Epigenetic erosion of H4K20me1 induced by inflammation drives aged stem cell ferroptosis

Abstract

Aging is characterized by a decline in the functionality and number of stem cells across the organism. In this study, we uncovered a mechanism by which systemic inflammation drives muscle stem cell (MuSC) aging through epigenetic erosion. We demonstrate that age-related inflammation decreases monomethylation of H4K20 in MuSCs, disrupting their quiescence and inducing ferroptosis, a form of iron-dependent cell death. Our findings show that inflammatory signals downregulate Kmt5a, the enzyme responsible for depositing H4K20me1, leading to the epigenetic silencing of anti-ferroptosis genes. This results in aberrant iron metabolism, increased reactive oxygen species levels and lipid peroxidation in aged MuSCs. Notably, long-term inhibition of systemic inflammation that is initiated at 12 months of age effectively prevents ferroptosis, preserves MuSC numbers and enhances muscle regeneration and functional recovery. These findings reveal an epigenetic switch that links chronic inflammation to MuSC aging and ferroptosis, offering potential therapeutic strategies for combating age-related muscle degeneration.

Fig. 1. Age-associated systemic inflammation correlates with chemokine signalling activity in skeletal muscles and chromatin remodeling in MuSCs.

a–f, Multiomics workflow and computational modeling of systemic inflammatory ligand signaling in skeletal muscle. Blood and plasma (n = 4 adult mice, n = 6 aged mice) were used for cell composition profiling (b) and cytokine analysis via proteomics and validation immunoassays (c; n = 3 mice per condition). Muscle transcriptomics were used to infer active signaling pathways (d), which were integrated with cytokine data through ligand–receptor prediction scoring (e). IL-17 and chemokine signaling were highly activated, with CCR2 as a key upstream node. f, Heatmap of differentially expressed chromatin organization genes (GO:0006325) in aged MuSCs. g,h, Representative pictures (g) and quantification (h) of H4K20me1 status in MuSCs on EDL single fibers isolated from adult and aged mice (n = 6 mice per age, with at least 30 Pax7⁺ cells counted per mouse). i,j, Immunoblot (i) and qPCR (j) assessing Kmt5a mRNA and protein levels from freshly sorted MuSCs (n = 3 mice per condition). Low Kmt5a correlated with decreased H4K20me1. For western blots, two aged mice were pulled together for one lane. Quantification is normalized to total H3 and relative to young values. k–m, Flow cytometry analysis for MACS pre-sorted cells stained for VCAM and Zombie dye prior to fixation and then fixed and permeabilized for H4K20me1 staining. Graph displays percentage of cells positive for VCAM and H4K20me1 for n = 3 mice per age. Two-dimensional plot of H4K20me1 in MuSCs for each replicate (l) and mean fluorescence intensity analysis (m) (n = 3 mice per age). For all violin plots, data represent biological replicates. Statistical significance was determined using two-sided Welch’s t-test, and exact adjusted P values are reported in the figure. FC, fold change; FDR, false discovery rate; WBC, white blood cell; MFI, mean fluorescence intensity; TLR, Toll-like receptor; SK.M., skeletal muscle; AGE-RAGE, advanced glycation end products–receptor for advanced glycation end products. Source data

Fig. 2. Kmt5a is required for quiescent MuSC homeostasis.

a,b, Schematic representation of the regeneration experiments and MuSC self-renewal. Mice were subjected to injury either immediately after tamoxifen injections or 3 weeks later. Injured muscles were harvested at 21 dpi, and MuSCs were counted by staining for Pax7. The relative number of MuSCs per area of regeneration is shown in b (n = 6 WT and n = 3 KO mice per timepoint; blue, male, and red, female). c,d, Schematic representation of long-term Kmt5a deletion and quantification of MuSC pool maintenance at homeostasis. Muscles were harvested at different times after knockout induction, and the number of Pax7⁺ cells was counted. The stochastic decline of MuSCs in the mutant mice is shown in d (n = 23 WT mice and n = 4–6 KO mice per timepoint; blue, male, and red, female). e, Percentage of activated MuSCs quantified by pS6 immunostaining 3 weeks and 6 weeks after knockout induction (n = 15 WT mice and n = 8–15 KO mice per timepoint). f–h, Relative number of MuSCs actively cycling (g; EdU⁺) or out of quiescence (h; Ki67⁺) in uninjured muscles (n = 6 WT mice and n = 6 KO mice). i, Quantification of precocious terminally committed MuSC progenitors (Pax7⁺MyoG⁺) 6 weeks after deletion of Kmt5a in MuSCs (n = 4 WT mice and n = 4 KO mice). j,k, Molecular profiling of DNA damage and DNA repair signatures in MuSCs 6 weeks after knockout induction. TUNEL was used to detect DNA strand breaks. The results are reported as the mean ± s.d. of Pax7⁺TUNEL⁺/Pax7⁺ (percentage) (j), and immunoblotting was used to measure the p53 signaling response (p-P53) to DNA damage (γH2AX) (n = 5 WT mice and n = 8 KO mice). For all violin plots, data represent individual biological replicates. Statistical significance was determined using two-sided Welch’s t-test (g–j) or one-way ANOVA (b,d,e), and exact P values and adjusted P values (q) are reported in the figure. qMuSC, quiescent MuSC; regen, regeneration; Tmx, tamoxifen; wks, weeks; WT, wild-type. Source data

Fig. 3. Kmt5a deletion in MuSCs causes ferroptosis.

a,b, GPX4 immunostaining in TA muscles of WT (a) and Kmt5a^KO (b) mice (n = 5 mice per group, one experiment). Arrows, MuSCs; yellow arrow, MuSCs in iron-rich pocket (~50%). c, Quantification of MuSCs with high levels of labile iron (Fe²⁺) (mean ± s.e.m., n = 5 mice per condition). d, Representative electron micrographs of a WT quiescent MuSC and a Kmt5a^KO MuSC showing features of activation and ferroptosis. e, ICP-MS quantification of elemental iron in MuSCs. Total iron was normalized to cell numbers. Data points are reported as average of replicate (n = 5 mice per condition). f,g, Quantification of lipid peroxidation in MuSCs. Flow cytometry plot shows a shift in 510-nm signals in mutant MuSCs. Inverted ratiometric signals of 590 nm/510 nm were calculated to report lipid peroxidation in each cell (g); data points are reported as average of replicate (n = 4 mice per condition). h, qPCR for Gpx4, Rgs4 and Ptgs2. i,j, Immunoblot and quantification for KMT5a, H4K20me1 and GPX4 with DMSO or Kmt5a catalytic inhibitor (Kmt5a_i). Kmt5a was normalized to GAPDH; H4K20me1 was normalized to histone H3; and GPX4 was normalized to actin (n = 3 mice per condition). Data are reported as normalized intensity ± s.d. k, qPCR quantification of ferroptosis markers Gpx4, Rgs4, Ptgs2 and Hmox1 in response to Kmt5a_i. For qPCR data, gene expression was normalized to the average levels of B2M, TBP and PPIA and is reported as normalized fold change ± s.d. (n = 3 mice per condition). l,m, Quantification of lipid peroxidation in response to drug treatments. Histogram shows the intensity of ratiometric signal (590 nm/510 nm) for the lipid peroxidation probe in live cells (l). Violin plots represent the inverted ratiometric signal normalized to vehicle (m). For all violin plots, data represent biological replicates. Statistical analyses were performed using two-sided Welch’s t-test (e,j,k) and one-way ANOVA (g,h,l,m), and exact P values and adjusted P values (q) are reported in the figure. wks, weeks; WT, wild-type; im, immediate. Source data

Fig. 4. Loss of H4K20me1 and Kmt5a in aged MuSCs is associated with pro-ferroptosis transcriptional signature.

a,b, H4K20me1 CUT&Tag in adult and aged MuSCs. a, Heatmap of RPKM-normalized sequencing reads centered on TSSs near H4K20me1 sites unique to young and aged MuSCs or shared between them. The numbers of TSSs in each group are labeled. b, Metaplot comparison of RPKM averaged in 50-bp bins around all TSSs. c, Transcriptional and H4K20me1 changes for ferroptosis genes. Heatmaps are organized by descending gene expression in aged MuSCs and are reported as fold change of z-score for RNA-seq and H4K20me1 CUT&Tag. RNA-seq and CUT&Tag were performed using different sets of mice and different times (n = 3 mice per condition, per experiment). d, H4K20me1 signal at Gpx4 gene. FC, fold change; RPKM, reads per kilobase per million mapped reads. Source data

Fig. 5. Identification and characterization of a ferroptotic subpopulation in aged MuSCs.

a–c, scRNA workflow: adult (blue) and aged (orange) MuSCs were identified and plotted with UMAP (a). Unbiased clustering separated adult and aged cells into four clusters (b) prior to running pseudotime analysis to assess cell trajectory (c). d, Enrichment analysis of 500 most altered genes as a function of velocity and pseudotime trajectory highlighting ‘ferroptosis’ and ‘glutathione metabolism’ (red underlined). Statistical significance: Fisherʼs exact test followed by Benjamini–Hochberg correction. e, Dot plot for Louvain metadata clusters of MuSCs showing average expression and percent expression per cell for key genes. f, Donut plot representing a selection of MuSC fate based on flow cytometry analysis (n = 5 mice per condition). Live cells were negative for all markers; senescent cells were SPiDER⁺; apoptotic cells were Annexin V⁺; ferroptotic cells were Lipid Peroxidation^High; other cells were DAPI⁺ but negative for other markers. g, Lipid Peroxidation^high MuSCs (Aged^Fe) were sorted for mRNA analysis to confirm a ferroptotic signature similar to cluster 3. h, Representative picture of an aged MuSC with intracellular labile iron and niche GPX4 (n = 2 separate experiments). i,j, Abundance of total elemental iron per cell (i) and lipid peroxidation (j) in adult and aged MuSCs. To avoid additional introduction of cell stress bias and to validate lipid peroxidation changes in aged cells, we used a glutathione depletion mouse model (Nrf2^KO;GCLC^KO) to naturally increase lipid peroxidation, therefore bypassing the need for a compound (that is, RSL3) or iron overload. k, Lipid peroxidation in MuSCs. Cells were isolated from adult and aged mice (n = 5 male mice per condition), plated for 4 hours and harvested 20 hours after treatment. l–o, MuSC survival (m), myogenic potential (o) and fusion competence (n) in response to radical trapping drug Fer1. For all violin plots, data represent biological replicates. Statistical analyses were performed using two-sided Welch’s t-test (g,i,j), one-way ANOVA (m–o) and two-way ANOVA (k), and exact P values and adjusted P values are reported in the figure. Avg., average; exp., expression; NS, not significant; Veh, vehicle. Source data

Fig. 6. Long-term inhibition of inflammation rejuvenates skeletal muscle and improves functional recovery.

a–d, Quantification of MuSC numbers at homeostasis (a,b) and at 21 days after regeneration (c,d) in aged mice. Vehicle (Veh) and bindarit (Bin; 30 mg kg⁻¹ per week, intraperitoneal) treated mice were injected weekly between 12 months and 24–30 months of age. e, Grip strength measurement after regeneration normalized to adult mice and histological evaluation of muscle regeneration. f,g, Quantification of the regenerated myofiber size (f) and size distribution (g) in the TA muscle. h,i, Quantification of lipid peroxidation (h) and single-cell average iron content (i). Two-sided Welch’s t-test. Each dot represents a mouse replicate. P values are reported in the figure. Source data

Extended Data Fig. 1. Acute systemic inflammation prime muscle stem cells and repress Kmt5a. Related to Fig. 1.

a, b) Muscle stem cell counts from adult tibialis anterior 24 hours post IP injection. CCR2-null mice were used as a negative control. c, d) Relative number of muscle stem cells (Pax7 + ) actively cycling (Edu + ) (C) or out of quiescence (Ki67 + ) (D). e-i) Plasma levels of cytokines assessed in the same cohort of mice before and after treatment. j) Percentage of circulatory monocytes in whole blood prior and after treatment measured by whole-blood count. k-m) Cycling status of MuSCs six weeks after treatment. n-q) Features of accelerated muscle regeneration at 4-day post-injury six weeks after treatment. Treated mice had larger fibers (N and O) and a higher number of MuSC progenitors (MyoD + ; P). Regenerating muscles from treated mice also displayed genetic signature of accelerated regeneration (Q). r, s) Epigenetic genes (R, writers; S, erasers) expression in MuSCs one day after treatment. Kmt5a is the only gene being significantly downregulated. t, u) Heatmap for Myogenic (T) and Cell cycle (U) genes expression Log2 fold change six weeks post-treatment. Data are reported as mean ± s.d‥ v, w) MuSC methylation status for Kmt5a-mediated mark H4K20me1 (V), and MyoD (W), six weeks after treatment. x) Flow cytometry mean fluorescence intensity for MuSCs six- and height-weeks post-treatment. y) Flow cytometry analysis showing detailed frequency of MuSCs with H4K20me1 low or high in relation to Ki67 status six weeks post-treatment. Welch’s t-test (L, M, O-S), one-way ANOVA E-J), or two-way ANOVA (B-D, X, Y). exact p values are reported in the figure. Source data

Extended Data Fig. 2. Loss of Kmt5a in muscle stem cells leads to genomic instability features. Related to Fig. 2.

a-d) Strategy for Kmt5a genetic deletion in muscle stem cells to assess immediate contribution to homeostasis and regeneration. Muscle stem cell counts from adult tibialis anterior muscles immediately after final tamoxifen injection (B, C). White arrows indicate Pax7+Kmt5a+ double-positive cells, while the orange arrows point to Pax7+Kmt5a- cells. e-k) Cell culture kinetics of wildtype and knockout muscle stem cells with quantification of H4K20me1 (F), cell survival (G), cell cycle features (H, I) and myogenic status (J, K). Cell density starts to decrease in the knockout as muscle stem cells loose H4K20me1, which correlates with terminal differentiation deficit. l, m) Relative number of muscle stem cells with aberrant morphologies and nuclear phenotypes (red arrows). n, Immunoblotting profiling of DNA damage response signature. Red dots represent females and blue dots represent males; we did not observe obvert sexual dimorphism. Welch’s t-test or one-way ANOVA. p values are reported in the figure. Source data

Extended Data Fig. 3. Kmt5a-null muscle stem cells prematurely enter an alert-like state. Related to Fig. 2.

a) Strategy to assess muscle stem cell myogenic fate and survival in response to a proximal injury (injured muscles), or to assess Galert and quiescence features in response to a distal injury (contralateral), 4-day post injury. Injuries were performed immediately upon last tamoxifen injection. b) Progression of muscle regeneration was indexed using embryonic myosin heavy chain (eMHC) at 4-, 7-, 30-, and 60dpi. Quantification of eMHC+ fiber per area of regeneration are reported on the representative pictures as mean ± s.e.m. c-e) Myogenic cell fate analysis throughout regeneration. Pax7 (C) was used for MuSCs post-regeneration, MyoD (D) quantified the number of myoblast, and MyoG (E) showed the number of terminally committed progenitors. f-k), Myogenic and cell cycle status of activated muscle stem cells in response to an acute injury at 4dpi. Myogenic progression was assessed by Pax7, MyoD and MyoG, while their cell cycle status was measured by EdU incorporation over 24 h prior to harvesting. l-q, Quiescence and activation status of muscle stem cells in uninjured muscles in response to a distal injury. Number of muscle stem cells was assessed with Pax7, while their activation status was assessed for active cycling (L, N; EdU), Galert (O, P; white arrow shows Pax7 + pS6+ (activated) cell, green arrow shows Pax7 + pS6- (quiescent) cell, orange arrow shows Pax7- pS6+ cell), and quiescence exit (Q; Ki67). Welch’s t-test or one-way ANOVA. p values are reported in the figure. Source data

Extended Data Fig. 4. Characterization of muscle stem cell sensitivity to ferroptosis and Kmt5a catalytic inhibition. Related to Fig. 3.

Muscle stem cells were freshly isolated and plated for 4 h prior being treated with the ferroptosis-inducing compounds along with Ferrostatin (Fer1), a potent and selective inhibitor of ferroptosis. a-d, Erastin (A, B) and RSL3 (C, D) dose and time effects on Muscle stem cells. Cell viability (A, C) was quantified by DAPI integration and increased ROS was quantified by MitoSOX CTCF (B, D). e-g, Optimal treatment for Erastin (10uM) and RSL3 (1uM) for 24 h caused decreased viability (E; data reported as mean of individual experiment), increased ROS (F; data reported as individual cell intensity [CTCF] from 3 different experiments) and lipid peroxidation (G; data reported as mean of individual experiment). Fer1 rescued the effects of Erastin and RSL3, as it traps lipid radicals and lipid ROS that are involved in the induction of ferroptosis in Muscle stem cells. h, i) Immunoblot and quantification for GPX4 relative to Actin following treatments. j) Optimal dose for Kmt5a catalytic inhibitor UNC0379 was determined over 24 hours period, with 4uM effectively reducing H4K20me1 beyond detectable levels. k) UNC0379 treatment (4uM) resulted in cell death initiating at 24 h with no detectable surviving cell past 72 h. Data are presented as mean ± standard deviation, n = 3 independent experiments. l) Transcript levels of ferroptosis-associated genes Gpx4, Rgs4, Ptgs2, Hmox1 in response to RSL3. Gene expression was normalized to the average levels of B2M, TBP, and PPIA, and are reported as normalized fold-change ± s.d. Source data

Extended Data Fig. 5. Kmt5a regulates GPX4 expression through H4K20me1 deposition and prevents ferroptosis in myoblasts. Related to Fig. 3.

(a, b) Quantitative PCR (A) and immunoblot (B) analysis showing the levels of Gpx4 mRNA and protein in myoblasts treated with Kmt5a siRNA, with or without Kmt5a re-expression. B shows the whole WB membrane Data are presented as mean ± s.d. (c, d) Analysis of Kmt5a (C) and H4K20me1 (D) occupancy around the Gpx4 transcription start site (TSS) in control and Kmt5a-silenced myoblasts, with or without Kmt5a re-expression. Data are presented as % input ± s.d. from three independent experiments. (e) Lipid peroxidation analysis in control and Kmt5a-silenced myoblasts. Data are presented as mean ± s.e.m. from three independent experiments. (f-i) Quantitative PCR (F), immunoblot (G), cell viability (H), and lipid peroxidation (I) analysis in myoblasts treated with a Kmt5a inhibitor, with or without Gpx4 overexpression. Statistical analyses were performed using one-way ANOVA (A, E, F, H and I), and exact p-values are reported in the figure. Source data

Extended Data Fig. 6. Chronic administration of Ccr2-ligands induces systemic inflammation and accelerates MuSC depletion, activation, and ferroptosis, resembling the aged MuSC phenotype. Related to Fig. 5.

a) Schematic representation of the experimental design. Young mice were administered a chemokine cocktail intraperitoneally over three weeks to induce systemic chronic inflammation. Mice were treated with saline (Veh.), Ccr2-ligands (chemokines), or a combination of Ccr2-ligands and neutralizing antibodies against Ccl2, Ccl7, and Ccl8 (chemokines + nAbs). (b) Quantification of the MuSC pool size in adult mice following chronic chemokine administration. Data are presented as mean ± s.e.m. with n = 5 mice per conditions. (c) EdU incorporation assay showing the accelerated cell cycle re-entry of MuSCs sorted from chemokine-treated mice. Data are presented as mean ± s.e.m. from three independent experiments. (d, e) Analysis of H4K20me1 (D) and qPCR analysis of Kmt5a and ferroptosis markers (E) in MuSCs derived from chronically treated mice. Data are presented as mean ± s.e.m. with n = 5 mice per conditions. (f) Lipid peroxidation analysis showing elevated levels in MuSCs from chemokine-treated mice, mirroring the aged MuSC phenotype. Data are presented as mean ± s.e.m. with n = 5 mice per conditions. (g) Survival analysis of MuSCs in culture exposed chronically –every 12hrs- to Ccr2-ligands. Data are presented as mean ± s.e.m. from three independent experiments. Statistical analyses were performed using one-way ANOVA, and p-values are indicated in the figure. Source data

Extended Data Fig. 7. Chronic exposure to aged plasma induces premature aging of MuSCs via Ccr2-ligands, recapitulating the Kmt5aKO phenotype. Related to Fig. 5.

a) Schematic representation of the experimental design for aged plasma transfusion. Young mice were intravenously injected with aged plasma over a specified period to mimic chronic exposure. b) Quantification of MuSC pool size following chronic aged plasma exposure. c-e) Analysis of MuSCs sorted from aged plasma-treated mice. Flow cytometry analysis showing the loss of H4K20me1 (C) and repression of Kmt5a (E). (D) Lipid peroxidation analysis indicating increased lipid peroxidation in aged plasma-treated MuSCs. (E) qPCR analysis showing the repression of Gpx4 in MuSCs exposed to aged plasma. All effects were rescued by neutralizing antibodies against Ccr2-ligands. Data are presented as mean ± s.e.m. from three independent experiments with n = 10 male mice per conditions. Statistical analyses were performed using one-way ANOVA and p-values are reported in the figure. Source data

Extended Data Fig. 8. Inflammatory profiling of aged mice in response to long-term Bindarit treatment. Related to Fig. 6.

a-f) HESKA to quantify whole blood cellular content with monocytes (A), lymphocytes (B), neutrophils (C), eosinophils (D), basophils (E), and total red blood cells (F). g-k, Multiplex quantification of circulatory pro-inflammatory molecules targeted by Bindarit: Ccl2 (G), Ccl7 (H), Ccl8 (I), TNF-α (J), and IL-6 (K). l, Kmt5a levels in muscle stem cells relative to adult cells. Source data

Extended Data Fig. 9. Short-term Bindarit treatment does not improve adult mouse skeletal muscle regeneration. Related to Fig. 6.

a) Strategy to assess Bindarit effect over adult skeletal muscle regeneration. Mice blood were collected pre-treatment, then post-treatment every week for whole blood count (WBC). Once levels of circulatory monocyte were significantly decreased, we stopped the treatment and proceeded with barium chloride injury. Mice rested for 21days before tissue harvest and analysis. b) WBC before and after 3-weeks Bindarit treatment versus vehicle treated mice. Twenty-four hours after the third round of treatment, Bindarit-treated mice showed significantly lower circulatory monocyte than before treatment. c) TA CSA from contralateral uninjured limb from mice treated for 3-weeks with vehicle or Bindarit. d) Regeneration assay for adult mice treated with vehicle or Bindarit for 3-weeks. e, f) Quantification of MuSC number in uninjured and injured TA (E) and regenerated fiber size average and frequency distribution (F) at 21dpi. For each violon plot, data points represent biological replicates (n = 5 mice per condition). Statistical significance was performed with ANOVA (B) or two-sided Welch’s t-test (E, F) and exact adjusted p values are reported in the figure. Source data

Extended Data Fig. 10. Plasma from Bindarit-treated aged mice reverses aging phenotypes in young recipient mice. Related to Fig. 6.

a) Schematic representation of the experimental design. Young mice were chronically injected with plasma derived from aged mice treated with Saline or Bindarit for 12 months to assess potential rejuvenating effects. b) Quantification of MuSC pool size in young mice following chronic plasma exposure. Plasma from Bindarit-treated aged mice maintained a normal MuSC pool comparable to control mice, unlike plasma from untreated aged mice. c) Assessment of skeletal muscle regeneration in young mice treated with aged Bindarit plasma, showing normal regeneration similar to controls. d-g) Analysis of MuSCs sorted from young mice exposed to aged Bindarit plasma. (D) Flow cytometry analysis showing levels of H4K20me1. (E) Lipid peroxidation analysis indicating normal lipid peroxidation levels. qPCR analysis showing normalized expression levels of Kmt5a (F) and Gpx4 (G). Data are presented as mean ± s.e.m. Statistical analyses were performed using one-way ANOVA (B-F), and p-values are indicated in the figure. Source data