic targets that could be exploited to develop more precise CNS therapeutics. For example, the selective enrichment of certain potassium channels in microglia or voltage‐gated calcium channels in neurons may offer new opportunities for modulating neuroinflammation or neuronal excitability in neuronal disease such as ALS, epilepsy or ischemia.
We used not only primary microglia but also BMDMs as a cell model for microglia for in vitro assays, and yielded similar results. BMDMs represent a practically accessible microglial model based on their morphological and functional similarities, despite not being true microglia [66, 67]. Transplanted bone marrow‐derived microglia‐like cells, which share ∼90% of their transcriptome with resident microglia [68], can ameliorate motor deficits and improve survival in ALS mice [69, 70, 71, 72], demonstrating significant functional conservation despite ontological differences.
In summary, our work screens out HVCN1 in microglia as an effective druggable target for ALS. HVCN1 is upregulated in microglia of ALS in both mouse model and in patients. Microglial HVCN1 deficiency enhances microglial migration and phagocytosis, modifies their activation state and improves their neuroprotection, consequently improves motor function and extends life span of the ALS mice. These results identify HVCN1 as a novel promising therapeutic target for ALS, opening a new avenue to develop specific inhibitor for HVCN1 to alleviates ALS.
Experimental Section
4
Animals
4.1
To specifically knockout HVCN1 in microglia of ALS mice, *SOD1^G93A^
- transgenic mice (Stock No. 002726) and Cx3cr1‐creERT mice (Stock No. 021160) were obtained from The Jackson Laboratory (Bar Harbor, ME, USA). *Hvcn1^flox/flox^
- mice (Stock No. T019403) were purchased from GemPharmatech (Nanjing, China). *SOD1^G93A^
- mice were crossed with *Hvcn1^flox/flox^
- mice to obtain *SOD1^G93A^; Hvcn1^flox/flox^
- mice. Cx3cr1‐creERT mice were crossed with *Hvcn1^flox/flox^
- mice to obtain *Cx3cr1‐creERT; Hvcn1^flox/flox^
- mice. *SOD1^G93A^; Hvcn1^flox/flox^
- mice were crossed with *Cx3cr1‐creERT; Hvcn1^flox/flox^
- mice to obtain *Cx3cr1‐creERT; Hvcn1^flox/flox^; SOD1^G93A^
- mice (*Hvcn1^ΔMG^; SOD1^G93A^
- mice). All mice were maintained on a C57BL/6J genetic background. Genotyping was performed by PCR analysis of genomic DNA extracted from mouse tails. Mice were housed under specific pathogen‐free environment with a 12‐h light/dark cycle and maintained at 22 ± 1°C. Food and water were provided ad libitum. All animal experiments were approved by the Ethics Committee of School of Medicine, Zhejiang University and performed in strict accordance with the National Guidelines for the Care and Use of Laboratory Animals in China (ETHICS number: 14660).
Tamoxifen Administration
4.2
To efficiently induce creERT‐dependent recombination, tamoxifen (Beyotime, ST1681) was dissolved in corn oil at a concentration of 20 mg/mL, and mice received daily intraperitoneal injections of tamoxifen at a dose of 100 mg/kg body weight for four consecutive days. Tamoxifen was administrated at different disease stages depending on the experimental endpoint: at age of 8 weeks (pre‐symptomatic and post‐development) for assessing pathology and symptom progression, whereas at age of 16 weeks (symptomatic) for evaluating microglial migration within the established disease environment.
Survival Test
4.3
To assess the survival period of ALS mice, the animals were monitored daily after the onset of bilateral hindlimb paralysis. Mice were laid on either side, and the inability to right themselves within 30 s was defined as the survival endpoint.
Neurological Score
4.4
To routinely and reliably assess neuromotor function in ALS mice, a neurological scoring system was adapted from the Amyotrophic Lateral Sclerosis Functional Disability Index (ALSTDI) scale with minor modifications [73]. Scoring criteria were defined as follows: 0, no apparent abnormalities in limbs or tail; 1, spontaneous tremor or obvious redness and swelling of hindlimb toes; 2, unilateral hindlimb dragging or persistent toe‐curling during walking; 3, bilateral hindlimb lameness, rigid paralysis, minimal joint movement, or foot not used for forward motion; 4, failure to right itself within 15 s when placed on either side.
Rotarod Test
4.5
To evaluate motor coordination in mice, the rotarod test was performed according to the standardized protocol from the International Mouse Phenotyping Consortium (IMPC, www.mousephenotype.org). During the acclimation phase, mice were placed on a rotarod apparatus at a constant speed of 4 rpm for 1 min. After a 15‐min rest period, mice were repositioned on the rotarod, and the apparatus was uniformly accelerated from 4 rpm to 40 rpm over a 300‐s period. The rotational speed at which the mouse fell off the rod was recorded. Each mouse underwent three consecutive trials with 15‐min intervals between trials, and the average speed across these trials was calculated as the final value.
Screen Test
4.6
To evaluate limb and hindlimb muscle strength and endurance in mice, Kondziela's inverted screen test was performed as previously described [74]. Briefly, mice were placed on a 40 cm × 30 cm wire mesh composed of 1 cm squares formed by 1‐mm diameter wires. The grid was then inverted upside down, and the latency to fall was recorded with a maximum cutoff time of 120 s to prevent exhaustion‐induced injury.
LPC/Myelin Injection
4.7
To assess microglial migration in vivo, mice were deeply anesthetized with 80 mg/kg sodium pentobarbital via intraperitoneal injection and fixed on a stereotactic apparatus. After disinfecting and exposing the target skull area, a hole was drilled above the injection site. A 10 µL Hamilton syringe was used to deliver lysolecithin (LPC, Sigma, L4129) or myelin. 1% LPC was sonicated and injected into the corpus callosum (coordinates: 0.5 mm anterior, 0 mm lateral, 2.3 mm ventral relative to the bregma). Denatured myelin was prepared as previously reported [23], and injected into the primary motor cortex (coordinates 1 mm anterior, 1 mm lateral, 1.5 mm ventral relative to the bregma). The needle was left in place for 5 min before injecting the solution at a rate of 1 µL every 5 min. After another 5‐min pause, the needle was withdrawn slowly. The scalp was sutured, and mice were placed on a heating pad for recovery. Mice were euthanized 4 h or 3 days post‐injection for tissue collection.
To assess cellular proliferation including microglia, 5‐bromo‐2'‐deoxyuridine (BrdU, Sigma, B5002) was applied in the myelin injury experiments. In the acute injury model, each mouse received a single intraperitoneal injection of 100 mg/kg BrdU immediately after myelin injection and were sacrificed 4 h post‐injury for tissue collection. In the prolonged injury model, mice were administered 50 mg/kg BrdU intraperitoneally every 24 h following myelin injection and were sacrificed 72 h post‐injury for tissue analysis.
Tissue Preparation
4.8
To prepare tissue sections, mice were deeply anesthetized via intraperitoneal injection of sodium pentobarbital at 80 mg/kg body weight and secured supine on a foam pad. The thoracic cavity was exposed using scissors, followed by transcardial perfusion with ice‐cold 0.01 M phosphate‐buffered saline (PBS) for 2–3 min and ice‐cold 4% paraformaldehyde (PFA) in 0.1 M PB for 7 min. The brains, spinal cords, and tibialis anterior muscles were collected and post‐fixed in 4% PFA at 4°C for 2 h. Then tissues were dehydrated in 30% sucrose in 0.1 M PB at 4°C for 3 days. For brain and spinal cord regions of interest, they were embedded in optimal cutting temperature compound (OCT, SAKURA, 4583), cut with a cryostat (Leica, CM1950) at a thickness of 12 µm, and stored at −80°C. For tibialis anterior muscles intended for weight measurement and neuromuscular junction (NMJ) imaging, the dehydrated muscles were blotted dry on filter paper and weighed, They were then embedded in OCT, sectioned at 70 µm thickness with the cryostat, and stored at −80 °C.
Immunofluorescence Staining on Tissues
4.9
For immunofluorescence staining, tissue sections were washed three times with PBS and blocked for 2 h in PBS containing 5% donkey serum and 0.3% Triton X‐100 at room temperature. Then, sections were incubated with primary antibodies diluted in PBS with 2.5% donkey serum and 0.3% Triton X‐100 at 4°C for 48 h, followed by three PBS washes. Subsequently, sections were incubated with fluorophore‐conjugated secondary antibodies (1:500) at 4°C for 2 h and washed three times with PBS. Nuclei were counterstained with 4′,6‐diamidino‐2‐phenylindole (DAPI, 1 µg/mL) at room temperature for 10 min, followed by three additional PBS washes. Excess liquid was carefully aspirated, and sections were mounted with FluorSave reagent (Millipore, 345789) under coverslips. Slides were cured in the dark at room temperature for 2 h. Images were taken by Olympus BX53 microscope, Olympus BX63 microscope or FV1000 confocal microscope, and analyzed by Image J.
To visualize neuromuscular junctions (NMJ), the concentration of Triton X‐100 was increased to 0.5% throughout the procedure to enhance permeabilization. Sections were incubated with CF488A‐conjugated α‐Bungarotoxin (Biotium, 00005; 1:500) diluted in PBS containing 2.5% donkey serum and 0.5% Triton X‐100 for 30 min at room temperature. After three washes with PBS, the sections were incubated with primary antibody. For BrdU immunostaining, sections underwent antigen retrieval in a microwave heating in citrate buffer (pH 6.0) for 10 min. This was followed by treatment with 2 M HCl for 30 min at room temperature and three PBS washes before blocking. For the detection of HVCN1, TGF‐β and SOX10, antigen retrieval was performed in a microwave heating in citrate buffer (pH 6.0) for 10 min, followed by three PBS washes prior to blocking.
Primary antibodies included goat anti‐Iba1 (Abcam, ab5076, 1:500), rabbit anti‐Iba1 (Wako, 019‐19741, 1:1000), rat anti‐CD11b (AbD serotec, MCA711, 1:100), chicken anti‐GFAP (Millipore, AB5541, 1:1000), mouse anti‐iNOS (BD Biosciences, 610329, 1:100), rabbit anti‐TGFβ (R&D Systems, AB‐100‐NA, 1:200), Rabbit anti‐Ki67 (Thermo, RM‐9106‐S, 1:500), rabbit anti‐HVCN1 (Origene, TA328862, 1:200), rabbit anti‐degraded MBP (Millipore, AB5864, 1:4000), mouse anti‐B8H10 (Medimabs, MM0070P, 1:200), goat anti‐ChAT (Millipore, AB144P), rat anti‐BrdU (Abcam, ab6326, 1:500), chicken anti‐NFH (abcam, ab4680, 1:5000), goat anti‐SOX10 (R&D Systems, AF2864, 1:200), guinea pig anti‐TMEM119 (Oasis Biofarm, OB‐PGP072, 1:500).
Fluorophore‐conjugated secondary antibodies included DyLight 405‐conjugated donkey anti‐rabbit (Jacksonimmunoresearch, 711‐475‐152), Alexa Fluor 488‐conjugated donkey anti‐rabbit (Jacksonimmunoresearch, 711‐545‐152), Alexa Fluor 488‐conjugated donkey anti‐rat (Jacksonimmunoresearch, 712‐545‐153), Alexa Fluor 488‐conjugated donkey anti‐mouse (Jackson ImmunoResearch, 715‐545‐150), Alexa Fluor 488‐conjugated donkey anti‐goat (Jackson ImmunoResearch, 705‐545‐003), Alexa Fluor 488‐conjugated donkey anti‐chicken (Jackson ImmunoResearch, 703‐545‐155), Alexa Fluor 488‐conjugated donkey anti‐guinea pig (Jackson ImmunoResearch, 706‐545‐148), Cy3‐conjugated donkey anti‐rat (Jackson ImmunoResearch, 712‐165‐153), Cy3‐conjugated donkey anti‐mouse (Jackson ImmunoResearch, 715‐165‐150), Cy3‐conjugated donkey anti‐rabbit (Jackson ImmunoResearch, 711‐165‐152), Cy3‐conjugated donkey anti‐goat (Jackson ImmunoResearch, 705‐165‐147), Cy3‐conjugated donkey anti‐chicken (Jackson ImmunoResearch, 703‐165‐155) Cy3‐conjugated donkey anti‐ guinea pig (Jacksonimmunoresearch, 706‐165‐148), Alexa Fluor 647‐conjugated donkey anti‐rabbit (Jacksonimmunoresearch, 711‐605‐152), Alexa Fluor 647‐conjugated donkey anti‐rat (Jacksonimmunoresearch, 712‐605‐153). Validation data for each antibody are given in the corresponding manufacturer's website.
Imaging Processing and Quantitative Analysis for In Vivo Assays
4.10
To assess ALS pathology or HVCN1 expression in the lumbar spinal cord of ALS mice, at least five randomly selected images from each region of interest per mouse were analyzed using ImageJ. The average value across these images is presented.
To evaluate the distribution of microglia around motor neurons in the lumbar spinal cord, z‐stack images (0.5 µm intervals) were acquired using a 60× oil‐immersion objective on an FV1000 confocal microscope and processed using Image J software. A z‐projected trace image was generated using non‐extended tracing format. A series of concentric circles with 10 µm radial increments were overlaid onto the center of motor neuron somata, identified by ChAT immunostaining co‐localized with DAPI. The number of microglial somata (Iba1⁺/DAPI⁺) within each ring was quantified. For each mouse, at least 20 randomly selected motor neurons were analyzed, and the average number of microglia per ring is presented.
Microglial migration was assessed in images acquired with a 40× objective on an Olympus BX63 microscope and processed using Image J software. The central lesion area was delineated based on regions of myelin or dMBP accumulation accompanied by adjacent clusters of amoeboid microglia/macrophages. The peripheral lesion area was defined as the region surrounding the core lesion exhibiting significantly elevated density of both DAPI intensity and microglia/macrophage staining. The somata of microglia (TMEM119⁺/Iba1⁺/DAPI⁺) and macrophages (TMEM119^−^/Iba1⁺/DAPI⁺) were quantified in each region. Cell density was determined via dividing the cell count by the corresponding area. A minimum of five randomly selected images per mouse were analyzed. and the average number of microglia per mouse is presented.
NMJ integrity was assessed as previously reported [75]. Briefly, z‐stack images (1 µm intervals) were acquired with a 40× oil‐immersion objective on an FV1000 confocal microscope and processed using Image J software. Z‐projected trace images were generated in non‐extended tracing format. NMJ innervations were quantified as the percentage of α‐bungarotoxin signal overlapping with the presynaptic signal. Endplates were classified as follows: fully innervated (≥60% overlap), partially innervated (20%–60% overlap), or fully denervated (<20% overlap). For each mouse, at least 30 randomly selected NMJs were analyzed.
Bone Marrow Derived Macrophage Isolation and Culture
4.11
To evaluated microglia phenotype by BMDMs in vitro, adult mice were euthanized with excess pentobarbital sodium and disinfected by immersion in 70% ethanol. Tibia and femur were isolated and washed three times in phosphate‐buffered saline (PBS). Both ends of the bones were clipped with scissors, and bone marrow cells were flushed out using a 1 mL syringe filled with PBS. The cell suspension was homogenized by pipetting with a 1 mL tip, filtered through a 300‐µm nylon mesh, and centrifuged at 300 g for 5 min. The supernatant was discarded, and the pellet was resuspended in 1 mL of red blood cell lysis buffer for 1–2 min at room temperature. Lysis was terminated by adding 3 mL Dulbecco's Modified Eagle Medium (DMEM, Vivacell, C3103), followed by centrifugation at 300 × g for 5 min. The cells were resuspended in DMEM medium supplemented with fetal bovine serum (FBS, Vivacell, C04001), 1% penicillin/streptomycin (P/S, Gibco, 15140122) and 10 ng/mL macrophage colony‐stimulating factor (M‐CSF, Sino biological, 51112‐MNAH) and plated in culture dishes. Cells were maintained at 37°C under 5% CO_2_. Half of the medium was replaced every 3 days. To induce HVCN1 knockout, tamoxifen was added to the culture medium at a final concentration of 4 µg/mL throughout the entire culture period. Cells were harvested on day 7 for downstream experiments.
Primary Glial Cell Culture and Purification
4.12
To obtain primary mouse glial cells, the P0 mice were disinfected with 75% alcohol and quickly decapitated. After cutting open the scalp and skull, the brain tissue was carefully removed with forceps and transferred to ice‐cold HBSS. Followed by peeling off the meninges and removing the olfactory bulb and cerebellum, the cerebral cortex was isolated on ice. Blood spots on the cortex were removed with forceps, and the cortex was transferred to a dish containing glia culture medium (DMEM/F12 supplemented with 20% FBS and 1% P/S). The tissue was minced with scissors and gently pipetted until no visible fragments remained. The mixture was transferred to poly‐D‐lysine (PDL, Sigma, P6407) precoated T75 flask and cultured at 37°C for 7–9 days with medium half‐changed every 3 days. For microglia isolation, the sealed T75 flask was placed on a shaker at 37°C and shaken at 200 rpm for 2 h. The supernatant was collected, centrifuged, and resuspended in DMEM containing 10% FBS, 1% P/S, and 10 ng/ml M‐CSF and microglia culture. For oligodendrocyte precursor cells (OPCs) and oligodendrocyte, the T75 flask was replenished with glia culture medium, sealed, and shaken at 250 rpm for 16 h. The supernatant was collected, centrifuged and resuspended in Neurobasal medium supplemented 1% N2, 2% B27, and 1% P/S. 10 ng/ml PDGFaa (R & D system, 221‐AA‐010) was added for OPC culture, whereas 40 ng/ml T3 (Sigma, T‐074) was added to induce differentiation into mature oligodendrocytes. For astrocytes, the T75 flask was washed once with PBS, digested with trypsin for 3 min. The cells were resuspended and cultured in DMEM containing 10% FBS and 1% P/S.
Primary Neuron Culture
4.13
To obtain embryonic hippocampal neuron, pregnant C57/BL6J mice at embryonic days 16–18 were euthanized via intraperitoneal injection of excess sodium pentobarbital. The uterine horns were excised and transferred to ice‐cold Hanks' Balanced Salt Solution. Under sterile conditions, embryos were decapitated, and fetal brains were rapidly isolated and placed in ice‐cold HBSS. After carefully removing the meninges, hippocampal tissues were dissected, minced, and digested in 0.25% trypsin‐EDTA at 37°C for 12 min. The digested tissues were made into single cell suspensions through fire polished 1 mL pipette in DMEM/F12 media (Gibco, 11320033) with 10% FBS (Vivacell, C04001). After filtering through a 70‐µm nylon mesh filter, Cell suspensions were counted with a hemocytometer, and adjusted to a density of 2 × 10^5^ cells/mL. Cells were seeded onto poly‐D‐lysine (PDL, Sigma, P6407)‐coated culture dishes. 12 h later, the media were replaced with neurobasal medium supplemented with 2% B27 (Thermo, 17504044) and 1% GlutaMAX (Thermo, 35050061), and maintained at 37°C. Half of the media were replaced every 3 days.
Isolation and Culture of Primary Microglia from Adult Mice CNS
4.14
Microglia were purified and cultured from ALS mice during disease progression according to previously described protocols with minor modifications [76, 77]. Briefly, 16‐week‐old symptomatic ALS mice received daily intraperitoneal injections of tamoxifen (100 mg/kg) for four consecutive days to induce HVCN1 knockout. 7 days post‐induction, mice were anesthetized with sodium pentobarbital (80 mg/kg) and transcardially perfused with ice‐cold PBS. Brains and spinal cords were rapidly dissected and placed in a pre‐chilled dish containing cold DMEM. The tissues were minced and mechanically dissociated using a Dounce homogenizer in supplemented with 1 U/mL DNase I (Thermo, EN0521) on ice. The homogenate was gently triturated with fire‐polished Pasteur pipettes, filtered through a 70‐µm cell strainer, and centrifuged at 400 × g for 5 min at 4°C. To remove myelin debris, an isotonic Percoll solution was prepared by mixing nine volume Percoll (GE Healthcare, 17‐0891‐09) with one volume 10× PBS, which was then diluted to 37% with DMEM. The cell pellet was resuspended in 37% Percoll and centrifuged at 400 × g for 40 min with slow acceleration and no brake. The upper myelin‐containing layer and Percoll was discarded, and the pelleted cells were washed twice with ice‐cold PBS. Cells were then incubated with rat anti‐mouse CD16/CD32 antibody (BD Biosciences, 553142; 1:200) in PBS containing 2% FBS for 10 min at 4°C to block Fc receptors. After washing in ice cold PBS, cells were stained with FITC‐conjugated anti‐CD45 (BD Biosciences, 561088; 1:200) and PE‐conjugated anti‐CD11b (BD Biosciences, 557397; 1:200) in PBS with 2% FBS for 30 min on ice, followed by two washes in ice cold PBS. Cells were resuspended in ice‐cold PBS with 2% FBS, and DAPI was added to a final concentration of 0.25 µg/mL immediately prior to sorting. Microglia (CD11b⁺CD45ˡ°ʷ) were sorted on a BD FACSAria II or Beckman MoFlo Astrios EQ using an 85‐µm nozzle. Sorted microglia were collected in DMEM supplemented with 20% FBS and 2% P/S and allowed to settle in an incubator at 37°C for 30 min. Cells were then centrifuged at 400 × g for 30 min at 4°C. For Western blot, cells were washed with PBS and lysed. For culture, cells were resuspended in DMEM containing 20% FBS, 2% P/S, and 10 ng/mL M‐CSF, and plated on poly‐D‐lysine‐coated dishes. After 24 h, the medium was replaced with DMEM containing 10% FBS, 1% P/S, and 10 ng/mL M‐CSF, with half of the medium being replaced every other day.
CCK8 Assay
4.15
For the in vitro assessment of microglial neuroprotection by cck8 assay, primary neurons were cultured in PDL coated 96 plate well for 8–10 days in vitro using Neurobasal medium supplemented with 2% B‐27 and 1% mM GlutaMAX. To minimize evaporation during culture, the outer perimeter wells were not used for cell culture but instead filled with 200 µL of PBS. BMDMs or microglia were prepared by discarding the original culture medium, washing the cells with warm PBS, and replacing the medium with fresh Neurobasal medium containing 2% B‐27 and 1× GlutaMAX. After 24 h, this conditioned medium was collected and centrifuged at 3000 × g for 5 min. The supernatant was then filtered through a 0.5 µm filter to obtain the final conditioned medium. Neurons in the 96‐well plates were treated with this conditioned medium along with 500 µM hydrogen peroxide for 24 h. Following treatment, the medium was replaced with Neurobasal medium containing 10% CCK‐8 reagent (APExBIO Technology, K1018) and incubated at 37°C in 5% CO_2_ for 4 h. Absorbance at 450 nm was measured using a SpectraMax iD5 microplate reader (Molecular Devices).
BMDM/Microglia‐Primary Neuron Co‐Culture
4.16
To evaluate the neuroprotective effect of microglia in vitro, primary neurons were cultured on PDL‐coated circular coverslips (13 mm) in 24‐well plates for 8–10 days in vitro using Neurobasal medium supplemented with 2% B‐27 and 1% mM GlutaMAX. Sterile 1 µm pore transwell inserts (Corning, 353104) were gently placed into the wells. BMDMs or microglia were harvested by scraping, centrifuged at 300 × g for 5 min, and resuspended in neurobasal medium supplemented with 2% B27 and 1% GlutaMAX. After cell counting, the density was adjusted to 2 × 10⁶ cells/mL. 200 µL of cell suspension was seeded into the upper transwell insert. To induce neuronal injury, hydrogen peroxide was added to the lower chamber at a final concentration of 500 µM to induce neuronal cell death. The coculture system was maintained at 37°C in 5% CO_2_ for 24 h. Following the treatment, the coverslips were washed with PBS and fixed with 4% PFA at room temperature for 15 min. After three additional washes with PBS, the samples were processed for standard immunofluorescence staining.
Immunofluorescence on Cells
4.17
For immunofluorescence staining of cells, cells plated on coverslips were washed with PBS, then fixed with 4% PFA for 15 min at room temperature. After three PBS washes, the coverslips were blocked with 5% donkey serum in PBS containing 0.3% Triton X‐100 for 1 h at room temperature. The coverslips were then incubated with primary antibodies for 12 h at 4°C, followed by three PBS washes. Subsequently, the coverslips were incubated with the fluorophore‐conjugated secondary antibodies (1:1000) for 1 h at 4°C. After three more PBS washes, the coverslips were stained with DAPI for 10 min at room temperature. Following three final PBS washes, the coverslips were mounted onto microscopy slides with FluorSave reagent. Images were captured using an Olympus BX53 or FV1000 confocal microscope and analyzed with ImageJ software.
Primary antibodies included chicken anti‐MAP2ab (Abcam, ab5392, 1:200), rabbit anti‐degraded MBP (Millipore, AB5864, 1:4000), rat anti‐Lamp2 (Abcam, ab13524, 1:500), biotin conjugate IB4 (Sigma, L2140, 1:1000), rabbit anti‐Iba1 (Wako, 019‐19741, 1:1000), guinea pig anti‐TMEM119 (Oasis Biofarm, OB‐PGP072, 1:500). Fluorophore‐conjugated secondary antibodies included Alexa Fluor 488‐ conjugated donkey anti‐chicken (Jacksonimmunoresearch, 703‐545‐155), Alexa Fluor 488‐conjugated donkey anti‐rabbit (Jacksonimmunoresearch, 711‐545‐152), Cy3‐conjugated donkey anti‐ guinea pig (Jacksonimmunoresearch, 706‐165‐148), Cy3‐conjugated donkey anti‐rat (Jacksonimmunoresearch, 712‐165‐153), Alexa Fluor 647‐conjugated Streptavidin (Jacksonimmunoresearch, 016‐600‐084). Validation data for each antibody are given in the corresponding manufacturer's website.
Quantification of Surviving Neurons and Dendritic Arborization
4.18
Surviving neurons were identified and quantified based on the co‐localization of DAPI with MAP2ab immunoreactivity and the presence of at least two neurites. In each experimental group, counts were performed on a minimum of eight randomly selected images, and the average value was presented.
Dendritic branching complexity was assessed using the Sholl analysis plugins in Image J software [78]. Briefly, z‐stack images (0.5 µm intervals) were acquired with a 60× oil‐immersion objective on an FV1000 confocal microscope. Non‐extended tracing was applied to generate a z‐projected 8‐bit trace image, wherein the soma and processes of individual neurons were carefully outlined to exclude adjacent cells and non‐specific immunoreactivity. A series of concentric circles with 10 µm radial increments were superimposed onto the center of the digitized soma. The number of intersections between dendritic processes and each concentric circle was counted. In each experimental group, at least 12 randomly selected neurons were analyzed.
Flow Cytometry
4.19
To assess cell polarization, cells were washed once with ice‐cold PBS, centrifuged, and resuspended in 100 µL of PBS containing flow cytometry antibodies diluted at a 1:200 ratio. Following gentle vortex, the samples were incubated at 4°C for 30 min in the dark. For ROS detection, cells were incubated with DCFH‐DA (Beyotime, S0033S, 1:500) in DMEM medium for 30 min at 37°C. To detect the lysosome function, cells were incubated with LysoTracker Red DND‐99 (Thermo, A66466, 1:20, 000) or LysoSensor Green DND‐189 (Thermo, A66463, 1:1000) in DMEM medium for 30 min at 37°C. After incubation, cells were washed twice with PBS, resuspended in 500 µL PBS, and filtered through a 70 µm nylon mesh. Finally, the cells were analyzed using a Cytoflex S or Cytoflex LX flow cytometer. Subsequent analysis of the acquired data was performed with FlowJo v10 software (BD Life Sciences). Flow cytometry antibodies included FITC rat anti‐CD11b (BD Biosciences, 557396), rat BV421 rat anti‐CD86 (BD Biosciences, 564198), APC rat anti‐CD206 (BD Biosciences, 565250).
Transwell Assay
4.20
To assess cell chemotaxis, 0.6 mL DMEM with 10% FBS and 10 ng/mL M‐CSF were carefully added to 24‐well plate without bubbles. Transwell inserts (Corning, 3422) with 8 µm pores were gently placed in the wells. 2 × 10^5^ viable cells were suspended in DMEM with 0.2% FBS and seeded into the upper chamber of each insert. The system was maintained at 37°C in 5% CO_2_ for 24 h. Then, the inserts were fixed in 4% ice‐cold PFA for 15 min and stained with 0.1% crystal violet solution. Cells above the insert were carefully wiped. Migrated cells below the insert were imaged using Olympus BX53 microscope and analyzed by Image J. In each experimental group, cell counts were performed on at least eight randomly selected images, and the average value was presented.
In Vitro Phagocytosis Assay
4.21
To assess phagocytosis ability in vitro, BMDMs were scraped, counted, and replated onto circular coverslips (13 mm) in 24‐well plates at a density of 30 000 cells per well. After 24 h, the culture medium was replaced with DMEM containing 5 µg/mL myelin debris and incubated at 37 °C for 1 h to allow phagocytosis occur. Following incubation, cells were washed three times with ice‐cold PBS, fixed with 4% PFA for 15 min, and washed again with PBS before being processed for standard immunofluorescence staining.
Western Blot
4.22
To quantify protein levels in vitro, cells were washed with ice‐cold PBS and lysed in Lysis III buffer (50 mM Tris‐Cl pH 7.4, 50 mM NaCl, 5 mM EDTA, 60 mM CHAPS, 0.5% sodium deoxycholate, 1% SDS, 1% NP‐40, 1% Triton X‐100) supplemented with 1 mM DTT, protease inhibitor cocktail (Roche life science, 4693132001), phosphatase inhibitor cocktail (Biomake, B15001), 2 µM MG132 (Millipore, 474790) on ice for 30 min. The lysate was centrifuging at 12 000 g for 10 min at 4°C. The supernatants were quantified using the BCA assay and denatured in 5×SDS buffer. Equal amounts of protein and a protein ladder were loaded onto a 10% SDS‐PAGE gel. The separated proteins were transferred onto 0.22 µm polyvinylidene difluoride membranes (Millipore, ISEQ00010), blocked with 5% milk in TBST for 2 h, and incubated with primary antibodies diluted in 3% milk in TBST overnight at 4 °C. After 6 TBST washes, the blots were stained with HRP‐conjugated secondary antibodies in 3% milk in TBST for 1 h at room temperature. Followed by 6 more TBST washes. The blots were visualized by ECL kit (FdBio science, FD8020). Images were acquired by ChemiDoc Touch Imaging System (Bio‐Rad).
Primary antibodies included Rabbit anti‐HVCN1 (Origene, TA328862, 1:1000), Rabbit anti‐Akt (Huabio, HA721870, 1:2000), Rabbit anti‐phospho‐Akt (Vazyme, RA2102, 1:2000). Fluorophore‐conjugated secondary antibodies included HRP conjugated Donkey anti‐rabbit (Jackson ImmunoResearch, 711‐035‐152), HRP conjugated Donkey anti‐mouse (Jackson ImmunoResearch, 715‐035‐151). Validation data for each antibody are given in the corresponding manufacturer's website.
Bulk RNA Sequencing
4.23
For bulk RNA sequencing, cells were seeded into 6‐well plates 24 h before RNA extraction. After removing the supernatant, cells were lysed in situ with 1 mL TRIzol reagent (Invitrogen, 15596‐018). Total RNA was isolated and measured by a Nanodrop 2000 spectrophotometer. RNA integrity was assessed by agarose gel electrophoresis and the Agilent 2100 Bioanalyzer. cDNA libraries were constructed and sequenced on the Illumina platform by Majorbio Biotech. The raw paired‐end reads underwent trimming and quality control using fastp software with default parameters. Subsequently, clean reads were aligned separately to the mouse reference genome GRCm38.p6 using HISAT2 software. StringTie was employed in a reference‐based approach to assemble the mapped reads from each sample. Differential expression analysis (DEGs) was performed using DESeq2 with significance thresholds set at p_adjust < 0.05 and |log_2_ fold change| ≥ 1). For network analysis, all DEGs (|log_2_FC| > 1, p_ajdust value < 0.05) was intersected with the major Gene Ontology term for cell migration (GO:0016477). The resulting gene set was imported into the STRING database (www.string‐db.org) to construct a protein‐protein interaction network, which was subsequently visualized and analyzed using Cytoscape 3.10.Three biological replicates from each group were used for RNA‐seq analysis.
RNA Isolation and Quantitative Real‐Time PCR
4.24
To quantify gene transcription level in vitro, total RNA was isolated from cells using TRIzol reagent (Invitrogen, 15596‐018) and the Quick RNA Purification kits according to the manufacturer's instructions, cDNA was synthesized using the HiScript II Q RT SuperMix kit (Vazyme, R223). qRT‐PCR analysis was performed on a Biorad CXF96 instrument using ChamQ SYBR qPCR Master Mix (Vazyme, Q311). The primers used for qPCR were listed in Supporting Table S3. The Gapdh gene was used as a reference gene for normalization, and relative expression levels were calculated using the 2^−ΔΔCT^ method. Each experiment was conducted in triplicate.
Quantification and Statistical Analysis
4.25
Data analysis for quantification was performed by using Image J and Graphpad prism 8.0 software. Statistical tests were Log‐rank (Mantel‐Cox) test, two way ANOVA with Sidak's multiple comparisons test, one‐way ANOVA with Tukey's multiple comparisons test, one way ANOVA with Dunnett's multiple comparisons test, unpaired two‐tailed t‐test, Data were presented as mean ± S.E.M., and p < 0.05 was considered as statistically significant. The details of statistical tests used are indicated in the figure legends.
Author Contributions
J.Z. perceived and supervised this project. J.Z. and F.W. designed the experiment. F.W., K.Z., L.Z., W.L., Y.W., X.M., J.W., Z.D., X.G., D.W., and X.Y. performed the experiments. F.W., L.Z., X.G., X.Y., Z.Z., S.W., Z.J., and L.H. analyzed the data. J.Z. and F.W. wrote the manuscript. All authors have read and approved the final manuscript.
Conflicts of Interest
The authors declare no conflicts of interest.
Supporting information
Supporting File 1: advs73679‐sup‐0001‐SuppMat.docx.
Supporting File 2: advs73679‐sup‐0002‐SuppMat.zip.
Supporting File 3: advs73679‐sup‐0003‐xlsx.zip.
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