Abstract

   RhoA, a small GTPase, plays a pivotal role in various diseases,
   including spinal cord injury (SCI). Although RhoA inhibition has been
   traditionally viewed as beneficial for SCI repair, recent clinical
   trials of RhoA inhibitors in SCI have failed to show significant
   therapeutic efficacy, suggesting functional heterogeneity across
   different cell types. The role of RhoA in microglia, the key immune
   cells involve in SCI, remains poorly understood. Using microglial RhoA
   conditional knockout mice, this study demonstrated that RhoA deficiency
   in microglia attenuates the morphological and functional repair of the
   SCI mice, and impairs the microglial biofunctions of proliferation,
   phagocytosis, and migration. Single-cell RNA sequencing, bulk RNA
   sequencing, and metabolomics revealed that RhoA deficiency can
   attenuate the microglial glycolytic enzyme expression, ATP production,
   ECAR and OCR levels through the Arhgap25/HIF-1α pathway. Overall, this
   is the first study to demonstrate that microglial RhoA is essential for
   SCI repair, the Arhgap25/HIF-1α pathway mediated glucose metabolism
   might enlighten a novel insight to enrich the understanding on the
   complex roles of RhoA and microglia in SCI repair. Moreover, this study
   highlights the importance of considering cell-specific roles of RhoA in
   SCI repair and provides a foundation for developing targeted therapies
   aimed at microglial metabolic reprogramming.

   graphic file with name 41419_2025_7947_Figa_HTML.jpg
   [48]Open in a new tab

   Schematic representation of the proposed mechanism by which microglial
   RhoA regulates glycolytic adaptation and spinal cord repair. (Created
   by Figdraw.com with permission of # wgq=r7c74c).

   Subject terms: Spinal cord diseases, Microglia

Introduction

   RhoA, a member of the small GTPase family, has garnered significant
   attention due to its marked upregulation or activation in various
   pathological conditions, including spinal cord injury (SCI) [[49]1,
   [50]2] and cancer [[51]3]. Historically, RhoA has been perceived as a
   detrimental molecule, with studies demonstrating that inhibiting its
   expression or activation can suppress tumor progression [[52]4] and
   promote tissue repair [[53]5, [54]6]. Such therapeutic benefits have
   been observed in conditions like erectile dysfunction [[55]7], diabetic
   nephropathy [[56]8], and atherosclerosis [[57]9]. In the nervous
   system, substantial evidences indicate that suppressing RhoA can
   enhance neuronal survival and promote axonal regeneration [[58]10,
   [59]11], making RhoA inhibitors promising candidates for treating SCI.
   However, recent multicenter, randomized, double-blind clinical trials
   have yielded unexpected results [[60]12, [61]13]. RhoA inhibitors
   failed to achieve the anticipated therapeutic benefits for SCI, leading
   to the premature termination of the study based on interim futility
   analyses. This discrepancy between preclinical success and clinical
   failure raises critical questions. Why have interventions targeting
   RhoA not translated into clinical efficacy?

   We hypothesize that the functional heterogeneity of RhoA across
   different cell types may underlie this disparity. For instance,
   neuron-specific RhoA knockout promotes SCI repair, while
   astrocyte-specific RhoA knockout appears to hinder recovery [[62]14].
   Similarly, our recent studies have demonstrated that RhoA conditional
   knockout in Schwann cells and macrophages exert opposing effects on
   peripheral nerve regeneration [[63]15, [64]16]. Importantly, microglia
   play a central role in SCI recovery, and our previous study revealed a
   significant and sustained upregulation of RhoA in microglia post-SCI,
   persisting longer than in neurons or astrocytes [[65]2]. This
   observation underscores the need to elucidate the specific role of RhoA
   in microglial function during SCI repair.

   Microglia, as the resident immune cells of the central nervous system
   (CNS), are pivotal in mediating injury repair through processes such as
   proliferation, phagocytosis, and migration [[66]17–[67]19]. These
   functions are crucial for SCI recovery, making microglial modulation a
   key therapeutic strategy. Previous studies have highlighted RhoA’s
   diverse roles in microglia, including its regulation of glutamate
   release in Alzheimer’s disease [[68]20], where RhoA deletion
   exacerbates disease progression, and ATP release [[69]21], which is
   promoted by increased RhoA activity. Additionally, the RhoA/ROCK
   pathway has been implicated in the M1/M2 polarization of microglia and
   macrophages [[70]22]. Beyond cytoskeletal dynamics, RhoA likely
   influences multiple biological processes through its involvement in
   diverse signaling pathways [[71]23]. Does the sustained high expression
   of RhoA in microglia enhance their reparative functions, or does it
   facilitate SCI recovery by modulating metabolic pathways? Current
   researches have yet to provide definitive answers to these questions,
   making this an area of critical importance for understanding SCI repair
   and guiding therapeutic development.

   In this study, we demonstrated the essential role of RhoA in microglia
   for regulating key functions such as proliferation, phagocytosis, and
   migration. Present findings revealed that RhoA modulates the glycolytic
   process in microglia via the Arhgap25/HIF-1α pathway, which offer a
   deeper understanding of its importance in SCI repair and indicate that
   targeted modulation of RhoA in microglia may therefore represent a
   promising strategy for improving clinical outcomes in SCI treatment.

Materials and methods

Animal procedures

   All procedures were performed in accordance with guidelines set in
   place by the Southern Medical University Animal Care and Use Committee.
   C57BL/6 adult female mice (aged 8–12 weeks, weighing 20–25 g) and
   neonatal mice (days 2–3) were provided by the Animal Center of Southern
   Medical University. RhoA^flox/flox and Cx3cr1^Cre mice (provided by
   Cyagen, Suzhou, China) were intercrossed to obtain RhoA cKO mice with
   genotype of RhoA^flox/flox; Cx3cr1^Cre, as previously described
   [[72]16]. The littermates with genotype of RhoA^flox/flox were utilized
   as negative controls for cKO mice.

Spinal cord injury model

   Mice spinal cord crush injury was performed at thoracic level 10-11
   (T10-T11) similar to that described before [[73]24]. Briefly, a midline
   incision was made over the thoracic vertebrae, followed by a T10-11
   laminectomy. Tips of forceps were carefully inserted on either side of
   the cord to include the full width of the cord and then to gently
   scrape them across the bone on the ventral side to not spare any tissue
   ventrally or laterally. The spinal cord was fully crushed for 3 s with
   0.1 mm forceps. The muscles were sutured, and the skin was closed with
   wound clips. Mice were placed on a warming pad after surgery until
   fully awake. Their bladders were manually expressed one times per day
   for the duration of the experiments.

Behavioral assessments

   The Basso Mouse Scale (BMS) was used to assess locomotor function at 1,
   3, 7, 14, and 28-days post-injury (dpi) as previously reported. The BMS
   scale ranges from 0 (paralysis) to 9 (normal locomotion), scored during
   a 2–3 min open-field test by three blinded observers [[74]25].

   At 28 dpi, mice were trained to swim across a water-filled tank, with
   performance scored on hindlimb movement, coordination, tail position,
   paw placement, and balance [[75]26]. The final score was the average of
   two trials, evaluated by two blinded investigators.

   Four hours later, footprint analysis was performed to assess hindlimb
   function, with forelimbs and hindlimbs inked in black and red,
   respectively [[76]27]. Mice ran along a paper-lined runway, and
   parameters like stride length, plantar stepping, and coordination were
   qualitatively analyzed.

Histomorphometry of the gastrocnemius muscle

   The harvested gastrocnemius muscles were weighted for calculating the
   wet weight [[77]27]. Then mid-belly of the muscle was transversally
   sectioned and executed routine hematoxylin-eosin (HE) staining to
   visualize the myofibers. The area of myofibers was measured by using
   ImageJ software 6.0.

Primary microglia culture

   Primary microglia were isolated and cultured as previously described
   [[78]28, [79]29]. In brief, a mixed glial cell culture was prepared
   from neonatal C57BL/6J mice and maintained for 10–21 d in DMEM (Thermo
   Fisher Scientific, MA, USA, C11995500BT) containing 10 ml of 10% FBS
   (Biochannel, Nanjing, China, BC-SE-FBS07) and 1% antibiotic-antimycotic
   (Thermo Fisher Scientific, MA, USA, 15240062) in a T75 flask. Microglia
   were collected by gentle shaking as the floating cells over the mixed
   glial cell culture. Then, primary microglia were transferred to a
   twelve-well or six-well plate for experiments.

Myelin debris preparation

   Myelin debris was prepared from the brains and spinal cords of mice as
   described previously [[80]15]. Briefly, brain and spinal cord tissues
   were harvested from 8–10-week-old wild-type mice following euthanasia
   by cervical dislocation. The tissues were disrupted into fine particles
   via sonication, and the resulting debris was washed three times with
   Milli-Q water by centrifugation at 14,462 × g for 15 min at 4 °C.
   Finally, the myelin debris was resuspended in Hank’s balanced salt
   solution (HBSS, Thermo Fisher Scientific, MA, USA, C14175500BT) at a
   concentration of 100 mg/mL and stored at −80 °C for later use.

Phagocytic capability assay

   Microglial phagocytosis of myelin debris was assessed by seeding cells
   at 1 × 10⁵ cells/well in 12-well plates. After 24 h, 1 mg/mL myelin
   debris was added for 0.5, 1, or 2 h. After three washes with HBSS,
   protein content was measured, and cells were fixed with 4% PFA and
   stained with IBA-1 to visualize microglia morphology. Myelin debris
   uptake was co-stained with MBP for visualization.

Migration assay

   Microglial migration was assessed using a Transwell assay [[81]15].
   Cells were seeded in the upper chamber, with the lower chamber
   containing 600 µL DMEM, 10% FBS, and 1 mg/mL myelin debris. After 48 h,
   cells were fixed with 4% PFA, non-migrated cells on the upper membrane
   surface were removed, and migrated cells on the lower surface were
   stained with 0.1% crystal violet for 30 min. Images from five random
   fields were captured and quantified.

Lipofectamine transfection for Arhgap25 knock down

   Primary microglia were cultured to 60% confluence in six-well plates.
   Fifty nM siRNA targeting Arhgap25 or a negative control was transfected
   using 5 μL of Lipofectamine RNAi (D-Nano Therapeutics, Beijing, China),
   following the manufacturer’s protocol. The siRNA oligonucleotides were
   synthesized by Beijing D-Nano Therapeutics Co., Ltd. Sequences are
   provided in Supplementary Materials Table [82]S1.

Measurements of glucose and ATP

   Primary microglia cells or culture medium were collected, and glucose
   and ATP were measured using glucose quantification colorimetric assay
   (Nanjing Jiancheng Bioengineering Institute, Nanjing, China, A154-1-1)
   and ATP bioluminescent assay kit (Beyotime Biotechnology, Shanghai,
   China, S0026). The contents of glucose and ATP were normalized by the
   total protein.

ECAR and OCR measurement

   After 24 h of treatment, the culture medium was removed, and ECAR
   (Elabscience, Wuhan, China, E-BC-F069) and OCR (Elabscience, Wuhan,
   China, E-BC-F068) were measured using commercial kits according to the
   manufacturer’s instructions. Fluorescence was recorded using a plate
   reader at 37 °C in dynamic mode. For ECAR, readings were taken every
   2.5 min for 120 min (excitation: 490 nm; emission: 535 nm), and for
   OCR, every 2 min for 90 min (excitation: 405 nm; emission: 675 nm).
   Slopes of fluorescence curves were used to quantify ECAR and OCR.

Rac1 activation assay

   Primary microglia were transfected with Arhgap25 siRNA as described in
   2.9 and then collected to detect the Rac1 activation using the Active
   Rac1 Detection Kit (NewEast Biosciences, China, 80501) following the
   manufacturer’s protocol.

RNA sequencing

   Primary microglia from the flox and cKO groups were sequenced using
   Solexa high-throughput sequencing (Biotree, Shanghai, China). RNA was
   extracted with Trizol reagent, and sequencing was followed by data
   analysis, including Venn analysis, GO, and KEGG enrichment analysis.

Targeted metabolomics

   Primary microglia were seeded at 5 × 10⁶ cells/dish. After 24 h, cells
   were washed with HBSS, collected, and subjected to metabolite
   extraction. The supernatant was analyzed by LC-MS using an ExionLC
   system and an AB Sciex Triple Quadrupole 6500 mass spectrometer in MRM
   mode. Data visualization was done using the BioDeep cloud platform.

Single-cell RNA sequencing and data analyses

   Raw data from the 10× Genomics platform were processed using Cell
   Ranger and imported into R for analysis with Seurat. Cells were
   filtered, and batch effects corrected using Harmony. PCA was performed
   with the top 2000 variable genes, and cell types were annotated using
   the SingleR package. Differentially expressed genes were identified
   using Seurat’s FindAllMarkers function. GO, KEGG, and GSEA were
   performed with the clusterProfiler package, considering terms with p
   value < 0.05 and q-value < 0.05.

Public microglia sequencing data analysis

   Microglia sequencing data were downloaded from the GEO database
   [[83]24, [84]30–[85]32]. Detailed information is provided in
   Supplementary Materials Table [86]S2.

Immunostaining

   After perfusing mice with 0.1 mol/L PBS and 4% PFA, spinal cords were
   fixed in 4% PFA for 24 h and then immersed in 30% sucrose until fully
   submerged. A 1 cm spinal cord segment was cut into 10-μm thick sections
   using a freezing microtome. For staining, sections were fixed for
   30 min, antigen-retrieved with Sodium Citrate Solution at 90 °C for
   30 min, and blocked in 5% BSA and 0.3% Triton X-100 for 1 h. Sections
   were incubated overnight with primary antibodies at 4 °C, then washed
   with PBS and incubated with secondary antibodies for 1 h at room
   temperature. Primary antibody details are in Supplementary Materials
   Table [87]S3. Sections were stained with DAPI to visualize the nucleus,
   and images were acquired with light and confocal microscopes, then
   analyzed using ImageJ and Photoshop.

Western blotting

   Microglia were lysed in ice-cold RIPA Buffer (FDbio, Hangzhou, China,
   FD009) with 100 mM PMSF, incubated at 4 °C for 30 min, and extracted
   with 5× loading buffer. Proteins were boiled at 100 °C for 10 min and
   separated using SDS-PAGE before being transferred to nitrocellulose
   membranes (Merck Millipore, Darmstadt, Germany, ISEQ00010). After
   blocking with 5% skim milk for 1 h, membranes were incubated overnight
   with primary antibodies at 4 °C, washed, and incubated with
   HRP-conjugated secondary antibodies for 2 h. The western blot was
   detected using the ECL detection kit (Biosharp, Anhui China, BL523B),
   and images were analyzed with ImageJ software.

Statistical analysis

   All statistical analyses were performed in Prism Version 8.0.2
   (GraphPad). Data were expressed as mean ± SD from at least three
   independent experiments. statistical significance was assessed by the
   Student’s t test for comparison between two groups or ANOVA with post
   hoc tests by Bonferroni correction for comparisons among three or more
   groups; p < 0.05 was considered statistically significant.

Results

RhoA expression is upregulated in microglia following spinal cord injury

   In the past, studies mainly focused on the activity of RhoA, but
   recently the total amount of RhoA changes has been found in several
   diseases. By analyzing the GEO databases, it was observed that RhoA
   expression is elevated following SCI and is significantly increased
   across multiple datasets (Fig. [88]1A). Western blotting assay on the
   injured spinal cord tissues at 1, 3, 7, 14, 28, and 42 days post injury
   (dpi) revealed an increase in RhoA protein level (Fig. [89]1B), which
   is consistent with previous reports, and indicated that RhoA is a key
   regulatory factor in SCI. By sorting out the [90]GSE196928 dataset,
   RhoA was found that extensively expressing in microglia after SCI (Fig.
   [91]1C–E). Immuno-staining illustrated that RhoA expression in
   microglia was increased after SCI (Fig. [92]1F). These findings suggest
   that the change of RhoA expression in microglia might be involved in
   SCI repair.

Fig. 1. RhoA expression in microglia was up-regulated by spinal cord injury.

   [93]Fig. 1
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   A Changes of RhoA mRNA expression in injured spinal cord obtained from
   different GEO databases. B Temporal changes of RhoA protein level in
   spinal cord following SCI (n = 5). C Major cell type classification in
   the [95]GSE196928 dataset. D Distribution map of RhoA expression. E
   RhoA expression levels across different cell types. F Immunostaining
   shows the RhoA expression in microglia (IBA-1^+) at different stages
   after SCI (n = 3). Data are presented as mean ± SD. Statistical
   significance: ∗p < 0.05, ∗∗p < 0.005, ∗∗∗p < 0.001 (vs. sham group).

Microglial RhoA deficiency delays the morphological and functional recovery
in the spinal cord injury mice

   To investigate the role of microglial RhoA in SCI, we created a
   microglial RhoA conditional knockout (cKO) mouse model. Western blots
   and immunostaining confirmed successful RhoA ablation in microglia in
   the cKO mice (Supplementary Fig. [96]S1). The cKO and RhoA-flox (flox,
   control) mice underwent SCI or sham surgery. At 28 dpi, in the SCI
   group, the cKO mice showed significantly worse BMS scores, swimming
   test results, and gait analysis scores compared to the flox group,
   while there were no significant differences between the two groups in
   the sham-treated mice (Fig. [97]2A–C). These results suggest that RhoA
   deficiency impairs functional recovery after SCI.

Fig. 2. Microglial RhoA deficiency delays the spinal cord injury recovery.

   [98]Fig. 2
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   A Representative foot print images of mice at 28 dpi. B BMS scores at
   different stages after SCI. C Swimming test was used to evaluate
   locomotor function in mice at 28 dpi. D Gastrocnemius muscle weight at
   28 dpi. E H&E staining of the gastrocnemius muscle at 28 dpi. F NF200
   Mean fluorescence intensity and the number of NeuN-positive cells at
   various distances from the SCI lesion center of the spinal cords
   present 1000 μm proximal (n = 3). Data are presented as mean ± SD; each
   dot represents an individual mouse. Statistical significance: n.s.
   nonsignificant; ∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001.

   The recovery of motor function after SCI depends heavily on
   gastrocnemius muscle atrophy. At 28 dpi, the gastrocnemius muscle
   weight was significantly lower in the cKO group compared to the flox
   group. HE staining further revealed a notable reduction in myofiber
   area in the cKO group (Fig. [100]2D, E).

   Immunostaining for NF200 (axon marker) and NeuN (neuron marker) was
   used to assess spinal cord morphology. Results showed a significant
   decrease in NF200 immuno-intensity and the number of neurons in the
   para-epicenter of the injured spinal cord in the cKO group (Fig.
   [101]2F). These findings suggest that microglial-specific RhoA knockout
   impairs both morphological and functional repair after SCI.

RhoA deficiency alleviates the microglial capabilities of proliferation,
phagocytosis, and migration

   Using IBA-1 immunostaining to label microglia, we found that the number
   of microglia in both injured and uninjured spinal cord sections at 28
   dpi was significantly lower in the cKO group compared to the flox group
   (Fig. [102]3A). This suggests that RhoA may influence microglial death
   or proliferation. To explore this, we examined markers for ferroptosis
   (SLC7A11, GPX4), cuproptosis (FDX1), autophagy (LC3B), and apoptosis
   (Bcl2/Bax) (Supplementary Fig. [103]S2). The results showed that RhoA
   knockout inhibited apoptosis and promoted autophagy, but had no
   significant effect on ferroptosis or copper-induced cell death. These
   findings suggest that the reduced microglial number in the cKO group is
   primarily due to impaired proliferation rather than increased cell
   death.

Fig. 3. RhoA knockout inhibits the microglial proliferation.

   [104]Fig. 3
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   A Immunostainings show the number of Microglia (Iba-1^+) cells in
   spinal cord tissue at 28 dpi. B Immunostainings show Ki67^+ (green) and
   IBA-1^+ (red) microglial cells in the SCI at 7 dpi. C Western blotting
   shows the related protein levels of proliferating and cell cycle in the
   primary cultured microglia. Immunostainings show the Ki67^+ cells (D),
   PCNA^+ cells (E) and Edu^+ cells (F) in primary cultured microglia.
   Data are presented as mean ± SD; each dot represents an individual
   mouse. Statistical significance: ∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001.

   To confirm this, we performed Ki67/IBA-1 double immunostaining at 7
   dpi, when microglial proliferation peaks. The data showed a
   significantly lower ratio of proliferating cells (Ki67+/IBA-1+) in the
   cKO group compared to the flox group (Fig. [106]3B). In primary
   microglial cultures, RhoA knockout significantly reduced the expression
   of proliferative markers (PCNA, PH3, Ki67, CDK1, CDK2) and increased
   cell cycle arrest markers (p-Chk1/Chk1) (Fig. [107]3C). Immunostaining
   also showed a significant decrease in Ki67+, PCNA+, and EdU+ microglia
   in cKO cultures compared to flox cultures (Fig. [108]3D–F).

   Microglial phagocytosis is critical for clearing debris following SCI,
   so we assessed this function in vivo and in vitro. IBA-1 and ORO
   staining revealed fewer ORO+ lipid droplets in cKO microglia in the
   injured spinal cord at 7 dpi (Fig. [109]4A). In vitro, the phagocytic
   capacity was evaluated by measuring myelin fragment uptake over time.
   CD68, a phagocytosis marker, showed significantly lower fluorescence
   intensity protein level in the cKO group (Fig. [110]4B, C).
   Additionally, MBP+ myelin debris content was reduced in the cKO group
   compared to the flox group (Fig. [111]4D, E). These findings indicate
   that RhoA is essential for maintaining microglial phagocytic function.

Fig. 4. RhoA knockout attenuates the microglial phagocytosis and migration.

   [112]Fig. 4
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   A Immunostainings show IBA-1^+ and ORO^+ on the cross sections of the 7
   dpi spinal cord tissue. B, C Immunostaining and Western blotting show
   the CD68 expression in the cultured microglia (IBA-1^+). D, E
   Immunostaining and Western blotting demonstrate microglia engulfed
   myelin debris (MBP^+) in the myelin debris treated microglial cultures.
   F Representative images of Transwell assay. Data are presented as
   mean ± SD; each dot represents an individual mouse. Statistical
   significance: ∗p < 0.05; ∗∗p < 0.01.

   Finally, since migration is important for microglial roles in SCI
   repair, we performed a Transwell assay and found that RhoA knockout
   significantly impaired microglial migration (Fig. [114]4F). This
   suggests that RhoA is crucial for microglial migration.

Multi-omics analysis reveals RhoA regulation of the glycolysis and oxidative
phosphorylation in microglia

   To investigate how microglial RhoA knockout affects SCI repair, we
   performed single-cell RNA sequencing (scRNA-Seq) at 7 dpi. After basic
   annotation and dimensionality reduction using t-SNE, we identified 12
   cell clusters, including microglia, macrophages, astrocytes, and
   oligodendrocytes (Fig. [115]5A and Supplementary Fig. [116]S3A). Given
   the importance of metabolic reprogramming in microglial activity, we
   focused on metabolism. GSEA analysis of microglial clusters revealed
   that oxidative phosphorylation and glycolysis were significantly
   downregulated in the cKO group compared to the flox group (Fig.
   [117]5B). Further analysis of microglia identified 10 subclusters and
   their markers (Fig. [118]5C, D). Pseudotime analysis showed the
   distribution of subclusters along the pseudotime trajectory (Fig.
   [119]5E, F). Supplementary Fig. [120]S4 displays the top 20 GSEA
   results for each subcluster. Notably, glycolysis was significantly
   altered in subclusters 2, 3, 4, and 7, while oxidative phosphorylation
   was affected in subclusters 0, 2, 3, 4, and 7. The UMAP plot
   highlighted clusters linked to these pathways (Fig. [121]5G, H).
   CellChat interaction analysis suggested that microglial subclusters
   with altered metabolism interact with astrocytes, endothelial cells,
   neurons, and oligodendrocytes (Fig. [122]5I and Supplementary Fig.
   [123]S5). Overall, scRNA-Seq data suggest that RhoA knockout modulates
   glycolysis and oxidative phosphorylation in microglia, influencing
   their function post-SCI.

Fig. 5. Multi-omics insights into RhoA-mediated regulation of glycolysis and
oxidative phosphorylation in microglia.

   [124]Fig. 5
   [125]Open in a new tab

   A t-SNE and unsupervised clustering of all cells based on biomarkers. B
   GSEA analysis highlighting differences in microglial subsets compared
   to other cell types. C High-resolution reclustering of microglia into
   ten distinct clusters. D Expression of key markers identified in the
   reclustered microglia. E Pseudotime trajectory analysis of microglial
   cells. F Visualization of ten microglial subclusters, each represented
   by a distinct color. Microglial subsets with significant differences in
   oxidative phosphorylation pathways (G subcluster 0, 2, 3, 4, 7) and
   glycolysis pathways (H subcluster 2, 3, 4, 7) identified through GSEA
   analysis. I CellChat analysis showing interactions between
   glycolysis-differentiated microglia (subcluster 2, 3, 4, 7) and other
   cell types. J PCA of bulk mRNA transcriptomics in primary microglial
   cells. K Heatmap illustrating the results of mRNA sequencing analysis.
   GSEA analysis of glycolysis-related (L) and oxidative
   phosphorylation-related (M) differences between flox and cKO groups. N
   PCA of metabolomic profiling data. O, P Comparative analysis of
   glycolysis and TCA cycle metabolic changes between cKO and flox groups.
   RhoA deletion decreases ATP production (Q), accompanied by reduced ECAR
   (R), OCR (S), and glucose consumption in the culture medium (T). Data
   are presented as mean ± SD; each dot represents an individual mouse.
   Statistical significance: ***p < 0.001.

   To explore RhoA’s role further, we isolated primary microglia and
   performed bulk mRNA sequencing. PCA analysis showed clear separation
   between the flox and cKO groups (Fig. [126]5J), indicating gene
   expression changes due to RhoA knockout. The heatmap highlighted
   significant gene expression differences (Fig. [127]5K). GSEA analysis
   confirmed that glycolysis and oxidative phosphorylation were
   downregulated in the cKO group (Fig. [128]5L, M), consistent with
   scRNA-Seq results. Targeted metabolomics revealed metabolic changes
   between the two groups, with PCA analysis showing clear separation
   (Fig. [129]5N). Alterations in glycolysis-related and TCA cycle
   metabolites are shown in Fig. [130]5O, with pathway mapping in Fig.
   [131]5P. ATP, ECAR, and OCR measurements revealed that ATP levels, as
   well as ECAR and OCR values, were reduced in cKO microglia (Fig.
   [132]5Q–S). In addition, glucose consumption was higher in the flox
   group, as indicated by the decreased glucose concentration in the
   culture medium (Fig. [133]5T). Together, scRNA-Seq, mRNA sequencing,
   and metabolomics data suggest that RhoA knockout in microglia alters
   glycolysis and oxidative phosphorylation, impacting their biological
   function.

Arhgap25/HIF-1α pathway is involved in RhoA regulating microglial
biofunctions

   The aforementioned results suggested that RhoA regulates microglial
   biofunction through metabolic reprogramming, however which signaling
   pathway is involved in this regulation remains unclear. To figure out
   this issue, our primary cultured microglia mRNA transcriptomic analysis
   revealed significant differential expression of Arhgap25, as shown in
   the volcano plot (Fig. [134]6A, B). Additionally, KEGG pathway
   enrichment analysis indicated that RhoA knockout affects the HIF-1
   pathway (Fig. [135]6C). Previous studies have demonstrated that
   Arhgap25 regulates the proliferation of human pancreatic cancer cells
   via HIF-1α [[136]33], leading us to hypothesize that RhoA modulates
   glycolytic processes through Arhgap25/HIF-1α pathway. Therefore, main
   components of this pathway were detected by Western blotting. The
   results showed a significant increase in Arhgap25 expression in the cKO
   group, accompanied by a decrease in HIF-1α protein levels and a
   corresponding increase in HIF1AN protein levels (Fig. [137]6D).
   Furthermore, the expression of key glycolytic enzymes, HK2, PKM2 and
   LDHA, were reduced. Immunostaining confirmed that Arhgap25 expression
   was significantly higher, while HIF-1α expression was markedly lower in
   the cKO group (Fig. [138]6E, F).

Fig. 6. RhoA regulates the microglial biofunctions by regulating the HIF-1
pathway through Arhgap25.

   [139]Fig. 6
   [140]Open in a new tab

   A Highlight Arhgap25 in volcano diagram of mRNA-sequence. B Arhgap25
   mRNA level in primary microglia in mRNA-sequence. C KEGG enrichment
   analysis of significant changes gene in mRNA-sequence. D Western blots
   show the Arhgap25, HIF-1α, HIF1AN, HK2, PKM2 and LDHA protein
   expression levels in primary cultured microglia. E, F Immunostainings
   show Arhgap25 and HIF-1α expression in primary cultured microglia. Data
   are presented as mean ± SD; each dot represents an individual mouse.
   Statistical significance: ∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001.

Arhgap25 knockdown reverses the effects of RhoA deficiency on the microglial
biological functions

   To validate if the Arhgap25/HIF-1α pathway is involved in RhoA
   regulation of microglial functions, we developed three siRNAs targeting
   Arhgap25. Among them, siArhgap25-1 showed the best knockdown efficiency
   in primary microglia (Supplementary Fig. [141]S6) and was used in
   further experiments. Arhgap25 knockdown enhanced cellular metabolism,
   as reflected by increased ATP production, ECAR, and OCR in the cKO
   group (Fig. [142]7A–C), suggesting it reversed the energy deficiency
   caused by RhoA knockout. Moreover, protein levels of HIF-1α, HK2, PKM2,
   and LDHA were significantly elevated in the cKO group after Arhgap25
   knockdown (Fig. [143]7D). These results suggest that RhoA regulates
   microglial metabolism through the Arhgap25/HIF-1α pathway.

Fig. 7. Arhgap25 knockdown reverses the regulatory effect of RhoA knockout on
the biological function of microglia.

   [144]Fig. 7
   [145]Open in a new tab

   ATP concentration (A), ECAR (B) and OCR (C) levels in primary cultured
   microglia after siArhgap25 transfection. D Western blotting shows the
   expression of Arhgap25, HIF-1α, HK2, PKM2 and LDHA after siArhgap25
   transfection. E Western blotting shows the expression of PCNA and PH3
   protein after siArhgap25 transfection. Immunostaining shows the
   PCNA^+(F), Ki67^+ (G) and Edu^+ (H) in primary cultured microglia after
   transfection with siArhgap25. Data are presented as mean ± SD; each dot
   represents an individual mouse. Statistical significance: ∗p < 0.05;
   ∗∗p < 0.01; ∗∗∗p < 0.001.

   We then assessed the effects of siArhgap25 on microglial proliferation.
   SiArhgap25 reversed the proliferation inhibition in the cKO group, as
   indicated by increased PCNA and PH3 expression (Fig. [146]7E), and
   consistent results from PCNA, Ki67, and Edu staining (Fig. [147]7F–H).
   For phagocytic function, CD68 and MBP staining and Western blotting
   showed that siArhgap25 restored phagocytosis in cKO microglia (Fig.
   [148]8A–C). Additionally, Transwell migration assays revealed that
   siArhgap25 reversed the migration impairment in the cKO group (Fig.
   [149]8D). In summary, these findings indicate that RhoA regulates
   microglial functions-proliferation, phagocytosis, and migration-via the
   Arhgap25/HIF-1α pathway and metabolic reprogramming.

Fig. 8. Arhgap25 knockdown reverses the effects of RhoA cKO on the
phagocytosis and migration of microglia.

   [150]Fig. 8
   [151]Open in a new tab

   A Immunostaining shows the CD68 expression in the cultured microglia
   (IBA-1^+). B Immunostaining shows the myelin debris (MBP^+) engulfed in
   the cultured microglia after the microglia were co-cultured with debris
   for 1 h. C Western blotting show the CD68 level in microglia (culture
   without debris) and MBP level in microglia (co-cultured with debris for
   1 h). D Representative images of Transwell assay. Data are presented as
   mean ± SD; each dot represents an individual mouse. Statistical
   significance: ∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001.

Discussion

   Research on SCI has spanned several decades, yielding numerous exciting
   breakthroughs. In particular, recent advancements in brain-spinal
   interface [[152]34–[153]36] and epidural electrical stimulation
   [[154]37–[155]39] have shown significant clinical efficacy. However, it
   is regrettable that despite extensive studies aimed at improving the
   microenvironment or the neuronal intrinsic regenerative capability to
   promote SCI repair, successful clinical applications remain rare
   [[156]12, [157]13, [158]40]. This underscores the inherent complexity
   of SCI pathophysiology.

   Following SCI, fragmented myelin sheaths, dead cells, and activated
   inflammatory factors release a variety of inhibitory molecules that
   impede neural regeneration [[159]41–[160]44]. Although previous
   approaches, such as antibodies or enzymes targeting chondroitin sulfate
   proteoglycans (CSPGs), have attempted to mitigate these inhibitors,
   they fail to comprehensively clear all inhibitory factors [[161]45].
   Subsequent researches identified RhoA signaling as a critical pathway
   mediating the effects of many inhibitory molecules in the SCI
   microenvironment [[162]45]. Up to now, the role of RhoA as a negative
   regulator of axonal regeneration has been well established [[163]14,
   [164]46, [165]47]. Our recent study also revealed that
   neuronal-specific RhoA deletion delays dendritic degeneration and
   promotes dendritic regeneration after the axonal injury [[166]48]. All
   the existing findings lead to the hypothesis that inhibiting RhoA could
   promote SCI repair, yet RhoA-targeted inhibitors have failed to meet
   expectations in clinical trials, emphasizing the urgent need to further
   explore the role and mechanisms of RhoA in SCI [[167]12, [168]13].

   Following SCI, RhoA upregulation and activation are not confined to
   neurons; these also occur in astrocytes and microglia [[169]2, [170]49,
   [171]50]. Recent studies with RhoA conditional knockout mice
   demonstrated that neuronal RhoA deficiency accelerates the SCI repair,
   while astrocyte RhoA depletion attenuates the SCI recovery [[172]14].
   Compared to neuron and astrocyte, microglia in the injured spinal cord
   exhibit the more significant and sustained upregulation of RhoA
   [[173]24, [174]51]. Given the pivotal role of microglia in SCI, this
   raises an intriguing question: whether RhoA plays pivotal role in
   microglial biofunctions, and how do they influence SCI repair?

   Herein, microglia-specific RhoA knockout mice were used to address
   above questions. Collected data revealed that RhoA deletion in
   microglia significantly impeded the structural and functional
   recoveries post-SCI. To our knowledge, this is the first study to
   reveal that microglial RhoA is essential for SCI repair. Even some
   studies mentioned the RhoA playing role in spinal cord injury is
   related to microglial inflammation [[175]52], their manipulations are
   not cell-specific, so their results might be interrupted by RhoA’s
   roles in other cells but not only microglia.

   Secondly, this study firstly reveals the role of RhoA in microglial
   capabilities of proliferation, migration and phagocytosis. Previous
   studies mainly focused on that RhoA regulates microglia producing
   inflammatory factors [[176]53], nobody reports the proliferation,
   migration and phagocytosis in spinal cord injury. While, these
   biofunctions are critical for microglia to play role in Wallerian
   degeneration after SCI and produce favorable microenvironment for
   axonal regeneration [[177]54–[178]57].

   Thirdly, this study provides a novel signaling pathway to explain the
   role of RhoA in microglia. Arhgap25/HIF-1α pathway mediated glucose
   metabolism might enlighten a novel insight to enrich our understanding
   on the complex role of RhoA. Although RhoA has long been regarded as an
   important switching molecule in regulating actin cytoskeleton
   reorganization [[179]58], recent studies have also shown that RhoA can
   affect microtubule dynamics [[180]59] or the expressions of certain
   transcription factors [[181]60, [182]61] independently of actin
   dynamics. Herein, a series results of sequence analyses and validation
   experiments revealed that RhoA deletion in microglia alters the
   expression of the cytoskeleton-related molecules (mDia and ROCK2)
   (Supplementary Materials Fig. [183]S7A, B). Which indicate cytoskeletal
   remodeling is also involved in the role of RhoA in microglia.
   Noticeably, we identified that RhoA depletion reduced glycolytic
   activity and ATP production through Arhgap25/HIF-1α signaling.
   Consequently, energy deficiency impairs microglial proliferation and
   migration, reducing the microglial population and diminishing
   phagocytic capacity. These impairments hinder the timely clearance of
   dead cells and debris in the SCI microenvironment, ultimately
   compromising structural and functional recoveries. Present data also
   illustrated that siArhgap25 resulted in a higher ratio of Rac1-GTP/
   total Rac1 in the primary cultured microglia, which means suppression
   of Arhgap25 promotes Rac1 activation (Supplementary Materials Fig.
   S[184]7C). This is consistent with the previous studies of that
   [[185]62–[186]64], Rac1-GTP has been shown to enhance cell
   proliferation, migration, and glycolytic activity. However, the exact
   role of Rac1 activation in RhoA modulating glucose metabolism in
   microglia remains unclear. Further investigation is needed to clarify
   the potential crosstalk between RhoA and Rac1 signaling in microglial
   function.

Conclusion

   In summary, this is the first study to reveal that the SCI resulted
   RhoA upregulation in microglia plays positive roles in SCI repair which
   may due to RhoA regulating microglial bio-functions of proliferation,
   migration and phagocytosis through the Arhgap25/HIF-1α pathway mediated
   modulating glycolysis. Even RhoA inhibition in neurons is widely
   regarded as a prospective strategy for promoting axonal regeneration
   after SCI [[187]65], recent study indicated astrocytic RhoA specific
   knockout alleviates SCI repair [[188]14] and the evidences of this
   study demonstrated microglial RhoA specific knockout is also not
   conducive to SCI repair, which suggests that RhoA inhibition for SCI
   repair has to consider the cell specific targeting. Moreover, this
   study hints that targeting Arhgap25/HIF-1α pathway to regulate
   microglial glucose metabolism might be another potential therapeutic
   strategy for SCI.

Supplementary information

   [189]supplementary data^ (973KB, pdf)
   [190]WB-RAW-DATA^ (385.3KB, pdf)

Acknowledgements