Zinc deficiency contributes to blunted myogenesis in chronic kidney disease

Abstract

Frailty in patients with chronic kidney disease (CKD) greatly exacerbates disease comorbidities and increases probability of death. Prior research underscores molecular alterations in skeletal muscle physiology that may underly frailty and poor intervention response in this patient population. CKD can negatively affect satellite cell abundance and function, reducing skeletal muscle injury resilience and adaptive capacity. Pathogenic drivers of compromised satellite cell abundance and activity in patients with CKD remain largely unknown. To address this gap in knowledge, we isolated primary myogenic progenitor cells (MPCs) from patients with CKD and control participants. We also sought to define cell-extrinsic and intrinsic processes that may underlie myogenic deficits. We performed RNA sequencing on MPCs from control participants cultured in control serum, MPCs from control participants cultured in CKD serum, and MPCs from CKD participants cultured in control serum. We identified zinc mishandling as a shared pathway between control cells treated with CKD serum and CKD cells treated with control serum. Consistent with these observations, we found zinc deficiency and attenuated myogenesis in MPCs from patients with CKD. Finally, we showed that zinc supplementation partially restores the myogenic capacity of MPCs from patients with CKD. Together, these data highlight the importance of zinc metabolism in myogenesis and identify a novel mechanism whereby CKD pathogenesis impedes MPC differentiation.

New & Noteworthy

Satellite cell abundance and function are negatively affected by chronic kidney disease (CKD). Using primary myogenic progenitor cells (MPCs) cultured from patients with late-stage CKD and matched controls, we expose cells to CKD or control serum and identify metallothionein-induced zinc deficiency as both a cell-autonomous and non-autonomous consequence of CKD on MPCs. We find zinc deficiency likely attenuates myogenesis through an AKT-FOXO1 signaling cascade, which can be partially rescued by supplementation of exogenous zinc.

Graphical Abstract

INTRODUCTION

Satellite cells are the intrinsic stem cell required for successful muscle regeneration. Depletion of the satellite cell pool severely compromises muscle regeneration after injury (1, 2) and adaptation to exercise (3). Our group has previously shown a progressive decline in satellite cell abundance that associates with estimated glomerular filtration rate (eGFR) in patients with chronic kidney disease (CKD) (4). Skeletal muscle health is severely compromised with CKD, and exercise intervention has shown only modest functional benefit (5–7), suggesting a need for investigation of molecular effectors that may underlie satellite cell deficits and muscle dysfunction in CKD.

Features of CKD-induced cachexia and impaired myogenesis appear to be retained in cultured myogenic progenitor cells (MPCs) (8, 9), supporting the use of primary in vitro culture to study MPC dynamics in this population. However, circulating factors also play an important role: dialysis treatment has been reported to restore satellite cell abundance in patients with CKD (4), and several uremic toxins known to accumulate in CKD can directly influence MPC activity (10, 11). These findings highlight the importance of both intrinsic (cell-autonomous) and extrinsic (e.g. non-autonomous or circulatory) signals in driving CKD-associated myopathy.

Here, we identify zinc mishandling as a convergent mechanism contributing to extrinsic and intrinsic alterations to MPCs in patients with CKD. Transcriptomic analyses identify induction of metallothioneins, a family of genes responsible for sequestering labile zinc, in MPCs from patients with CKD and control MPCs treated with serum from patients with CKD. C2C12 myoblasts undergo large increases in intracellular zinc during differentiation (12), and whole-body metallothionein knockout models increase muscle size through increasing the cellular zinc pool (13). Thus, metallothionein-driven zinc sequestration may underlie the impaired myogenic capacity observed in CKD. Consistent with this, we find that zinc supplementation partially restores myogenic potential in MPCs derived from CKD patients. Together, these data support zinc supplementation as a novel therapeutic adjuvant for supporting muscle health with CKD.

MATERIALS AND METHODS

Study Population

Ethical approval was received by the institutional review boards at Albert Einstein College of Medicine and the University of Kentucky, and the study was conducted following the Declaration of Helsinki. All participants provided written informed consent. All patients with CKD were recruited from the nephrology clinics and faculty practice at Montefiore Medical Center. Participants with CKD were recruited between March 2022 and January 2025 from two prospective cohort studies of patients with an eGFR <30 ml·min⁻¹·1.73 m⁻² (stages 4 and 5 CKD) (Table 1). Eligible patients were ≥21 years of age and able to provide written informed consent for study participation. Exclusion criteria included lower extremity amputation, use of immunosuppressive medication in the previous 6 months, current cancer diagnosis and/or treatment, peripheral vascular disease requiring lower extremity vascular stenting, and additionally, for muscle biopsy, the use of anticoagulant medications or antiplatelet agents that could not be stopped for at least one week. CKD participant study visits occurred in the Clinical Research Center at Albert Einstein College of Medicine. eGFR was calculated by the Chronic Kidney Disease Epidemiology Collaboration (CKD-EPI) equation (14). Study data were collected and managed using REDCap (Research Electronic Data Capture) electronic data capture tools hosted at the Albert Einstein College of Medicine (15).

Table 1.

Participant Characteristics

	CKD	Control
N	6	6
Age (years)	57.7±14.5	56.2±23.6
Gender – n (%)
Female	4 (66.7%)	5 (83.3%)
Male	2 (33.3%)	1 (16.7%)
BMI (kg/m2)	25.8±5.4	23.6±3.0
Hypertension – n (%)	6 (100%)	0 (0%)
Diabetes Mellitus – n (%)	2 (33.3%)	0 (0%)
Coronary Artery Disease – n (%)	1 (16.7%)	0 (0.0%)
Peripheral Vascular Disease – n (%)	0 (0.0%)	0 (0.0%)
eGFR (mL/min/1.73m2)	18.2±6.6	N/A
Etiology of kidney disease
Diabetes	2 (33.3%)	N/A
Hypertension	1 (16.7%)	N/A
Glomerular disease	2 (33.3%)	N/A
Polycystic kidney disease	1 (16.7%)	N/A

Data are presented as mean ± standard deviation for continuous variables.

Abbreviations: CKD: chronic kidney disease, BMI: body mass index, eGFR: estimated glomerular filtration rate.

Generally healthy, sedentary individuals without diagnosis of kidney disease were recruited as Control participants from the University of Kentucky. Sedentary was defined as not participating in physically strenuous work, brisk physical leisure activity, or formal exercise more than once per week in the previous 3 months. Combined patient demographics are displayed in table 1.

Serum Collection, Muscle Biopsies, and Tissue Processing

Blood samples were acquired following an overnight fast and processed for serum prior to storage at −80°C. Biopsy procedures have been previously described (16, 17). CKD subjects were admitted to the Albert Einstein College of Medicine Clinical Research Center following an overnight fast. Through an incision site 15 cm proximal to the superior border of the patella, approximately 100–150 mg of muscle tissue was collected from vastus lateralis using a 12-gauge biopsy needle (Bard Monopty, Bard Biopsy Systems, Tempe, AZ). A ~30mg portion of the muscle sample was quickly and gently blotted to remove excess blood and then placed in sterile 1% glucose and kept on wet ice prior to overnight shipment to the University of Kentucky for cell isolation. Control participants were admitted to the University of Kentucky Center for Clinical and Translational Science following an overnight fast. Percutaneous muscle biopsies from the vastus lateralis were performed using a Bergström 5mm muscle biopsy needle with suction (18). A ~50mg portion of the muscle sample was quickly and gently blotted to remove excess blood and then placed in sterile 1% glucose and kept at 4°C overnight prior to cell isolation.

Primary isolation of MPCs

MPCs were purified from muscle biopsies from Control participants or patients with CKD following overnight incubation in sterile 1% glucose at 4°C. Biopsy samples collected at Albert Einstein College of Medicine were shipped overnight on wet ice to the University of Kentucky; samples collected from both institutions were incubated ~24hr in sterile 1% glucose at 4°C prior to isolation per our prior methods (19). Briefly, muscles were minced in F10 containing 10% fetal bovine serum, dispase II (2.4 U/ml; Roche Applied Science, Indianapolis, IN, USA) and collagenase D (1 mg/ml; Sigma-Aldrich, St. Louis, MO, USA). The muscle slurry was incubated with agitation for 60 minutes at 37°C. Enzymes were subsequently refreshed and the muscle slurry was incubated with agitation for an additional 30 minutes. The muscle slurry was then filtered through a 40μm filter and then spun at 500 g for 5 min to pellet cells. Cells were then plated on 10cm cell culture plates (Corning Primaria) and MPCs were isolated through multiple rounds of pre-plating until purity was reached. MPC purity was assessed by counting desmin+ cells relative to total nuclei at the time of experimentation (Control: 92.0±6.9% desmin+; CKD: 95.3±5.7% desmin+, P = 0.55). MPCs were maintained in growth medium composed of Ham’s F10 (cat#11550043), 20% fetal bovine serum, 1% pen-streptavidin, and 5ng/ml basic fibroblast growth factor (bFGF). MPCs for RNA sequencing experiments were cultured in 10% Control or CKD human serum (Figure 1A). To reduce the influence of participant serum heterogeneity, serum samples were pooled across groups (e.g. serum samples for all Control participants were pooled and serum samples for all CKD participants were pooled) prior to MPC treatment. Serum samples were matched for age and sex between groups (CKD vs. Control). During early expansion, MPCs were kept under 60% confluency, and MPCs were used between passages 6–9 for all experiments. Media was refreshed every other day.

Figure 1. Altered metallothionein expression in primary myogenic progenitor cells from patients with chronic kidney disease.

(A) Experimental design. (B-C) Volcano plots depicting DEGs between control MPCs treated with CKD vs control serum (B) and between CKD vs control MPCs (C). (D-E) GO: REACTOME pathways from DEGs in serum (D) and cell (E) comparisons. MT2A (F), MT1X (G), MTIE (H), and SLC30A1 (I) expression in control MPCs cultured in control serum (CC CS), control MPCs cultured in CKD serum (CC CKDS), and CKD cells cultured in control serum (CKDC CS). MT2A (J), MT1X (K), and MTIE (L) expression obtained from bulk RNA sequencing data from patients with CKD. Statistical significance was denoted as FDR p < 0.05. N=3 independent biological samples (B-I), n=7 Control and CKD (J-L). Data are presented as mean ± SD with individual points plotted.

Myotube differentiation

MPCs were plated at high density (200,000 cells/well) in 6 well plates in growth medium. Twenty-four hours after plating, media was switched to differentiation medium (Minimum Essential Medium (MEM) (Thermo cat#11095080, 2% fetal bovine serum, 1% pen-streptavidin). MPCs were differentiated into myotubes over 7 days with media changes every other day prior to RNA or protein isolation. MPCs used for immunocytochemistry and imaging were plated in chamber slides (Nunc Lab-Tek Chamber Slide System, #177445, ThermoFisher) at 20,000 MPCs/well prior to addition of differentiation medium and culture for 7 days.

Protein isolation and western blotting

Cells were washed once in 1x PBS and scraped in 200 μL of RIPA buffer (Sigma Aldrich, USA) supplemented with 10 μL of HALT protease inhibitor cocktail (Halt Protease and Phosphatase Inhibitor Cocktail, #78440, ThermoFisher). Protein concentration was determined using the Bradford assay (20). 30 μg of protein were diluted 1:3 in sample buffer (4x Laemmli Sample Buffer, Bio-Rad, #1610747) boiled at 100°C for five minutes, and loaded onto a 4–20% gradient gel (Bio107 Rad, #5678094, TGX Stain-Free Protein Gels) and ran electrophoretically at 150V until the protein bands reached the bottom of the gel (~70min). Following electrophoretic separation, stain free gels were activated on a ChemiDoc MP for 1 minute under UV illumination. Protein was then transferred to PVDF membranes (Bio-Rad, #1620175) at 50V for one hour in ice cold transfer buffer. Equivalent loading and efficient transfer was verified using stain free imaging of the PVDF membranes. Membranes were subsequently blocked for one hour at room temperature in a 2% bovine serum albumin solution and then incubated in primary antibody overnight with gentle agitation: an anti-p-AKT (cat#9271), anti-t-AKT (cat#9272), anti-p-FOXO1 (cat#9461), anti-t-FOXO1 (cat#2880), and anti-pan myosin heavy chain (cat#) at 1:1000. A horse radish peroxidase (HRP)-linked donkey anti rabbit IgG secondary antibody (#GENA934; Sigma Aldrich) or goat anti-rabbit AF555 (cat#A-21428) secondary antibody was used for AKT and FOXO1 blots. Anti-mouse IgG1 AF647 (cat#A-21240) was used to probe myosin heavy chain. Membranes were imaged using optical density measurements on a ChemiDoc MP (Bio-Rad) and quantified using Image Lab software (Image Lab, Bio-Rad). Density values obtained from quantification of bands of interest were normalized to total protein via stain-free gels. Comparison across membranes was further considered by scaling all density values to an internal control present on all membranes.

RNA isolation and sequencing

MPCs used for RNA sequencing were cultured for 4 days in 10% pooled serum obtained from control participants or from patients with stage 4 or 5 CKD. Total RNA was extracted from primary MPCs by pipetting Trizol TRI Reagent (#R2050–1-200, Zymo Research) onto wells and manual scraping. RNA was isolated using the Direct-zol RNA Miniprep kit (#R2051, Zymo Research) per manufacturer’s instructions. RNA concentration and quality (average RIN: 9.7) was assessed with an RNA Nano chip kit on a bioanalyzer by the University of Kentucky Genomics Core Laboratory. Six hundred nanograms of total RNA was sent to Novogene for library construction and sequencing on a NovaSeq PE150 system using a paired-end 150 bp dual-indexing protocol. Raw FASTQ files underwent pre-alignment quality control and alignment at novagene using the GRCh38/HG38 reference genome. For previously published data, an aligned RNAseq count matrix containing raw counts from vastus lateralis biopsies in control participants and patients with CKD was downloaded from GEO series GSE157712. Differential gene expression analysis was performed in R (v4.4.2) using the DeSeq2 package was calculated comparing CKD and control samples (21). Raw p-values were adjusted for multiple testing using the Benjamini-Hochberg false discovery rate (FDR) step-up method. FDR-adjusted p values less than 0.05 and log2 fold change values <0.5 or >0.5 were used as significance cutoffs. RNA-sequencing data are deposited in Gene Expression Omnibus: GSE310640.

Immunocytochemistry

Following 7 days of differentiation, myotubes were fixed in 4% PFA for 15 minutes. Cells were then washed in PBS containing 0.1% triton-x for permeabilization and blocked for 60 minutes in 1% bovine serum albumin (BSA) for 1 hour. Cells were incubated overnight in anti-desmin (Abcam cat#ab15200) 1:100 and anti-myosin heavy chain (Santa Cruz, cat# sc-376157) 1:100. The next day, sections were washed in PBS prior to incubating in the appropriate secondary antibody (1:250 anti-rabbit Alexa Fluor 555 cat#A-21428 and 1:250 anti-mouse Alexa Fluor 488 cat# A-21121) for one hour and subsequently co-stained for DAPI with washes in PBS being performed in-between each secondary step.

Fluozin-3 incubation and analysis

Fluozin-3 AM (Thermo fisher, cat# F24195) is a cell-permeable zinc indicator with high zinc affinity that is unperturbed by calcium. For live imaging, all MPCs were incubated in 2 μM of Fluozin-3 for 40 minutes prior to imaging in a gas and temperature-controlled chamber (5% CO₂, 37°C) on a Nikon AXR confocal microscope (Nikon Instruments, Tokyo, Japan). Fluorescent intensity was quantified using ImageJ in at least 30 cells per sample by manually tracing the cell body and capturing the mean integrated density of the GFP channel by a single investigator blinded to status. Images were taken in the GFP channel corresponding to Fluozin-3 and brightfield to identify the cell body periphery. For flow cytometry, all samples underwent the same Fluozin-3 incubation time and dosages prior to trypsinization, resuspension in sorting buffer, and sorting using an iCyt FACS machine (Sony Biotechnology). Mean raw GFP fluorescent intensity was quantified on a minimum of 10,000 cells for each sample and replicate using FlowJo (v10.9.0).

Zinc supplementation

20 μM of zinc was determined as an efficacious dose from a combination of prior literature (22). Myotubes receiving zinc were cultured in differentiation medium containing 20 μM Zinc Chloride starting on day 1 of through day 7 of differentiation.

Image acquisition and analysis

Myotubes were visualized at 20x or 40x magnification (Plan-Apochromat 20x, NA: 0.8, air; Plan-Apochromat 40x, NA: 1.15, water) using a Zeiss M2 Axioimager with an automatic stage (AxioImager M2; Zeiss, Oberkochen, Germany) or a Nikon AXR confocal microscope (Nikon Instruments, Tokyo, Japan). Fusion index was calculated as number of nuclei in myotubes containing 2 or more nuclei, divided by the total number of desmin+ nuclei.

Statistical analysis

Statistical analysis of the RNA-sequencing data was performed using R (version 4.4.2), as detailed in the RNA isolation and sequencing methods. Unpaired t-tests were used for CKD vs. control comparisons, while paired t-tests were used for CKD+zinc vs. CKD comparisons. Statistical significance was denoted as P < 0.05 (*) and P < 0.01 (**). No data points were excluded from any analysis. Statistical testing and sample size are denoted in all figure legends. Comparisons that failed normality assumptions were natural log transformed and subsequently analyzed with appropriate parametric tests (Figure 2E, 3I).

Figure 2. Zinc deficiency attenuates AKT signaling and differentiation in myogenic progenitor cells from patients with CKD.

(A) Fluorescent intensity of Fluozin-3 traced from live cell images between control and CKD MPCs (p<0.05). (B) Representative images of from control and CKD MPCs treated with Fluozin-3. (C) Mean fluorescent intensity of Fluozin-3 via FACS and (D) representative histograms demonstrating cell counts and Fluozin-3 fluorescent intensity of control and CKD MPCs. (E) Myosin heavy chain expression in primary myotubes from controls or patients with CKD. (F) Representative western blot for MHC between control and CKD myotubes. (G) Representative image depicting multinucleation in control myotubes and not CKD myotubes. Scale bar=50μm. (H) Fusion index of CTRL and CKD myotubes after 7 days of differentiation. (I) phospho, (J) total, and (K) phospho/total AKT protein in primary myotubes from controls or patients with CKD. (L) Representative western blot for phospho and total AKT between control and CKD participants. (M) phospho, (N) total, and (O) phospho/total FOXO1 protein in primary myotubes from controls or patients with CKD. (P) Representative western blot for phospho and total FOXO1 between control and CKD participants. Scale bar=50μm. Statistical significance was denoted as p < 0.05 (*) and p < 0.01 (**) via independent t-test. N=3 independent biological samples (A-D), n=6 independent biological samples (E-H, M-P), n=5–6 independent biological samples (I-L). AU: Arbitrary Units.

Figure 3. Zinc supplementation partially restores AKT signaling and differentiation of myogenic progenitor cells from patients with CKD.

(A) Phospho-AKT, (B) total-AKT, and (C) phospho/total AKT protein from CKD primary myotubes with and without zinc supplementation. (D) Representative western blot for phospho and total-AKT between CKD myotubes with or without zinc supplementation. (E) phopho-FOXO1, (F) total-FOXO1, and (G) phospho/total FOXO1 protein from CKD primary myotubes with and without zinc supplementation. (H) Representative western blot for phospho- and total-FOXO1 between CKD myotubes with or without zinc supplementation. (I) MHC protein from CKD primary myotubes with and without zinc supplementation. (J) Representative western blot for MHC between CKD myotubes with or without zinc supplementation. (K) Fusion index between CKD and CKD+zinc conditions after 7 days of differentiation. Scale bar=50μm. Statistical significance was denoted as p < 0.05 (*) via paired t-test (CKD+zinc vs. CKD). N=6 paired biological samples (A-I), n=3 paired biological samples (K).

RESULTS

Zinc mishandling links extrinsic and intrinsic CKD-induced changes in MPCs

RNA sequencing identified a total of 11 differentially expressed genes (DEGs) between control muscle progenitor cells (MPCs) treated with CKD or control serum (Figure 1B), and 56 DEGs (FDR-adjusted P < 0.05) between CKD MPCs and control MPCs (Figure 1C). Gene ontology (GO: Biological Process) pathway analysis using Enrichr (23) revealed intracellular zinc ion homeostasis as the top enriched pathway in both comparisons (serum: Figure 1D; cells: Figure 1E). This pathway was driven by a cysteine-rich family of proteins that sequester labile zinc, termed metallothioneins, in tandem with Znt1, a plasma membrane transporter that is a major exporter of zinc (PMID: 7882967). Specifically, MT2A, MT1X, MT1E, and SLC30A1, were significantly upregulated (FDR-adjusted P < 0.05) in both CKD serum-treated control MPCs and CKD MPCs (Figure 1F–I).

To determine whether this signature is also present in vivo, we analyzed a previously published RNA-sequencing dataset of skeletal muscle biopsies from CKD patients and age-matched controls (4). Consistent with our in vitro findings, MT1X and MT1E were significantly upregulated in skeletal muscle of patients with CKD (FDR-adjusted P < 0.05), with a modest increase in MT2A expression as well in CKD (Figure 1J–L).

CKD MPCs exhibit zinc deficiency and impaired myogenesis

To assess intracellular labile zinc levels, we used Fluozin-3, a zinc-sensitive, cell-permeable fluorescent dye. Live-cell imaging and FACS analysis revealed significantly reduced free zinc levels in CKD MPCs (Figure 2A–D). In C2C12 myoblasts, labile zinc increases during differentiation, and exogenous zinc promotes myotube formation (12). To test whether myogenesis is impaired in CKD MPCs, we differentiated control and CKD MPCs for 7 days. CKD MPCs exhibited significantly reduced myosin heavy chain (MHC) expression (Figure 2E–F) and decreased fusion (Figure 2G–H).

We sought then to explore putative effectors of impaired myogenesis given the reduced intracellular zinc we observed. Zinc can directly support phosphorylation and activation of AKT (24), and AKT–mTORC1 signaling is essential for myogenesis (25). CKD myotubes showed significantly reduced phosphorylation of AKT at Ser473 (Figure 2I–K, P < 0.01) and reduced phospho/total AKT ratio (Figure 2K, P < 0.05), with no change in total AKT levels (Figure 2J; representative blot, Figure 2L). We next evaluated phosphorylation of FOXO1 at Ser256, a key AKT phosphorylation target that is also required for myogenesis (26). Phospho-FOXO1 was significantly decreased in CKD myotubes (Figure 2M), with no changes in total FOXO1 (Figure 2N) or phospho/total FOXO1 ratio (Figure 2O; representative blot, Figure 2P).

Zinc supplementation enhances AKT signaling and myogenesis in CKD MPCs

Given our observed zinc deficits in primary MPC’s from patients with CKD, we sought to evaluate the restorative effect(s) of supplemental zinc. MPCs from patients with CKD were differentiated in standard media or media supplemented with 20 μM zinc chloride. Zinc significantly increased phospho-AKT levels (Figure 3A), with no change in total AKT (Figure 3B), resulting in a significantly increased phospho/total AKT ratio (Figure 3C; representative blot, Figure 3D). Zinc supplementation also significantly increased phospho-FOXO1 (Figure 3E), without affecting total FOXO1 (Figure 3F) or phospho/total FOXO1 ratio (Figure 3G; representative blot, Figure 3H). Finally, zinc-treated CKD MPCs displayed significantly increased MHC protein expression (Figure 3I–J) and numerically (but not statistically) enhanced myotube fusion (Figure 3K), indicating partial restoration of myogenic capacity.

Discussion

Here, we identify metallothionein expression in MPCs from patients with CKD as a contributor to free zinc deficiency that may result in reduced AKT phosphorylation and blunted myogenic capacity. We partially rescue myogenesis in CKD MPCs through zinc supplementation during differentiation. Zinc has been identified as a potent activator of AKT that acts through increasing ERK signaling cascades and reducing phosphatase activity (27). Zinc deficiency is also common in patients with CKD due to multiple factors (28, 29).

Cellular pools of free zinc are delicately regulated to avoid heavy metal toxicity, redox damage, and aberrant protein activity, as zinc binds an estimated 10% of the proteome (30) and is heavily concentrated in skeletal muscle (31). Metallothioneins, a family of small cysteine-rich proteins, function as heavy-metal cellular buffers with a particularly high affinity for zinc, cadmium, and copper ions (32). They play a critical role in maintaining the labile zinc pool and keeping free zinc concentrations low (33). Notably, metallothionein knockout significantly increases myofiber size and Myog expression, suggesting a zinc-dependent enhancement of myogenesis (13). This effect is mirrored in C2C12 differentiation experiments that directly show accumulation of free zinc during differentiation (12), and zinc supplementation increases myotube multinucleation (22). Metallothioneins are typically induced by zinc toxicity, making the substantial upregulation of metallothionein gene expression puzzling, given that CKD patients are predominantly zinc deficient (29). However, chemicals that induce oxidative stress are known to increase metallothionein expression (34) and metallothioneins are known to act as antioxidants (35). Accumulation of oxidative stress in patients with CKD is well-characterized both systemically and in skeletal muscle (36). Perhaps metallothionein expression is a protective response to uremia, inadvertently sequestering zinc and contributing to its deficiency, as metallothionein’s redox state is intimately tied to zinc binding and release (37). Like metallothioneins, ZnT1 expression typically increases in response to increased intracellular zinc (38), indicating that cellular zinc homeostasis requires cooperative control of both intracellular sequestration and zinc export. Thus, it is remarkable that coordinated upregulation of metallothioneins and ZnT1 occurs in the presence of intracellular zinc depletion, underscoring the extent to which zinc homeostasis is markedly disordered in CKD MPCs.

This study was not without limitations. Firstly, all experiments were carried out in vitro, isolating cells from environmental cues that greatly influence cellular behavior. Further, chronic kidney disease is not a genetic myopathy; thus, retainment of disease phenotype in vitro is not guaranteed. However, multiple studies have suggested CKD disease patterns are observed in myogenic progenitor cells in vitro (8, 9). Secondly, zinc mishandling was only partially confirmed in bulk RNAseq performed on patient biopsies, suggesting this phenomenon may partially be restricted to MPCs or attributed to the in vitro nature of the study. Additionally, differentiation medium contains no zinc. It is possible the use of differentiation medium exacerbates zinc deficiency in differentiating MPCs, partially contributing to the large myogenesis deficits we observed. Further, MPCs were purified using pre-plating, which is less rigorous than fluorescent or magnetic cell sorting via CD56 to establish purity. While pre-plating is a common and gentle method for enriching myogenic cells, it is less precise than fluorescent or magnetic cell sorting. However, while cell sorting approaches often achieve higher purity, they may also introduce cell stress or loss due to harsher processing conditions. Pre-plating was utilized to maximize MPC yield given the small tissue allotment for primary cell isolation, and still resulted in highly pure MPCs. All cells in the current study were between passages 6–9. Given myogenic purity variably declines with passage number, fibroblast contamination is a possible source of the biological variance observed between replicates (39). Additionally, restrictions in patient serum availability unfortunately prevented us from performing zinc live-imaging and myotube differentiation experiments in control cells treated with CKD serum.

Small fluctuations in zinc availability could meaningfully impact MPC myogenic capacity, potentially limiting skeletal muscle regenerative capacity and adaptation to exercise (40). Given patients with CKD often exhibit poor responsiveness to exercise interventions (5–7) and the salience of satellite cells for exercise adaptation (41–44), skeletal muscle zinc deficiency may represent an underexplored contributing factor to this phenomenon. Beyond skeletal muscle, zinc supplementation has garnered increasing attention as a therapeutic strategy in CKD (29), particularly for its role in modulating renal sodium retention and systemic hypertension (45). Given its broad physiological importance and established safety profile, zinc supplementation presents a practical, low-risk intervention with potential benefits across multiple organ systems. Despite this, few, if any, clinical trials have directly tested zinc supplementation in CKD, making this an exciting and highly promising avenue for future research.

Preserving Physical Function in CKD (PPF-CKD) Investigators

Albert Einstein College of Medicine/Montefiore Medical Center

Matthew K. Abramowitz, MD, MS

Laith Alzyood, MBBS

Maureen E. Brogan, MD

Moazam M. Cheema, MBBS

Mary J. Dominguez, MD

Sally Duran, BA

Molly C. Fisher, DO, MS

Ladan Golestaneh, MD, MS

Rimon Golovey, MD

Sonali Gupta, MBBS

Tanya S. Johns, MD, MHS

Michele H. Mokrzycki, MD, MS

Rahul Maheshwari, MD

Tina M. Neade, MD

Joel J. Neugarten, MD

William Paredes, BS

Qi Peng, PhD

Amanda C. Raff, MD

Stuart O. Rosenberg, MD

Gracy Sebastian, NP

Deep Sharma, MBBS

Milagros D. Yunes, MD

University of Kentucky

Christopher S. Fry, PhD

Alexander R. Keeble, PhD

Allison M. Owen, PhD

Lehman College, City University of New York

Brad J. Schoenfeld, PhD

University of Maryland School of Medicine

Christopher W. Ward, PhD

University of California, Berkeley

Moriel Vandsburger, PhD