PRMT6 is required for initiating and amplifying macrophage-induced inflammation in heterotopic ossification by increasing CCL2 expression
Wenxiang Chu, Weilin Peng, Zhengqiang Wu, Yu Xiong, Zhongya Gao, Yang Li, Bangke Zhang, Liang Wang, Haibin Wang, Chaofeng Han, Xuhua Lu
TL;DR
PRMT6 promotes inflammation in a bone disorder called heterotopic ossification by increasing CCL2, offering a potential early treatment target.
Contribution
PRMT6 is identified as a novel epigenetic driver of macrophage-induced inflammation in HO.
Findings
PRMT6 deletion or inhibition reduces macrophage accumulation and HO without affecting tendon repair.
PRMT6 enhances CCL2 expression via epigenetic modification of the Ccl2 promoter.
CCL2 supplementation partially restores HO in PRMT6-deficient mice.
Abstract
Heterotopic ossification (HO) is a debilitating disorder marked by ectopic bone formation in soft tissues, frequently triggered by inflammation after trauma. While macrophage-driven inflammation plays a critical role in HO pathogenesis, the molecular mechanisms governing its initiation, amplification and resolution remain elusive. Using a trauma/burn injury (TBI)-induced mouse model of HO, we identified rapid and sustained macrophage accumulation at the injury site during the early inflammatory phase, and macrophage depletion markedly suppressed HO formation. Transcriptomic profiling identified a pronounced upregulation of protein arginine methyltransferase 6 (PRMT6) in macrophages following injury. Genetic deletion or macrophage-targeted knockdown of Prmt6 reduced macrophage accumulation and significantly attenuated HO, without impairing tendon repair. Consistently, pharmacological…
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Figure 9- —https://doi.org/10.13039/501100001809National Natural Science Foundation of China (National Science Foundation of China)
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Taxonomy
TopicsHeterotopic Ossification and Related Conditions · Genetic Syndromes and Imprinting · Amyotrophic Lateral Sclerosis Research
Introduction
Heterotopic ossification (HO) is characterized by the aberrant formation of bone within soft tissues, typically triggered by traumatic events such as surgical interventions, neurological damage, or severe burns.^1^ This pathological condition affects more than 10% of patients undergoing invasive surgeries, 30% of those following hip arthroplasties, and up to 65% of seriously injured military personnel. The prevalence soars above 90% in cases of severe traumatic amputations.^2–4^ The clinical impact of HO is profound, often leading to chronic pain, joint stiffness, mobility impairment, and recurrent infections.^2^ Current therapeutic approaches, including NSAIDs, radiation, and surgical excision, remain largely ineffective, with high recurrence rates post-treatment.^3^ These limitations underscore a critical need for deeper insights into the molecular and cellular mechanisms driving HO pathogenesis.
Although the precise mechanisms underlying HO remain incompletely understood, it is well-established that the pathological process mimics endochondral ossification, encompassing phases of inflammation, chondrogenesis, osteogenesis, and bone maturation.^5^ Trauma or surgical injury initiates a localized inflammatory response, recruiting immune cells such as neutrophils, monocytes, and lymphocytes to the site of injury.^6,7^ These immune cells release a cascade of inflammatory mediators, which subsequently drive the recruitment and differentiation of skeletal progenitor cells into chondrocytes. These chondrocytes undergo hypertrophy, vascularization, and mineralization, eventually leading to ectopic bone formation.^8–10^ Among the immune cells involved, macrophages play a pivotal role, particularly those derived from monocytes in bone marrow cells. Macrophages are central to both normal tissue repair—such as bone remodeling and angiogenesis—and the pathology of inflammatory disorders like rheumatoid arthritis and HO.^11–14^ Their critical role in HO is linked to the inflammatory responses that promote aberrant bone formation.^6,15,16^ Despite this knowledge, the intricate cellular and molecular mechanisms that regulate the initiation, amplification and resolution of macrophage-induced inflammation in HO remain poorly defined.
Protein arginine methyltransferase 6 (PRMT6), a type 1 PRMT, has emerged as a key epigenetic regulator, catalyzing the asymmetric dimethylation of arginine residues on a range of protein substrates.^17,18^ PRMT6 methylates histone H3 at arginine 17 (H3R17me2a) and arginine 42 (H3R42me2a), modifications that facilitate transcriptional activation. This is in contrast to repressive modifications, such as H3R2me2a and H2AR26me2a, which restrict chromatin accessibility.^19–22^ Beyond its enzymatic function, PRMT6 modulates cellular processes through non-catalytic pathways.^23^ Previous studies have highlighted the involvement of PRMT6 in various biological contexts, including tumorigenesis and neurodegeneration.^24–26^ In the context of HO, PRMT6 expression is markedly elevated in injured tissues, as observed in mouse models of TBI, suggesting a potential role in the progression of HO. However, the detailed mechanisms by which PRMT6 modulates monocyte/macrophage behavior in response to inflammation during HO development remain to be fully elucidated.
In this study, we investigated the indispensable role of macrophages within the inflammatory landscape of HO and identify PRMT6 as a critical enhancer in this context. Upon initial inflammatory stimuli, PRMT6 upregulation triggers a positive feedback loop that enhances macrophage recruitment to the injury site by promoting the secretion of CCL2, a chemokine essential for initiating and amplifying inflammation in ectopic bone formation. We further demonstrate that PRMT6 activates NF-κB signaling, working in concert with this transcription factor to upregulate CCL2 transcription. Additionally, PRMT6 augments CCL2 transcription through H3R17me2a modifications at the CCL2 promoter. Importantly, PRMT6 inhibition in the early inflammatory phase effectively attenuates HO, offering promising avenues for therapeutic intervention by targeting early inflammatory responses, potentially improving outcomes for patients with HO.
Results
Macrophages are essential drivers of heterotopic ossification in a TBI-induced mouse model
To define the cellular events underlying heterotopic ossification (HO), we employed a well-established trauma/burn injury (TBI)-induced HO model that combines Achilles tendon transection with dorsal partial-thickness burn injury. μCT analysis revealed the emergence of ectopic bone at the injury site by 6 weeks post-injury, followed by marked expansion by 12 weeks (Fig. S1a, b). Consistently, histological analyses using hematoxylin and eosin (H&E) and Safranin O/Fast Green (SOFG) staining demonstrated a progressive transition from injured soft tissue to cartilage formation and ultimately mature trabecular bone, recapitulating endochondral ossification (Fig. S1c, d).
To characterize the involvement of macrophages during HO progression, we quantified CD68^+^ macrophages at multiple time points following injury. While CD68^+^ cells were nearly absent in sham controls, macrophages accumulated robustly at the injury site by 1 week post-TBI and remained detectable, albeit at lower levels, within cartilage and ossified regions up to 12 weeks (Fig. S1e, f). This macrophage accumulation coincided spatially and temporally with the recruitment of Nestin^+^ progenitor cells, encompassing mesenchymal and endothelial precursor populations,^15^ as well as CD31^+^ endothelial cells, both of which are critical for osteogenesis and angiogenesis during HO (Fig. S1g–i). These data indicate that macrophages rapidly infiltrate the injury site during the early inflammatory phase and persist throughout HO progression in concert with stromal and vascular components.
To directly assess the functional contribution of macrophages to HO formation, we depleted macrophages using PLX73086 in the TBI model (Fig. S2a). μCT analysis demonstrated a marked reduction in ectopic bone volume in macrophage-depleted mice compared to vehicle-treated controls (Fig. S2b, c). Histological examination at 9 weeks post-injury revealed well-developed trabecular bone with marrow cavities in control mice, whereas macrophage-depleted mice exhibited only sparse cartilage-like regions with minimal bone formation (Fig. S2d). Immunofluorescence analysis confirmed a substantial reduction in CD68^+^ macrophages within ossification areas following depletion (Fig. S2e, f), accompanied by a pronounced decrease in Nestin^+^ cells and CD31^+^ endothelial cells (Fig. S2g–i).
Collectively, these findings establish macrophages as indispensable orchestrators of the inflammatory microenvironment that supports stromal and vascular cell recruitment and drives ectopic bone formation following traumatic injury.
PRMT6 upregulation in macrophages during TBI
To gain deeper insights into the molecular mechanisms driving HO during the inflammatory phase, we conducted transcriptomic analysis on soft tissues from the injury site 3 days post-TBI, a time point corresponding to early inflammatory activation, and compared them with sham controls. Differential gene expression analysis revealed 4 940 significantly upregulated and 1 786 downregulated genes in injured tissues (Fig. 1a). Gene Ontology (GO) enrichment analysis of these differentially expressed genes highlighted several key biological processes involved in molecular regulation, including “Regulation of response to stimulus,” “Regulation of gene expression,” “Nucleic acid metabolic process,” and “Cellular protein metabolic process” (Fig. 1b). Venn analysis of these enriched GO terms identified 16 co-expressed genes (Fig. 1c), 13 of which were upregulated and 3 downregulated (Fig. 1d). Notably, Prmt6 emerged as one of the most highly upregulated genes, alongside canonical inflammatory mediators such as Tnf (Fig. 3d, e). To Validate these findings, we conducted q-PCR and Western blot analysis, confirming a substantial increase in Prmt6 mRNA (Fig. 1f) and protein expression (Fig. 1g, h) in the injured tissues at 3 days post-TBI. Immunofluorescence staining further demonstrated strong co-localization of PRMT6 with CD68^+^ macrophages (Fig. 1i), suggesting macrophages as a major cellular source of PRMT6 during early HO development.Fig. 1. Upregulation of PRMT6 in macrophages during trauma/burn injury. a Volcano plot showing differentially expressed genes (fold change > 2, q < 0.01) in soft tissues from the injury site of sham and trauma/burn injury (TBI) mice at 3 days post-injury. b Gene Ontology (GO) enrichment analysis of differentially expressed genes, highlighting biological processes related to injury response and molecular regulation. c Venn diagram analysis identifying co-expressed genes shared among the enriched GO biological processes shown in (b). d Heatmap of the co-expressed genes identified in (c), with Prmt6 among the most significantly upregulated genes. e Transcriptomic analysis showing significant upregulation of Prmt6 expression in injured tissues compared with sham controls. f Quantitative PCR analysis confirming increased Prmt6 mRNA expression in injured tissues at 3 days post-TBI. Western blot analysis (g) and quantitative densitometric analysis (h) demonstrating increased PRMT6 protein expression in injured tissues following TBI. i Immunofluorescence staining showing upregulated PRMT6 expression co-localizing with CD68^+^ macrophages at the injury site. j Single-cell RNA sequencing analysis of the Achilles tendon injury site, identifying 10 distinct cell clusters. k Quantification of Prmt6^+^ cells and mean Prmt6 expression across cell clusters at 3 days post-injury, showing predominant enrichment of Prmt6 expression in macrophages during the early stage of HO. Representative immunofluorescence images (l) and quantitative analysis (m) of lineage-specific co-immunostaining performed using markers for macrophages (F4/80), mesenchymal cells (PDGFRα), monocytes (Ly6C), and neutrophils (Ly6G) at the injury site 3 days post-injury (arrows indicate double-positive cells) (n = 4)
To further delineate tissue and cell-type specificity, we reanalyzed a previously published single-cell RNA sequencing dataset generated from the similar TBI-induced HO model at 3 days post-injury.^6,27^ Unsupervised clustering identified multiple cell populations within the injury site (Fig. 1j and Fig. S3). Among these populations, macrophages exhibited the highest Prmt6 expression, whereas mesenchymal cells contributed a comparatively smaller proportion of Prmt6^+^ cells (Fig. 1k). To experimentally validate these findings, we performed lineage-specific co-immunostaining using markers for macrophages (F4/80), mesenchymal cells (PDGFRα),^28^ monocytes (Ly6C), and neutrophils (Ly6G). Quantitative analysis revealed that the majority of PRMT6^+^ cells co-localized with macrophages, followed by PDGFRα^+^ mesenchymal cells, while minimal PRMT6 expression was detected in Ly6C^+^ monocytes and was nearly absent in Ly6G^+^ neutrophils at 3 days post-injury (Fig. 1l, m). Importantly, PRMT6 expression remained predominantly localized to macrophages at 7 days post-injury (Fig. S4a, b), indicating sustained macrophage-associated PRMT6 expression during early HO progression.
To determine whether inflammatory stimuli directly regulate PRMT6 expression in macrophages, we stimulated bone marrow–derived macrophages (BMDMs) with lipopolysaccharide (LPS), a classical macrophage activator,^29^ and high-mobility group box 1 (HMGB1), a damage-associated molecular pattern implicated in TBI-induced HO.^30^ LPS stimulation significantly increased Prmt6 mRNA expression (Fig. S5a) and PRMT6 protein levels (Fig. S5b, c). Similarly, HMGB1 induced a dose-dependent increase in PRMT6 protein expression, as confirmed by Western blot (Fig. S5d, e) and immunofluorescence staining (Fig. S5f, g).
Collectively, these findings identify PRMT6 as a macrophage-enriched inflammatory regulator during the early phase of TBI-induced HO, supporting its potential role as a therapeutic target for early intervention aimed at preventing pathological ossification.
Prmt6 deficiency impairs macrophage accumulation and suppresses heterotopic ossification
To determine the role of PRMT6 in regulating macrophage-driven inflammation and its contribution to heterotopic ossification (HO), we employed the TBI model in Prmt6 knockout mice and wild-type littermate controls. During the early inflammatory phase, Prmt6-deficient mice exhibited a marked reduction in macrophage accumulation at the injury site compared with wild-type controls, indicating that PRMT6 is required for efficient macrophage recruitment following injury (Fig. 2a, b). At later stages, wild-type mice developed extensive heterotopic bone formation by 15 weeks post-injury, whereas Prmt6 knockout mice showed a pronounced reduction in ectopic bone volume, as assessed by μCT analysis (Fig. 2c, d). Histological examination further revealed well-organized, marrow-containing ectopic bone structures in wild-type mice, in contrast to minimal and poorly developed ossification in Prmt6-deficient animals (Fig. 2e). Consistent with these findings, Prmt6 knockout mice displayed significantly fewer CD68^+^ macrophages within ossification regions (Fig. 2f, g), accompanied by a reduction in Nestin^+^ stromal cells (Fig. 2h) and CD31^+^ endothelial cells (Fig. 2i), indicating impaired inflammatory niche formation and disrupted stromal–vascular coupling.Fig. 2Prmt6 deficiency impairs macrophage recruitment and suppresses heterotopic ossification. a, b Quantitative analysis of CD68^+^ macrophages in the injury region at 4 days post–trauma/burn injury shows a significant reduction in Prmt6 knockout mice compared with wild-type controls, indicating impaired macrophage accumulation during the early inflammatory phase. c, d μCT reconstructions at 15 weeks post-injury demonstrating marked suppression of heterotopic ossification in Prmt6 knockout mice, with a significant reduction in ectopic bone volume compared with wild-type controls. e Representative histological staining (H&E and SOFG) at 15 weeks post-injury illustrating diminished ectopic bone formation in Prmt6 knockout mice relative to wild-type controls. Immunofluorescence staining and quantitative analysis showing significantly reduced numbers of CD68^+^ macrophages (f, g), Nestin^+^ stromal cells (h), and relative fluorescence intensity of CD31^+^ endothelial cells (i) in the injury regions of Prmt6 knockout mice compared with wild-type controls (n = 4). j Representative H&E-stained sagittal sections of the injury site at 10 weeks post-injury. k Modified Movin’s score assessing tendon repair based on sagittal H&E-stained sections (n = 4). Representative immunofluorescence images of Fibronectin and Collagen I (l) and corresponding quantitative analyses (m, n) at the injury site (n = 4)
Given that HO represents a pathological outcome of injury repair, we next assessed whether reduced HO in Prmt6-deficient mice was associated with impaired tissue restoration. Histological evaluation of the injured Achilles tendon revealed that Prmt6 deficiency did not compromise tendon repair. Instead, H&E staining and tendon histological scoring demonstrated a reduction in local defect area and improved tissue organization in Prmt6^−/−^ mice compared with wild-type controls (Fig. 2j, k, and Fig. S6).^31^ Consistent with these morphological changes, immunofluorescence analysis showed a significant reduction in Fibronectin^+^ and Collagen I^+^ scar tissue deposition within the injury site of Prmt6^−/−^ mice (Fig. 2l–n). Together, these findings indicate that loss of PRMT6 attenuates pathological ossification without impairing tendon repair, and may promote a more organized regenerative response following injury.
Macrophage-intrinsic PRMT6 drives inflammatory amplification and HO initiation
To directly assess whether the observed in vivo phenotype is attributable to macrophage-intrinsic PRMT6, we generated a myeloid/macrophage-targeted Prmt6 knockdown model using AAV9-LysM-EGFP-miR30-shRNA targeting Prmt6 or a negative control sequence (Fig. 3a).^32,33^ Selective knockdown of Prmt6 in macrophages significantly reduced CD68^+^ macrophage accumulation at the injury site during the early inflammatory stage (Fig. 3b–d) and resulted in a marked attenuation of HO formation at later time points (Fig. 3e–g). Macrophage-targeted Prmt6 knockdown also resulted in reduced CD68^+^ macrophage accumulation within ossification regions (Fig. 3h, i), accompanied by decreased Nestin^+^ stromal cells (Fig. 3j) and CD31^+^ endothelial cells (Fig. 3k). Importantly, macrophage-specific Prmt6 knockdown did not adversely affect tendon healing. Histological analyses revealed a smaller tendon defect area and higher histological repair scores in mice with macrophage-targeted Prmt6 knockdown compared with control mice (Fig. 3l, m, and Fig. S7). In parallel, immunofluorescence staining demonstrated a marked reduction in Fibronectin^+^ and Collagen I^+^ scar formation within the injury region following macrophage-specific Prmt6 knockdown (Fig. 3n–p). These results indicate that suppression of macrophage-intrinsic PRMT6 signaling limits pathological inflammation and ectopic ossification while preserving—and potentially facilitating—effective tendon repair.Fig. 3. Macrophage-specific Prmt6 knockdown attenuates heterotopic ossification and promotes tendon repair. a Schematic illustration of macrophage-specific Prmt6 knockdown using AAV9-LysM-EGFP-miR30-shRNA and the experimental timeline. Representative immunofluorescence images showing CD68, EGFP, and PRMT6 expression (b) and quantitative analysis of PRMT6 fluorescence intensity in macrophages (c) at the injury site 3 days post-injury, confirming efficient macrophage-specific Prmt6 knockdown (n = 4). d Quantification of activated macrophages in the injury region, showing reduced macrophage accumulation following macrophage-specific Prmt6 knockdown (n = 4). e, f μCT reconstructions at 10 weeks post-injury demonstrating significant attenuation of heterotopic ossification in mice with macrophage-specific Prmt6 knockdown, with reduced ectopic bone volume compared with controls (n = 4). g Representative histological staining (H&E and SOFG) at 10 weeks post-injury showing diminished ectopic bone formation in mice with macrophage-specific Prmt6 knockdown relative to controls. Immunofluorescence staining and quantitative analysis showing reduced numbers of CD68^+^ macrophages (h, i), Nestin^+^ stromal cells (j), and relative fluorescence intensity of CD31^+^ endothelial cells (k) in the injury regions of mice with macrophage-specific Prmt6 knockdown at 10 weeks post-injury (n = 4). l Representative H&E-stained sagittal sections of the injury site. m Histological scoring of tendon healing between groups (n = 4). Representative immunofluorescence images of Fibronectin and Collagen I (n) and corresponding quantitative analyses (o, p) at the injury site (n = 4)
Early-phase PRMT6 inhibition suppresses heterotopic ossification within a restricted therapeutic window
Given the critical role of PRMT6 in TBI-induced HO, we next assessed the temporal requirement and therapeutic potential of pharmacological PRMT6 inhibition. Mice were treated daily with the PRMT6 inhibitor EPZ020411 for 3 consecutive weeks starting at distinct post-injury time points: day 1 to week 3 [EPZ(d1–w3)], week 3 to week 6 [EPZ(w3–w6)], week 6 to week 9 [EPZ(w6–w9)], or week 12 to week 15 [EPZ(w12–w15)]. Vehicle-treated mice received injections from day 1 to week 15 [Veh(d1–w15)].
μCT analysis demonstrated that only early PRMT6 inhibition during the initial inflammatory phase [EPZ(d1–w3)] markedly reduced ectopic bone volume compared with vehicle-treated controls (Fig. 4a, b). In contrast, initiation of PRMT6 inhibition at later stages failed to significantly alter HO development, indicating a restricted temporal window during which PRMT6 activity is required for HO initiation. Histological analyses (H&E and SOFG) corroborated these findings: early EPZ020411 treatment substantially reduced ectopic trabecular bone structures, whereas delayed intervention produced no appreciable impact on ossification progression (Fig. 4c, d). Consistent with these structural changes, immunofluorescence analysis revealed that early PRMT6 inhibition significantly reduced CD68^+^ macrophage accumulation within the ossification regions, an effect not observed with later treatment regimens (Fig. 4e, f). Moreover, early EPZ020411 administration decreased the abundance of Nestin^+^ stromal progenitor cells and CD31^+^ endothelial cells, populations associated with vascularization and osteogenic niche formation, whereas delayed PRMT6 inhibition did not significantly affect these cellular components (Fig. 4g–i). Importantly, early PRMT6 inhibition during the inflammatory phase not only suppressed heterotopic ossification but also improved tendon repair quality, as reflected by significantly higher modified Movin’s scores compared with vehicle-treated controls (Fig. S8). In contrast, initiation of PRMT6 inhibition at later stages exerted minimal effects on repair morphology. These findings further support the notion that early PRMT6 activity contributes to pathological inflammation and ossification, whereas its timely inhibition uncouples aberrant bone formation from normal tissue repair.Fig. 4. Early administration of the PRMT6 inhibitor EPZ020411 suppresses heterotopic ossification. Mice received EPZ020411 daily for 3 weeks starting at day 1 [EPZ(d1–w3)], week 3 [EPZ(w3–w6)], week 6 [EPZ(w6–w9)], or week 12 [EPZ(w12–w15)] post–trauma/burn injury (TBI). Vehicle-treated mice received injections from day 1 to week 15 [Veh(d1–w15)]. All mice were analyzed at 15 weeks post-injury. a Representative μCT three-dimensional reconstructions showing heterotopic bone formation across treatment regimens. b Quantification of ectopic bone volume by μCT analysis. Representative histological staining (H&E, c; SOFG, d) showing reduced ossification following early EPZ020411 treatment. Representative immunofluorescence images (e) and quantification (f) of CD68^+^ macrophages within ossification regions. g Representative immunofluorescence staining showing Nestin^+^ stromal cells and CD31^+^ endothelial cells in ossification regions. Quantification of Nestin^+^ cells (h) and relative fluorescence intensity of CD31^+^ cells (i) within ossification regions. (n = 4)
PRMT6 regulates macrophage recruitment and HO initiation via CCL2
Given the reduced macrophage accumulation and attenuated heterotopic ossification (HO) observed in Prmt6^−/−^ mice, we next investigated how PRMT6 modulates macrophage inflammatory programs under activating conditions. Transcriptomic profiling of LPS-stimulated Prmt6^−/−^ and wild-type macrophages revealed broad suppression of chemotactic signaling in the absence of PRMT6, including downregulation of the “cytokine–cytokine receptor interaction” and “chemokine signaling” pathways (Fig. 5a, b). Gene set enrichment analysis further confirmed significant repression of the chemokine signaling pathway (Fig. 5c). Consistent with these global changes, qPCR analysis validated reduced expression of multiple chemokines, including Ccl2, Ccl3, and Ccl6, indicating impaired chemokine-driven recruitment capacity in PRMT6-deficient macrophages (Fig. 5d, e).Fig. 5PRMT6 promotes CCL2 expression in macrophages under inflammatory stimulation. a, b Prmt6^+/+^ and Prmt6^−/−^ macrophages were stimulated with LPS and subjected to transcriptome sequencing. KEGG pathway enrichment analysis of downregulated (a) and upregulated (b) differentially expressed genes (DEGs) shows the top 10 significantly enriched pathways altered by Prmt6 deficiency. c GSEA indicates significant repression of chemokine/chemotactic signaling pathways in Prmt6^−/−^ macrophages compared with wild-type cells under inflammatory conditions. d Heatmap showing significantly altered chemokines and receptors (P < 0.05) between Prmt6^−/−^ and wild-type macrophages after LPS stimulation. e qPCR confirms reduced expression of chemokine genes in Prmt6^−/−^ macrophages. Western blot (f) and densitometric quantification (g) show induction of CCL2 protein in Prmt6^+/+^ macrophages after LPS stimulation, which is markedly reduced in Prmt6^−/−^ macrophages at 12 and 24 h. Immunofluorescence staining (h) and semi-quantification (i) demonstrate reduced CCL2 protein levels in Prmt6^−/−^ macrophages at 12 and 24 h after LPS stimulation. j ELISA shows decreased CCL2 secretion from Prmt6^−/−^ macrophages at 12 and 24 h post-LPS stimulation. Immunohistochemistry (k) and quantification (l) show fewer CCL2^+^ cells in the injury zones of Prmt6^−/−^ mice at day 4 post-injury compared with Prmt6^+/+^ controls (n = 3). Representative immunofluorescence of CD68 and CCL2 (m) and quantification of CCL2 fluorescence intensity in macrophages (n) at 3 days post-injury show reduced CCL2 expression after macrophage-targeted Prmt6 knockdown using AAV9-LysM-shRNA (Prmt6) (n = 4)
Among these targets, we focused on CCL2, given its established role as a key driver of monocyte/macrophage recruitment.^34^ PRMT6 deficiency markedly reduced CCL2 expression in LPS-stimulated macrophages, as confirmed by immunoblotting (Fig. 5f, g) and immunofluorescence (Fig. 5h, i). Similarly, HMGB1, a damage-associated molecular pattern implicated in TBI-induced HO, induced a dose-dependent increase in CCL2 in wild-type macrophages, whereas Prmt6^−/−^ macrophages maintained low CCL2 levels (Fig. S5h). ELISA further demonstrated reduced secretion of CCL2 from LPS-stimulated Prmt6^−/−^ macrophages (Fig. 5j). In vivo, immunohistochemistry at the early injury site revealed fewer CCL2-positive cells in Prmt6^−/−^ mice (Fig. 5k, l) and reduced CCL2 fluorescence intensity after macrophage-targeted Prmt6 knockdown (Fig. 5m, n), supporting a role for PRMT6 in shaping the early chemokine landscape in injured tissues.
Next, we analyzed published single-cell datasets from the early inflammatory phase (3 days post-injury) to identify the major cellular sources of CCL2. Temporal and cell-type–resolved analyses showed that macrophages were the predominant source of CCL2 at this stage, followed by mesenchymal cells (Fig. S9a, b). Immunofluorescence staining further corroborated these findings (Fig. S9c, d). To directly test whether macrophage-derived CCL2 is required for early inflammatory amplification and HO progression, we generated a myeloid/macrophage-targeted Ccl2 knockdown model using AAV9-LysM-EGFP-miR30shRNA(Ccl2) (Fig. S10a). Selective loss of CCL2 in macrophages significantly reduced macrophage accumulation at the injury site during the early inflammatory phase (Fig. S10b–d) and attenuated HO formation at later stages (Fig. S10e–g). Moreover, Ccl2 knockdown decreased the accumulation of CD68^+^ macrophages, Nestin^+^ cells, and CD31^+^ endothelial cells within the ectopic bone area at late time points (Fig. S10h–k). Together, these findings establish macrophage-derived CCL2 as a key functional relay downstream of PRMT6 that drives macrophage recruitment and HO initiation.
We next assessed whether PRMT6-dependent CCL2 production is sufficient to regulate macrophage migratory behavior. Using conditioned media from LPS-stimulated macrophages, scratch assays showed that wild-type conditioned media promoted macrophage migration, whereas conditioned media from Prmt6^−/−^ macrophages markedly impaired migration; this defect was rescued by supplementation with recombinant CCL2 (Fig. 6a–c). Transwell assays yielded similar results: wild-type conditioned media robustly induced macrophage migration, which was diminished with Prmt6^−/−^ conditioned media but restored upon CCL2 supplementation (Fig. 6d–f). Consistent with these in vitro findings, local CCL2 supplementation in Prmt6^−/−^ mice increased CD68^+^ macrophage accumulation in the injury region (Fig. 6g, h) and partially restored HO formation, as evidenced by μCT (Fig. 6i, j) and histological analyses (HE and SOFG staining) (Fig. 6k, l). Collectively, these results identify CCL2 as a necessary and functionally sufficient effector downstream of PRMT6 that drives macrophage recruitment during the early inflammatory stage, thereby promoting HO initiation.Fig. 6PRMT6 facilitates macrophage recruitment and HO via CCL2 secretion. a Schematic representation of the experimental setup showing the impact of conditioned media from LPS pre-stimulated Prmt6^+/+^ and Prmt6^-/-^ macrophages on macrophage migration. b, c Scratch assay and its semi-quantitative analysis showing enhanced macrophage migration with Prmt6^+/+^media, reduced with Prmt6^-/-^ media, and restored by CCL2 supplementation. d Schematic of Transwell co-culture system assessing macrophage recruitment with LPS pre-stimulated Prmt6^+/+^and Prmt6^-/-^ macrophages. e, f Transwell assays and their semi-quantitative analysis show that LPS pre-stimulated Prmt6^+/+^ macrophages effectively promote macrophage migration, a capacity significantly diminished in LPS pre-stimulated Prmt6^-/-^ cells. This diminished effect can be reversed by adding CCL2 to the culture media. g, h Immunofluorescence quantification at 7 days post-TBI reveals significantly reduced CD68^+^ macrophage density in the injured site of Prmt6^-/-^ mice compared to Prmt6^+/+^ controls, restored by local CCL2 supplementation. i–l Micro-CT analysis and histological staining (HE and SOFG) demonstrate partial recovery of ectopic bone volume and microstructure in CCL2-supplemented Prmt6^-/-^ mice 9 weeks after. (n = 4)
PRMT6 and NF-κB coactivation complex enhances macrophage Ccl2 transcription
Exploring the molecular mechanisms by which PRMT6 modulates CCL2 expression in macrophages, we focused on the NF-κB pathway, known to be an important transcription factor for the Ccl2 gene. GSEA indicated a significant downregulation of the NF-κB pathway in Prmt6-deficient macrophages upon LPS stimulation (Fig. 7a). Western blot analysis supported this finding, showing reduced NF-κB activation in the absence of PRMT6 (Fig. 7b–d). Immunofluorescence demonstrated that activated NF-κB in wild-type macrophages prominently translocated to the nucleus, an effect markedly reduced in Prmt6-deficient cells (Fig. 7e, f). Co-immunoprecipitation experiments confirmed that PRMT6 interacts with activated NF-κB to form the PRMT6/NF-κB complex, suggesting direct interaction of PRMT6 and NF-κB (Fig. 7g, h). Ch-IP assays further demonstrated that the PRMT6/NF-κB complex binds to the Ccl2 promoter, enhancing transcriptional activity (Fig. 7i–k).Fig. 7PRMT6 regulates macrophage CCL2 transcription through the NF-κB Pathway. a GSEA reveals reduced activation of the NF-kB signaling pathway in Prmt6-deficient macrophages compared to wild-type following LPS stimulation. b Western blot confirms increased PRMT6 expression and p-p65 activation in wild-type macrophages under LPS stimuli, with significant suppression in Prmt6-deficient cells. Semi-quantitative analysis indicates significant suppression of p-p65 activation in Prmt6-deficient macrophages at 12 and 24 h post-LPS stimulation (c), along with an increase in PRMT6 expression observed in wild-type macrophages (d). e Immunofluorescence staining shows enhanced p-p65 fluorescence intensity localized mainly in the nuclei of wild-type macrophages post-LPS stimulation, which is significantly reduced in Prmt6-deficient cells. f Semi-quantitative analysis confirms significantly lower p-p65 fluorescence intensity in Prmt6-deficient macrophages at 12 and 24 h post-stimulation compared to wild-type. LPS stimulation of wild-type mouse primary macrophages followed by immunoprecipitation using anti-PRMT6 (g) or anti-p-p65 antibodies (h). i Schematic representation of the Ccl2 gene transcription start site (TSS) and NF-kB binding sites on its promoter. ChIP-qPCR analysis of p-p65 (j) and PRMT6 (k) enrichment at the Ccl2 promoter in LPS-stimulated Prmt6^+/+^ and Prmt6^-/-^ macrophages. (n = 3)
PRMT6 catalysis at the Ccl2 promoter via H3R17me2a modulation enhances transcription
To elucidate the specific contributions of PRMT6’s enzymatic functions, we contrasted the effects of overexpressing the catalytically active PRMT6 with a catalytically inactive PRMT6-dead mutant. Overexpression of active PRMT6 significantly enhanced CCL2 expression and NF-κB activation following LPS stimulation compared to the scramble control (Fig. 8a–c). Interestingly, overexpression of PRMT6-dead also promoted CCL2 expression and NF-κB activation relative to the scramble control, but to a lesser extent for CCL2 expression when compared to the active PRMT6. Notably, there was no significant difference in NF-κB activation between cells overexpressing PRMT6 and PRMT6-dead (Fig. 8a–c). This differential impact underscores the role of PRMT6’s enzymatic activity in specifically enhancing CCL2 expression beyond its structural contribution to NF-κB activation.Fig. 8PRMT6-mediated methylation at the Ccl2 promoter enhances transcription via H3R17me2a modification. a Western blot analysis following overexpression of vehicle (Veh), PRMT6, or a catalytically inactive PRMT6-dead mutant in primary mouse macrophages, showing levels of CCL2 and p-p65 before and 24 h after LPS stimulation. b Semi-quantitative analysis demonstrates that PRMT6 overexpression increases CCL2 protein levels under inflammatory conditions, whereas PRMT6-dead overexpression results in significantly lower CCL2 levels compared to active PRMT6. c The levels of p-p65 protein are elevated in cells overexpressing PRMT6 or PRMT6-dead compared to controls; however, PRMT6-dead does not significantly reduce p-p65 levels compared to active PRMT6 overexpression. Western blot analysis (d) and its semi-quantitative evaluation (e) reveal that overexpression of PRMT6, not PRMT6-dead, significantly increases overall H3R17me2a modification levels in macrophages following LPS stimulation. Immunofluorescence (f) and semi-quantitative analysis (g) indicate that H3R17me2a modifications in the nuclei of wild-type macrophages are significantly higher than in Prmt6 knockout cells under inflammatory conditions. h ChIP-qPCR analysis of H3R17me2a enrichment at the Ccl2 promoter in LPS-stimulated Prmt6^+/+^ and Prmt6^-/-^ macrophages, demonstrating differential promoter occupancy. (n = 3)
Further investigations through Western blot and immunofluorescence analyses revealed that only the active PRMT6, not the PRMT6-dead, significantly increased the levels of H3R17me2a modifications within macrophages, a histone mark associated with transcriptional activation (Fig. 8d–g). This selective enhancement of histone modification by enzymatically active PRMT6 points to its direct role in modulating chromatin dynamics to facilitate transcription. Moreover, ChIP-PCR analysis confirmed a higher enrichment of H3R17me2a at the Ccl2 promoter in wild-type macrophages compared to Prmt6 knockout cells (Fig. 8h). This finding highlights the specific dependency of Ccl2 transcriptional enhancement on the catalytic activity of PRMT6, reinforcing its critical role in the transcriptional regulation of inflammatory mediators under inflammatory conditions.
Discussion
Macrophage-driven inflammation is increasingly recognized as a critical determinant of heterotopic ossification (HO) initiation and progression.^6,15^ However, how macrophages orchestrate the early inflammatory microenvironment and amplify local immune responses following trauma has remained incompletely understood. In this study, we provide comprehensive in vivo and mechanistic evidence identifying PRMT6 as a central regulator of macrophage-mediated inflammatory amplification during the early phase of HO, acting through transcriptional and epigenetic control of the chemokine CCL2 (Fig. 9).Fig. 9. Schematic of PRMT6-driven heterotopic ossification via CCL2-mediated macrophage recruitment. Trauma-induced inflammation upregulates PRMT6 in CD68^+^ macrophages, co-activating NF-kB p65 and catalyzing H3R17me2a modifications at the Ccl2 promoter, together driving CCL2 expression to recruit macrophages, which coordinate with CD31^+^/Nestin^+^ cells for ectopic bone formation
Recent studies have increasingly acknowledged the crucial role of macrophages in the development and progression of heterotopic ossification (HO).^35^ Our findings demonstrate that macrophages robustly respond to trauma, persisting at ossification centers to facilitate the transition from soft tissue injury to bone pathology, indicating their involvement across multiple stages of HO, particularly during the initial inflammatory response. This notion is further supported by our observation of reduced ectopic bone formation in macrophage-depleted mice, which is consistent with findings in Csf1-deficient models.^15^ As key mediators of immune response, macrophages secrete various cytokines and growth factors—including VEGF, TGF-β1, BMP, Act A, OSM, SP, and NT-3—that promote angiogenesis and osteogenesis, thus driving the processes of vascularization and ossification in HO.^35^ Additionally, Nestin^+^ cells, a subset of mesenchymal progenitor cells in adult bone marrow,^36,37^ and CD31^+^ endothelial cells contribute to this complex microenviroment.^15^ Our data indicate an increased presence of CD68^+^ macrophages, Nestin^+^ cells, and CD31^+^ endothelial cells at the injury site, with macrophage depletion significantly disrupting this cellular assembly, underscoring the role of macrophages in orchestrating a microenvironment conducive to bone formation.
The observed elevation of PRMT6 in macrophages post-trauma/burn injury highlights its crucial regulatory function in the inflammatory cascade that is essential for HO. This upregulation, which aligns with reports that PRMT6 interacts with proinflammatory transcription factors,^38^ emphasizes its significance in orchestrating the inflammatory response. Elevated levels of PRMT6 in LPS-stimulated macrophages further substantiate its essential role in inflammation. The use of Prmt6 knockout mice and Prmt6-targeted Prmt6 knockdown mice provided definitive in vivo insights, demonstrating a marked reduction in macrophage recruitment at the injury site and bone formation. These results indicate the indispensable role of PRMT6 in macrophage-mediated inflammation and HO progression.
Our findings further elucidate the critical timing of PRMT6’s role in orchestrating the inflammatory cascade leading to HO. Systematic inhibition of PRMT6 following TBI reveals a pivotal window in which the inflammatory response can be effectively modulated to prevent pathological bone formation. This temporal sensitivity underscores the importance of PRMT6 in macrophage recruitment and activation. The significant reduction in ectopic bone formation in early treatment group, compared to negligible effects in later interventions, suggests that PRMT6’s influence on HO predominately hinges on the macrophage-mediated inflammatory response. Once the inflammatory and cellular recruitment processes are established, they become less dependent on PRMT6 activity. Identifying this therapeutic window for PRMT6 inhibition could guide clinical strategies for managing trauma-induced ossification, potentially shifting the focus from symptom management to prevention during the acute phase post-injury.
Chemokine-mediated recruitment of macrophages to injury sites is a critical driver of the inflammatory response across various diseases, including HO pathogenesis.^39–41^ Our study demonstrates that PRMT6 deficiency significantly reduces the expression of key chemokines, including CCL2, CCL3, and CCL6, as well as their receptors (e.g., CCR5, CCR3), highlighting PRMT6’s crucial role in modulating chemotactic gradients to orchestrate immune cell recruitment in TBI model. CCL2, also known as MCP-1, is a principal chemokine for macrophage recruitment^42–45^ and has been implicated in both neurogenic and trauma-induced HO.^6,46^ In the TBI model, where early-infiltrating macrophages amplify local inflammation, we show that PRMT6 directly regulates CCL2 secretion, enhancing macrophage mobility and inflammatory center formation. Macrophage-specific Ccl2 knockdown phenocopied Prmt6 deficiency, resulting in reduced macrophage recruitment, impaired inflammatory niche formation, and attenuated HO. Rescue experiments validate CCL2’s functional significance: exogenous CCL2 restores CD68^+^ macrophage infiltration and ectopic bone formation in Prmt6^-/-^ mice, confirming CCL2 as a key mediator of PRMT6-driven inflammation and HO progression. Therapeutically, targeting PRMT6 could modulate the CCL2/CCR2 axis, a strategy validated in inflammatory diseases and HIV treatment,^47,48^ offering a precise approach to mitigate HO progression compared to broad chemokine inhibition.
Further elucidation of the molecular mechanisms revealed that PRMT6 modulates CCL2 expression in macrophages predominantly through the NF-κB pathway. Remarkably, PRMT6 controls the activation of NF-κB independent of its methyltransferase activity, as indicated by similar NF-κB activation levels in macrophages overexpressing both the catalytically active and inactive forms of PRMT6. Furthermore, PRMT6 directly interacts with activated NF-κB, facilitating its translocation to the nucleus. This interaction is reasonable, considering PRMT6’s predominant nuclear localization,^49^ which suggests that newly synthesized cytoplasmic PRMT6 could guide p-NF-κB into the nucleus upon activation. PRMT6’s influence on NF-κB pathway regulation is critical for CCL2 transcription, as evidenced by the significant enrichment of PRMT6 and p-NF-κB at the CCL2 promoter in response to inflammatory stimuli. Additionally, PRMT6 extends its influence beyond transcriptional regulation to encompass epigenetic modifications. Intriguingly, despite no significant differences in NF-κB activation between the overexpression of catalytically mutated and wild-type PRMT6, the expression of CCL2 was considerably higher in cells expressing the latter, pointing to a potential active epigenetic modification by PRMT6 at this locus. Histone arginine residues, which are critical catalytic sites for PRMT6, feature both repressive (H3R2me2a, H2AR26me2a) and activating (H3R17me2a, H3R42me2a) modifications.^18^ H3R17me2a, known as a transcriptionally activating histone mark, has been identified as a shared catalytic site for CARM1 and PRMT6.^19^ Our data demonstrate that PRMT6 catalyzes activating histone marks such as H3R17me2a, enhancing CCL2 transcription. This dual role of PRMT6, involving both NF-κB signaling and epigenetic modifications, presents new therapeutic avenues for conditions marked by excessive inflammation or aberrant macrophage activity.
In conclusion, this study elucidates the fundamental role of PRMT6 in modulating the early inflammatory responses post-trauma, primarily through CCL2-mediated macrophage recruitment, thereby promoting the development of heterotopic ossification. By delineating the molecular interactions through which PRMT6 regulates CCL2 expression—including its engagement with the NF-κB signaling pathway and its influence on epigenetic modifications—our research advances the understanding of the inflammatory cascade in HO. These findings underscore the feasibility of intervening in the early inflammatory processes through targeted inhibition of PRMT6, aiming to curtail the cascade of events that lead to undesired bone formation.
Materials and methods
Animals
All animal experiments were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals, and received approval from the Ethics Committee on Animal Research at the Naval Medical University.^23^ All mice were bred and housed in specific-pathogen-free conditions.
Generation of Prmt6-deficient mice
Prmt6-deficient mice on a C57BL/6 J background were generated using the CRISPR-Cas9 gene-editing system.^23^ Two guide RNAs (5′-ACG AAT CCC AGC AGG CCC CG-3′ and 5′-GAG ATC GCC TAT GCA AGT TG-3′) were designed to target the single exon of Prmt6 on chromosome 3. Cas9 mRNA and the aforementioned gRNAs were microinjected into fertilized eggs from C57BL/6 J mice to produce the F0 generation. These F0 mice were sequence-validated and then crossed with C57BL/6 J mice to produce F1 generation heterozygous mice. The heterozygotes were further bred to produce homozygous offspring. Tail genomic DNA PCR was conducted to determine the genotypes of the offspring using the following primers: 5′-TTTCGCCGTCTGGTTTCA-3′ and 5′-GGTCAGGGATGCTCACTTTT-3′ for the wild type allele, and 5′-AGGCTACCCATACGTTCT-3′ and 5′-CCTTTCTCCCAGTTTCAT-3′ for the Prmt6 knockout allele. Representative genotyping results are shown in Fig. S11.
Trauma/Burn injury HO model
The mouse model of heterotopic ossification (HO) was established using a combination of Achilles tenotomy and a dorsal burn injury, as previously described.^50^ Briefly, male C57BL/6 J mice, aged 10 weeks, were anesthetized with 2.5% isoflurane. A longitudinal incision (~0.5 cm) was made along the Achilles tendon, followed by a mid-tendon tenotomy without subsequent suturing to induce HO. Concurrently, a partial-thickness burn injury was inflicted by applying a 60 °C heated aluminum block to the shaved dorsal skin, covering 30% of the total body surface area, for 17 s. In sham controls, the Achilles tendon was exposed without tenotomy, and the dorsal skin was contacted with an unheated aluminum block for the same duration.
Macrophage-targeted knockdown of Prmt6 or Ccl2 in vivo
Gene knockdown in peripheral macrophages was achieved using AAVs, which have been shown to effectively target these cells.^32,33^ AAVs were administered via tail vein injection into 10-week-old male C57BL/6 J mice using a sterile syringe. Two weeks post-injection, the trauma/burn injury HO model was established. The AAV constructs used in this study include: AAV9-LysM-EGFP-miR30shRNA(Prmt6)-WPRE (1 × 10^13^ genomic copies/ml in sterile saline), AAV9-LysM-EGFP-miR30shRNA(Ccl2)-WPRE (1 × 10^13^ genomic copies/ml in sterile saline), and AAV9-LysM-EGFP-miR30shRNA(NC)-WPRE (1 × 10^13^ genomic copies/ml in sterile saline), all designed and generated by PackGene Biotech (Guangzhou, China).
Micro-computed tomography (μCT) analysis
Mouse hind limbs were harvested, fixed overnight in 10% formalin, and subsequently analyzed using a high-resolution μCT system (SkyScan 1076, Bruker, Saarbrücken, Germany). The scanner settings were adjusted to a voltage of 60 kV and a resolution of 9 μm per pixel. Images were reconstructed, and the volume of ectopic bone was quantified within a defined region of interest, spanning from the proximal knee joint to the distal hind paw.
Histology, immunohistochemistry and immunofluorescence
Mouse hind limbs were decalcified in 10% EDTA, sequentially dehydrated in ethanol concentrations ranging from 70% to 100%, embedded in paraffin, and sectioned into 5-μm slices. These sections were stained using H&E and SOFG. For immunohistochemistry (IHC), antigen retrieval was performed by immersing the sections in sodium citrate buffer at 99 °C for 20 min. The sections were blocked for 1 h at room temperature using 5% normal goat serum and 0.2% Triton X-100 (both from Sigma-Aldrich, St. Louis, USA), followed by overnight incubation at 4 °C with primary antibodies against CCL2 (Abcam, Cambridge, UK). Subsequently, the sections were incubated with HRP-conjugated secondary antibodies. Color development was achieved using a DAB kit (Beyotime Biotech, Shanghai, China). For fluorescence immunohistochemistry, sections were incubated with primary antibodies against PRMT6 (Santa Cruz, California, USA), CD68 (CST, Boston, USA), Nestin (CST, Boston, USA), CD31 (Abcam, Cambridge, UK), F4/80 (CST, Boston, USA), Ly6G (Abcam, Cambridge, UK), Ly6C (Abcam, Cambridge, UK), PDGFRα (Abcam, Cambridge, UK), Fibronectin (Millipore, Massachusetts, USA), and Collagen I (Abcam, Cambridge, UK) followed by incubation with fluorescently labeled secondary antibodies (Invitrogen, California, USA) for 2 h at room temperature. After three washes with 0.01 mol/L PBS, the sections were examined under a fluorescence microscope.
H&E-stained sagittal slides were assessed using a modified Movin’s score,^31^ a semiquantitative grading scale evaluating various aspects of tendon tissue, including fiber structure, fiber arrangement, nuclear morphology, cellularity, vascularity, collagen stainability, and hyalinization. Each variable was scored from 0 to 3, where 0 indicated normal tissue, 1 indicated slight abnormality, 2 indicated moderate abnormality, and 3 indicated marked abnormality. The total histologic score ranged from 0 (normal tendon) to 21 (severe abnormality). Immunofluorescence intensity for DAPI, PRMT6, CD31, CCL2, Fibronectin, and Collagen I was quantified using ImageJ software.
Macrophages depletion
To selectively deplete monocyte-derived macrophages, mice were administered PLX73086 (Plexxikon, California, USA), incorporated into AIN-76A chow at a dosage of 200 mg/kg daily.^51^ This regimen was initiated three weeks prior to and continued for nine weeks following trauma induction. Mice in the control group were maintained on a standard AIN-76A diet concurrently.
Cell culture
Mouse bone marrow cells (BMCs) were isolated from the tibiae and femurs of 4-week-old Prmt6^+/+^ and Prmt6^-/-^ littermates by flushing the bone marrow. To generate BMDMs, BMCs were cultured in α-MEM (Hyclone, Logan, USA) supplemented with 10% fetal bovine serum (FBS), 1% penicillin-streptomycin (both from Gibco, Grand Island, USA), and 30 ng/mL macrophage colony-stimulating factor (M-CSF; Peprotech, New Jersey, USA). The cultures were maintained at 37 °C in a 5% CO₂ atmosphere, and the medium was refreshed every two days. To induce an inflammatory response in vitro, BMDMs were treated with lipopolysaccharide (LPS; 500 ng/mL) or high mobility group box 1 protein (HMGB1; 1 or 2 μg/mL) for indicated durations.
Plasmid construction and transfection
cDNA fragments encoding mouse wild-type Prmt6 and a catalytically inactive Prmt6 with point mutations at the 89^th^ valine and the 91^st^ aspartic acid residues [Prmt6(dead)] were synthesized and confirmed by sequencing.^52^ Plasmids containing Prmt6, Prmt6(dead), or a scramble control were transfected into mouse macrophages using the jetPEI-Macrophage DNA Transfection Reagent, following the manufacturer’s instructions.
Quantitative RT-PCR
Total RNA from cultured cells was extracted using TRIzol (Invitrogen, California, USA) following the manufacturer’s instructions, and quantified using a NanoDrop2000 (Thermo Fisher Scientific, California, USA). Complementary DNA (cDNA) was synthesized using the PrimeScript RT Reagent Kit (Takara, Osaka, Japan). Quantitative RT-PCR (qRT-PCR) was performed on a QuantStudio 3 RT-PCR System (Thermo Fisher Scientific, California, USA) using SYBR Premix Ex Taq II (Takara, Osaka, Japan) with specific primers. Relative gene expression levels were calculated using the 2-ΔΔCt method.
Western blot analysis
Cell samples were first washed with ice-cold phosphate-buffered saline (PBS) and then lysed in RIPA buffer supplemented with protease and phosphatase inhibitors (Epizyme, Shanghai, China). The lysates underwent ultrasonic disruption followed by centrifugation at 14 000 r/min at 4 °C for 15 min. A small aliquot of the lysate was used to measure the total protein concentration using a BCA Protein Assay Kit (Epizyme, Shanghai, China). The majority of the lysate was then mixed with 5× sodium dodecyl sulfate (SDS) loading buffer and heated at 95 °C for 10 min. Equal amounts of total protein from each sample were separated on 10% SDS-polyacrylamide gels and subsequently transferred onto PVDF membranes using a wet transfer system. Membranes were blocked using protein-free rapid blocking buffer (Epizyme, Shanghai, China) for 30 min at room temperature and then incubated overnight at 4 °C with primary antibodies. After a two-hour incubation with HRP-conjugated goat anti-rabbit or anti-mouse IgG secondary antibodies at room temperature, the immunoreactive bands were detected using enhanced chemiluminescence (Epizyme, Shanghai, China) following the manufacturer’s protocol.
Cell immunofluorescent staining
For immunofluorescent analyses, BMDMs were cultured on chamber slides. Following stimulation with 500 ng/mL LPS (MedChemExpress, New Jersey, USA) for indicated hours, cells were fixed with cold methanol, permeabilized with PBS containing 0.1% Triton X-100, and blocked with 5% normal goat serum (Sigma-Aldrich, Saint Louis, USA). Cells were then incubated with primary antibodies specific for NF-κB (CST, Boston, USA), CCL2 (Abcam, Cambridge, UK), and H3R17me2a (Abcam, Cambridge, UK). Following the primary antibody incubation, cells were washed three times with PBS containing 0.1% Tween-20 (PBS-T), then incubated with fluorophore-conjugated secondary antibodies and counterstained with DAPI. Stained cells were visualized under a laser-scanning confocal microscope.
Enzyme-linked Immunosorbent Assay (ELISA)
The concentrations of CCL2 in the supernatants from Prmt6^+/+^ and Prmt6^-/-^BMDMs were quantified before and after stimulation with 500 ng/mL LPS (MedChemExpress, New Jersey, USA). Measurements were conducted using a CCL2 ELISA kit (SAB Biotherapeutics, Boston, USA), strictly following the manufacturer’s protocol.
Cell migration assay
To assess the chemotactic capacity of secretions from Prmt6^-/-^ BMDMs under inflammatory stimulation, both scratch and Transwell assays were utilized.
Scratch assay
BMDMs from Prmt6^+/+^ and Prmt6^-/-^ mice were pre-stimulated with 500 ng/mL LPS or PBS for 12 h. After replacing the media with LPS-free media and an additional 24 hours of culture, the supernatants were collected and stored at 4 °C. For the rescue experiments, CCL2 (HY-P7764, MedChemExpress, New Jersey, USA) was added to the supernatant from Prmt6^-/-^ BMMs. Concurrently, confluent wild-type BMMs underwent a scratch using a 100 μL pipette tip. The cells were rinsed with PBS to remove detached cells, and the media were replaced with the aforementioned supernatants. After 24 h, bright-field images were captured using an Olympus X53 microscope both before and after incubation. The cell migration rate was quantified using the equation: Cell migration rate = (Area_0h_−Area_24h_)/Area_0h_ × 100% where Area_0h_ is the initial scratch cell-free area, and Area_24h_ is the area after 24 h of incubation.
Transwell assay
BMDMs from Prmt6^+/+^ and Prmt6^-/-^ mice were seeded in the lower chambers of Transwell plates (Corning, Lowell, MA, USA) and stimulated with LPS for 12 h. Post-stimulation, the LPS-containing media were replaced with LPS-free media and cultured for an additional 24 h. For rescue experiments, CCL2 was added to the media in the lower chamber containing Prmt6^-/-^ BMDMs. Simultaneously, wild-type BMDMs were placed in the upper chamber and cocultured for 24 h. Cells that migrated to the lower chamber were stained with crystal violet, and the positively stained cells were counted.
Co-immunoprecipitation
Cell samples were harvested and lysed using an immunoprecipitation kit (Thermo Fisher Scientific, California, USA) for 5 min at 4 °C. The lysates were then subjected to ultrasonication followed by centrifugation to separate the supernatant (input control) from the pellet. For the immunoprecipitation, 500 µL of the supernatant was incubated with 5 µL each of Protein A and Protein G magnetic beads for 1 h at 4 °C to pre-clear the lysate. The pre-cleared supernatant was then incubated with specific primary antibodies and control IgG overnight at 4 °C.The following day, an additional 5 µL each of Protein A and Protein G magnetic beads were added to the antibody-lysate mixture and incubated for 3 h at 4 °C to capture the antibody-antigen complexes. The beads were subsequently collected by centrifugation and washed three times with 1× wash buffer to remove non-specifically bound proteins. Finally, the bead-bound complexes were eluted by resuspending in 1× SDS sample buffer, denatured at 99 °C for 10 min, and prepared for analysis by Western blot.
Chromatin Immunoprecipitation (ChIP)
The ChIP assay was conducted using the SimpleChIP® Enzymatic Chromatin IP Kit (CST, Boston, USA), following the manufacturer’s instructions. Cells were treated with 1% formaldehyde for 10 min at room temperature to crosslink chromatin. The reaction was quenched by adding glycine for 5 min. Nuclei were then isolated using a cold buffer containing DTT and a cocktail of protease inhibitors for 10 minutes. Chromatin was enzymatically digested with Micrococcal Nuclease at 37 °C for 20 min and subsequently sheared by ultrasonic sonication to generate fragments ranging from 150 to 900 base pairs, as verified by agarose gel electrophoresis. For immunoprecipitation, the sheared chromatin was incubated with primary antibodies specifically targeting histone H3 (included in the kit), H3R17me2a (Thermo Fisher Scientific, California, USA), PRMT6 (CST, Boston, USA), NF-κB (CST, Boston, USA), and control IgG (also included in the kit). The immune complexes were captured using magnetic beads, washed, and the DNA was eluted. The purified DNA was then quantified by quantitative real-time PCR (qRT-PCR), with results expressed as a percentage of the input DNA to account for sample loading variations.
Bioinformatics analysis and visualization of scRNA-Seq
Previously published single-nuclei RNA-seq data from mouse tendon (GEO accession no. GSE126060) were obtained from the NCBI Gene Expression Omnibus (GEO).^6,27^ Analysis of scRNA-seq data, as shown in Fig. 1, Figs. S3 and S9, was performed on samples from HO as described in the original publication.
For quality control, cells with fewer than 500 detected genes, more than 60 000 unique molecular identifiers (UMIs), or a mitochondrial UMI fraction greater than 0.2 were excluded. Genes expressed in fewer than three cells per set were discarded. Datasets were merged and re-clustered, with minimal batch effect after correction using the Harmony package (v1.2.4). Downstream analysis was conducted using R (v4.4.3) and the Seurat package (v5.3.1, SatijaLab/Seurat). Cell annotation was performed according to the original publication.^27^ To investigate the cellular expression patterns of Prmt6 and Ccl2 after traumatic brain injury (TBI) at 3 days post-injury (dpi), the counts of Prmt6^+^ and Ccl2^+^ cells and the mean expression of Prmt6 and Ccl2 were compared across cell clusters.
RNA sequencing (RNA-seq)
To investigate potential targets driving heterotopic ossification (HO) during the inflammatory phase, soft tissue samples were collected from the injury site 3 days post-trauma/burn injury and from sham controls for RNA-seq. To evaluate the impact of Prmt6 deficiency on macrophage behavior under inflammatory conditions, BMDMs from both Prmt6^+/+^ and Prmt6^-/-^ mice were stimulated in vitro with LPS for 6 h prior to RNA extraction.
Total RNA was extracted using the TRIzol reagent (Invitrogen, California, USA) and mRNA was enriched with Oligo(dT) beads. The mRNA was then fragmented into short sequences, which were reverse transcribed into cDNA using random primers. The resulting cDNA fragments were purified using the QIAquick PCR Extraction Kit, subjected to end repair, A-tailing, and ligation with Illumina sequencing adapters. The ligated products were size-selected via agarose gel electrophoresis, amplified by PCR, and sequenced on an Illumina HiSeq 2500 system.Quality control of raw sequence data involved filtering out adapter sequences, reads containing more than 10% unknown nucleotides, and reads where over 50% of bases had a quality score (Q-value) of 20 or less. Ribosomal RNA (rRNA) was subtracted using Bowtie2. Subsequently, the remaining reads were aligned to the reference genome using TopHat2 (version 2.0.13). Gene expression levels were quantified with RSEM and normalized using the Transcripts Per Million (TPM) method.
For the identification of differentially expressed genes (DEGs), the DESeq2 package in R was employed, applying significance thresholds appropriate for each sample type. For tissue samples, criteria included Q < 0.01 and a fold change (FC) ≥ 2, while for cell samples, thresholds were set at P < 0.05 and a FC ≥ 1.5. GO annotation and KEGG pathway enrichment analyses were performed using the clusterProfiler package in R. The RNA sequencing data have been deposited in the Sequence Read Archive (SRA) with the accession number PRJNA (1166647, 1166660).
Statistical analysis
All data were analyzed using SPSS 24.0 (IBM, Armonk, USA). Statistic difference was analyzed by unpaired two-tailed Student’s t test or by a one-way analysis of variance (ANOVA) followed by the lest significant difference test. A P < 0.05 was considered statistically significant.
Supplementary information
Fig. S1 Dynamic progression of heterotopic ossification (HO) in a TBI-induced mouse model Fig. S2 Macrophage depletion suppresses heterotopic ossification Fig. S3 Cell-type marker expression in single-cell RNA sequencing analysis Fig. S4 Cell-type specificity of PRMT6 expression during TBI-induced HO Fig. S5 PRMT6 upregulation in macrophages following LPS and HMGB1 stimulation in vitro Fig. S6 Prmt6^-/-^ mice shows promoted tendon healing after TBI (Modified Movin’s score) Fig. S7 Improved tendon repair after macrophages-specific Prmt6 knockdown (Modified Movin’s score) Fig. S8 Early PRMT6 inhibition improves Achilles tendon repair Fig. S9 Cell-type specificity of CCL2 expression during TBI-induced HO Fig.S10 Macrophages-specific Ccl2 knockdown mitigates heterotopic ossification Fig. S11 Genotype identification via PCR amplification Supplementary figure caption (Figs. S1-S11)
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