APOE-mediated lipid transport prevents C1q-related synaptic loss after spinal cord injury via attenuating macrophage lipid stress.

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Learn more: PMC Disclaimer | PMC Copyright Notice J Neuroinflammation . 2026 Mar 6;23:120. doi: 10.1186/s12974-026-03756-9 Search in PMC Search in PubMed View in NLM Catalog Add to search APOE-mediated lipid transport prevents C1q-related synaptic loss after spinal cord injury via attenuating macrophage lipid stress Yersen Mulat Yersen Mulat 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Yersen Mulat 1, # , Zun Ren Zun Ren 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Zun Ren 1, # , Chaocao Nong Chaocao Nong 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Chaocao Nong 1, # , Chuanliang Fu Chuanliang Fu 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Chuanliang Fu 1 , Yilin Huo Yilin Huo 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Yilin Huo 1 , Mai Zhao Mai Zhao 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Mai Zhao 1 , Yahui Dai Yahui Dai 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Yahui Dai 1 , Canyu Chen Canyu Chen 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Canyu Chen 1 , Peilin Wang Peilin Wang 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Peilin Wang 1 , Renyuan Wang Renyuan Wang 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Renyuan Wang 1 , Hao Zhang Hao Zhang 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Hao Zhang 1 , Yi Hu Yi Hu 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Yi Hu 1 , Peng Lai Peng Lai 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Peng Lai 1 , Ruoyi Guo Ruoyi Guo 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Ruoyi Guo 1 , Dongsheng Jiang Dongsheng Jiang 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China 2 Precision Research Center for Refractory Diseases, Pioneer Research Institute for Molecular and Cell Therapies, Shanghai General Hospital, Shanghai Jiao Tong University, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Dongsheng Jiang 1, 2, ✉ , Ying Peng Ying Peng 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Ying Peng 1, ✉ , Haodong Lin Haodong Lin 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China Find articles by Haodong Lin 1, ✉ Author information Article notes Copyright and License information 1 Trauma Center, Shanghai General Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China 2 Precision Research Center for Refractory Diseases, Pioneer Research Institute for Molecular and Cell Therapies, Shanghai General Hospital, Shanghai Jiao Tong University, Shanghai Jiao Tong University School of Medicine, Shanghai, 201620 China ✉ Corresponding author. # Contributed equally. Received 2025 Nov 25; Accepted 2026 Feb 25; Collection date 2026. © The Author(s) 2026 Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if you modified the licensed material. You do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by-nc-nd/4.0/ . PMC Copyright notice PMCID: PMC13077989  PMID: 41792769 Abstract Following spinal cord injury (SCI), neuroinflammation driven by lipid-laden macrophage foam cells is a key pathology, yet how these cells manage their lipid homeostasis is unclear. We delineate a neuroprotective axis in which macrophages deploy apolipoprotein E (APOE) to transfer intracellular lipids to neighboring cells, especially fibroblasts. Genetic ablation of Apoe disrupts this intercellular lipid transport, culminating in pathological lipid retention that activates the Hippo signalling cascade and transcriptionally induces complement component C1q. This excess C1q aberrantly tags intact synapses for excessive microglial pruning, leading to significant synaptic loss and impaired locomotor function recovery. Direct blockade of C1q using neutralizing antibodies recapitulated these neuroprotective effects, confirming C1q as the critical mediator. Crucially, macrophage-specific APOE re-expression reverses this entire cascade, preserving synapses and restoring locomotor function (BMS score: 4.81 ± 0.21 ( Apoe ) vs. 1.75 ± 1.28 ( NC ); Incline plane: 69.24° ± 2.33° ( Apoe ) vs. 51.66° ± 5.14° ( NC ) in Apoe −/− mice). These findings identify the APOE-Hippo-C1q pathway in macrophages as a novel therapeutic target for SCI. Supplementary Information The online version contains supplementary material available at 10.1186/s12974-026-03756-9. Introduction Macrophages, pivotal effectors of innate immunity, execute dual functions in pathogen clearance via phagocytosis and immunomodulation through cytokine signaling [ 1 ]. Following spinal cord injury (SCI), peripheral circulating monocytes infiltrate the damaged site through the disrupted blood-spinal barrier and differentiate into mature macrophages [ 2 , 3 ]. Macrophages are responsible for clearing cellular debris post SCI, including lipid-rich myelin fragments [ 4 , 5 ]. After phagocytizing myelin debris, macrophages degrade it in lysosomes and produce a large amount of lipid droplets (LDs), leading to the formation of foam cells [ 6 , 7 ]. In studies of human autopsy tissues, it was found that lipid-rich macrophages still existed at the lesion site one year after SCI [ 8 ]. In peripheral chronic inflammatory diseases such as atherosclerosis, the formation of foam cells is generally considered a pathological hallmark that drives disease progression [ 9 , 10 ]. However, in the unique context of central nervous system (CNS) injury, the functional consequences of this myelin-induced foam cell formation, and how these cells dispose of their immense internal lipid load, remain unclear. Apolipoprotein E (APOE), as the primary lipid transporter in the CNS, plays a central role in regulating the transport and metabolism of cholesterol and phospholipids in the brain [ 11 , 12 ] In humans, the APOE gene has three primary alleles: ε2 , ε3 , and ε4 [ 11 , 13 ]. APOE ε4 is the most significant genetic determinant of risk for neurodegenerative disorders, notably Alzheimer’s disease. APOE plays a pivotal role in pathogenesis by modulating key pathological pathways, such as Aβ and tau, neuroinflammatory responses, and brain lipid metabolism [ 14 – 16 ]. Although the function of APOE has been extensively studied in neurodegenerative diseases, its specific role and molecular mechanisms in regulating macrophage lipid metabolism after SCI, and its effects on synaptic plasticity and locomotor function recovery, remain to be elucidated. This study aims to fill this critical knowledge gap. We found that after SCI, macrophages actively transfer their intracellular LDs to adjacent cells, especially fibroblasts, through an APOE-dependent pathway. We further demonstrate that the absence of APOE disrupts this process, leading to the pathological retention of LDs within macrophages. Crucially, this intracellular lipid stress drives a significant upregulation in the expression of C1q by activating the Hippo signaling pathway. Excess C1q, in turn, tags intact synapses in the peri-lesional area, inducing their excessive engulfment and clearance by microglia, ultimately leading to a substantial loss of synapses and severe locomotor dysfunction. Finally, by specifically re-expressing APOE in the macrophages of Apoe −/− and wild type mice, we successfully reversed lipid retention, suppressed C1q upregulation, preserved synaptic density, and significantly promoted functional recovery. Our work reveals a signaling pathway connecting macrophage lipid metabolism, complement-mediated synaptic pruning, and locomotor function recovery. This suggests that targeting the APOE-Hippo-C1q pathway in macrophages could be a promising novel therapeutic strategy to promote repair after SCI. Results Macrophages-to-fibroblasts LDs transport dominates after SCI To determine the fate of LDs within macrophages following SCI, we established an in vivo LDs tracing model (Fig. 1 A). In the experiment group, bone marrow-derived macrophages (BMDMs) were differentiated into foam cells by treating them with complete medium (CM) containing 1 mg/mL spinal cord homogenate (Hom) for 24 h in vitro. The intracellular LDs of these foam cells were fluorescently labeled with BODIPY (green), and their cell membranes were labeled with DiD (blue) to track them post-transplantation (BMDM@DiD@LDs@BODIPY). BMDMs in the control group were treated with CM containing PBS and stained with BODIPY and DiD (BMDM@DiD). These cells were then immediately injected into the lesion site of C57BL/6J mice following a T10 spinal cord crush model, with sham mice serving as uninjured controls. To validate the successful engraftment and labeling strategy, spinal cord sections at 3 and 7 days post-injury (dpi) were stained for the macrophage marker F4/80 [ 17 ]. As expected, tissues from the control group exhibited F4/80 + DiD+ cells, whereas tissues from the experiment group displayed F4/80 + BODIPY + DiD+ cells, confirming the presence of lipid-laden macrophages (Fig. 1 B). Fig. 1. Open in a new tab Macrophages transfer lipid droplets to fibroblasts following spinal cord injury. A Schematic of the in vivo LDs tracing experiment. BMDMs were labeled with DiD (cell membrane, blue) and BODIPY (LDs, green) before being transplanted into the lesion site of mice after T10 crush injury. Spinal cord tissues were collected at 3 and 7 dpi. B Representative immunofluorescence images confirming the successful engraftment of DiD-labeled control BMDMs (BMDM@DiD, upper) and DiD and BODIPY-labeled foam cells (BMDM@DiD@LDs@BODIPY, lower) at the lesion site. Cells were co-stained for the macrophage marker F4/80 (red). Scale bar, 100 μm. C Immunofluorescence images showing BODIPY + LDs (green) within PDGFRα + fibroblasts (red) in the lesion at 3 and 7 dpi. Recipient cells are identified as BODIPY + DiD-. Scale bars, 100 μm (upper), 10 μm (lower). D Quantification of the percentage of PDGFRα + fibroblasts containing BODIPY + LDs at 3 and 7 dpi. Data are presented as mean ± SD, n = 3 per group, two-tailed Student’s t-test. E Schematic of the in vitro transwell co-culture system designed to assess LDs transfer from BMDMs to L929 fibroblasts without direct cell contact. F Representative images showing significant accumulation of BODIPY + LDs (green) in L929 fibroblasts (PDGFRα, red) after 24 h co-culture with BMDM@LDs@BODIPY, but not with control BMDMs. Scale bars, 100 μm (upper), 10 μm (lower). G Quantification of the BODIPY+ area within L929 fibroblasts. Data are presented as mean ± SD, n = 5 per group, two-tailed Student’s t-test. BMDMs Bone marrow derived macrophages. LDs Lipid droplets. dpi days post injury We next sought to identify recipient cells of the labeled LDs, which would be characterized by the BODIPY + DiD- cells. We observed that PDGFRa+ fibroblasts within the lesion clearly contained BODIPY + LDs at both 3 and 7 dpi (Fig. 1 C) [ 18 ]. Notably, the accumulation of these lipids within fibroblasts was more pronounced at 7 dpi (28.04%±2.48%) compared to 3 dpi (19.48%±1.18%) (Fig. 1 D). LDs transfer was also observed in other cells, including GFAP+ astrocytes (6.47%±0.28% at 3dpi, 10.62%±3.5% at 7dpi) and NeuN+ neurons (3.33%±5.77% at 3dpi, 10.36%±2.91% at 7dpi), at these time points (Supplementary Fig. 1A-D). To determine the baseline profile of lipid trafficking in the absence of injury, we performed the same adoptive transfer of BMDM@LDs@BODIPY into the spinal cords of Sham mice. In this uninjured environment, we observed a basal level of lipid transfer from macrophages to resident neural cells. Specifically, quantitative analysis revealed that fibroblasts contained macrophage-derived lipid droplets at both 3 dpi (20.57% ± 3.31%) and 7 dpi (19.77% ± 3.33%) (Supplementary Fig. 1E, F). We also detected lipid uptake in GFAP+ astrocytes (9.91% ± 0.93% at 3 dpi; 8.39% ± 2.62% at 7 dpi) (Supplementary Fig. 1G, H) and NeuN+ neurons (10.68% ± 5.86% at 3 dpi; 12.01% ± 4.23% at 7 dpi) (Supplementary Fig. 1I, J). These findings provide a quantitative baseline for macrophage-mediated lipid redistribution, serving as a control for the injury-induced changes observed in the SCI model [ 19 , 20 ]. However, these cells internalize a markedly smaller proportion of LDs compared with fibroblasts. To further confirm that macrophages can directly transfer LDs to fibroblasts, we designed an in vitro co-culture experiment using a transwell system with 0.4 μm pores, preventing direct cell-to-cell contact (Fig. 1 E). L929 fibroblasts were seeded in the bottom chamber, while the top chamber was seeded with either control BMDMs or BMDM@LDs@BODIPY that had been thoroughly washed to remove extracellular LDs [ 21 ]. After 24 h of co-culture, immunofluorescence staining revealed that L929 fibroblasts co-cultured with BMDM@LDs@BODIPY showed significant intracellular accumulation of BODIPY + LDs. In contrast, no LDs was detected in fibroblasts co-cultured with control BMDMs (Fig. 1 F and G). Collectively, these in vivo and in vitro results demonstrate that foam cells effectively transfer their intracellular LDs to adjacent fibroblasts and other neural cells after SCI. APOE mediates the transport of LDs from macrophages to fibroblasts Given that APOE is the primary lipid transporter in the CNS, we hypothesized that it mediates the lipid transfer from macrophages to fibroblasts [ 22 , 23 ]. To test this hypothesis, we first performed immunofluorescence staining on spinal cord sections from our in vivo tracing model (Fig. 1 A). We observed that extracellular BODIPY + LDs were frequently co-localized with APOE protein, suggesting that the LDs are packaged into APOE-containing lipoprotein particles for transport (Fig. 2 A). Furthermore, PDGFRa+ fibroblasts also co-expressed the low-density lipoprotein receptor (LDLR), a key receptor for APOE-lipoproteins, and were situated in close proximity to APOE (Fig. 2 B) [ 11 ]. These findings implicate APOE in the intercellular transport of LDs from macrophages to fibroblasts. Fig. 2. Open in a new tab APOE is essential for the transport of LDs from macrophages to fibroblasts. A Representative image showing co-localization of extracellular BODIPY + LDs (green) with APOE protein (red) in the BMDM@DiD@LDs@BODIPY group. Scale bars, 100 μm (left), 10 μm (right). B Image showing PDGFRα + fibroblasts (blue) expressing the LDLR (green) in close proximity to APOE (red) at 11 dpi. Scale bar, 100 μm (left), 10 μm (right). C Time-course analysis of Apoe RNA (red) and APOE protein (blue) expression in relation to F4/80 + macrophages (green) at 7, 11, and 14 dpi. Dotted lines delineate the lesion epicenter and penumbra. Scale bars, 200 μm (upper), 100 μm (lower). D Quantification of APOE protein IntDen and Apoe mRNA IntDen expression ratios between the penumbra and epicenter. Data are presented as mean ± SD, n = 5, two-tailed Student’s t-test. E Images showing significantly higher accumulation of BODIPY + LDs (green) within F4/80 + macrophages (red) in Apoe −/− mice compared to WT controls at 11 dpi. Scale bar, 1 mm. F Quantification of the BODIPY+ area within the F4/80 + macrophage area. Data are presented as mean ± SD, n = 6, two-tailed Student’s t-test. G Images showing reduced accumulation of BODIPY + LDs (green) within PDGFRα + fibroblasts (red) in Apoe −/− mice compared to WT mice at 11 dpi. Scale bar, 100 μm. H Quantification of the BODIPY+ area within the PDGFRα + fibroblast area. Data are presented as mean ± SD, n = 6, two-tailed Student’s t-test. Intden, Integrated Density We next investigated the temporal and spatial dynamics of APOE expression after SCI. Quantitative analysis showed that APOE protein levels began to increase at 7 dpi, peaked at 11 dpi, and subsequently declined by 14 dpi (Supplementary Fig. 2A, B). To identify the cellular source of APOE, we combined RNA scope for Apoe mRNA with immunofluorescence for F4/80 and APOE protein. Apoe mRNA was predominantly localized within F4/80 + macrophages in the lesion epicenter at 7, 11, and 14 dpi, with the epicenter-to-penumbra ratio remaining stable at 1.63 ± 0.36 (7 dpi), 1.61 ± 0.27 (11 dpi), and 1.62 ± 0.43 (14 dpi). In contrast, the ratio of APOE protein in the penumbra versus the epicenter progressively increased over time, indicating that APOE is primarily synthesized by macrophages at the epicenter and secreted to the penumbra (Fig. 2 C and D). To directly test the functional necessity of APOE in this process, we performed SCI surgeries in both wild-type (WT) and Apoe knockout ( Apoe −/− ) mice. At 11 dpi, the peak of APOE expression, we observed the significant higher accumulation of BODIPY + LDs within F4/80 + macrophages in Apoe −/− mice compared to WT controls (Fig. 2 E and F). This suggests an impaired lipid efflux from macrophages in the absence of APOE. We confirmed this conclusion in vitro using our transwell co-culture system (Supplementary Figs. 2 C). After 24 h of culture, foam cells derived from Apoe −/− BMDMs retained substantially more intracellular LDs than their WT counterparts (Supplementary Figs. 2D and 2E). Critically, this lipid retention in donor macrophages resulted in a failure of lipid delivery to recipient cells. Immunofluorescences experiment at 11 dpi revealed a concomitant reduction in LDs accumulation within fibroblasts of Apoe −/− mice compared to WT mice (Fig. 2 G and H). Additionally, we found that in penumbra, astrocytes and neurons in Apoe −/− mice contained fewer intracellular LDs than those in WT mice (Supplementary Figs. 2 F-I). Taken together, these findings provide strong evidence that macrophages utilize APOE to package and transport intracellular LDs, which are then transferred primarily to neighboring fibroblasts in the lesion. APOE deficiency leads to a significant upregulation of complement C1q in macrophages To investigate the pathological consequences of APOE deficiency in macrophages, we performed a transcriptomic analysis of the spinal cord lesion at 11 dpi from both WT and Apoe −/− mice. We employed a dual strategy combining 10x Visium HD spatial transcriptomics with validation by single-cell RNA sequencing (scRNA-seq) and immunofluorescence experiments (Fig. 3 A). For the spatial analysis, we utilized the Bin2cell algorithm, which specifically analyzes nuclear transcripts to minimize cross-contamination between adjacent cells [ 24 ]. This analysis confirmed the absence of Apoe expression in the Apoe −/− sections and its localization to the lesion epicenter in WT mice (Supplementary Fig. 3A). Fig. 3. Open in a new tab APOE deficiency drives upregulation of complement C1q in macrophages. A Schematic overview of the 10x Visium HD spatial transcriptomics workflow. B Spatial distribution of six major cell types identified in spinal cord sections from WT and Apoe −/− mice at 11 dpi. n = 1 for the WT group and n = 1 for the Apoe −/− group ( C )Spatial plot highlighting the distribution of macrophages in WT and Apoe −/− sections. D Heatmap of differentially expressed genes between macrophages from Apoe +/+ and Apoe −/− mice, showing significant upregulation of complement component genes ( C1qa , C1qb ) in the absence of APOE. E Gene Ontology (GO) analysis of upregulated genes in Apoe −/− macrophages, revealing enrichment for complement-related pathways and “synaptic pruning.” F Representative immunofluorescence images showing increased C1q protein expression (red) within F4/80 + macrophages (green) in Apoe −/− mice compared to WT controls. Scale bar, 100 μm. G Quantification of C1q IntDen normalized to the macrophage area. Data are presented as mean ± SD, n = 6, two-tailed Student’s t-test Using canonical cell-type markers, we identified six major cell types (Fig. 3 B and Supplementary Fig. 3B). Due to we focus on the macrophages, we compared macrophages from Apoe −/− mice ( Apoe −/− Mac) with those from WT mice ( Apoe +/+ Mac) (Fig. 3 C). Differential gene expression analysis revealed a striking upregulation of complement component genes, particularly C1qa and C1qb , in Apoe −/− Mac compared to their WT counterparts (Fig. 3 D). Correspondingly, Gene Ontology (GO) analysis of the upregulated genes in Apoe −/− Mac showed significant enrichment for C1q-related biological processes, such as “synaptic pruning”, In contrast, Apoe +/+ Mac upregulated genes were predominantly associated with lipid metabolic processes, including cholesterol transport (Fig. 3 E). To validate these findings at single-cell resolution, our scRNA-seq analysis robustly confirmed the spatial transcriptomic results. After dimensionality reduction and clustering, a comparison between macrophage populations again showed significantly higher expression of C1qa , C1qb , and C1qc in Apoe −/− Mac, with GO analysis similarly highlighting C1q-related functional pathways (Supplementary Figs. 3 C-H). Finally, to confirm this upregulation at the protein level, we performed immunofluorescence staining on 11 dpi spinal cord sections. Co-staining for F4/80 and C1q demonstrated a marked increase in C1q protein expression within macrophages in Apoe −/− mice relative to WT controls (Fig. 3 F and G).In summary, our combined transcriptomic and protein analyses reveal that a key pathological consequence of APOE deficiency in foam cells is the significant upregulation of the complement component C1q. APOE deficiency promotes C1q-mediated synaptic loss and impairs locomotor function recovery after SCI Next, we sought to explore the functional role of the C1q upregulation in Apoe −/− mice. As our transcriptomic analysis highlighted “synaptic pruning”, we hypothesized that excess C1q drives pathological synapse elimination by microglia [ 25 – 27 ]. This canonical pathway involves C1q tagging synapses for removal, which then activates C3, leading to microglial engulfment of the C3b-opsonized synapse [ 28 – 30 ]. To investigate this cascade, we performed immunofluorescence analysis on spinal cord sections from WT and Apoe −/− mice at 11 dpi within a 1 mm peri-lesion region. We first assessed the key steps of complement-mediated pruning. We found a significant increase in the co-localization of C1q and C3 with the postsynaptic marker Homer1 in Apoe −/− mice compared to WT, indicating enhanced complement tagging of synapses (Fig. 4 A and B) [ 31 ]. To determine whether the upregulated C1q in Apoe −/− mice directly targets synapses for elimination, we examined the colocalization of C1q with synaptic markers (Homer1) on cholinergic neurons (ChAT+) and interneurons (PV+) in both the perilesional area and the anterior horn of the lumbar walking executive center. We found that C1q labels neuronal synapses indiscriminately across different neuronal subtypes (Fig. 4 C-E, Supplementary Fig. 4A-C). Subsequently, we observed a greater degree of triple co-localization between the microglial marker IBA-1, C1q, and Homer1 in Apoe −/− mice [ 32 ]. This finding, further supported by 3D reconstruction and rendering, provides direct evidence of increased engulfment of C1q-tagged synapses by microglia (Fig. 4 F and G). Fig. 4. Open in a new tab APOE deficiency exacerbates C1q-mediated synaptic loss and impairs locomotor function recovery after SCI. A Immunofluorescence staining showing increased co-localization of C1q (blue) and C3 (red) with the postsynaptic marker Homer1 (green) in the peri-lesion area of Apoe −/− mice compared to WT at 11 dpi. B Quantification of the ratio of C3-positive tagged synapses (C1q+C3 + Homer1+) to total C1q-tagged synapses (C1q+Homer1+). C Representative immunofluorescence staining for ChAT (cholinergic neurons, green), C1q (red), and Homer1 (blue) in the perilesional area and the anterior horn of the lumbar walking executive center. D , E Quantification of the proportion of C1q-tagged synapses on ChAT+ neurons (ChAT+C1q+Homer1+ / ChAT+Homer1+) in the perilesional area ( D ) and the lumbar anterior horn E . F Images and 3D rendering showing increased engulfment of C1q-tagged (blue) synapses (Homer1, red) by IBA-1 + microglia (green) in Apoe −/− mice compared to WT mice. G Quantification of the synaptic engulfment index (volume of internalized C1q+Homer1 + puncta per microglia). H Representative immunofluorescence images of excitatory synaptic density. Upper panels: Perilesional area stained for VGLUT2 (green) and Homer1 (red). Lower panels: lumbar walking executive center’s anterior horn stained for ChAT (green), VGLUT2 (red), and Homer1 (blue). I , J Quantification of VGLUT2 + Homer1+ excitatory synaptic density in the perilesional area (I) and lumbar walking executive center’s anterior horn J . K-M Assessment of locomotor functional recovery over 28 days. (K) Representative images of mice during BMS scoring. (L) BMS scores, and (M) Inclined plane test results, all showing significantly poorer functional recovery in Apoe −/− mice. Data are presented as mean ± SD, n = 6 biologically independent animals for histological analyses ( A - J ), two-tailed Student’s t-test; n = 8 biologically independent animals for behavioral assessments( K - M ), two-way ANOVA with Tukey’s post-hoc test. Scale bars are indicated in the images This exacerbated synaptic pruning led to a significant loss of synapses in the peri-lesion area and the anterior horn of the lumbar walking executive center. By quantifying co-localized puncta of the excitatory presynaptic marker VGLUT2 and postsynaptic marker Homer1 in both the perilesional area and the anterior horn of the lumbar walking executive center, we confirmed that the overall synapse density was significantly lower in Apoe −/− mice compared to WT controls (Fig. 4 H, I and J) [ 33 ]. We also found that the density of inhibitory synapses (VGAT + and Gephyrin+) was significantly reduced in both the perilesional area and the anterior horn of the lumbar walking executive center in Apoe −/− mice (Supplementary Fig. 4D-F). We found that the synaptic loss in Apoe −/− mice is non-selective, involving both excitatory and inhibitory synapses. To determine if the synapse loss related to locomotor deficits, we assessed locomotor function for 28 days following SCI. At 28 dpi, WT mice had regained sufficient hind-limb strength to consistently support their own body weight, reflected by an average BMS of 4.5 (4.813 ± 0.214), whereas Apoe −/− mice still lacked appreciable large-range ankle movement and scored < 2 (1.75 ± 1.28). On the inclined-plane task the majority of WT animals maintained their position at angles close to 70° (69.24 ± 2.33), but Apoe −/− mice rarely exceeded 50° before sliding (51.66 ± 5.14). (Fig. 4 K-M). Concurrently, we treated Apoe −/− mice with anti-C1q antibodies following SCI and found that this treatment effectively rescued the reduction in synaptic density and poorer locomotor function recover caused by APOE deficiency (Supplementary Fig. 5). Macrophage-specific APOE overexpression restores synaptic density and locomotor function recovery after SCI To definitively establish a causal link between the lack of macrophage-derived APOE and the observed deficits, we conducted a rescue experiment. Specifically, we asked whether selective re-expression of APOE in macrophages of SCI Apoe −/− mice is sufficient to prevent synaptic loss and restore locomotor function. To achieve this, we injected an adeno-associated virus 9 (AAV9) engineered to overexpress APOE and an mCherry reporter under the control of the macrophage-specific F4/80 promoter (AAV9-F4/80- Apoe -mCherry) into the lesion epicenter of Apoe −/− mice at 3 dpi. A control AAV9 expressing only mCherry (AAV9-F4/80- NC -mCherry) was used for comparison. Behavioral phenotypes were monitored for 28 days post-injection, with tissue collection at 31 dpi. Immunofluorescence experiments revealed that over 60%(75.73 ± 6.975% in AAV9-F4/80- NC -mCherry and 72.86 ± 7.418% in AAV9-F4/80- Apoe -mCherry) of F4/80 + macrophages expressing mCherry in the lesion core (Fig. 5 A, Supplementary Fig. 5A, B). Fig. 5. Open in a new tab Macrophage-specific APOE re-expression rescues synaptic loss and promotes functional recovery in Apoe −/− mice. A Schematic of the rescue experiment using AAV9-F4/80- Apoe -mCherry or AAV9-F4/80- NC -mCherry injected into the lesion site of Apoe −/− mice. B , C APOE re-expression significantly reduced LDs (BODIPY, green) accumulation in macrophages (F4/80, red) of Apoe −/− mice at 31 dpi. D , E Concurrently, LDs accumulation was increased in surrounding fibroblasts (PDGFRα, red). F , J APOE re-expression decreased the colocalization of C1q/C3 (blue/red) with synapses (Homer1, green). G , K , L APOE re-expression significantly reduced the number of C1q (red)-tagged synapses (Homer1, blue) on ChAT+ cholinergic neurons(green) in the perilesional area (K) and in the lumbar walking executive center’s anterior horn (L). H , M Engulfment of C1q-tagged (blue) synapses (Homer1, red) by microglia (IBA-1, green) was also reduced. I , N , O APOE re-expression significantly rescued excitatory synaptic density in the perilesional area (N) and in the lumbar walking executive center’s anterior horn (O). Quantifications are shown in (J), (K), (L), (M), (N) and (O). Data are presented as mean ± SD, n = 6, two-tailed Student’s t-test. Scale bars are indicated in the images First, we examined whether APOE re-expression could attenuate LDs accumulation. Indeed, in Apoe −/− mice treated with AAV9-F4/80- Apoe -mCherry, we observed a significant reduction in LDs accumulation within lesion-core macrophages, coupled with a corresponding increase in LDs within adjacent fibroblasts. This demonstrates that the APOE-mediated lipid transfer pathway was successfully reactivated (Fig. 5 B-E). Crucially, this restoration of APOE function suppressed the downstream pathological cascade. APOE overexpression significantly attenuated the activation of the C1q/C3 axis and reduced C1q tagging on cholinergic synapses, thereby suppressing microglial phagocytosis (Fig. 5 F-H, J-M). This mitigation of synaptic pruning resulted in a significant preservation of synaptic density. Synaptic density was markedly increased in the peri-lesion area and lumbar walking executive center’s anterior horn regions of AAV9-F4/80- Apoe -mCherry-treated Apoe −/− mice. (Fig. 5 I, N and O). The results indicate that treating Apoe −/− mice with AAV9-F4/80-Apoe-mCherry rescued the reduction in excitatory synaptic density caused by APOE deficiency. Concurrently, the density of inhibitory synapses was also rescued in both the perilesional area and the anterior horn of the lumbar walking executive center’s anterior horn (Supplementary Fig. 6C-E). Locomotor function analysis revealed that Apoe −/− mice receiving the AAV9-F4/80- Apoe -mCherry treatment exhibited significantly better locomotor performance compared to the control group (Supplementary Fig. 6F-H). To determine whether the protective effects of macrophage-derived APOE are restricted to APOE-deficient animals or also extend to the normal genotype, WT mice were also subjected to SCI and injected with AAV9-F4/80- Apoe -mCherry or NC control at 3dpi. Similar beneficial effects of APOE overexpression were also noted in WT mice (Supplementary Figs. 7 and 8). This implies that, in most SCI patients, endogenously produced APOE remains quantitatively insufficient, and that exogenous supplementation of APOE could provide an additional therapeutic avenue to augment functional recovery. In conclusion, macrophage-specific APOE overexpression is sufficient to reverse the pathological cascade of lipid retention, C1q-mediated synaptic loss, and locomotor deficits in both Apoe −/− and WT mice, establishing macrophage-derived APOE as a critical neuroprotective factor in SCI. APOE deficiency promotes C1q expression in macrophages via Hippo pathway To determine whether the elevation of C1q in APOE-deficient macrophages stems directly from the loss of APOE-dependent signaling or indirectly from the resulting lipid stress, we established an in vitro foam cell model using BMDMs from WT and Apoe −/− mice either separately or in co-culture (Fig. 6 A). In a single-culture condition, where APOE-dependent lipid export to other cells is absent, 24-hour stimulation with Hom induced comparable levels of intracellular LDs accumulation and C1q expression between WT and Apoe −/− macrophages (Supplementary Fig. 9A and B). However, in a co-culture system, Hom-stimulated Apoe −/− macrophages exhibited significantly higher intracellular LDs levels and elevated C1q expression than WT macrophages (Fig. 6 B and C). These data indicate that, within the co-culture, fibroblasts accept macrophage-derived LDs, thereby relieving intracellular lipid stress and reducing C1q expression in an APOE-dependent manner. Notably, under single-culture conditions, C1q expression did not differ between Hom-stimulated Apoe −/− and WT macrophages, suggesting that elevated C1q levels are driven by intracellular lipid stress rather than by APOE deficiency-induced activation of unknown signaling pathways. To determine whether lipid efflux regulates macrophage C1q expression, we treated HOM-stimulated BMDMs with the small-molecule inhibitor Glyburide for 24 h to block lipid efflux. BODIPY fluorescence revealed that Glyburide treatment significantly increased intracellular lipid droplets in BMDMs (Supplementary Fig. 9C and D). Western blot analysis revealed that the inhibition of lipid efflux significantly increased C1q expression in BMDMs (Fig. 6 D and E). Fig. 6. Open in a new tab Intracellular lipid stress activates the Hippo pathway to drive C1q expression. A Schematic of the in vitro co-culture or sigle-culture model used to investigate the mechanism of C1q upregulation. B , C In a co-culture system with L929 fibroblasts, Apoe −/− macrophages exhibited significantly higher intracellular LDs (BODIPY, green) and elevated C1q expression (red) compared to WT macrophages. Data are presented as mean ± SD, n = 6. Statistical significance was determined by one-way ANOVA with Tukey’s post-hoc test. D , E Western blot analysis revealed that inhibiting lipid efflux with Glyburide in HOM-stimulated BMDMs significantly increased C1q protein levels, as shown by Western blot (D) and quantification (E). Data are presented as mean ± SD, n = 3. Statistical significance was determined by one-way ANOVA with Tukey’s post-hoc test. F , G Pharmacological inhibition of the Hippo signaling pathway significantly reduced C1q protein levels in foam cells, as shown by Western blot (D) and quantification (E). Data are presented as mean ± SD, n = 3. Statistical significance was determined by one-way ANOVA with Tukey’s post-hoc test. H , I In vivo analysis of SCI lesions revealed a significantly higher percentage of nuclear active-YAP1 (red) in F4/80 + macrophages (green) of Apoe −/− mice compared to WT mice. Data are presented as mean ± SD, n = 6. Statistical significance was determined by two-tailed Student’s t-test. J Schematic diagram illustrating the experimental protocol for tail vein injection of Verteporfin in Apoe −/− mice following SCI. I.v., intravenous. Q.d, once a day. K , L Intravenous administration of Verteporfin significantly attenuated C1q(red) levels in macrophages (F4/80, blue) at the lesion site. Data are presented as mean ± SD, n = 5. Statistical significance was determined by two-tailed Student’s t-test. Normality and homogeneity of variances were verified by Shapiro–Wilk and Levene’s tests, respectively, before one-way ANOVA. Scale bars are indicated in the images To identify the signaling pathways linking lipid stress to C1q expression, we performed bulk RNA sequencing on Hom-stimulated BMDMs. KEGG analysis identified several highly enriched signaling pathways (Supplementary Fig. 9E and F). We then systematically targeted these top-ranked pathways using specific inhibitors in our foam cell model. Notably, pharmacological inhibition of the Hippo signaling pathway resulted in a significant reduction in C1q protein expression (Fig. 6 F and G), implicating it as a critical mediator. A canonical feature of Hippo pathway activation is the nuclear translocation of its primary downstream effector, the transcriptional co-activator active-YAP1 [ 34 ]. We therefore examined its subcellular localization. Immunofluorescence staining in co-cultured foam cells demonstrated a marked increase in nuclear active-YAP1 in Apoe −/− macrophages relative to WT controls (Supplement Fig. 9G and H). Crucially, this finding was validated in vivo; analysis of SCI lesions revealed a significantly greater abundance of nuclear active-YAP1 in Apoe −/− mice compared to WT mice (Fig. 6 H and I). We administered the active YAP1 inhibitor Verteporfin (10 mg/kg) to Apoe −/− mice via daily tail vein injection starting from 3 dpi. Tissues were harvested at 11 dpi for immunofluorescence analysis (Fig. 6 J). The results showed that Verteporfin treatment significantly reduced the nuclear accumulation of active YAP1 (colocalization with DAPI) in the lesion center (Fig. 6 K and Supplementary Fig. 9I). Furthermore, C1q expression in macrophages within the lesion core was significantly suppressed (Fig. 6 L). Collectively, these data demonstrate that macrophage APOE deficiency impairs the resolution of intracellular lipid stress, leading to the activation of the Hippo signaling pathway. This, in turn, promotes the nuclear translocation of active-YAP1, ultimately driving the pathological upregulation of C1q. Taken together, our findings delineate a clear pathological cascade following spinal cord injury: the absence of macrophage-derived APOE leads to intracellular lipid stress, which in turn activates the Hippo-YAP1 pathway to drive C1q upregulation. This excess C1q subsequently triggers excessive microglial pruning of synapses, ultimately impairing locomotor functional recovery. The key elements of this signaling axis are summarized in the graphical abstract (Fig. 7 ). Fig. 7. Open in a new tab The Neuroprotective Mechanism of the APOE-Mediated Lipid Transport Pathway Following Spinal Cord Injury. This schematic summarizes the central findings of our study. (Left) Following spinal cord injury (SCI), macrophages engulf myelin debris and transform into lipid-laden foam cells. (Center) Under physiological conditions, these macrophages secrete Apolipoprotein E (APOE) to transport excess intracellular lipids to adjacent cells, such as fibroblasts, thereby alleviating their own lipotoxic stress. (Right) In the absence or deficiency of APOE, lipids pathologically accumulate within macrophages. This lipotoxicity activates the Hippo signaling pathway, leading to the nuclear translocation of its downstream effector, YAP1, which in turn upregulates the expression and secretion of the complement protein C1q. Excess C1q acts as an “eat-me” signal, aberrantly tagging viable synapses in the peri-lesional area. Ultimately, microglia recognize these C1q/C3-opsonized synapses via complement receptors (e.g., CR3) and subject them to excessive phagocytic elimination. This results in substantial synaptic loss and contributes to poorer neurological recovery Discussion This study uncovers a mechanism by which macrophages, following SCI, transfer intracellular LDs to neighboring cells via APOE to alleviate excessive lipid stress. Further investigation revealed that lipid stress can activate C1q expression through the Hippo signaling pathway, thereby triggering downstream cascades that promote microglial phagocytosis of normal synapse both adjacent to and distant from the lesion site, ultimately impairing locomotor function recovery after SCI. The findings of this study build upon and are interconnected with previous research, providing key answers to several unresolved questions in the field. For instance, previous studies have found that promoting lipid degradation in foam cells within the lesion improves SCI prognosis, hypothesizing that the mechanism was likely related to mitigating foam cell-mediated inflammation, though the specific molecular pathways for lipid clearance were not investigated [ 35 – 38 ]. Concurrently, other research has observed that macrophages and microglia upregulate complement C1q expression following CNS injury, leading to poorer functional recovery [ 39 – 41 ]. However, the upstream triggers for C1q upregulation and its specific downstream pathological effects remained unclear. Furthermore, studies on APOE have also shown that Apoe −/− mice exhibit impaired locomotor functional recovery after SCI, attributing this to a more severe inflammatory response and increased permeability of the blood-spinal cord barrier, but without elucidating the underlying cellular mechanisms driving these pathologies in the absence of APOE [ 42 – 44 ]. Our central finding is that the absence of APOE, the primary lipid transporter in the CNS, impairs this lipid efflux pathway and triggers a pathological cascade. APOE deficiency leads to the pathological accumulation of lipids within macrophages. This intracellular lipid stress, in turn, activates the Hippo signaling pathway. Although the Hippo pathway did not exhibit the highest gene ratio in our initial KEGG enrichment analysis, our functional validation identified it as a critical regulatory hub. We demonstrated that activation of the Hippo-YAP1 axis serves as a potent driver for the transcriptional upregulation of C1q, outweighing its simple enrichment proportion. Excess C1q aberrantly tags synapses in the peri-lesional area, inducing their excessive engulfment by microglia, which ultimately leads to a substantial loss of synaptic networks and severe functional impairment. It is also worth noting that human APOE exists in three major isoforms (APOE2, APOE3, and APOE4), which differ significantly in their lipid transport efficiency and amyloid-β clearance capabilities. Clinical and preclinical studies have firmly established APOE4 as a major risk factor for poor outcomes in neurodegenerative diseases like Alzheimer’s disease and traumatic brain injury. While our current study utilized wild-type mice expressing a single murine APOE isoform, it is plausible that human APOE4 carriers may exhibit exacerbated macrophage lipid accumulation and more severe synaptic loss following SCI compared to APOE2 or APOE3 carriers. Exploring these isoform-specific effects using APOE-targeted replacement mice will be a crucial direction for our future research to enhance the translational value of these findings. Furthermore, given that C1q-mediated synaptic elimination is a well-documented phenomenon in neurodevelopment and neurodegenerative diseases, the “Macrophage APOE–Hippo–C1q” axis we uncovered likely represents a conserved pathological mechanism across various CNS injuries. Similar to SCI, conditions such as Traumatic Brain Injury and stroke involve significant tissue damage, lipid debris accumulation, and macrophage infiltration. We hypothesize that targeting this axis could offer broad neuroprotective benefits in these brain injuries as well. Crucially, we reversed this entire pathological process by specifically re-expressing APOE in the macrophages of Apoe −/− mice. APOE reconstitution successfully restored lipid transport, suppressed C1q overexpression, rescued synaptic loss, and significantly promoted motor function recovery. Additionally, our use of C1q neutralizing antibodies in vivo provided direct evidence that blocking this downstream effector is sufficient to rescue synaptic density, confirming the necessity of C1q in this pathological cascade. This definitive evidence of causality establishes macrophage-derived APOE as a critical neuroprotective factor in the SCI recovery process. However, our study is not without limitations. First, this research utilized global Apoe knockout mice rather than conditional knockout (cKO) models. Therefore, while the observed pathologies and functional outcomes are strongly linked to macrophage dysfunction (as evidenced by our rescue experiments), we cannot entirely rule out the potential confounding effects stemming from the absence of APOE in other cell types. Second, although our study clearly demonstrates the transfer of lipids from macrophages to fibroblasts, astrocytes, and neurons, the subsequent physiological or pathophysiological responses within these recipient cells remain an open question. Third, regarding the mechanism of C1q regulation, our current conclusions rely heavily on pharmacological inhibition of the Hippo pathway (e.g., Verteporfin). While our in vivo and in vitro data are consistent, off-target effects of small molecule inhibitors cannot be completely excluded. Future studies utilizing macrophage-specific Yap1 conditional knockout mice are warranted to definitively dissect the cell-autonomous role of YAP1. Finally, our assessment of motor function recovery was primarily based on BMS scores and inclined plane tests. Future investigations should incorporate more sensitive automated gait analysis systems (e.g., Rotarod or CatWalk) to capture subtle changes in locomotor coordination. In summary, this research not only deepens our understanding of the interplay between neuroinflammation and lipid metabolism after SCI but also identifies a novel therapeutic target: the “macrophage APOE-Hippo-C1q axis”. Modulating this pathway, for instance by enhancing APOE function in macrophages or inhibiting downstream C1q-mediated synaptic pruning, may offer a promising new therapeutic strategy to promote neural repair following SCI. Methods Ethical Statement All animal procedures were performed in strict accordance with the guidelines approved by the Institutional Animal Care and Use Committee (IACUC) of Shanghai General Hospital (IACUC Number: 2025AW037). Animals Wild-type C57BL/6J mice and Apolipoprotein E knockout ( Apoe −/− ) mice were purchased from Shulaibao (Wuhan) Biotechnology Co. Ltd. All mice were housed in a specific-pathogen-free (SPF) facility with a 12-hour light/dark cycle and had ad libitum access to food and water. Female mice, aged 8–10 weeks, were used for the experiments. Spinal cord injury (SCI) Model Mice were anesthetized using an RWD small animal isoflurane anesthesia machine (isoflurane concentration: 3–4% for induction, 1.5-2% for maintenance). After achieving a surgical level of anesthesia, a laminectomy was performed at the T10 vertebral level. A complete crush injury was induced by compressing the spinal cord with FST [Cat# 11252-20] forceps for 2 s. The sham group underwent laminectomy only, without spinal cord compression. The muscle layers and skin were sutured sequentially. Post-operatively, animals received subcutaneous injections of saline for hydration and meloxicam (20 mg/kg) for analgesia for 3 consecutive days. Manual bladder expression was performed twice daily until bladder function was restored. Cell Culture Bone Marrow-Derived Macrophages (BMDMs): Bone marrow cells were flushed from the femurs and tibias of WT or Apoe −/− mice. Cells were cultured in DMEM [Gibco, Cat# 11965092] supplemented with 10% fetal bovine serum (FBS), 1% penicillin-streptomycin, and 40 ng/mL macrophage colony-stimulating factor (M-CSF) [HUABIO, Cat# HA210725 ] for 7 days to differentiate into BMDMs. Foam Cell Induction To induce foam cell formation, BMDMs were incubated for 24 h with a complete medium containing 1 mg/mL spinal cord homogenate (Hom). Fibroblasts The L929 fibroblast cell line was cultured in DMEM with 10% FBS and 1% penicillin-streptomycin. Preparation of mouse spinal cord homogenate Eight-week-old male C57BL/6J mice were deeply anesthetized and immediately euthanized by cervical dislocation. After surface sterilization with 75% ethanol, the vertebral column was exposed by a dorsal midline incision and transected along the ribs. The intact spinal cord was rapidly dissected, placed into pre-weighed, autoclaved 1.5 mL tubes, and suspended in ice-cold DMEM at a final concentration of 100 mg /mL. Two sterile stainless-steel beads were added, and the sample was homogenized at 70 Hz for 300 s using a bead-beater. The homogenate was then heat-inactivated at 95 °C for 10 min to abolish endogenous cytokine activity, followed by three freeze–thaw cycles (− 80 °C to room temperature) to ensure complete cell lysis. All procedures were performed under strict aseptic conditions [ 45 ]. In Vivo and In Vitro Lipid Droplets (LDs) Tracing For in vivo tracing, BMDMs or foam cells were labeled with 1 µM BODIPY 493/503 (for LDs, green; [Invitrogen, Cat# D3922]) and 5 µM DiD (for cell membrane, blue; [Beyotime, Cat# C1039]) for 15 min. After washing, 2 µL of cell suspension containing 3 × 10^6 cells were immediately injected into the lesion epicenter following SCI with Hamilton microsyringe (Cat# 7635-01). Injections were performed with a mouse brain/spine stereotaxic apparatus. The needle was advanced 1 mm into the target site, the payload was infused at 0.4 µL/min for 5 min, and the cannula was left in place for an additional 5 min to prevent back-flow before slow withdrawal. For in vitro tracing, BMDMs were treated and labeled as described above. A transwell co-culture system with 0.4 μm pores [Beyotime, Cat# FTW031] was used. L929 fibroblasts were seeded in the bottom chamber, and labeled BMDMs (control or BMDM@LDs@BODIPY) were seeded in the top insert. After 24 h of co-culture, cells were fixed for imaging. Immunofluorescence staining Mice were euthanized and transcardially perfused with PBS followed by 4% paraformaldehyde (PFA). Spinal cord segments were dissected, post-fixed in 4% PFA overnight, and cryoprotected in 15% and 30% sucrose. Tissues were then embedded in Optimal Cutting Temperature (OCT) compound and sectioned 6 μm thick slices using a cryostat. Sections were permeabilized with 0.3% Triton X-100, blocked with 10% goat serum for 1 h, and incubated overnight at 4 °C with the following primary antibodies: F4/80 [1:1000, Servicebio, Cat# 113373], PDGFRα [1:100, HUABIO, Cat# ET1702-49], APOE [1:100, CST, Cat# 49285], C1q [1:100, Abcam, Cat# ab182451], Homer1 [1:100, Synaptic Systems, Cat# 160003], VGLUT2 [1:100, Synaptic Systems, Cat# 135403], IBA-1 [1:200, Abclonal, Cat# A19776], active-YAP1 [1:200, Abcam, Cat# EPR19812 ], C3 [1:100, Abcam, Cat# EPR19394 ], GFAP [1:200, Affinity, Cat# DF6040], NeuN [1:200, Abclonal, Cat# A19086], and LDLR [1:100, Proteintech, Cat# 82724], VGAT [1:200, Abcam, Cat#ab308062], Gephyrin [1:200, Abcam, Cat#ab177154], ChAT [1:200, HUABIO, Cat#HA723705], PV [1:200, Abcam, Cat#ab181086]. After washing, sections were incubated with corresponding Alexa Fluor-conjugated secondary antibodies [1:1000, Invitrogen] for 1 h at room temperature. Nuclei were counterstained with DAPI. For multiplex immunofluorescence staining, a Tyramide Signal Amplification (TSA) kit was used [Abclonal, Cat# RK05903]. The procedure was as follows: Sections were treated with an endogenous peroxidase quencher to block endogenous peroxidase activity. Non-specific sites were blocked with goat serum for 1 h at room temperature. Sections were incubated with the first primary antibody overnight at 4 °C. On the following day, after washing, sections were incubated with an HRP-conjugated secondary antibody. The signal was developed by incubating the sections with a specific TSA fluorophore (e.g., 440, 488, 555, or 647) for 15 min. To prepare for the next primary antibody, the existing antibodies were removed by incubating the sections in an antibody stripping buffer at 37 °C for 20 min. Steps were repeated for each subsequent primary antibody until all proteins of interest were labeled. After the final staining, sections were washed, counterstained with DAPI, and mounted with an anti-fade mounting medium. Image acquisition and analysis Images were captured using a Leica SP8 confocal microscope and Zeiss Axio Imager 2. For quantitative analysis, images were processed using ImageJ/Fiji software. Synaptic density was quantified by measuring the co-localized area of VGLUT2 and Homer1, VGAT and Gephyirn puncta. Protein expression levels were quantified by measuring the integrated density (IntDen). Cell counting and co-localization analyses were performed on at least three randomly selected fields per section from at least three different animals per group. 3D rendering was performed using ImageJ/Fiji and Blender software. RNAscope In situ hybridization for Apoe mRNA was performed on spinal cord sections using the RNAscope Multiplex Fluorescent Reagent Kit v2 [ACD, Cat# 323100]. For RNAscope, Apoe RNA is first exposed by heat-induced retrieval and protease digestion; 20- to 40-mer double-“Z” probe pairs are then hybridized at 40 °C for 2 h, generating specific landing sites only when adjacent probes bind. Sequential 40 °C incubations with branching pre-amplifiers and HRP/AP-conjugated amplifiers (15–30 min each) create a 400-fold signal tree, visualized with fluorescent or chromogenic substrates that render one punctum per RNA molecule. This was followed by immunofluorescence staining for F4/80 and APOE protein as described above. Transcriptomic analysis (spatial and single-cell) Spatial transcriptomics and single-cell RNA sequencing were performed by Beijing SeekGene Biosciences Co., Ltd. 5 μm sections of WT and Apoe −/− spinal cord samples were taken from the FFPE tissue blocks with a microtome (RWD Life Science Co., Ltd). Five-micrometer FFPE sections were placed on capture areas of the immunocytochemistry slides. After incubating for 2 h at 42℃ and keeping drying at room temperature. The isolation of total RNA from FFPE tissue blocks was achieved using the RNeasy FFPE kit (73504, Qiagen). The quality assessment of the extracted RNA was performed by calculating DV200. Tissue sections passing the quality control (DV200 > 30%) were subjected to ST assay. The Visium HD workflow requires new HD slides that require thawing, washes, and equilibration in appropriate buffers. Visium HD slides also feature high-resolution fiducials for subpixel image alignment, and a dispensing pad and spacer for CytAssist compatibility. We first placed FFPE tissue sections on plain glass slides for deparaffinization, H&E staining and imaging following the Visium HD FFPE Tissue Preparation Handbook ( CG000684 ). Subsequently, the sections were stained with H&E and imaged at 20× magnification in brightfield using PANNORAMIC MIDIⅡDigital Scanner (3DHISTECH). Probe hybridization, probe ligation, slide preparation, probe release, extension, library construction, and sequencing followed the Visium HD Spatial Gene Expression Reagent Kits User Guide ( CG000685 ). Sequencing was performed on an Illumina NovaSeq 6000 with paired-end reads (43 cycles Read 1, 10 cycles i7, 10 cycles i5, 50 cycles Read 2). We used Space Ranger v3.0 to map FASTQ files to the human reference, detect the tissue section, align the sequencing data to the microscope image and the CytAssist image, and output gene-barcode matrices for further analysis. Spatial transcriptomics data preprocessing and quality control Following genomic sequence alignment and generation of the expression matrix, genes expressed in fewer than three bins were filtered out. To address the issue of fixed-size bins failing to match the true morphology and size of cells, and to mitigate the effects of RNA diffusion in spatial transcriptomics, the bin2cell package was employed under stringent parameters. This approach identified the nucleus of each cell, and only the expression levels from bins within the nucleus were extracted. These were then aggregated to define a total of 23,051 cells. The cells were assigned to either the Apoe −/− or WT sample groups based on their spatial coordinates. Following this, cells expressing fewer than 100 genes were excluded, and subsequently, genes expressed in fewer than three cells were removed from the analysis. Cell type annotation After quality control, cell type annotation was performed using the scanpy package. Gene expression counts were normalized and log-transformed using default parameters. The 2000 most highly variable genes were identified, and Principal Component Analysis (PCA) was performed on the expression of these genes except for Apoe . A nearest-neighbor graph was computed based on the PCA results, which was then used for clustering at a resolution of 1. Cell types for each cluster were identified using a predefined list of marker genes. For clusters expressing markers for multiple cell types, the score_genes function was used to calculate a gene set score for each cell based on the different cell type marker lists. This allowed for the separation of distinctly classified cells, while cells with ambiguous types were discarded. Finally, the reliability of the cell type annotations was verified by examining the highly expressed genes for each type, as identified by the rank_genes_groups function, against the marker list. Differential gene expression analysis Differential gene expression analysis was conducted on the macrophage population using the Wilcoxon rank-sum test implemented in Scanpy. Macrophages were grouped into Apoe −/− and control samples, and differentially expressed genes (DEGs) were identified. To control for multiple comparisons, p-values were adjusted using the Benjamini-Hochberg procedure to calculate the False Discovery Rate (FDR). Genes with an FDR-adjusted p-value < 0.05 were considered statistically significant. Subsequently, Gene Ontology (GO) enrichment analyses were performed using the enrichr API from the gseapy package. The results of the differential gene analysis were visualized using a custom function. Differential expression analysis was performed between control and SCI scRNA-seq samples using Scanpy. Genes exhibiting a |log₂FC|>2 and a Benjamini–Hochberg-adjusted P-value < 0.05 were considered statistically significant. KEGG pathway enrichment analysis was conducted using the gseapy package, with expressed genes selected from the intersection of the sample gene lists serving as the background. Bulk RNA sequencing RNA sequencing was performed by OE Biotech (Shanghai, China). Total RNA was isolated using TRIzol reagent (Invitrogen, Cat. No. 15596026), and RNA integrity was verified on an Agilent 2100 Bioanalyzer (RIN ≥ 7.0). Strand-specific libraries were generated using the VAHTS Universal V5 RNA-seq Library Prep Kit (Vazyme) following the manufacturer’s instructions and sequenced on an Illumina NovaSeq 6000 platform (2 × 150 bp paired-end reads). Raw reads were aligned to the mouse reference genome (mm10) using HISAT2, and gene-level counts were obtained using HTSeq. FPKM (Fragments Per Kilobase of transcript per Million mapped reads) values were calculated for data visualization. Principal Component Analysis (PCA) was performed to visualize transcriptomic relationships among samples. Differential expression analysis was performed using DESeq2 (v1.38.3) based on raw read counts. To account for multiple testing, P-values were adjusted using the Benjamini–Hochberg procedure to control the False Discovery Rate (FDR). Genes meeting the criteria of an adjusted P$-value < 0.05 and an absolute log2-fold change |< log2FC>| 1 were considered significantly differentially expressed genes (DEGs). Functional interpretation, including Gene Ontology (GO) annotation, KEGG pathway enrichment, and Gene Set Enrichment Analysis (GSEA), was conducted using clusterProfiler (v4.10.0). Similarly, P-values for enrichment analyses were corrected for multiple comparisons using the Benjamini–Hochberg method, with an adjusted P-value < 0.05 considered statistically significant. Western blot To screen for relevant pathways, Hom-stimulated foam cells were treated with specific inhibitors for 24 h. Cells were lysed, and protein concentrations were determined. Equal amounts of protein were separated by SDS-PAGE, transferred to a PVDF membrane, and incubated with primary antibodies against C1q [1:1000, Thermo, Cat# PA5-51241], B-tubulin [1:10000, Affinity, Cat# T0023]. On the following day, after washing, sections were incubated with an HRP-conjugated secondary antibody. Protein bands were visualized using an Tanon imaging system. The inhibitor used were Resatorvid [1µM, Sparkjade, Cat# SJ-MX0293], R-7050 [1µM, Sparkjade, Cat# SJ-MX1573], Aristolochic acid A [10µM, Sparkjade, Cat# SJ-MN0173], Verteporfin [1.328µM, Selleck, Cat# S1786], Tretinoin [1µM, Sparkjade, Cat# SJ-MV0063], GW9662 [10µM, Abmole, Cat# M2748], GSK2033 [1µM, Abmole, Cat# M6768], Glyburide [100µM, Selleck, Cat# S1716]. AAV9 administration experiments An adeno-associated virus serotype 9 (AAV9) carrying the Apoe gene and an mCherry reporter under the control of the macrophage-specific F4/80 promoter (AAV9- F4/80-Apoe-mCherry ) and a control virus (AAV9- F4/80-NC-mCherry ) were constructed by HANBIO. At 3 dpi, 2 µL of AAV (titer: 1*10^13) was injected into the lesion epicenter of Apoe −/− or WT mice with Hamilton microsyringe (Cat# 7635-01). Injections were performed with a mouse brain/spine stereotaxic apparatus. The cannula was lowered to 1 mm and 0.7 mm, respectively, and 1 µL was delivered at each depth at 0.4 µL/min; after each infusion the needle remained in situ for 5 min to prevent back-flow before slow withdrawal. Locomotor functional assessments Locomotor function was assessed at 1, 3, 7, 14, 21, 28 dpi by two blinded observers. Basso Mouse Scale (BMS): Mice were observed for 4 min in an open field, and hindlimb locomotor function was scored on a scale of 0 to 9. [ 46 ]. Inclined Plane Test: The maximum angle at which a mouse could maintain its position for 5 s on an inclined plane was recorded [ 47 ]. Verteporfin treatment To inhibit YAP1 activity, the small-molecule inhibitor Verteporfin (Cat# S1786, Selleck, Shanghai, China) was prepared in a vehicle mixture consisting of 5% DMSO, 40% PEG300, 5% Tween 80, and 50% ddH2O. Mice in the treatment group received Verteporfin via tail vein injection at a dosage of 10 mg/kg daily (q.d.) from 3 to 11 dpi. The vehicle control group was administered the same solvent mixture (containing DMSO without Verteporfin) following the identical regimen. Anti-C1q antibody treatment For C1q neutralization experiments, a specific anti-C1q antibody (Cat# A2580, Selleck) or a non-immune IgG isotype control (Cat# A2052, Selleck) was administered intravenously. Treatments were initiated at 3 dpi via tail vein injection. Mice received the antibody injections once a week (q.w.) until 28 dpi. Statistical analysis All quantitative data are presented as mean ± standard deviation (SD). Statistical analyses were performed using GraphPad Prism 10.2. A two-tailed Student’s t-test was used for comparisons between two groups. One-way ANOVA followed by Tukey’s post-hoc test was used for multiple group comparisons. Two-way ANOVA was used for behavioral data analysis over time. Normality and homogeneity of variances were verified by Shapiro–Wilk and Levene’s tests, respectively, before one-way ANOVA or two-way ANOVA. A p-value of less than 0.05 was considered statistically significant. Supplementary Information Supplementary Material 1. (7.1MB, pdf) Supplementary Material 2. (14.9MB, docx) Acknowledgements This work was supported by the National Natural Science Foundation of China (NSFC 82271407, NSFC 82501639) and the Excellent Academic Leader Project of Shanghai Science and Technology Commission (22XD1402600) to H.L. and Y.P.; We acknowledge BioRender.com for the schematic illustration elements used in this study. The publication licenses and export codes for the corresponding figures are as follows: Figure 1A (OU29FQQU9W), Figure 1E (QX29FQQKXE), Figure 3A (QU29FQR4F5), Figure 5A (PO29FQQXG6), and Supplementary Figure 9E (QX29FQQKXE). All other schematic diagrams in this manuscript were generated in Adobe Illustrator utilizing elements derived from these five BioRender exports. Authors’ contributions Y.P. were responsible for the concept and experimental design. Y.M., R.Z., C.N. and Y.P. performed the experiments, data analysis, and statistical analysis. Z.R., C.N., C.F., Y.H., M.Z., Y.D., C.C., P.W., R.W., H.Z. and Y.H. provided technical and material support. Y.M. was involved in drafting and revision of the manuscript. D.J., Y.P. and H.L. supervised this study. All authors discussed the results and commented on the manuscript. All authors read and approved the final manuscript. Data availability No datasets were generated or analysed during the current study. Declarations Competing interests The authors declare no competing interests. Footnotes Publisher’s Note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Yersen Mulat, Zun Ren and Chaocao Nong contributed equally to this work. Contributor Information Dongsheng Jiang, Email: [email protected]. Ying Peng, Email: [email protected]. Haodong Lin, Email: [email protected]. References 1. Locati M, Curtale G, Mantovani A. Diversity, Mechanisms, and Significance of Macrophage Plasticity. 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