Aged muscle displays a blunted α-Klotho response to injury

To determine whether α-Klotho is upregulated locally in response to an acute muscle injury, we performed immunofluorescence analysis of α-Klotho in the skeletal muscle of young (4–6 months) and aged (22–24 months) male mice under conditions of homeostasis and following a cardiotoxin-induced injury. α-Klotho was virtually undetectable in healthy, uninjured muscle, regardless of age (Fig. 1a–e). In contrast, strong expression of α-Klotho was observed at the regenerating site of young muscle 14 days post injury (dpi) (Fig. 1c, e; confirmation of antibody specificity is presented in Supplementary Fig. 1). Aged muscle, however, displayed no appreciable increase in α-Klotho expression following an acute injury (Fig. 1d, e). Serum α-Klotho levels followed a similar expression pattern according to age and injury status (Fig. 1f). RT-qPCR findings revealed that Klotho transcript expression increases significantly at 3 and 7 dpi injury in the skeletal muscle of young mice (Fig. 1g). Despite the fact that α-Klotho protein is still detected in young muscle at 14 dpi (Fig. 1c, e), gene expression approached baseline levels at this later time point. On the other hand, aged counterparts display unaltered gene expression across all the time points tested (Fig. 1g). The α-Klotho response to injury was not unique to a cardiotoxin injury, as we found that young mice exposed to a severe contusion injury displayed a robust α-Klotho response at the protein level 14 days after injury (Supplementary Fig. 2). Young female mice displayed a similar, yet blunted, increase in Klotho expression in response to injury, but, like males, the response is lost with aging (Supplementary Fig. 3).

Fig. 1 α-Klotho is increased in young muscle after injury, but the response is lost with increased age. a−d Immunofluorescent imaging of α-Klotho and F-actin in skeletal muscle from uninjured young (UIY; 4–6 months) and old mice (UIO; 22–24 months) as well as 14 days post injury (dpi) in young (YI) and old (OI) mice. Scale: 50 µm. e Quantification of α-Klotho across the four comparison groups, UIY, UIO, YI, and OI. Experimental cohorts were performed in duplicate, n = 3/group/cohort. ****p < 0.0001. f ELISA analysis of serum obtained from UIY, UIO, YI, and OI mice (n = 8–11/group; ****p < 0.0001). g RT-qPCR analysis of α-Klotho in young and old muscles at 0 (control), 3, 7, and 14 dpi (n = 4/age/timepoint). h MSPCR analysis of young and old muscle at 0 (control), 3, 7, and 14 dpi (n = 4/age/timepoint). i ChIP analysis of DNMT3a in young and old muscle at 0 (control), 3, 7, and 14 dpi (n = 3-4/age/timepoint). j ChIP analysis of H3K9me2 in the Klotho promoter in young and old muscle at 0 (control), 3, 7, and 14 dpi (n = 4/age/timepoint) (*p < 0.05 compared to young, sex-matched uninjured muscles; #p < 0.05 indicates a significant difference between young and aged groups at the respective timepoint). e−g One-way ANOVA with Tukey’s post-hoc test. h−j Two-way ANOVA with Tukey’s post-hoc test. Data represented as mean ± SEM Full size image

Epigenetic silencing of Klotho contributes to the impaired regenerative potential of dystrophic skeletal muscle, and a differentially methylated region (DMR) of 110 nucleotides within the Klotho promoter region was identified in the muscles of aged mdx mice26. Therefore, we measured methylation levels of the DMR after injury in young and aged muscle. An acute injury to young muscle triggered demethylation of the DMR in the Klotho promoter 3 and 7 dpi (Fig. 1h). Injury-induced demethylation was, however, absent in the Klotho promoter within aged muscle (Fig. 1h).

To examine whether modifying enzymes for DNA methylation contribute to the observed methylation changes, we used a chromatin immunoprecipitation (ChIP) assay to measure the enrichment of DNA methyltransferases 3a (DNMT3a) in the Klotho promoter region26. There was a decrease of DNMT3a binding to the Klotho promoter in the injured young muscle that, at the timepoints evaluated, reached a nadir 3 days after injury and was increased by day 14 (Fig. 1i). In contrast, the injury-induced decrease in DNMT3a binding was delayed and blunted in injured aged muscle (Fig. 1i). Consistent with the finding that dimethylated histone H3 lysine 9 (H3K9me2) bound to the Klotho promoter in dystrophic skeletal muscle represses Klotho26, H3K9me2 binding to the Klotho promoter region was decreased after injury only in young muscle (Fig. 1j). Taken together, these findings suggest that acute injury drives the reactivation of Klotho by reducing DNA methylation and H3K9 dimethylation in the promoter of young muscle, but that Klotho remains epigenetically repressed after injury in aged muscle.

Genetic inhibition of α-Klotho impairs muscle regeneration

To directly implicate a functional role for α-Klotho in skeletal muscle regeneration, we next evaluated the regenerative response to acute muscle injury in adult mice that are heterozygously deficient for Klotho (Kl+/− mice). As expected, Kl+/− mice displayed a significantly decreased local α-Klotho expression at the site of injury (Fig. 2a, b). Kl+/− mice also displayed a decreased regenerative index, smaller myofiber cross-sectional area, and increased fibrosis when compared to age- and sex-matched wild-type counterparts (Fig. 2a−e). These findings are consistent with a recent study of muscle regeneration in α-Klotho hypomorphs27. Together, the data demonstrate that α-Klotho is necessary for effective skeletal muscle regeneration after injury.

Fig. 2 Genetic and muscle-specific loss of α-Klotho impairs skeletal muscle regeneration. a Immunofluorescence of α-Klotho, laminin, F-actin, and Sirius red stain in wild-type and Kl+/− mice 14 dpi. Scale: 50 µm. b Quantitation of α-Klotho in wild-type versus Kl+/− mice 14 dpi (n = 3–4/group; *p < 0.05, one-tailed Student’s t test). c−e Quantitation of the % of centrally nucleated fibers (n = 4/group; *p < 0.05, Mann−Whitney U test), fiber cross-sectional area (n = 4/group; *p < 0.05, one-tailed Student’s t test) and collagen (Sirius red) deposition (n = 4/group; *p < 0.05, Welchʼs t test). f Representative hematoxylin and eosin stain of non-targeting control (NTC) and shRNA to α-Klotho (0.2–3.82×106 TU/TA) Scale: 50 µm. g, h Quantification of the % centrally nucleated fibers and ratio of myofiber area to total area, respectively, in NTC and Klotho shRNA-treated mice at 14 dpi (n = 3–8/group; *p < 0.05, Mann−Whitney U test). i Representative immunofluorescence imaging of lipid in NTC and α-Klotho shRNA-treated muscle at 14 dpi. Scale: 50 µm. j Quantification of lipid in NTC and Klotho shRNA-treated muscle 14 dpi (n = 4–6/group; *p < 0.05, one-tailed Student’s t test). k Quantification of collagen deposition (Sirius red) in NTC and Klotho shRNA groups (n = 4–9/group; p > 0.05, Student’s t test). l Quantification of fiber cross-sectional area of regenerating muscle fibers in NTC and α-Klotho shRNA-treated muscle (n = 5–7/group; ****p < 0.0001, one-tailed Student’s t test). m Representative second harmonic generation (SHG) images of tibialis anterior (TA) muscles injected with NTC or α-Klotho shRNA. Scale: 30 µm. n SHG quantification of the regeneration index in NTC and Klotho shRNA-treated mice at 14 dpi (n = 6–8/group; ****p < 0.0001, one-tailed Student’s t test). o Hang impulse (calculated as hanging time × mouse weight) at 14 dpi represented as a fold change from baseline score pre-injury (n = 6/group; *p < 0.05, one-tailed Student’s t test). Data represented as mean ± SEM Full size image

Given our findings of increased Klotho expression within the injured muscles of young mice (Fig. 1c,g,e), we next evaluated the contribution of local α-Klotho in functional muscle regeneration. The tibialis anterior (TA) muscles of young mice were injected with GFP-tagged SMARTpool® lentiviral particles carrying shRNA to α-Klotho, whereas control counterparts were injected with an equal volume of the non-targeting control (NTC) lentivirus. After 4 weeks, TAs were injured via a local cardiotoxin injection, and regeneration was evaluated 2 weeks after injury. As expected, muscles treated with shRNA to α-Klotho displayed a significant decrease in α-Klotho expression at the site of injury (Supplementary Fig. 1B, C). Circulating α-Klotho was also significantly decreased (Supplementary Fig. 1D). This suggests that muscle-derived α-Klotho may contribute to the increased circulating levels observed in young mice after injury (Fig. 1f).

Histological analysis revealed that knockdown of α-Klotho expression resulted in a decreased number of regenerating fibers, a decrease in the percentage of myofiber area/total area, and increased adiposity (Fig. 2f−j). There was, however, no difference in the fibrosis across groups (Fig. 2k). The cross-sectional area of regenerating (centrally nucleated) fibers in muscles treated with shRNA to α-Klotho was also significantly smaller than NTC controls (Fig. 2l), further confirming a defective regenerative response. Unexpectedly, we also observed the presence of a number of large, nonregenerating myofibers at the injury site of muscles treated with shRNA to α-Klotho (Fig. 2f). To evaluate the structural integrity of these myofibers, we performed second harmonic generation (SHG) imaging, which allows for three-dimensional visualization of myofiber structure and organization. Consistent with histological findings, SHG analysis revealed that muscles treated with shRNA to α-Klotho contain a decreased number of centrally nucleated fibers (Fig. 2m, n). This decreased evidence of active regeneration was concomitant with pathologic myofiber architecture and integrity (Fig. 2m and Supplementary Movies 1 and 2). Most importantly, the impaired regenerative response and disrupted myofiber structure was concomitant with a decreased functional recovery after injury (Fig. 2o).

α-Klotho is expressed by MuSCs and their progeny

The fact that α-Klotho expression in young muscle is elevated at 3 days after injury (Fig. 1g)—a time point that corresponds with MuSC activation—led us to next investigate whether MuSCs express α-Klotho and whether α-Klotho is necessary for the MuSC response to injury. We accessed RNAseq data from a recent study28, which is stored on the Gene Expression Omnibus (GEO) publicly accessible database. Analysis of archived data revealed a ten-fold increase in Klotho expression of freshly sorted MuSCs as compared to whole muscle lysates (Fig. 3a). Structured illumination microscopy (SIM) confirmed robust α-Klotho in MPCs isolated from young mice (Fig. 3b, c). MPCs were cultured for no more than three passages prior to analysis and were confirmed to be >90% MyoD+. MPCs isolated from aged muscle, however, displayed a markedly decreased α-Klotho protein expression (Fig. 3b, c). We also evaluated α-Klotho expression in MuSCs isolated by fluorescence-activated cell sorting29. As observed in MPCs, we found that young MuSCs displayed a robust α-Klotho expression, but that α-Klotho expression was decreased in aged MuSCs (Supplementary Fig. 4).

Tissues previously shown to express high levels of α-Klotho, such as the kidney, contain a membrane-bound form of α-Klotho (~120 kDa), which functions as an obligate co-receptor for fibroblast growth factor-23 (FGF23)30. However, upon proteolytic cleavage of the extracellular domain, α-Klotho is released from the cells, where it functions as a humoral factor15. To determine whether MPCs secrete α-Klotho, we performed ELISA of the conditioned media from MPCs isolated from young and old muscle. Conditioned media derived from young MPCs contained significantly more α-Klotho than the conditioned media from aged MPCs (Fig. 3d), suggesting that MuSCs secrete α-Klotho but that secretion declines with age.

Fig. 3 α-Klotho expression in quiescent and activated MuSCs. a Klotho expression in isolated MuSCs versus whole muscle lysates as per RNAseq analysis in transcripts per million (TPM)27 (n = 4/group; ****p < 0.0001, one-tailed Student’s t test). b Representative structured illuminescent microscopy of α-Klotho in young and old MPCs. Scale: 5 µm. c Quantification of α-Klotho in young and old MPCs (*p < 0.05, one-tailed Student’s t test). d ELISA analysis of α-Klotho in culture media alone, as well as conditioned media from young and old MPCs (n = 3/group; **p < 0.01, ****p < 0.0001, one-way ANOVA with Tukey’s post-hoc test). e Immunofluorescent colocalization of MyoD, F-actin, and α-Klotho 3 dpi. Scale: 50 µm. f Heat-map representation of α-Klotho as well as markers of MuSC activation (MyoD1, Fos, Jun, Myf5) in quiescent and activated cells from RNASeq analysis of publicly archived data from a recent report27. g Immunofluorescence staining of α-Klotho and DAPI in sorted MuSCs and fibroadipogenic progenitors (FAPs) fixed immediately after isolation (Day 0) or after activation in culture (Day 3). Scale: 12.5 µm. h Quantification of α-Klotho expression in MuSCs and FAPs at Day 0 and Day 3 of culture (****p < 0.0001, two-way ANOVA with Tukey’s post-hoc test). i Quantification of α-Klotho in the conditioned media of MuSCs and FAPs sorted from uninjured muscle. Conditioned media was obtained after 3 days in culture (n = 3/group; ****p < 0.0001, one-way ANOVA with Tukey’s post-hoc test). j Quantification of α-Klotho in MuSCs and FAPs isolated from uninjured muscle and muscle 3 dpi (**p < 0.01, ***p < 0.001, two-way ANOVA with Tukey’s post-hoc test). k Immunofluorescence imaging of α-Klotho and DAPI in MuSCs and FAPs sorted from uninjured muscle and muscle 3 dpi. Scale: 50 µm. l Immunofluorescent imaging of Pax7 and MyoD in MPCs isolated from wild-type and Kl+/− mice. Scale: 50 µm. m Quantification of the % of MyoD+ cells in MPCs from wild-type and Kl+/− mice (*p < 0.05, one-tailed Student’s t test). n Quantification of the % of Pax7+ cells in MPCs from wild-type and Kl+/− mice (One-tailed Student’s t test). o Immunofluorescent imaging of MyoD and DAPI in the injured muscles of nontargeting control (NTC) and shRNA to α-Klotho 14 dpi. Scale: 25 µm. p Quantification of the percentage of MyoD+ nuclei within the injured muscles of nontargeting control (NTC) and shRNA to α-Klotho 14 dpi (n = 4–6/group; ****p < 0.0001, one-tailed Student’s t test). Data represented as mean ± SEM Full size image

Immunohistochemical analysis of young muscle three days after injury revealed that 96.7% of MyoD+ cells express α-Klotho (Fig. 3e). MyoD is a marker of activated MuSCs and is required for myogenic lineage progression31. We accessed and analyzed MuSCs RNAseq data housed on the GEO repository28 and found that activated MuSCs display significantly increased Klotho expression when compared to quiescent MuSC counterparts that were isolated from 1% PFA-perfused mice (Fig. 3f). These findings suggest that α-Klotho expression is increased in the transition from MuSC quiescence to activation. To investigate this more directly, we compared α-Klotho expression in freshly sorted (quiescent) MuSCs and MuSCs that were maintained in culture for 3 days, thereby promoting activation in vitro32. Activated MuSCs displayed a 4.5-fold increase in α-Klotho, when compared to quiescent counterparts (Fig. 3g, h). We also isolated MuSCs from uninjured young muscle and young muscle 3 dpi, yielding a population of quiescent and activated MuSCs, respectively (Supplementary Fig. 4C). As observed when MuSCs were activated in vitro, there was a significant increase in α-Klotho expression in MuSCs that were activated in vivo (Fig. 3j, k). As a comparison, fibroadipogenic progenitor cells (FAPs), which also play a critical role in the skeletal muscle regenerative cascade33, displayed no change in α-Klotho over 3 days of activation in culture, nor was α-Klotho expression increased when FAPS were isolated from acutely injured muscle (Fig. 3g–k, Supplementary Fig. 4D). The conditioned media from activated FAPs also contained significantly less α-Klotho when compared to the conditioned media from activated MuSCs (Fig. 3i). Therefore, at least some of the α-Klotho protein detected in muscle after injury could come from MPCs themselves, although other neighboring cell populations may also express and secrete α-Klotho in response to an acute injury. Indeed, it was recently demonstrated that α-Klotho derived from macrophages promotes muscle regeneration34.

We next asked whether α-Klotho is necessary for normal MuSC lineage progression. MPCs isolated from the skeletal muscle of Kl+/− mice displayed a small, but significant, decrease in the percentage of MyoD+ cells when compared to age-matched wild-type counterparts. There was, however, no difference in Pax7 expression across groups (Fig. 3l–n). These findings are consistent with a previous report demonstrating that α-Klotho hypomorphs display a decreased number of MyoD+ cells as compared to control counterparts27. In vivo, lentiviral shRNA inhibition of α-Klotho resulted in a decreased MyoD expression at the site of injury (Fig. 3o, p). Taken together, these data suggest that a loss of α-Klotho disrupts MuSC lineage progression.

Loss of α-Klotho in MPCs drives mitochondrial dysfunction

Previous studies have implicated α-Klotho as being involved in multiple processes associated with inhibition of senescence24. Senescence is actively repressed in young MuSCs, but this capacity declines with aging, resulting in a failure of MuSCs to activate and proliferate in response to injury35. Therefore, we investigated whether the decreased MuSC activation in α-Klotho-deficient muscles may be attributed to cellular senescence. To confirm an inhibitory role for α-Klotho in myogenic cell senescence, we used small interference RNA (siRNA) inhibition, which resulted in a ~3-fold decrease in α-Klotho (Supplementary Fig. 1G, H). As expected, knockdown of α-Klotho induced a senescent phenotype in young MPCs, as evidenced by the percentage of senescence associated (SA)-βgal-positive cells, increased cytosolic HMGB1 levels (an indicator of cellular stress), and decreased cellular proliferation (Supplementary Fig. 5). These findings mimicked the phenotype of MPCs isolated from aged muscle (Supplementary Fig. 5).

Given that senescent cells are known to have higher levels of DNA damage36, we used LXRepair multiplex technology37,38 to evaluate the DNA base excision repair (BER) enzyme activities of OGG1 and APE1, which work on two common oxidative DNA lesions, 8-oxodG and abasic sites, respectively. When compared to scrambled siRNA-treated young MPCs, there was no significant decline in BER in the nucleus of cells treated with siRNA to α-Klotho (Supplementary Fig. 5G, H). However, just 48 h of α-Klotho supplementation to siRNA-treated MPCs from young animals dramatically stimulated the activity of these BER enzymes (Supplementary Fig. 5G, H). These data suggest that α-Klotho’s role in MPC senescence may be attributed to induction of key enzymes in the BER pathway, which is rapid and can occur even within a couple of hours39.

While α-Klotho’s role in cellular senescence has been demonstrated in multiple systems, the mechanisms underlying this role are incompletely understood. Cellular senescence is associated with altered mitochondrial morphology and dysregulated bioenergetics that can lead to aging and aging-associated pathologies40,41. Thus, regulation of MPC senescence by α-Klotho raised the intriguing possibility that α-Klotho may regulate MPC mitochondrial structure and function. Using an antibody against the mitochondrial membrane protein, Tom20, we observed no difference in mitochondrial morphology, as determined by sphericity, the number of mitochondria per cell, or mitochondrial volume according to age or α-Klotho levels (Supplementary Fig. 6A, C–F). This was further supported qualitatively using STimulation Emission Depletion (STED) microscopy to visualize the mitochondrial network (Supplementary Fig. 6B). These findings suggest that a loss of α-Klotho does not affect mitochondrial morphology or mass. However, detailed analysis by transmission electron microscopy (TEM) revealed a striking alteration of ultrastructural integrity of the mitochondrial cristae and endoplasmic reticulum in addition to lipid droplet accumulation when young MPCs were treated with siRNA to α-Klotho (Fig. 4a).

Fig. 4 Loss of α-Klotho drives mitochondrial dysfunction and disrupts mitochondrial DNA integrity. a TEM images of young, old, young + scramble and young + siRNA MPCs showing mitochondria (M), lipid droplet accumulation (L), as well as endoplasmic reticuli (ER). Scale: 400 nm. b, c Seahorse analysis of young, old, young + scramble and young + siRNA MPCs quantifying the basal oxygen consumption rate (OCR). d Seahorse analysis of basal OCR of MPCs isolated from wild-type and Kl+/− mice. e, f Seahorse analysis of reserve capacity (calculated as the difference between basal and maximum OCR) of young, old, young + scramble and young + siRNA MPCs. g Seahorse analysis of reserve capacity of MPCs isolated from wild-type and Kl+/− mice. h, i RT-qPCR based analysis of mtDNA damage in young, old, young + scramble and young + siRNA MPCs. j RT-qPCR analysis of mtDNA damage in MPCs isolated from wild-type and Kl+/− mice (*p < 0.05, **p < 0.01, ****p < 0.0001, one-tailed Student’s t test). Data represented as mean ± SEM Full size image

In light of the loss of mitochondrial ultrastructure resulting from decreased α-Klotho expression, we next evaluated whether loss of α-Klotho drives MPC mitochondrial dysfunction. Indeed, recent reports have shown a critical role for mitochondria in MuSC function42,43. The bioenergetic profiles of young and aged MPCs were studied using a Seahorse XFe96 Flux analyzer, which quantifies the oxygen consumption rate (OCR), a measure of oxidative phosphorylation. OCR was measured again after sequential injection of oligomycin, FCCP, 2-deoxyglucose (2-DG), and rotenone. These data demonstrate that, when normalized to total number of cells, MPCs from older animals display dramatically decreased levels of basal OCR as compared to young counterparts (Fig. 4b; Supplementary Fig. 7A, B). A similar age-associated deficit in the OCR of freshly isolated and cultured MuSCs was recently reported44. When young MPCs were treated with siRNA to α-Klotho, basal OCR was reduced to ~25% values of scramble-treated counterparts (Fig. 4c, Supplementary Fig. 7A, B). MPCs isolated from uninjured Kl+/− mice display a similarly blunted bioenergetic profile (Fig. 4d). Though there was no appreciable decrease in reserve capacity in aged MPCs, young MPCs treated with siRNA to α-Klotho and MPCs isolated from Kl+/− mice both showed a ~25% decrease in the reserve capacity (Fig. 4e–g; Supplementary Fig. 8A). Reserve capacity represents the spare bioenergetic capacity, is calculated as the difference between the basal and maximal OCR, and indicates the ability of a cell to respond to stress45. Taken together, these data support the hypothesis that age-related declines in α-Klotho drive impaired MPC mitochondrial bioenergetics.

One factor that could contribute to decreased mitochondrial function with decreased α-Klotho is oxidative damage, which could manifest itself as mtDNA damage46. Therefore, we next examined mtDNA integrity. Using a qPCR-based assay that we have successfully applied to different cell types including mouse tissue and human peripheral lymphocytes47,48,49, we evaluated mtDNA damage in MPCs isolated from young or aged mice. The method used is based on the principle that a wide variety of types of DNA damage have the propensity to block DNA polymerase progression46. Therefore, this assay detects numerous kinds of base DNA damage or DNA repair intermediates such as abasic sites, as well as single and double DNA strand breaks.

There was no difference in steady-state mtDNA copy number across groups (Supplementary Fig. 7C, D), consistent with immunocytochemical analysis showing no difference in mitochondrial mass (Supplementary Fig. 6C). However, we found that MPCs isolated from aged mice displayed higher levels of mtDNA damage as compared to young counterparts (Fig. 4h). Accordingly, young MPCs receiving siRNA to α-Klotho and MPCs isolated from Kl+/− mice displayed increased numbers of mtDNA lesions when compared to scramble-treated and wild-type controls, respectively (Fig. 4i, j).

Loss of α-Klotho impairs mitochondrial bioenergetics

What is the mechanism by which α-Klotho regulates mitochondrial function? The disrupted mitochondrial cristae structure within cells displaying decreased α-Klotho expression (Fig. 4a) suggests that α-Klotho is necessary for the maintenance of mitochondrial matrix integrity. Loss of matrix integrity disrupts mitochondrial respiration and induces the accumulation of reactive oxygen species (ROS). ROS accumulation, in turn, drives mtDNA damage and, ultimately, cellular senescence50. This led us to ask whether α-Klotho may preserve mitochondrial function through maintenance of mitochondrial ultrastructure. TEM analysis reveals a significantly greater number of vacuolated mitochondria with disrupted cristae in the MPCs of Kl+/− mice (Fig. 5a, b).

Fig. 5 Mitochondrial structure and function are impaired in Kl+/− mice, but are rescued with SS-31. a, b Representative TEM images and analysis of damaged mitochondria of wild-type (WT), Kl+/− and Kl+/−+ SS-31 groups (10–20 images were analyzed to quantify >100 mitochondria/group; ****p < 0.0001, one-way ANOVA with Tukey’s post-hoc test). Scale: 500 nm. c, d Representative immunofluorescent images and quantification of cardiolipin content, by Nonyl Acridine Orange staining (NAO, green) in WT, Kl+/− and Kl+/−+ SS-31 MPCs (*p < 0.05, one-way ANOVA with Tukey’s post-hoc test). Scale: 50 µm. e, f Representative immunofluorescent images from live imaging and quantification of ROS as determined by MitoSox staining (red) on live cells from WT, Kl+/− and Kl+/−+ SS-31 groups (****p < 0.0001, one-way ANOVA with Tukey’s post-hoc test). Scale: 50 µm. g RT-qPCR-based analysis of mtDNA damage on WT, Kl+/− and Kl+/−+ SS-31 MPCs (n = 3/group; *p < 0.05, one-way ANOVA with Tukey’s post-hoc test). h, i Seahorse analysis of basal OCR and reserve capacity of WT, Kl+/− and Kl+/−+ SS-31 MPCs (n = 4–6/group; *p < 0.05, one-way ANOVA with Tukey’s post-hoc test). j, k Quantification of MyoD+ cells at the site of injury of TA muscles from WT, Kl+/− and Kl+/−+ SS-31 groups (n = 3–6/group; ***p < 0.001, one-way ANOVA with Tukey’s post-hoc test). Scale: 25 µm. l, m Representative SHG images and analysis of the percentage of centrally nucleated fibers from WT, Kl+/− and Kl+/−+ SS-31 mice at 14 dpi (n = 3–4/group; *p < 0.05, **p < 0.01, one-way ANOVA with Tukey’s post-hoc test). Scale: 50 µm. n Quantification of myofiber cross-sectional area from WT, Kl+/− and Kl+/−+ SS-31 groups (n = 3–4/group; *p < 0.05, **p < 0.01, one-way ANOVA with Tukey’s post-hoc test). o Hang impulse (calculated as hanging time × mouse weight) at 14 dpi, represented as a fold-change from 1 dpi score during the wire hang test (n = 3–6/group; *p < 0.05, ***p < 0.001, two-way ANOVA with Tukey’s post-hoc test). Each mouse was used as its own control in order to account for baseline variability (shown in Supplementary Figure 8). Data represented as mean ± SEM Full size image

Cardiolipin is an anionic phospholipid that is confined almost exclusively to the inner mitochondrial membrane where it is synthesized. It is required for the mitochondrial respiratory supercomplex assembly and activity, the disruption of which enhances the generation of mitochondrial ROS25. We found that cardiolipin content is depleted in MPCs isolated from Kl+/− mice, as compared to wild-type counterparts (Fig. 5c, d). However, treatment with SS-31, a mitochondrially targeted peptide that mitigates cardiolipin peroxidation25, restored Kl+/− MPC ROS generation, mtDNA damage and cellular bioenergetics to levels comparable to wild-type counterparts (Fig. 5a–i, Supplementary Figure 8). Administration of SS-31 to Kl+/− mice also resulted in an increased number of MyoD+ cells at the site of injury, an enhanced regenerative index, and an increased myofiber cross-sectional area (Fig. 5j–n). Accordingly, Kl+/− mice treated with SS-31 display an increased strength recovery after injury that parallels that of wild-type counterparts (Fig. 5o). Of note, treatment of wild-type mice with SS-31 did not alter strength recovery after injury when compared to saline-injected counterparts. Taken together, these findings suggest that the defect in MPC mitochondrial bioenergetics and the accumulation of mtDNA lesions accompanying a loss of α-Klotho are mediated by disruption of mitochondrial ultrastructure, ultimately resulting in dysfunctional muscle regeneration.

Muscle regeneration is enhanced by α-Klotho supplementation

Given the established hormonal role of α-Klotho, we next investigated whether supplementation of α-Klotho may restore mitochondrial function in aged MPCs. We found that when MPCs isolated from aged skeletal muscle were cultured in the presence of recombinant α-Klotho for 48 h, the aged mitochondrial phenotype was improved, as determined by decreased mtDNA damage, increased OCR, and increased reserve capacity (Fig. 6a–c).

Fig. 6 α-Klotho supplementation improves aged MPC bioenergetics and muscle regeneration. a RT-qPCR-based analysis of mtDNA damage in old and MPCs and in old MPCs that received supplementation with recombinant α-Klotho in the culture medium for 48 h (n = 3/group; *p < 0.05, one-tailed Student’s t test). b, c Seahorse analysis of basal OCR and reserve capacity of age and aged+ α-Klotho MPCs (n = 6–8/group; **p < 0.01). d, e Representative immunofluorescent images and quantification of α-Klotho expression in aged muscle 14 dpi after systemic supplementation of α-Klotho via an osmotic pump, as compared to saline-infused control muscles (n = 5/group; ****p < 0.0001). Scale: 50 µm. f Quantification of the percentage of centrally nucleated fibers as per histological analysis across saline and α-Klotho infused animals 14 dpi (n = 3/group; *p < 0.05). g, h Representative SHG imaging and quantification of the percentage of centrally nucleated fibers of saline versus α-Klotho infused animals at 14 dpi (n = 5–6/group; *p < 0.05). Scale: 35 µm. i, j Representative images and quantification of MyoD+ cells at the site of injury 14 dpi in animals receiving osmotic pump delivery of saline or α-Klotho (n = 5/group; ***p < 0.001). Scale: 25 µm. k Representative immunofluorescent images showing laminin and DAPI in animals receiving i.p. administration of saline, α-Klotho 1–3 dpi, and α-Klotho 3–5 dpi. Scale: 50 µm. l Quantification of percentage of centrally nucleated fibers in aged animals receiving i.p. administration of saline, α-Klotho 1–3 dpi, and α-Klotho 3–5 dpi (n = 4/group; *p < 0.05, **p < 0.01). m Quantification of fiber cross-sectional area across the three i.p. injection groups (n = 4/group; ***p < 0.001, ****p < 0.0001). n Fold change hang impulse score over baseline scores across the three i.p. injection groups (n = 6/group). o Force-frequency curves obtained from in situ contractile testing analysis of specific force (n = 6–8/group, *p < 0.05 when comparing α-Klotho 3–5 dpi with saline control, #p < 0.05 when comparing α-Klotho 3–5 dpi with α-Klotho 1–3 dpi group; two-way ANOVA with repeated measures). a−j One-tailed Student’s t test. k−n One-way ANOVA with Tukey’s post-hoc test. Data represented as mean ± SEM Full size image

These encouraging in vitro findings led us to next probe whether α-Klotho supplementation may enhance skeletal muscle regeneration in vivo. To do this, α-Klotho was administered to aged mice via osmotic pump. Osmotic pumps were implanted three days prior to injury and were maintained for 14 dpi. At the dose tested, we observed a significant increase in local α-Klotho within the injured muscle areas (Fig. 6d, e). Systemic administration of α-Klotho in aged muscle resulted in an increased number of regenerating fibers after injury, as determined both by histology and SHG imaging (Fig. 6d–h). These findings were consistent with an approximately 3.5-fold increase in the number of MyoD+ cells at the site of injury in animals that received supplementation with α-Klotho (Fig. 6i, j). However, osmotic pump delivery of α-Klotho yielded no significant increase in myofiber cross-sectional area or total muscle area when compared to saline counterparts.

Given that osmotic pump administration delivers α-Klotho continuously, we next tested the hypothesis that the timing of α-Klotho administration may be critical for functional tissue regeneration. To do this, we performed daily intraperitoneal injections of recombinant α-Klotho to aged mice either from 1–3 dpi or from 3–5 dpi (i.e. the timepoints at which we found Klotho to be highly expressed in young muscle (Fig. 1g)). Similar to our findings using osmotic pumps, α-Klotho delivery both from 1–3 dpi and 3–5 dpi resulted in an increased number of regenerating fibers when compared to vehicle controls (Fig. 6k, l). Both treatment groups also displayed an increased myofiber area, though the magnitude of the improvement was greater in mice receiving α-Klotho over 3–5 dpi (Fig. 6m). Importantly, only animals receiving α-Klotho over 3–5 dpi displayed an improved functional recovery (Fig. 6n, o, Supplementary Table 1).

Taken together, the data suggest that α-Klotho is required for an adequate regenerative response to an acute injury, and that supplementation with α-Klotho via the circulation promotes MuSC commitment and myofiber regeneration in aged mice when administered at the appropriate timepoint. These findings implicate declines in this longevity protein as a contributor to a defective muscle regenerative response with aging and raise the possibility of systemic administration of α-Klotho as a therapeutic approach to promote the healing of aged skeletal muscle after injury.