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. 2026 Oct 8;604(20):8360–8378. doi: 10.1113/JP291356

Mitochondrial dynamics and repair differences in two mouse models of Friedreich's ataxia

Lili Salinas 1, Francisco Figueroa 1, Claire B Montgomery‐Bowie 1, Garrett P Cox 1, Gino Cortopassi 1, Elena N Dedkova 1,2,✉
PMCID: PMC13674193  PMID: 42850823

Abstract

Abstract

Friedreich's ataxia (FA) is an inherited disorder caused by reduced expression of the mitochondrial protein frataxin (FXN). Although defined as a neurodegenerative disease, cardiomyopathy is the leading cause of mortality. Cardiac manifestations of FA span from concentric hypertrophic cardiomyopathy with preserved systolic function and impaired diastolic relaxation, resembling heart failure with preserved ejection fraction (HFpEF), to systolic heart failure with reduced ejection fraction (HFrEF). The molecular mechanisms driving progression towards these cardiac phenotypes remain poorly understood. We examined mitochondrial content, dynamics and mitophagy in two FA mouse models: cardiac/skeletal muscle‐specific MCK‐Cre FXN knockout (FXN‐cKO) mice, which develop HFrEF, and inducible FXN knockdown (FXNKD) mice, which develop HFpEF‐like cardiomyopathy. FXN‐cKO hearts exhibited increased mitochondrial transcription factor A expression, an elevated mitochondrial‐to‐nuclear DNA ratio and greater mitochondrial density by electron microscopy, consistent with enhanced mitochondrial biogenesis. In contrast, FXNKD hearts showed no changes in mitochondrial content or biogenesis markers. Mitochondrial dynamics were markedly altered in FXN‐cKO hearts, with reduced mitofusin‐2 and increased FIS1 expression, accompanied by smaller, fragmented mitochondria. FXNKD hearts exhibited preserved fission, modestly increased fusion and mild mitochondrial enlargement. Markers of mitophagy were increased in FXN‐cKO hearts, but a reduced LC3‐II/I ratio suggested impaired autophagosome maturation and defective mitochondrial clearance. No changes in mitophagy were observed in FXNKD hearts. These findings demonstrate that FA‐associated HFrEF is characterized by increased mitochondrial biogenesis coupled with impaired mitochondrial quality control and fragmentation, whereas FA‐associated HFpEF‐like phenotype shows preserved mitochondrial architecture and turnover. Mitochondrial remodelling programmes may underlie divergent cardiac phenotypes in FA and represent therapeutic targets for disease modification.

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Key points

  • Friedreich's ataxia (FA) cardiomyopathy develops as heart failure with reduced ejection fraction (HFrEF) or heart failure with preserved ejection fraction (HFpEF)‐like disease, but mechanisms leading to different cardiac phenotypes are unclear.

  • We compared mitochondrial remodelling in FA mouse models with divergent cardiac phenotypes.

  • Frataxin knockout (FXN‐cKO) mice with HFrEF phenotype showed increased mitochondrial biogenesis, fragmentation and impaired mitochondrial quality control.

  • FXN knockdown (FXNKD) mice with HFpEF‐like phenotype showed largely preserved mitochondrial structure and turnover.

  • Distinct mitochondrial adaptations may drive phenotype‐specific cardiac outcomes in FA.

Keywords: cardiomyopathy, frataxin, Friedreich's ataxia, mitochondria, mitochondrial biogenesis, mitochondrial fission, mitochondrial fusion, mitophagy

Abstract figure legend Differential mitochondrial remodelling in Friedreich's ataxia (FA)‐associated cardiomyopathy. Schematic summary of mitochondrial alterations in two mouse models of FA exhibiting distinct cardiac phenotypes. In cardiac/skeletal muscle‐specific MCK‐Cre frataxin knockout (FXN‐cKO) mice, which develop heart failure with reduced ejection fraction (HFrEF), mitochondrial biogenesis is increased, as indicated by elevated mitochondrial transcription factor A (TFAM) expression, increased mitochondrial‐to‐nuclear DNA ratio and higher mitochondrial density. This is accompanied by reduced mitochondrial fusion (↓ mitofusin‐2), increased fission (↑ FIS1) and a fragmented mitochondrial network. Despite increased markers of mitophagy, a reduced LC3‐II/I ratio suggests impaired autophagosome maturation and defective mitochondrial quality control. In contrast, frataxin knockdown (FXNKD) mice, which exhibit a heart failure with preserved ejection fraction (HFpEF)‐like phenotype, display preserved mitochondrial content, relatively balanced dynamics with modestly increased fusion, unchanged fission and elongated mitochondrial morphology. Mitophagy markers remain largely unchanged, indicating intact basal mitochondrial turnover. Together, these findings highlight phenotype‐specific differences in mitochondrial biogenesis, dynamics and quality‐control pathways that distinguish FA‐associated HFrEF from HFpEF‐like cardiomyopathy.

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Introduction

Friedreich's ataxia (FA) is an autosomal recessive disorder caused by a GAA trinucleotide repeat expansion in the first intron of the FXN gene, which encodes the mitochondrial protein frataxin (FXN) (Campuzano et al., 1996, 1997). FXN plays a critical role in iron–sulphur cluster biogenesis and mitochondrial iron homoeostasis (Pastore & Puccio, 2013). Reduced FXN expression leads to mitochondrial iron accumulation, increased reactive oxygen species (ROS) production and impaired oxidative phosphorylation (Bidichandani et al., 1993). At the systemic level, these cellular disturbances manifest as progressive cerebellar ataxia, neurodegeneration, skeletal muscle atrophy and sensory loss (Bidichandani et al., 1993). Importantly, cardiomyopathy represents the leading cause of mortality in individuals with FA (Pousset et al., 2015; Tsou et al., 2011).

Given that FA originates from a primary mitochondrial defect, understanding mitochondrial homoeostasis is central to elucidating disease pathophysiology. Previous studies from our and other laboratories demonstrated dysregulation of mitochondrial biogenesis in FA fibroblasts (Garcia‐Gimenez et al., 2011; Jasoliya et al., 2017), but less is known about mitochondrial biogenesis and mitochondrial quality control (mitophagy) in FA hearts. Mitochondrial biogenesis refers to the synthesis of new mitochondria, whereas mitophagy is the selective removal of damaged or dysfunctional mitochondria (Dorn, 2019; Rimbaud et al., 2009). Maintenance of an appropriate balance between these processes is essential for meeting cellular energetic demands and preventing the accumulation of dysfunctional mitochondria (Thai et al., 2019). Mitochondrial fission and fusion are additional key regulators of mitochondrial homoeostasis and are particularly vulnerable to disruption in FA due to impaired iron–sulphur cluster formation and oxidative stress. Fusion promotes the mixing of mitochondrial contents and functional complementation, whereas fission facilitates mitochondrial redistribution, segregation of damaged organelles and targeting of dysfunctional mitochondria for degradation via mitophagy (Adebayo et al., 2021).

Clinically, FA‐associated cardiomyopathy presents with heterogeneous phenotypes. A longitudinal retrospective study following FA patients for over a decade identified two principal forms of cardiac involvement (Legrand et al., 2022; Pousset et al., 2015). One subgroup developed concentric hypertrophic cardiomyopathy characterized by increased left ventricular (LV) wall thickness with preserved systolic function, similar to heart failure with preserved ejection fraction (HFpEF). A second subgroup exhibited a dilated hypertrophic cardiomyopathy with marked LV enlargement, wall thinning and progressive systolic dysfunction, consistent with heart failure with reduced ejection fraction (HFrEF) (Lane et al., 2013; Legrand et al., 2020; Pousset et al., 2015; Weidemann et al., 2013). Patients who progressed to HFrEF exhibited longer GAA repeat expansions, lower FXN levels and significantly poorer survival outcomes (Pousset et al., 2015).

To model these divergent cardiac phenotypes, we employed two complementary mouse models of FA. FXN knockdown (FXNKD) mice, originally developed in the Geschwind laboratory (Chandran et al., 2017), display increased LV wall thickness without changes in total cardiac mass and maintain preserved ejection fraction, similar to the HFpEF phenotype. In contrast, cardiac/skeletal muscle‐specific MCK‐Cre FXN knockout mice (Fxnflox/null::MCK‐Cre; named here as FXN‐cKO) develop a dilated cardiomyopathy characterized by increased LV mass without proportional wall thickening, increased total cardiac mass and a severe reduction in ejection fraction, consistent with HFrEF (Salinas et al., 2024, 2025). Despite their relevance to human disease, the mitochondrial mechanisms underlying these divergent cardiac phenotypes remain incompletely understood. In particular, the contributions of mitochondrial biogenesis, mitochondrial dynamics and mitophagy to disease progression have not been systematically compared between these models. Direct evaluation of these pathways may reveal mitochondrial determinants that distinguish adaptive remodelling associated with preserved cardiac function from maladaptive remodelling that culminates in heart failure, thereby providing new insights into the pathogenesis of FA cardiomyopathy.

Materials and methods

Animal model and ethical approval

All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) protocol #23613 at the University of California, Davis, and were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. All experiments are reported in compliance with ARRIVE 2.0 guidelines. The experimental unit was the individual animal.

Mouse models and experimental design

Cardiac/skeletal muscle‐specific MCK‐Cre FXN knockout mice (Fxnflox/null::MCK‐Cre; hereafter FXN‐cKO) were generated and bred as previously described and studied alongside littermate control animals (control or CTRL) (Salinas et al., 2024, 2025). FXN‐cKO mice and controls were maintained on standard rodent chow (Teklad Global 18% Protein Rodent Diet; Inotiv Inc, West Lafayette, Indiana, USA) and killed at 8 weeks of age, corresponding to the established onset of cardiac dysfunction in this model.

FXNKD mice were generated as previously described (Chandran et al., 2017; Montgomery et al., 2025). Heterozygous transgenic mice were bred in‐house with C57BL/6J wild‐type (WT) mice. To induce FXNKD, transgenic mice were administered doxycycline‐containing chow (200 ppm; Teklad 2018 base; Inotiv Inc, West Lafayette, Indiana, USA). Mice were killed 12 weeks after doxycycline induction, which was designated as a cross‐sectional experimental time point. WT control mice were also maintained on doxycycline chow to control for potential off‐target effects of doxycycline treatment.

Animals were allocated to experimental groups based on genotype. Where feasible, investigators were blinded to genotype during data acquisition and analysis. No animals were excluded from analysis unless pre‐established humane end‐points were met or tissue quality was insufficient for downstream analyses.

Housing and husbandry

All mice were housed in a temperature‐ and humidity‐controlled animal facility under a 12 h:12 h light–dark cycle, with ad libitum access to food and water. Health status and food intake were monitored daily by trained animal care staff.

Euthanasia

The mice were killed by pentobarbital overdose, followed by exsanguination under deep anaesthesia to ensure death prior to tissue collection.

Sex as a biological variable

Only male mice were used in this study, as male animals exhibit more severe and reproducible cardiac phenotypes in both FXN‐cKO and FXNKD models (Montgomery et al., 2025; Perfitt et al., 2024; Salinas et al., 2024, 2025). Sex was therefore controlled to reduce biological variability and enhance interpretability of mitochondrial phenotyping.

Echocardiography

Cardiac structure and function were assessed using two‐dimensional (2‐D) transthoracic echocardiography with a VisualSonics Vevo 2100 imaging system (VisualSonics, Toronto, ON, Canada) equipped with an MS‐550D high‐frequency linear‐array transducer (22–55 MHz). Mice were anaesthetized with 1%–1.5% isoflurane delivered in 100% oxygen during image acquisition, as previously described (Salinas et al., 2024, 2025; Thai et al., 2018; Thai et al., 2021), and then quick recordings were performed in conscious animals to exclude potential anaesthetic effects on cardiac function. Parasternal short‐axis views were used for 2‐D and M‐mode imaging. LV dimensions, including end‐diastolic diameter, end‐systolic diameter and posterior wall thickness, were measured from 2‐D‐guided M‐mode tracings. Measures of systolic function, including ejection fraction (EF), stroke volume and cardiac output, were calculated from M‐mode measurements using standard formulas. LV mass was determined using established M‐mode–derived equations, as previously described (Diloretto et al., 2026; Thai et al., 2018; Thai et al., 2021).

Morphometric measurements

After euthanasia, hearts were rapidly excised, rinsed in ice‐cold buffer and weighed. Cardiac tissue was then either flash‐frozen for downstream biochemical analyses or fixed in Karnovsky's fixative for transmission electron microscopy. Sample processing, embedding and ultrathin sectioning were performed by the University of California, Davis Biological Electron Microscopy Core Facility. Sections were imaged using a FEI L120C transmission electron microscope. Digital micrographs were analysed using FIJI ImageJ (National Institutes of Health) for quantitative morphometric assessment. Briefly, mitochondrial size was assessed by setting the scale according to the scale bar on the image, then mitochondria were traced, and the area was output. Aspect ratio was assessed by measuring the longest part of the mitochondria (width) and dividing by the section perpendicular to the width (height). Cristae density was measured using Trainable Weka Segmentation by training the software to recognize cristae within a mitochondrion, then calculating the number of cristae per mitochondria (Segawa et al., 2020).

Quantitative real‐time PCR

Genomic DNA was extracted using the DNeasy Blood and Tissue Kit, and total RNA was isolated using the RNeasy Fibrous Tissue Kit (Qiagen, 74704, Germantown, MD, USA), according to the manufacturer's instructions. RNA concentration and purity were assessed spectrophotometrically, and cDNA was synthesized using the iScript cDNA Synthesis Kit (Bio‐Rad Laboratories, 1725038, Hercules, CA, USA). Quantitative reverse transcription PCR (qRTPCR) was performed using PowerUp SYBR Green Master Mix (Thermo Fisher Scientific, Waltham, MA, USA, A25742) on a QuantStudio 7 Pro Real‐Time PCR System (Thermo Fisher Scientific, Waltham, MA, USA). Gene expression levels were calculated using the comparative ΔΔCT method, with ribosomal protein S18 (Rps18) used as the reference gene.

Conventional and capillary western blotting

Total protein was extracted from frozen, crushed cardiac tissue using CellLytic MT Cell Lysis Reagent (Sigma‐Aldrich, C3228) supplemented with PhosSTOP phosphatase inhibitor and Complete EDTA‐free Mini protease inhibitor cocktail (Roche Diagnostics, 4906845001 and 04693159001). Samples were homogenized with stainless steel beads and agitated for 30 min at 4°C, followed by centrifugation at 5000 rpm for 10 min to remove insoluble debris. Supernatants were collected for total protein analysis. Mitochondrial fractions were isolated from frozen, crushed tissue using the Mitochondria Isolation Kit for Tissue (Abcam, ab110168), according to the manufacturer's instructions. Protein concentration was determined using the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific, 23225). For conventional western blotting, equal amounts of protein were resolved on NuPAGE 4%,–12% Bis‐Tris gels using NuPAGE LDS sample buffer (Invitrogen, WG1403 and NP0008) and transferred onto nitrocellulose membranes (Bio‐Rad Laboratories, 12023945). Membranes were imaged using an Odyssey CLx imaging system (LI‐COR Biosciences), and band intensities were quantified using Image Studio software. For capillary western blotting (Jess system), equal amounts of protein were loaded into the manufacturer‐provided assay plates along with appropriate reagents, according to the manufacturer's instructions and analysed using the Jess automated capillary immunoblotting system (ProteinSimple, Bio‐Techne, Minneapolis, MN, USA).

Antibodies

Primary antibodies: TFAM (Cell Signalling Technology, 8076), DRP1 (Cell Signalling Technology, 8570), FIS1 (Cell Signalling Technology, 32525), MFN2 (Abcam, ab56889), OPA1 (Cell Signalling Technology, 8047), Pink1 (Abcam, 23707), Parkin (Cell Signalling Technology, 32833), p62 (Abcam, ab56416), FUNDC1 (Cell Signalling Technology, 49240), BNIP3L (Cell Signalling Technology, 12396), LC3 (Cell Signalling Technology, 12 741), ATG7 (Cell Signalling Technology, 8558), ATG12–5 (Cell Signalling Technology, 4180), LAMP1 (Abcam, 24170), CD63 (Cell Signalling Technology, 10112), Cathepsin D (Cell Signalling, 74089), GAPDH (Abcam, ab8245), αTubulin (Millipore Sigma, T6199). Secondary antibodies: IRDye 800CW Donkey anti‐Rabbit IgG Secondary Antibody (LICOR, 926–32213), IRDye 680RD Goat anti‐Mouse IgG Secondary Antibody (LICOR, 926–68070).

Statistical analysis

All data were analysed for normality using Shapiro–Wilk test. If normal, data were analysed using unpaired t tests. If data were not normally distributed, Mann–Whitney test was used to compare groups. Data were analysed using Prism version 10.0 (Graph‐Pad Software Inc., San Diego, CA, USA), and P < 0.05 was considered statistically significant. Data are shown as mean ± SD.

Results

FXN‐cKO and FXNKD mice develop distinct types of cardiomyopathies

To determine differences in cardiac structure and function between FXN‐cKO, FXNKD and their respective control mice, we performed in vivo echocardiographic assessment. As shown in Fig. 1A , there were striking differences in overall heart size, LV dimensions, wall thickness and contractile function between the two FA mouse models. Quantitative analysis demonstrated that FXN‐cKO hearts were significantly enlarged compared to WT controls (+126%, P = 0.01; n = 4 per genotype), whereas FXNKD hearts (+8.5%, P = 0.4658; n = 5 per genotype) were comparable in size to controls (Fig. 1B ). LV posterior wall thickness at diastole (LVPW,d) was not changed significantly in FXN‐cKO mice (+20%, P = 0.8358; n = 4) but was increased in FXNKD (+70%, P = 0.0023; n = 5) mice (Fig. 1C ). In contrast, LV posterior wall thickness at systole (LVPW,s) was significantly reduced in FXN‐cKO mice (−38%, P = 0.009; n = 4), with no detectable changes observed in FXNKD (+11%, P = 0.237; n = 5) hearts (Fig. 1D ). These findings indicate that FXN‐cKO mice develop a dilated cardiomyopathy characterized by ventricular enlargement and excessive wall thinning during contraction. In contrast, FXNKD hearts maintain normal chamber dimensions but exhibit increased wall thickness, consistent with impaired ventricular relaxation. Although stroke volume was reduced in both FXN‐cKO (−30%, P = 0.033; n = 4) and FXNKD (−45%, P = 0.0008; n = 5) mice (Fig. 1E ), cardiac output was decreased only in FXNKD (−45%, P = 0.0008; n = 5) mice (Fig. 1F ), whereas EF was significantly decreased only in FXN‐cKO (−67%, P = 0.033; n = 4) hearts (Fig. 1G ). In summary, our data demonstrate that the FXN‐cKO model recapitulates a severe dilated cardiomyopathy consistent with HFrEF, whereas the FXNKD model develops a restrictive cardiomyopathy resembling the features of HFpEF.

Figure 1. Cardiac function in frataxin knockout (FXN‐cKO) and frataxin knockdown (FXNKD) mice.

Figure 1

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A, representative B and M‐mode echocardiography images recorded in control (CTRL) versus FXN‐cKO mice and WT+Doxy (WT) versus FXKD mice. B–G, summary of cardiac outcomes in FXN‐cKO and FXNKD mice compared to the corresponding control mice. B, left ventricle (LV) mass corrected (mg); C, LV posterior wall (LVPW) thickness in diastole (mm); D, LVPW thickness in systole (mm); E, stroke volume (µl); F, cardiac output (ml/min); G, LV ejection fraction (%). N = 4–5 mice per genotype. Significance was determined using t test, with P‐value indicated above the brackets.

Mitochondrial biogenesis is increased in FXN‐cKO but not FXNKD mice

Mitochondrial transcription factor A (TFAM) is a key regulator of mitochondrial biogenesis. It binds mitochondrial DNA (mtDNA) and plays a critical role in mtDNA transcription and replication, processes essential for maintaining mitochondrial number and function (Kang et al., 2018; Picca & Lezza, 2015). Western blot analysis demonstrated that TFAM protein expression was significantly increased in FXN‐cKO hearts (+72%, P = 0.0036; n = 4 per genotype) compared to CTRL mice (Fig. 2A ).

Figure 2. Mitochondrial biogenesis in frataxin knockout (FXN‐cKO) and frataxin knockdown (FXNKD) mice.

Figure 2

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A, protein expression of mitochondrial transcription factor A (TFAM) normalized to GAPDH in CTRL versus FXN‐cKO mice. B, mitochondrial‐to‐nuclear DNA ratio of Nd2/Rps18 in CTRL versus FXN‐cKO hearts. C, mitochondrial number per µm2 in CTRL versus FXN‐cKO hearts. D, protein expression of TFAM normalized to GAPDH in wild‐type (WT) versus FXNKD mice. E, mitochondrial‐to‐nuclear DNA ratio of Nd2/Rps18 in WT versus FXNKD hearts. F, mitochondrial number per µm2 in WT versus FXNKD hearts. A, D, N = 4 mice per genotype. B, E, N = 8 mice per genotype. C, F, N = 20 measurements from three different mouse hearts. Numbers above the brackets indicate the exact P‐value.

To further assess mitochondrial biogenesis, we quantified the mitochondrial‐to‐nuclear DNA ratio (mt/nuc DNA) in FXN‐cKO and CTRL heart tissues. ND2 (NADH dehydrogenase subunit 2), encoded by mtDNA, was used as a marker of mitochondrial DNA content, whereas Rps18, a nuclear‐encoded gene, served as a reference for nuclear DNA. The mt/nuc DNA ratio was increased by 53% in FXN‐cKO hearts (P = 0.0018; n = 8 per genotype) (Fig. 2B ). Rps18 levels remained unchanged, whereas ND2 levels were elevated, indicating a relative increase in mtDNA compared to nuclear DNA, consistent with enhanced mitochondrial biogenesis.

Transmission electron microscopy (TEM) further confirmed these findings. FXN‐cKO hearts exhibited a 57% increase in mitochondrial density (number of mitochondria per µm2) compared to CTRL hearts (P = 1.6E‐11; n = 20 measurements from 4300× images obtained from three hearts per genotype) (Fig. 2C ).

In contrast, FXNKD hearts showed no significant change in TFAM protein expression (P = 0.9104) relative to controls (Fig. 2D ). Similarly, no differences were detected in the mt/nuc DNA ratio (P = 0.7831) or mitochondrial density (P = 0.8017) as assessed by TEM (Fig. 2E , F ).

Collectively, these findings indicate that mitochondrial proliferation is markedly increased in FXN‐cKO hearts exhibiting HFrEF but is absent in FXNKD hearts with HFpEF‐like phenotype, suggesting phenotype‐specific mitochondrial remodelling. The enhanced mitochondrial biogenesis observed in FXN‐cKO hearts may represent a compensatory response to reduced contractile output, whereas FXNKD hearts, which retain preserved systolic function, may not require such mitochondrial adaptation.

FXN‐cKO, but not FXNKD, mice exhibit unbalanced mitochondrial fusion and fission

Analysis of mitochondrial dynamics revealed distinct remodelling patterns between FXN‐cKO (HFrEF) and FXNKD (HFpEF) hearts. In FXN‐cKO mice, fission‐promoting signals were dominant: Fis1 transcript levels were elevated (+36%, P = 0.018; n = 8 per genotype) (Fig. 3A ), and both FIS1 and DRP1 proteins were increased (+87%, P = 0.022 and +27%, P = 0.0723; n = 4 per genotype) (Fig. 3B , C ), demonstrating the activation of mitochondrial fragmentation pathways. Fusion capacity, on the other hand, was reduced, as evidenced by substantial decreases in Mfn2 and Opa1 gene expression (−75%, P = 1.8E‐8, −26%, P = 0.0005, respectively; n = 8 per genotype) (Fig. 3A ). MFN2 protein expression was decreased in FXN‐cKO hearts (−42%, P = 0.039), and although total OPA1 protein was paradoxically increased (+52%, P = 0.0048) (Fig. 3C ), the decreased OPA1‐l/OPA1‐s ratio (−31%, P = 0.0041; n = 4 per genotype) (Fig. 3D ) demonstrated a functional shift away from fusion towards fission. OPA1‐l (long) is the fusion‐permissive form of OPA‐1 compared to OPA1‐s (short), which tends to favour fission (Wang et al., 2021). Therefore, an increase in OPA1‐l/OPA1‐s would indicate a shift towards fusion, and a decrease would indicate a decrease in fusion and a potential increase in fission (Anand et al., 2014). These findings indicate impaired mitochondrial network integrity and increased mitochondrial fragmentation in the FXN‐cKO HFrEF model.

Figure 3. Disbalance of fission and fusion in frataxin knockout (FXN‐cKO) and frataxin knockdown (FXNKD) mice.

Figure 3

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A, gene expression of fission and fusion markers Fis1, Drp1, OPA1 and MFN2 in FXN‐cKO and FXNKD hearts normalized to controls. B, western blot images of the corresponding proteins involved in mitochondrial fission–fusion processes. C, protein expression of fission and fusion markers Fis1, Drp1, OPA1 and MFN2 in FXN‐cKO and FXNKD mice normalized to controls. D, protein expression of OPA1‐l/OPA1‐s ratio in FXN‐cKO hearts. E, OPA1‐l/OPA1‐s ratio in FXNKD hearts. F, schematic representation of fusion and fission. N = 8 mouse hearts per genotype for gene expression and n = 4 mouse hearts per genotype for protein expression. Numbers above the brackets indicate the exact P‐value.

In contrast, FXNKD mice showed trends of increased expression of both Mfn2 (+52%, P = 0.077) and fission‐associated Fis1 and Drp1 transcripts (+150%, P = 0.006; +49%, P = 0.085; n = 8 per genotype) (Fig. 3A ). MFN2 protein expression was increased (+26%, P = 0.027), with no major changes observed in FIS1 and DRP1 protein expression (n = 4 per genotype) (Fig. 3B , C ), indicating a shift towards fusion. Notably, despite reduced total OPA1 protein (−38%, P = 0.023), the OPA1‐l/OPA1‐s ratio increased (+50%, P = 0.0051) (Fig. 3E ). This suggests that mitochondrial networks in FXNKD HFpEF hearts favour structural preservation and adaptive fusion rather than fragmentation. Together, these findings demonstrate that FXN‐cKO hearts shift towards mitochondrial fission and fragmentation, whereas FXNKD hearts promote mitochondrial fusion, revealing fundamentally different mitochondrial remodelling strategies in HFrEF versus HFpEF.

Mitochondrial morphology is differentially altered in FXN‐cKO and FXNKD hearts

TEM revealed pronounced mitochondrial structural abnormalities in FXN‐cKO hearts (Fig. 4A ). Mitochondria exhibited reduced aspect ratio (−13%, P = 0.00001), reflecting increased circularity and the loss of elongated morphology (Fig. 4B ), along with a significant reduction in mitochondrial size (−17%, P = 1.6E‐22; n = 300 individual mitochondria from three mouse hearts) (Fig. 4C , D ). Cristae density is decreased in FXN‐cKO hearts as well (−25%, P = 6.4E‐9; n = 100 mitochondria from three hearts per genotype) (Fig. 4E ), which can indicate further damage in the mitochondria. These findings are consistent with enhanced mitochondrial fragmentation.

Figure 4. Changes in mitochondrial morphology in frataxin knockout (FXN‐cKO) hearts.

Figure 4

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A, control (CTRL) and FXN‐cKO mitochondrial electron microscopy (EM) images. B, CTRL versus FXN‐cKO mitochondrial aspect ratio. C, D, average mitochondrial size and size distribution in CTRL versus FXN‐cKO hearts. E, cristae density in CTRL and FXN‐cKO hearts. F, wild‐type (WT) and frataxin knockdown (FXNKD) mitochondrial EM images. G, WT versus FXNKD mitochondrial aspect ratio. H, I, average mitochondrial size and size distribution in WT versus FXNKD hearts. J, cristae density in FXNKD hearts. B, C, D, G, H, I, N = 300 mitochondria from three different mouse hearts per genotype. I, J, N = 100 mitochondria from three different mouse hearts per genotype. Numbers above the brackets indicate the exact P‐value.

In FXNKD hearts, morphological changes were more modest (Fig. 4F ). Aspect ratio was reduced by 7% (P = 3.5E‐12) (Fig. 4G ), whereas mitochondrial size was increased by 9% (P = 0.0005; n = 300 mitochondria from three hearts per genotype) (Fig. 4H , I ). The increase in size may reflect mitochondrial swelling or altered fusion–fission balance. Cristae density was unchanged in FXNKD hearts (Fig. 4J ).

Cardiac mitochondria were further classified into interfibrillar, subsarcolemmal and perinuclear populations. In FXN‐cKO hearts, mitochondrial size was reduced in interfibrillar (–43%, P = 4.3E‐17) and perinuclear regions (–19%, P = 0.0486), whereas subsarcolemmal mitochondrial size was unchanged (–9%, P = 0.2009) (Fig. 5A , C , E ). Aspect ratio was decreased in interfibrillar (–18%, P = 8.9E‐10) and subsarcolemmal mitochondria (–19%, P = 4.4E‐8) but not in perinuclear mitochondria (+0.4%, P = 0.8746, n = 100 mitochondria from three hearts per genotype).

Figure 5. Mitochondrial size and aspect ratio (length‐to‐width shape factor) vary significantly depending on their subcellular location and functional microdomain within cells.

Figure 5

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Interfibrillar mitochondrial size and aspect ratio in (A) frataxin knockout (FXN‐cKO) and (B) frataxin knockdown (FXNKD) hearts. Perinuclear mitochondrial size and aspect ratio in (C) FXN‐cKO and (D) FXNKD hearts. Subsarcolemmal mitochondrial size and aspect ratio in (E) FXN‐cKO and (F) FXNKD hearts. N = 100 mitochondria per subpopulation from three different mouse hearts per genotype. Numbers above the brackets indicate the exact P‐value.

In contrast, FXNKD hearts exhibited increased mitochondrial size within interfibrillar (+13%, P = 0.0023) and subsarcolemmal (+10%, P = 0.068) compartments but not in perinuclear (+2.6%, P = 0.6292) regions (Fig. 5B , F ). Also, the mitochondrial aspect ratio was reduced in these regions: interfibrillar (–7%, P = 0.0272) and subsarcolemmal regions (–12%, P = 1.4E‐5, n = 100 mitochondria from three hearts per genotype).

Collectively, FXN‐cKO hearts exhibit smaller, rounder and regionally depleted mitochondria, particularly within interfibrillar and perinuclear compartments. In contrast, FXNKD hearts demonstrate larger and more spherical mitochondria, particularly in interfibrillar and subsarcolemmal compartments, suggestive of enhanced fusion or compensatory remodelling that may help sustain calcium handling and contractility.

Mitophagy is impaired in FXN‐cKO hearts but preserved in FXNKD hearts

To determine whether mitochondrial clearance differed between the two FA models, we assessed key components of canonical and non‐canonical mitophagy pathways (Fig. 6A ). In FXN‐cKO hearts, Parkin protein expression was markedly increased (+127%, P = 0.0156; n = 4 per genotype), whereas no change was observed in FXNKD hearts (Fig. 6B , C ). Pink1 levels were unchanged in both models. The autophagy adaptor p62 was strongly elevated in FXN‐cKO hearts (+301%, P = 0.0008) but reduced in FXNKD hearts (−18%, P = 0.0022), suggesting divergent handling of ubiquitinated cargo.

Figure 6. Mitophagy process was halted at the level of autophagosome formation in frataxin knockout (FXN‐cKO) hearts.

Figure 6

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A, mitophagy schematic. B, western blot images. C, protein expression of Parkin, Pink1, p62, FUNDC1, BNIP3L, LC3‐I and II in FXN‐cKO and frataxin knockdown (FXNKD) hearts normalized to their respective control group. D, LC3‐II/I ratio in CTRL versus FXN‐cKO hearts. E, LC3‐II/I ratio in wild‐type (WT) versus FXNKD hearts. F, western blot images. G, protein expression of ATG7, ATG5‐12, LAMP1, CD63 and cathepsin D in FXN‐cKO and FXNKD hearts normalized to their respective control group. N = 4 per genotype. Numbers above the brackets indicate the exact P‐value.

Non‐canonical mitophagy receptors were also examined. FUNDC1 (FUN14 domain containing1) and BNIP3L (BCL2 Interacting protein 3 Like) expression was increased in FXN‐cKO hearts (+99%, P = 0.0123; +126%, P = 0.0296; n = 4 per genotype), with no change observed in FXNKD hearts (Fig. 6B , C ). These data indicate the activation of mitophagy signalling pathways in FXN‐cKO hearts at the receptor level.

To assess autophagosome formation, LC3 processing was evaluated. FXN‐cKO hearts exhibited a reduced LC3‐II/LC3‐I ratio (–43%, P = 0.0011; n = 4 per genotype), attributable to increased LC3‐I with unchanged LC3‐II (Fig. 6B , D ). This pattern is consistent with impaired LC3 lipidation and reduced autophagosome maturation. In contrast, FXNKD hearts showed only a non‐significant trend in LC3 processing reduction (–13%, P = 0.0679; n = 4 per genotype) (Fig. 6E ).

Because LC3 lipidation requires autophagy‐related 7 (ATG7) and the complex of autophagy‐related 12 and 5 (ATG12–ATG5) conjugation system, we assessed these components. ATG7 and ATG12–ATG5 expressions were markedly decreased in FXN‐cKO hearts (–67%, P = 0.0005; −52%, P = 0.03; n = 4 per genotype), whereas no changes were observed in FXNKD hearts (Fig. 6F , G ), providing a mechanistic explanation for impaired autophagosome formation in the FXN‐cKO model. Finally, lysosomal capacity was evaluated. LAMP1 (lysosome‐associated membrane protein‐1) expression was significantly reduced in FXN‐cKO hearts (–70%, P = 0.0002), as well as CD63 (−54%, P = 0.0256) and cathepsin D (−60%, P = 0.01376) (Fig. 6F,H ), suggesting compromised lysosomal handling in the FXN‐cKO model. LAMP1 was unchanged in FXNKD hearts; however, both CD63 and cathepsin D were significantly upregulated (+92%, P = 0.0309 and +166%, P = 0.0158, respectively) (Fig. 6F , H ), indicating potential increased lysosomal presence and function in FXNKD hearts compared to WT hearts.

Taken together, these findings indicate that mitophagy signalling is activated in FXN‐cKO hearts, as reflected by increased protein expression in the Pink1/Parkin mitophagy pathway, as well as an increase in direct mitophagy receptors FUNDC1 and BNIP3L. However, downstream autophagosome formation and lysosomal processing appear to be impaired, suggesting a disruption in mitophagy flux. In contrast, FXNKD hearts display preserved autophagic and mitophagic markers, consistent with maintained mitochondrial turnover.

Discussion

Mitochondria occupy approximately one‐third of cardiomyocyte volume, reflecting the extraordinary energetic demands of the heart (Nguyen et al., 2019). Because FXN deficiency is central to FA pathophysiology, mitochondrial dysfunction has long been considered a primary driver of cardiac disease. Understanding how FXN loss alters mitochondrial quality control, turnover and morphology in the heart is essential for identifying therapeutic strategies aimed at preventing cardiac failure in FA.

Mitochondrial dysfunction and oxidative stress

Mitochondrial dysfunction is a central hallmark of FA, underscoring the importance of effective mitochondrial quality‐control mechanisms. Clinical studies in patients with FA have demonstrated impaired ATP production and reduced activity of mitochondrial enzymes (Jimenez‐Jimenez et al., 2025). Similar mitochondrial abnormalities have been extensively characterized in both the FXN‐cKO and FXNKD mouse models. In FXN‐cKO hearts, our prior studies have reported decreased ATP production, reduced aconitase activity, impaired oxygen consumption rates and diminished activity of multiple electron transport chain complexes (Salinas et al., 2024, 2025). FXNKD mice exhibit many of the same mitochondrial defects, including increased ROS generation and impaired activation of the antioxidant transcription factor Nrf2. Deficient Nrf2 signalling is a well‐established feature of FA and likely contributes to inadequate cellular antioxidant defences. Although both models display comparable reductions in complex II activity, aconitase activity appears to be more severely impaired in FXN‐cKO hearts than in FXNKD hearts, suggesting a greater degree of iron–sulphur cluster dysfunction in the former model (Hui et al., 2021; Salinas et al., 2024, 2025).

Consistent with the reduced Nrf2 activity, our previous studies demonstrated impairment of downstream antioxidant defence pathways in both FA models, albeit with distinct molecular signatures. In FXN‐cKO hearts, expression of the antioxidant enzymes superoxide dismutase 2 (SOD2; −56%) and glutathione S‐transferase pi (GSTP; −66%) was significantly reduced, whereas NAD(P)H quinone dehydrogenase 1 (NQO1) expression remained unchanged (Salinas et al., 2024, 2025). In contrast, FXNKD hearts exhibited a marked reduction in NQO1 expression (−50%), whereas SOD2 and GSTP levels were not significantly altered (Montgomery et al., 2025). These findings suggest that oxidative stress is more severe in FXN‐cKO hearts than in FXNKD hearts. The greater oxidative burden in FXN‐cKO mice may contribute to the more pronounced disturbances in mitochondrial dynamics and quality‐control pathways observed in this model, potentially reflecting a more advanced stage of mitochondrial dysfunction and disease progression.

Mitochondrial biogenesis in FXN deficiency

Mitochondrial biogenesis is a critical mechanism for maintaining mitochondrial homoeostasis. Although FA is fundamentally a mitochondrial disease, the adaptive response observed in our models differs from that described in traditional HFpEF and HFrEF. FXN‐cKO mice exhibited increased mitochondrial number and elevated TFAM levels, likely representing a compensatory attempt to sustain ATP production in the setting of mitochondrial dysfunction. Although previous studies have reported reduced mitochondrial biogenesis in FA (Jasoliya et al., 2017; Lynch & Farmer, 2021), our data indicate enhanced biogenic signalling in the FXN‐cKO model.

In classical cardiomyopathies, TFAM‐driven mitochondrial biogenesis is generally beneficial, generating functional organelles (Kunkel et al., 2016). In FA, however, proliferation of structurally and metabolically compromised mitochondria may exacerbate ROS production and cellular stress. In contrast, FXNKD hearts showed no increase in mitochondrial biogenesis, consistent with a phenotype in which mitochondrial remodelling is not a dominant pathogenic mechanism.

Mitochondrial morphology and subpopulation‐specific remodelling

Comparative analyses of mitochondrial morphology in HFpEF and HFrEF remain limited. In FXN‐cKO hearts, we observed reduced mitochondrial aspect ratio and decreased mitochondrial size, consistent with the accumulation of fragmented and damaged mitochondria. The presence of enlarged swollen mitochondria (Fig. 4D ) further suggests impaired mitochondrial quality control and defective clearance of dysfunctional organelles. In contrast, mitochondria in FXNKD hearts were modestly larger and exhibited a reduced aspect ratio, indicative of mild swelling but less severe fragmentation than that in FXN‐cKO mice.

Beyond alterations in mitochondrial size and shape, FXN‐cKO hearts demonstrated a marked reduction in cristae density (Fig. 4E ). Electron micrographs further revealed disrupted cristae architecture, with control mitochondria exhibiting highly organized and uniformly distributed cristae, whereas FXN‐cKO mitochondria displayed irregular cristae orientation and heterogeneous cristae spacing throughout individual organelles. Such structural abnormalities are commonly associated with impaired oxidative phosphorylation, reduced bioenergetic capacity and mitochondrial dysfunction. Notably, reduced cristae density has been linked to alterations in optic atrophy protein 1 (OPA1)‐mediated inner mitochondrial membrane remodelling, although the specific contributions of individual OPA1 isoforms remain incompletely understood (Fry et al., 2024; Lee & Yoon, 2018). Mitochondria in FXNKD hearts do not show any change in cristae density Fig. 4J ), maintaining cristae structure and organization more than FXN‐cKO hearts.

Subpopulation analysis revealed that interfibrillar mitochondria exhibited the most pronounced morphological disruption in FXN‐cKO hearts (Fig. 5), consistent with their central role in supporting contractile ATP demand and calcium cycling. Given their primary function in providing ATP for sarcomeres’ contraction and ATP‐dependent calcium (Ravindran & Rau, 2024; Voglhuber et al., 2022), these structural abnormalities align with impaired contractile performance and disrupted calcium homoeostasis rather than compensatory mitochondrial expansion per se. Subsarcolemmal mitochondria, which regulate signalling pathways and facilitate substrate exchange at the cell membrane (Dedkova & Blatter, 2013; Holmuhamedov et al., 2012; Ravindran & Rau, 2024), displayed a reduced aspect ratio in FXN‐cKO hearts without significant changes in the overall size. Perinuclear mitochondria, implicated in nuclear signalling, protein homoeostasis and DNA repair processes (Lu et al., 2019), were reduced in size while maintaining a relatively preserved aspect ratio.

In FXNKD hearts, mitochondrial structural alterations were less severe but remained most evident in interfibrillar and subsarcolemmal populations, indicating partial impairment of contractile energetics and membrane‐associated signalling, with relative preservation of perinuclear mitochondrial structure. Together, these findings highlight the spatial specialization of mitochondrial subpopulations within cardiomyocytes and underscore the vulnerability of interfibrillar mitochondria in the context of FXN deficiency and contractile dysfunction.

Mitochondrial dynamics: fission–fusion imbalance

Mitochondrial fission and fusion constitute core quality‐control processes that regulate mitochondrial morphology, function and turnover (Youle & van der Bliek, 2012). Fission, mediated primarily by dynamin‐related protein 1 (DRP1) and its recruitment to the outer mitochondrial membrane via FIS1, enables the segregation of damaged mitochondrial components for removal. In contrast, fusion is co‐ordinated by mitofusin 2 (MFN2) at the outer membrane and optic atrophy 1 (OPA1) at the inner membrane, promoting intermitochondrial content mixing and functional complementation (Adebayo et al., 2021; Dorn, 2019; Green et al., 2022).

FXN‐cKO hearts demonstrated a clear shift towards enhanced fission, with increased expression of fission‐associated markers at both transcript and protein levels, accompanied by pronounced mitochondrial fragmentation. This was paralleled by reduced MFN2 expression and an OPA1 isoform shift consistent with impaired inner membrane fusion capacity, collectively indicating a loss of fusion competence. Together, these changes define a fission–fusion imbalance characterized by excessive mitochondrial division and insufficient structural recovery, leading to a fragmented mitochondrial network.

Functionally, this imbalance is typically associated with mitochondrial quality‐control activation and downstream mitophagy (Dorn, 2019; Green et al., 2022). However, in FXN‐cKO hearts, increased fission does not appear to translate into efficient mitochondrial clearance, suggesting uncoupling between fragmentation and removal of damaged organelles. In contrast, FXNKD hearts exhibit a more balanced fission–fusion profile, with a relative preservation of fusion capacity that may help maintain mitochondrial network integrity and limit fragmentation.

FXN‐cKO hearts demonstrated increased fission marker expression at both transcript and protein levels, accompanied by morphological fragmentation. Decreased MFN2 expression and a shift towards short‐isoform OPA1 suggest impaired fusion capacity (Chandhok et al., 2018; Dorn, 2019; Green et al., 2022; Wang et al., 2021). Collectively, these findings indicate a shift towards excessive fission and reduced fusion, resulting in fragmented mitochondrial networks. In heart failure, increased fission often accompanies mitochondrial dysfunction and may precede mitophagy or cell death. In FXN‐cKO hearts, however, enhanced fission appears uncoupled from effective mitochondrial clearance, potentially promoting the accumulation of dysfunctional organelles. In contrast, FXNKD hearts displayed a relative shift favouring fusion, which may buffer against excessive fragmentation and preserve mitochondrial integrity.

Mitophagy dysfunction in FXN‐cKO hearts

Mitophagy is essential for the selective removal of depolarized or dysfunctional mitochondria and is initiated through both canonical PINK1–Parkin–p62 signalling and receptor‐mediated pathways involving FUNDC1 and BNIP3L (Jin & Youle, 2012). In the canonical pathway, mitochondrial depolarization stabilizes PINK1 on the outer mitochondrial membrane, promoting Parkin recruitment and ubiquitination of mitochondrial substrates, which are subsequently recognized by the autophagy adaptor p62. This facilitates recruitment of LC3‐positive membranes and autophagosome formation (Kawajiri et al., 2010). LC3 lipidation (LC3‐I to LC3‐II conversion) is mediated by ATG7, ATG3 and the ATG12–ATG5–ATG16 complex, enabling autophagosome expansion and closure (Kawajiri et al., 2010).

FXNKD hearts showed no significant change in LC3‐II/I ratio, consistent with preserved basal mitophagic flux. In contrast, FXN‐cKO hearts exhibited evidence of mitophagy initiation, including increased upstream signalling markers, but a reduced LC3‐II/I ratio accompanied by the accumulation of LC3‐I and decreased LC3‐II levels. This was further supported by reduced expression of ATG7 and ATG12–ATG5 complex components, indicating impaired LC3 lipidation and defective autophagosome formation.

Collectively, these findings suggest that mitophagy is initiated in FXN‐cKO hearts but fails to progress to completion, resulting in inefficient clearance of damaged mitochondria. This defect, in combination with enhanced fission and reduced fusion capacity, provides a mechanistic basis for the accumulation of structurally and functionally compromised mitochondria in FXN‐cKO cardiomyopathy.

P62 then binds to ubiquitin and acts as a mitophagy receptor (Dorn, 2019; Wang et al., 2023). Non‐canonical mitophagy pathways include FUNDC1 and BNIP3L. Both are mitophagy receptors. FUNDC1 and BNIP3L are typically recruited during hypoxia (Chen et al., 2016; Li, Li et al., 2021; Li, Zheng et al., 2021; Marinković & Novak, 2021). Once a mitophagy receptor is engaged on the outer mitochondrial membrane, it recruits LC3 to initiate autophagosome formation. LC3 exists initially in its cytosolic, non‐lipidated form known as LC3‐I. Upon activation, LC3‐I undergoes conjugation with phosphatidylethanolamine (PE), generating the lipidated form LC3‐II, which associates with the autophagosomal membrane (Kawajiri et al., 2010). This lipidation process requires a cascade of ATGs, including ATG7 (E1‐like enzyme), ATG3 (E2‐like enzyme) and the ATG12–ATG5–ATG16 complex, which functions analogously to an E3 ligase (Tanida et al., 2008). After lipidation, LC3‐II binds directly to mitophagy receptors through LC3‐interacting regions (LIRs), facilitating expansion and closure of the autophagosome. The mature autophagosome subsequently fuses with a lysosome to form an autolysosome, enabling the degradation of the engulfed mitochondrion.

FXNKD hearts showed no significant alterations in LC3II/I ratio, suggesting preserved mitophagy. In contrast, FXN‐cKO hearts exhibited elevated upstream mitophagy markers but a decreased LC3‐II/I ratio, with the accumulation of inactive LC3‐I and reduced LC3‐II. Decreased ATG7 and ATG12–5 further indicate impaired LC3 processing and stalled autophagosome formation. These findings suggest that mitophagy initiation may occur in FXN‐cKO hearts, but completion is defective, resulting in the accumulation of damaged mitochondria.

Fibrosis, inflammation and arrhythmogenesis

Excessive mitochondrial fragmentation, ROS production and defective clearance can drive fibroblast activation and fibrosis (Salinas et al., 2025). This along with an apparent halting of mitophagy is likely contributing to the fibrosis formation in the heart. DRP1‐mediated fission has been linked to fibroblast activation and collagen synthesis in the heart (Lin et al., 2023). Impaired mitophagy may also activate inflammatory pathways, including NLRP3‐mediated IL‐18 signalling, which contributes to fibrotic remodelling (Cui et al., 2024; Lin et al., 2023).

Although FXNKD hearts demonstrated less dramatic mitochondrial remodelling, both models ultimately developed fibrosis, likely via chronic inflammatory signalling (Chandran et al., 2017). Fibrotic remodelling disrupts electrical conduction and contractile mechanics and is a major contributor to lethal arrhythmias in FA patients (Hanson et al., 2019). These arrhythmias arise from impaired electrical propagation secondary to structural remodelling and fibrosis, rather than solely from autonomic dysfunction.

Arrhythmias in FA are due to alterations in the autonomic nervous system and more often, due to cardiac remodelling and fibrosis that inhibit those electrical propagations (Figueroa et al., 2025). Restoring mitochondrial quality control by limiting excessive fission, enhancing fusion and promoting effective mitophagy represents a promising therapeutic strategy to reduce fibrosis, preserve electrical stability and improve cardiac outcomes in FA (Lin et al., 2023).

Limitations

Some of the findings in FXN‐cKO hearts differ from those reported in a prior study (Huang et al., 2013), which examined the same mouse model at 3, 6 and 10 weeks of age, whereas our analyses were performed specifically at 8 weeks. Discrepancies in LC3, Parkin and MFN2 expression may therefore reflect age‐dependent differences in mitochondrial remodelling and disease progression. Additionally, mitochondrial fission–fusion protein expression in our study was assessed in mitochondrial fractions, whereas other markers were measured in whole‐heart lysates, which may influence relative protein detection and quantification. Importantly, our upstream assessment of ATGs in the lipidation pathway supports the interpretation that mitophagy is impaired at the autophagosome formation step, providing mechanistic alignment despite methodological differences. Nonetheless, caution should be used when comparing studies with differing sample preparation approaches and time points, as mitochondrial dynamics are highly context dependent and evolve over disease trajectory. Additionally, the different mouse models are assessed at different ages. It has been shown that mitochondrial dynamics can be alerted in aged mice. Although mice in our different mouse models are of different ages, we believe that their age difference is quite modest, at 4 months, and age‐matched controls are used to directly assess what is occurring in healthy versus disease state.

Conclusion

FXN‐cKO hearts with HFrEF exhibit profound disruption of mitochondrial dynamics, characterized by excessive fission and impaired mitophagy resulting from defective autophagosome maturation. This combination likely promotes the accumulation of fragmented, dysfunctional mitochondria and contributes to energetic collapse in systolic heart failure. In contrast, FXNKD hearts with HFpEF‐like phenotype demonstrate relatively preserved mitochondrial turnover, with only modest morphological changes, including slightly enlarged and more spherical mitochondria, consistent with a milder remodelling phenotype and maintained mitochondrial quality control. These divergent alterations in mitochondrial structure and clearance pathways suggest fundamentally different energetic states and stress adaptations between the two FA cardiac mouse models. In FXN‐cKO hearts, severe FXN loss appears to overwhelm quality‐control systems, whereas partial FXN deficiency in FXNKD hearts may allow compensatory mechanisms to preserve mitochondrial homoeostasis.

To determine whether these mechanisms translate to human disease, future studies should incorporate myocardial biopsy and post mortem analyses from FA patients to directly evaluate mitochondrial morphology, mitophagy markers and respiratory function across HFpEF and HFrEF phenotypes. Establishing these mechanistic distinctions in patients could inform the development of targeted therapeutic strategies and enable precision‐based interventions tailored to the specific mitochondrial defects underlying each cardiac presentation of FA.

Additional information

Competing interests

The authors declare no conflicts of interest, financial or otherwise.

Author contributions

E.N.D. and L.S.: conceptualization, writing – original draft and methodology. E.N.D., G.P.C. and L.S.: formal analysis. L.S., F.F., C.B.M. and G.P.C.: investigation. E.N.D., L.S., F.F., C.B.M. and G.C.: writing – review and editing. E.N.D.: supervision.

Funding

This project was supported by the National Institute of Health 1R01HL155907‐1 (to E.N.D.), FARA Research Award (to E.N.D.), American Heart Association Transformational Research Project 25TPA1478903 (to E.N.D.), University of California Davis CRCF Pilot & Feasibility Award 181031 (to E.N.D.), University of California Innovative Development Award (to E.N.D.), predoctoral fellowship from NIH R01HL155907‐02S1 Diversity Supplement and postdoctoral fellowship from NIH T32 HL007013 in Comparative Lung Biology and Medicine Training Grant (to L.S.), predoctoral fellowship from NIH T32 HL086350 Training Grant in Basic & Translational Cardiovascular Science and FARA predoctoral fellowship (to F.F.).

Generative AI statement

During the preparation of this manuscript, Grammarly was used to assist with grammar and language editing. All scientific content, interpretations and conclusions were generated, reviewed and approved by the authors.

Supporting information

Peer Review History

TJP-604-8360-s001.pdf (426.2KB, pdf)

Acknowledgements

We thank Dr. Nipavan Chiamvimonvat and the University of California, Davis, Department of Cardiology, for providing access to and general use of their echocardiography facility.

Biography

Lili Salinas is a postdoctoral researcher in the laboratory of Dr. Elena N. Dedkova at the University of California, Davis. She earned her bachelor's degree in chemistry from Willamette University, where she first developed a passion for biomedical research. Lili subsequently completed her PhD at the University of California, Davis, under the mentorship of Dr. Dedkova. Her research focuses on the molecular mechanisms underlying cardiac dysfunction in Friedreich's ataxia, a devastating inherited neurodegenerative disorder in which cardiomyopathy is the leading cause of mortality. Lili is particularly passionate about advancing research on rare metabolic and mitochondrial disorders, with the goal of developing new therapeutic strategies for patients.

graphic file with name TJP-604-8360-g002.gif

Handling Editors: Bjorn Knollmann & T Alexander Quinn

The peer review history is available in the Supporting Information section of this article (https://doi.org/10.1113/JP291356#support‐information‐section).

Data availability statement

The data underlying this article will be shared upon reasonable request to the corresponding author.

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