Resilience of the Mitochondrial Reticulum in Aging

Abstract

Resting and maximal exercise respiratory rates (${\dot{\mathrm{V}}\mathrm{O}}_2$) decline in aging. Those losses have been attributed to impaired mitochondrial function, but the data are inconsistent with healthy aging. To interrogate the hypothesis of mitochondrial dysregulation in aging, we studied hind limb skeletal muscles from young and older, male and female, NIA C57BL/6JN mice. We observed no age-associated changes in coupling efficiency (ADP:O) of mitochondrial reticulum preparations, but respiratory control (RCR) was decreased in older mice. Additionally, older skeletal muscle exhibited subtle yet significant reductions in the expression of proteins functionally related to substrate uptake and oxidation (mMCT1, mPC1, CPT1b, HADH). While there were no differences in mitochondrial contents per mg of muscle in older mice, there were significant losses of muscle, and hence mitochondrial mass as well as proteins associated with membrane dynamics (DRP1, FIS1, and MFN2). Further, 2D and 3D, cross- and longitudinal muscle sections showed alterations in mitochondrial reticulum organization in muscles of older mice. Therefore, aging is associated with subtle, but significant changes in the organization and functioning of muscle mitochondrial reticula.

NEW & NOTEWORTHY

We interrogated numerous structural and functional aspects of the mitochondrial reticulum using a standard mouse model of aging. We observed no age-associated changes in coupling efficiency of mitochondrial preparations, but respiratory control decreased and there were numerous subtle changes in mitochondrial morphology in aging mouse muscles. Overall mitochondrial functioning is well preserved in aging indicating the performance decrements are related to loss of muscle mass and cardiovascular function.

INTRODUCTION

While the relationship between muscle wasting and incidences of lifestyle diseases is strong in aging (1), controversy exists concerning the primary drivers of sarcopenia and mitochondrial atrophy. Notable comorbidities in aging include Type II Diabetes (2), cardiovascular disease (3, 4), renal failure (5, 6), and neurodegeneration. Regardless of age- or disease-related alterations in metabolism, it is reasonable to hypothesize that changes in metabolism are rooted in structural and functional alterations in the muscle mitochondrial reticulum (7).

As the major site of energy substrate disposal, the skeletal muscle mitochondrial reticulum serves as a primary determinant of body carbon energy flux. Often misrepresented as “bean-shaped” organelles, the cellular respiratory apparatus should instead be viewed as a complex and intertwined network dynamically undergoing fusion and fission events, which in turn preserve ATP homeostasis, redox balance, apoptotic signaling, calcium regulation and other aspects of mitochondrial function (8–11). Accordingly, the mitochondrial reticulum has been characterized as the intracellular energy highway (8), or power grid (12). Regulation of the mitochondrial reticulum, its structure and function, is necessary to preserve ATP homeostasis and cell function (13–16).

Classic and contemporary models for the decline of the cellular respiratory apparatus in aging include the free radical theory (17–20), dysregulated mitophagy (21, 22), perturbed ATP kinetics (23, 24), and decline in tissue mitochondrial content (25). Considering the importance of cellular ATP homeostasis, reduced ATP production capacity has been observed in the skeletal muscles of older animals (26–28). In contrast, others have found preserved efficiency of oxidative phosphorylation (23, 29). Further, there are inconsistent findings regarding changes in the transport, uptake, and utilization of energy substrates in aging (30–33). The same can be said for mitochondrial content, with data indicating either decreased (34), or similar mitochondrial densities in aging (35, 36). Understandably, discrepancies in the aging literature are subject to multiple factors, including the species investigated, samples utilized (i.e., whole muscle homogenate vs. isolated mitochondria preparations), instrumentation, isolation protocols, presence of comorbidities, and age categorization.

In the context of mitochondrial membrane remodeling, researchers have postulated that perturbations in synchronization of fusion and fission processes result in metabolic inadequacies, ultimately leading to a decline in muscle mass and functionality. Optic Atrophy 1 (OPA1), Mitofusin 1 (MFN1), and Mitofusin 2 (MFN2) control mitochondrial membrane fusion (37). Studies examining these proteins in relation to age have yielded mixed results, with some demonstrating no changes (38) while others indicate increased levels of expression (39). Responses of these proteins are believed to vary depending on the type of muscle fiber (40). In contrast, studies on fission-related events are associated with elevated mitochondrial membrane fragmentation. Fission is primarily regulated by Dynamin-Related Protein 1 (DRP1) and Fission 1 (FIS1) (41). Considering its role in the control of mitochondrial biogenesis, other investigators have proposed that mitochondrial membrane remodeling is under selective control of PGC1α. In contrast, others have postulated that mitochondrial membrane trafficking is a limiting factor in the translocation of dynamic proteins in aging (42). Importantly, exercise training has been successfully utilized as an intervention to restore age-associated deficits to mitochondrial dynamics (43–46). Those studies demonstrated partial or complete recovery of mitochondrial protein abundances in aged muscles post-training. Most recently, mitochondrial fragmentation has been associated with loss of physical capacity in aging (47). overall, those experiments reassert what has previously been observed in classic experiments linking mitochondrial biogenesis to endurance training (23, 48, 49).

In the context of the myriad of reductionist studies of aging effects on mitochondrial functioning are results of experiments on muscular efficiency in healthy young and older, exercise-trained and untrained, men and women that showed no age-related effects on the efficiency of energy coupling (50). To resolve differences in the interpretation of discordant results obtained at different levels of physiological organization, we studied young and older NIA C57Bl/6JN mice using classic and contemporary techniques to interrogate the hypothesis that aging results in disorganization and functioning of the muscle mitochondrial reticulum. Overall results show resilience of mitochondrial structure and function in aging; nonetheless many subtle, but significant age-related mitochondrial effects were found.

MATERIAL AND METHODS

Experimental model and animal characteristics

Experimental procedures involving animals were approved prior to experimentation by the Institutional Animal Care and Use Committee of The University of California, Berkeley (UCB) (AUP-2018–07-11294–2). C57Bl/6JN Mice were bred and aged at the National Institute of Aging (NIA) and were transferred to the University of California, where they were allowed to acclimatize for 1 week prior to any experimentation. Cages were maintained at a constant temperature and humidity with a 12-hour light-dark cycle (Light: 7:00 am to 7:00 pm). Young mice were 3–5 months, and old mice were 21–24 months old. Animals were grouped according to sex and age as follows: Young Male Mice (YM), Young Female Mice (YF), Old Male Mice (OM), and Old Female Mice (OF). The animals were primarily euthanized via CO₂ inhalation and secondarily via cervical dislocation, in accordance with the University Animal Use Committee. Following euthanasia, the mice were weighed and mixed muscles (MM) (gastrocnemius and quadriceps) from the hind limbs were isolated from both legs. One set of muscles was immediately placed in an isolation buffer (IB) to be respired the same day, while the other set of MM was flash frozen and stored at −80°C for future biochemical analysis.

Muscle Mitochondrial Isolation

As previously described, mixed muscle (MM) from a total of 8 mice per group was isolated from the hind limbs (gastrocnemius and quadriceps) (48). Briefly, MM was placed into ice-cold (IB: 250 mM mannitol, 10 mM EDTA, 45 mM Tris-HCI, 5 mM Tris(hydroxymethyl)aminomethane, and 0.15% protease inhibitor, pH 7.4). Proteases were not used in this study. To obtain a muscle homogenate (MU), muscles were weighed, minced in a 1:10 dilution of fresh IB, and homogenized using a Potter-Elvehjem Teflon pestle on glass homogenizer. Mitochondrial (MI) fragments were isolated from the muscle homogenate by differential centrifugation at 4°C. The homogenate was centrifuged at 600 × g for 10 min to pellet debris and nuclei. The supernatant was then transferred and centrifuged at 10,000 × g for 10 min to pellet the mitochondrial fraction. Mitochondrial fragments were resuspended in 150 μL of IB and kept on ice until respiratory measurements. Mitochondrial protein concentration was determined using a BCA kit with bovine serum albumin (BSA) as a standard (Pierce Biotechnology, Radford, IL).

Mitochondrial Preparation Respiration

Mitochondrial oxygen (O₂) consumption was measured using a dual-chambered Clark-Type Oxygen Electrode (Rank Bros., Cambridge, UK), and data were collected using a High-Resolution Data Logger (Picolog Systems). All measurements were performed at a constant temperature of 25°C instead of 37°C to avoid the possibility of loss, respiratory control, or partial uncoupling, as previously observed (51). Prior to data collection, the respiration chamber was filled with 1 mL of respiration medium (RM) that contained (15 mM KCI, 15 mM K₂HPO₄, 15 mM KH₂PO₄, 25 mM Tris base, 45 mM sucrose, 12 mM mannitol, 5 mM MgCI₂, 7 mM EDTA, 0.2% BSA, and 20 mM glucose at pH 7.4) and continuously stirred with a magnetic stir bar. After the RM was allowed to equilibrate with the environmental conditions, 100 μg of the mitochondrial preparation was added to both chambers. The substrates utilized in separate chambers included 10 mM of pyruvate + 2.5 mM of malate to simulate carbohydrate-derived respiration and, in the other chamber, 40 μM of palmitoyl-L-carnitine + 1 mM of malate to simulate fatty acid-derived respiration. ADP was added at a final concentration of 100 μM to initiate state 3 (ADP-stimulated) respiration (S3), and state 4 (S4) respiration was defined as the rate of O₂ consumption in the absence of ADP phosphorylation. Oxygen consumption was measured in nmol O₂·μg·mitochondria-1·min-1. The respiratory control ratios (RCR) S3/S4, ADP/O, and solubility factor of O₂ using a factor of 240 nmol O₂·mL⁻¹ in the chamber were calculated as described previously (52).

Immunoblotting

The second aliquot of MM from a total of 12 mice, 3 per group, was processed in the same manner as previously described (53–56). Briefly, the tissue was partitioned so that a portion of the sample could be used for immunoblotting and another for enzymatic kinetics. After the first 600 × g centrifugation, a fraction of the supernatant was aliquoted to represent the whole muscle (MU) fraction. The remaining supernatant of the MU was centrifuged at 10,000 × g, and the supernatant [(Cytosolic) (C)] from this step was removed and stored for immunoblotting. The remaining mitochondrial protein pellet (MI) was resuspended in 200 μL RIPA buffer and mixed on ice for 15 min. Protein concentration was determined using a BCA protein assay kit (Pierce Biotechnology, Rockford, IL, USA). Samples were subsequently flash-frozen until further analysis.

Western blotting was performed as previously described (53–55) utilizing 20 μg of isolated mitochondria. Samples were diluted in 10x LDS Sample Buffer, 4x Sample Reducing Agent, distilled and deionized (dd) H₂O, and incubated for 10 min at either 50 or 70°C based on the protein target. Samples were loaded onto 4–12% BT gels containing a molecular weight standard (Invitrogen), separated at 150 V for 45 min, and transferred to a low-fluorescent PVDF membrane (Azure). The membrane was then probed using a Total Protein Stain (TPS) solution (Licor) to ensure equal loading of the proteins. The membrane was blocked using commercial TTBS Blocking Buffer (Licor) for 1 hr at room temperature and incubated overnight at 4°C with a primary antibody diluted in blocking buffer + 0.02% Tween. The next day, the membrane was washed 3 times for 10 min using TBS+0.02% Tween, then incubated with a secondary antibody Licor IR 800 in TBS blocking buffer + 0.02% Tween for 1 hr at room temperature. The membrane was washed 3 times for 10 min using TBS+0.02% tween and 1 time with 1XTBS. Finally, membranes were imaged on a NIR Azure 500C system prior to densitometry analysis using the NIH Image J software. A list of all the antibodies used can be found in a supplemental file (see Supplemental Table S1, available at Figshare: [10.6084/m9.figshare.29669213]).

Enzyme Activity Assays

Enzyme activities of both MI and MU fractions from an n = 5 mice in each group included Citrate Synthase (CS), Lactate Dehydrogenase (LDH), 3-hydroxyacyl-CoA dehydrogenase (HADH), and Cytochrome Oxidase (COx). Samples were prepared similarly to respirometry experiments but were further resuspended using buffers based on previous investigations or supplied by the company. CS activity was assayed by utilizing a kit from Sigma-Aldrich (MAK193), which utilizes GSH as a standard and measures the appearance of thionitorbenzoate anion (TNB) in the sample at an absorbance of 412 nm. COx activity was assayed by a kit from Abcam (ab109911) to measure the oxidation rate of reduced Cytochrome c in the sample at an absorbance of 550 nm. For both CS and COx activities, 10 μg of samples were loaded into each well of a Molecular Devices SPECTRAMAX 340 spectrophotometer plate reader.

Lactate Dehydrogenase activity was measured in 250 μL aliquots instead of 1 mL, as described previously (56). Briefly, 10 μg of MI or MU were diluted in 50 mM potassium phosphate (pH 7.0) +1 mM of EDTA. Prior to measuring enzymatic activity, 100 μM of NADH and 2.1 mM of pyruvate were rapidly added to start the reaction. The decrease in absorbance at 340 nm was followed for a minimum of 10 min, allowing adequate time to calculate the highest amount of activity in each fraction and animal group.

3-hydroxyacyl-CoA dehydrogenase (HADH) activity was adapted to a microplate-based assay according to (57). Briefly, assay buffer (50 mM Tris-HCl buffer (pH 7.0), 250 μM NADH, 2 mM EDTA, and 0.2% Triton X-100 mixed in dd H2O) was warmed to 30°C prior to filling the spectrophotometer wells. At the same time, a 20 μg aliquot of either MI or MU was individually added into each well, followed by 186 μL of warmed assay buffer, and finally, using a multichannel pipette, 4 μL of 5 mM acetoacetyl-CoA to initiate the reaction. The plate was immediately placed in a spectrophotometer and read every 15s for a minimum of 4 min at 340 nm to measure the rate of NADH disappearance. Enzymatic activity was calculated according to (57) and values were normalized to protein concentration and reported as nmol·mg-1·min-1 for MU and nmol·μg-1·min-1 for MI, allowing us to calculate mitochondrial content, as previously described (48).

Pyruvate Dehydrogenase (PDH) activity was only assayed in the MI fraction according to the manufacturer’s instructions (Sigma-Aldrich MAK 183).

Immunohistochemistry, Confocal Laser Scanning Microscopy, and Three-Dimensional Reconstruction

Samples were freshly collected from the gastrocnemius washed in 1X PBS and fixed overnight in 4% paraformaldehyde before long-term storage in 70% EtOH. Tissues were then washed in 1X PBS, followed by 15% and 30% sucrose gradient incubations. Immediately following sucrose preservation tissues were placed in OCT and frozen utilizing chilled isopentane. Tissue blocks were then allowed to equilibrate in the cryostat at −20°C prior to collecting 10 um sections, which were directly placed onto positively charged microscope slides. Following this, sections were washed with 1XPBS, permeabilized with 0.5% Triton in PBS, and antigens were retrieved in 10 mM citrate buffer (pH 6.0) for 10 min. Sections were then blocked using a commercial high background blocking buffer (ThermoFisher, 00–4952-54), washed with PBS, and incubated overnight with 1:50 TOMM20-AF488 nm. The slides were again washed, mounted onto charged microscope slides (Fisher Scientific 1358A) using Vectashield Plus Antifade Mounting Medium with DAPI (Vector Laboratories, H1200), and sealed. Confocal Laser Scanning Microscopy (Zeiss 880) was utilized for immunofluorescent detection of TOMM20 (488 nm) and DAPI (405 nm). The exposure conditions were optimized to visualize the mitochondrial network using an objective of 100X. Detection wavelengths remained the same for all the groups. The digital gain was individually adjusted to avoid pixel oversaturation. Z-stacks of 16 um were taken in steps of 0.40 um with a resolution of 372×372 pixels with twofold line averaging. Mitochondrial network analysis for 2D images was conducted on 3–4 muscle fibers from 5 mice. For Two-dimensional images were analyzed utilizing the mitochondrial analyzer plugin of ImageJ to optimize the background and threshold conditions for each section, followed by an automated analysis to calculate the Mitochondrial Form Factor (MFF), Count, Branch Count, Average Branch Length, Average Branch Diameter, and Branch Area. The Imaris software was utilized for three-dimensional modeling. For 3D imaging, for each plane we examined images from 3 mice in each group. Surfaces were reconstructed utilizing the smooth function and were built with the machine learning function to generate voxels with a lower limit of approximately 0.5 micrometers and an upper limit of approximately 1.2 micrometers for the mitochondrial networks. The channel utilized for 3D modeling was reconstructed using a built-in automated background subtraction function included in Imaris software. Similar procedures were conducted for the transverse sections with the exception that TOMM20 conjugated to Alexa Fluorescence 488 nm (AF 488 nm) (1:200) was utilized to stain the mitochondrial networks. Hematoxylin and eosin (H&E) sections were fixed overnight in 4% PFA and prepared by the University of California, San Francisco Gladstone Institutes. Images were acquired using a Zeiss Axio Z1 Slide Scanner at 20x magnification. Quantification analyses were conducted using the Zen Blue software by outlining each muscle fiber with the contour assist tool. Data are presented as mean ± SD.

Statistical Analyses

Statistical significance was assessed using an ordinary two-way ANOVA for (Age × Sex), with Bonferroni correction applied for multiple comparisons to control the type I error rate, ensuring an appropriate balance between avoiding false positives and maintaining power. Post hoc comparisons were conducted where the main effects or interactions were significant. All statistical analysis was performed using GraphPad Prism version 10.0 for Windows. Data are expressed as mean ± SD, and significance was determined at (p < 0.05).

RESULTS

Coupling Efficiency of Mitochondrial Preparations from Older Mice is Preserved Despite Reductions in the Respiratory Control Ratio

Physical characteristics of the young (4 months) and older (24 months) C57Bl/6JN mice that were studied are displayed in (Table 1). Not surprisingly, older mice were significantly heavier than the younger mice F (1, 16) = 88.17, p < 0.0001) and possessed relatively less muscle mass relative to total body mass F (1, 16) = 15.69, p = 0.0011). In addition, muscle fiber diameter, analyzed using transverse H&E preparations, indicated that fibers in older mice were significantly smaller when normalized to total muscle area F (1, 28) = 6.50, p ≤ 0.05) (Figure 1), which was also demonstrated by muscle mass weighed prior to respirometry assays F (1, 16) = 29.81, p < 0.0001).

Table 1.

Characteristics of the old and young mice utilized for the study.

Sex (n)	Male (n=8)	Female (n=8)	Male (n=8)	Female (n=8)
Age (Months)	4 ± 1	4 ± 1	24 ± 1*	24 ± 1*
Body Weight (g)	26.8 ± 1.8	21.6 ± 1.9	31.2 ± 1.1**	27.4 ± 0.9*
Muscle Weight (mg)	393.8 ± 97.9	344.1 ± 42.5	273.8 ± 67.7**	232.8 ± 32.6*
mg muscle g body weight− 1	15.4 ± 2.2	16.7 ± 2.9	9.4 ± 2.1**	8.6 ± 1.3***

Data are reported as mean ± SD and reporting changes that are associated with aging. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction for multiple comparisons. Significance was determined at p < 0.05 (*p < 0.05, **p < 0.01, ***p < 0.001).

Figure 1. H&E Transverse Sections of mouse skeletal muscle indicating smaller fibers in aged muscle.

(A) Transverse muscle sections (20x magnification) utilizing haematoxylin and eosin (H&E) in young (n = 200 fibers, 100M and 100F) and old (n = 200 fibers, 100M and 100F) from n=5 mice in each of the 4 groups. (B) Illustration of reductions in the average muscle fiber diameter of older mouse skeletal muscles. (C) Age-associated decline in cross-sectional area (CSA) of the entire muscle normalized to the total muscle fiber diameter in each group. Data are presented as mean ± SD. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction for multiple comparisons. Significance was determined at p < 0.05, (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001). The scale bar is 20 μm. The units used to analyze the distance and length of the images were reported in micrometers. Images were collected utilizing the Zeiss Axio Scan. Z1 Slide Scanner. Quantification analyses were conducted using the Zen software, and the contour assist tool was used to outline each muscle fiber. = Males = Females.

To evaluate the effects of aging on energy substrate oxidation, we measured rates of O₂ consumption in mitochondrial preparations using carbohydrate- and fatty acid-derived substrates. Regardless of the substrate used, preparations from older mice exhibited significantly reduced State 3 (S3) respiration and elevated State 4 (S4) respiration compared to young mice. With pyruvate and malate (P+M), S3 respiration was significantly lower (F (1, 28) = 17.23, p = 0.0003), while S4 respiration was higher (F (1, 28) = 10.99, p = 0.0025) (Figure 2a, b). Similarly, with palmitoylcarnitine and malate (Pc+M), S3 respiration was markedly reduced (F (1, 28) = 35.77, p < 0.0001), and S4 respiration was elevated (F (1, 28) = 17.37, p = 0.0003) (Figure 2e, f). Noteworthy was that incubation with either substrate resulted in significantly lower respiratory control ratios (RCR = S3/S4) in mitochondrial preparations from older compared to those from young mice when using P+M (F (1, 28) = 31.60, p < 0.0001) (Figure 2d) and Pc+M (F (1, 28) = 26.92, p < 0.0001) (Figure 2h). These results indicate a mild loosening of mitochondrial respiratory control with age. In contrast, ADP:O ratios (coupling efficiencies) remained consistent between young and older mice, regardless of the substrate used, showing no significant effect of Age for P+M (F (1, 28) = 0.082, p = 0.777) and Pc+M (F (1, 28) = 1.201, p = 0.281) (Figure 2c, g).

Figure 2. Respirometry data depicting preserved mitochondrial efficiency (ADP/O) and simultaneous loss of membrane integrity (RCR).

Oxygen consumption in isolated mitochondria from skeletal muscle in 4-month-old (Young) (n = 16, 8 male and 8 female) and 24-month-old (Old) (n = 16, 8 male and 8 female) mice displaying (A) State 3 pyruvate + malate (P+M) derived respiration and (B) State 4 pyruvate + malate (P+M) derived respiration. (C) Ratios of ADP to oxygen consumption (ADP:O) illustrating preserved mitochondrial efficiency when utilizing (P+M) (D) Respiratory Control Ratios (RCR) utilizing (P+M) highlighting loss of mitochondrial membrane integrity following state 3 and state 4 respiration. (E) State 3 pyruvate + malate palmitoyl carnitine + malate (PC+M) and (F) State 4 palmitoyl carnitine + malate (PC+M). (G) (ADP:O) again illustrates preserved mitochondrial efficiency when utilizing (PC+M) (D) Respiratory Control Ratios (RCR) utilizing (PC+M), highlighting the loss of mitochondrial membrane integrity following the addition of (PC + M). Data are presented as mean ± SD. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction for multiple comparisons. Significance was determined at p < 0.05. Results indicated a significant reduction in respiration rates in aged mitochondria compared to young (*p < 0.05, **p < 0.01, ***p < 0.001). = Males = Females.

Altered Mitochondrial Protein Profiles Linked to Fuel Uptake and Oxidation

Following measurements of O₂ consumption, we assessed abundance of proteins that affect the uptake, reduction, and subsequent oxidation of fatty acid- and carbohydrate-derived energy substrates. In isolated mitochondrial fragments per unit mitochondrial remnant protein, there were no differences in the relative abundances of citrate synthase (CS) F (1, 8) = 0.1674, p = 0.6932) or cytochrome oxidase (COx) F (1, 8) = 0.7026, p = 0.4262) (Figure 3a, b). Moreover, other than a small yet, significantly lower abundance of CPT2 in old compared to young females, as a group, the abundance of proteins utilized for fatty acid transport did not differ between young and older preparations (Figure 3d, e). However, our analyses observed significantly reduced levels of 3-hydroxyacyl-CoA dehydrogenase (HADH) in older mice, F (1, 8) = 38.34, p = 0.0001) (Figure 3c).

Figure 3. Mitochondrial Transporter and Enzyme Expression Decline with Age.

Immunoblots, normalized to total protein stain (TPS), of key mitochondrial transporters utilizing MI from young (n = 6, 3M and 3F) and old (n = 6, 3M and 3F) mice. Protein expression of (A) Citrate Synthase (CS) and (B) Cytochrome Oxidase (COx) did not decline with age indicating preserved mitochondrial content. Lower abundances of (C) 3-hydroxyacyl-CoA dehydrogenase (HADH)and (D) the mitochondrial Pyruvate Carrier 1 (mPC1) indicate age-associated loss of substrate handling. At the same time there were no changes to either (e) Carnitine palmitoyltransferase 1b (CPT1b) abundances for males or (F) overall Carnitine palmitoyltransferase 2 (CPT2) suggesting a small but significant limitation to fatty acid transport in MI from female mice. Data are expressed as mean ± SD. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction applied. Significance was determined at p < 0.05. Transporter levels were significantly reduced in aged tissues compared to young (*p < 0.05, **p < 0.01). = Males = Females. For full immunoblots see Supplemental Figure S1, available at Figshare: [10.6084/m9.figshare.29669213.

Following our analysis of the proteins involved in β-oxidation, we analyzed the limitations of carbohydrate-derived transport. Aging had a small, but significant effect on the abundance of the mitochondrial pyruvate carrier (mPC1), which showed a significant effect of age F (1, 8) = 63.93, p < 0.001 (Figure 3f). As well, there was a complementary decline in abundance of mitochondrial lactate transporters (mMCT1) (F(1,8) = 61.47, p < 0.001) (Figure 3g) (56).

In view of reductions in the relative abundances of enzymes involved in energy substrate uptake and use, we assayed the activities of key enzymes for energy substrate disposal.

Similar Enzymatic Activities in Mitochondrial Preparations of Young and Older Mice

To test age-related effects on the activities of selected mitochondrial enzymes, we analyzed muscle (MU) homogenates, and mitochondrial (MI) preparations derived from the same animals. There were no effects of age on the rates of cytochrome oxidase (COx) activities in either MU or MI preparations; F (1, 16) = 0.009974, p= 0.9217; F (1, 16) = 0.002992, p = 0.9571) (Figure 4a, d). Similarly, there were no significant differences in citrate synthase (CS) activities in either MU or MI fractions; (F (1, 16) = 2.659, p = 0.1225; F (1, 16) = 0.8064, p = 0.3825) (Figure 4b, e). For HADH activity, no differences were observed between young and older mice in either MU or MI fractions; F (1, 16) = 1.396, p = 0.2546; F (1, 16) = 0.1582, p = 0.6961) (Figure 4c, f). However, the rates of lactate dehydrogenase (LDH) activity were significantly lower in both MU and MI fractions of older mice; (F (1, 16) = 10.66, p = 0.0049; F (1, 16) = 38.69, p < 0.0001) Figure 4g, h). In addition, measurements of PDH activity assayed on MI preparations were also significantly lower in older mice F (1, 16) = 30.98, p < 0.0001) (Figure 4i). Given that there were similar changes to enzymatic rates of LDH and PDH, the correlative strength between both data sets were assessed and found to be significantly related (n = 20, r = 0.8814, p<0.0001) (Figure 5). This was not surprising considering we had previously determined that, compared to young individuals, older participants exhibited delayed and diminished post-absorptive and postprandial capacities to oxidize lactate (Arevalo. 2024). Having assessed most enzymatic activity rates in both the MU and MI fractions, we also quantified mitochondrial content by normalizing MI enzymatic rates to those of the MU fraction. These ratios indicated similar mitochondrial content in older and young skeletal muscle (Figure 6 a–d). Finally, correlations between enzymatic activities showed that the classic markers of mitochondrial content (CS and COx) were strongly associated (Figure 6 e). In comparison, the weaker correlation between LDH and other enzyme activities may reflect the dynamic role of LDH in both mitochondrial and cytosolic compartments, making it a less reliable marker of mitochondrial content in aging muscle.

Figure 4. Enzyme Activity in Aging Mitochondria is limited upstream of the Electron Transport Chain.

Specific activity of mitochondrial enzymes involved in the TCA cycle and ETC in MI and whole Muscle Homogenates (MU) from young (n = 10, 5M and 5F) and old (n = 10, 5M and 5F) mice. Aging had no effect on skeletal muscle COx activity in (A) MU or (D) MI fractions. Enzymatic activity of CS in (B) MU and (E) MI fractions illustrating no differences. Enzymatic analysis of HADH in (C) Mu and (F) MI fractions, again displaying no age-associated declines. With the exception for the female MU fractions, Lactate Dehydrogenase (LDH) enzymatic analysis revealed impaired rates of activity in (G) Male MU and (H) in both sexes for MI fractions and Pyruvate Dehydrogenase activity in (I) in both sexes for MI fraction. Data are presented as mean ± SD. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction for multiple comparisons. Significance was determined at p < 0.05. In addition to LDH kinetics, enzyme activity was similar between old and young mice (*p < 0.05, **p < 0.01, ***p < 0.001). = Males = Females.

Figure 5. Positive Correlation between LDH and PDH Enzyme Activity in Young and Older Mitochondria.

Pearson r was assessed on activity of mLDH and PDH in MI from young (n = 10, 5M and 5F) and old (n = 10, 5M and 5F) mice. Significance was determined at p < 0.05. (*p < 0.05, **p < 0.01, ***p < 0.001). = Young = Old.

Figure 6. Mitochondrial Content does not decline with age in mixed muscles from mice.

The ratios of mitochondrial (MI) and muscle (MU) enzymatic activities resulted in similar mitochondrial content when analyzing (A) COx (B) CS (C) HADH and (D) LDH. (E) Correlation plot comparing enzymatic activities to one another. Data are expressed as mean ± SD. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction applied. Significance was determined at p < 0.05. Results indicated a significant reduction in mitochondrial content in aged tissues compared to young (*p < 0.05, **p < 0.01). = Males = Females.

Diminished Abundance of Proteins Affecting Mitochondrial Dynamics in Preparations from Older Mice

Following measurements of mitochondrial content and function, we examined proteins associated with mitochondrial dynamics. In older mice, immunoblot analysis revealed a significant loss of proteins responsible for maintaining mitochondrial membrane integrity. Considering the proteins associated with mitochondrial fusion, there were no significant changes in abundances of MFN2 (F (1, 8) = 0.2069, p = 0.6613), OPA1 (F (1, 8) = 4.163, p = 0.1279), or phosphorylated (DRP1) SER637 (F(1, 8) = 8.945, p = 0.1732) (Figure 7 b, d, f). In contrast, there were significantly lower MFN1 abundances (F (1, 8) = 50.25, p < 0.0001) (Figure 7 a), indicating impaired mitochondrial fusion. Analogous to findings on MFN1, we observed significantly lower abundances of proteins that regulate mitochondrial membrane fission, specifically DRP1 (F (1, 8) = 66.46, p < 0.001), and FIS1 (F (1, 8) = 69.50, p < 0.0001) in preparations from older mice (Figure 7 c, e, f). Having found age-associated reductions to the protein abundances responsible for maintaining the dynamic nature of the mitochondrial reticulum logically, we moved on to visualizing the networks utilizing 2- and 3- dimensional reconstructions.

Figure 7. Mitochondrial Dynamic Proteins decline with age.

(A) Western blot images, normalized to total protein stain (TPS), in young (n = 6, 3M and 3F) and old (n = 6, 3M and 3F) mice showing lower levels of Mitofusin1 (MFN1) protein and similar amounts of (B) Mitofusin 2 (MFN2) and (D) Optic-Atrophy1 (OPA1), the proteins responsible for mitochondrial membrane fusion. Alternatively, quantification of membrane protein expression levels revealed significantly lower amounts of (C) Fission1 (FIS1) and (E) Dynamin-Related Protein1(DRP1). (F) Aging was associated with a non-significant loss of Phosphorylated DRP-1 SER637 (P-DRP1) abundance in older mice. Data are presented as mean ± SD. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction applied. Significance was determined at p < 0.05. Significant differences were observed in the expression levels of dynamic proteins in aged samples (*p < 0.05, **p < 0.01). = Males = Females. For full immunoblots see Supplemental Figure S2, available at Figshare: [10.6084/m9.figshare.29669213].

Does Mitochondrial Network Organization Deteriorate with Age?

To examine age-related changes in mitochondrial network organization, we utilized immuno-histological analyses, including both longitudinal and transverse sections of mouse skeletal muscles. The mitochondrial reticulum was co-stained using TOMM20 to visualize the network in both 2- (2D) and 3-dimensions (3D).

From a two-dimensional (2D) perspective, quantification of mitochondrial networks in the longitudinal plane revealed significant age-related changes in mitochondrial form factor (MFF), a measurement of morphology (F (1, 16) = 34.75, p < 0.0001) (Figure 8 e, Table 2). Mitochondrial counts showed significantly more fragmented networks in older mice compared to younger mice (F (1, 16) = 40.90, p < 0.0001) (Figure 8 g). Using the average length of each branch, we found that younger mice had both longer and thicker mitochondrial networks compared to older mice (F (1, 16) = 15.41, p < 0.0001) (Figure 8 f). In transverse sections, cross-sectional analyses showed a similar pattern of significant age-related changes. Average branch diameter was smaller in older mice (F (1, 16) = 14.54, p = 0.0015) (Figure 9 f, Table 3). The mean form factor was also significantly lower in older mitochondrial networks (F (1, 16) = 24.74, p = 0.0001) (Figure 9 e). The 2D results indicated that mitochondrial networks were less elongated and more fragmented in aged muscles, which was corroborated by means of 3D analyses.

Figure 8. Increased mitochondrial fragility is associated with age-related reductions in membrane diameter.

(A) Longitudinal muscle sections (100x magnification) stained with Translocase of Outer Mitochondrial Membrane 20 (TOMM20) conjugated Alexa Fluorescence 488 nm (AF488 nm-Green) for mitochondria and DAPI (Nuclei) ( 81) from young (Y) (n = 20 fibers, 10 male, M and 10 female, F) and old (O) (n = 20 fibers, 10M and 10F) n=5 mice in each of the 4 groups. (B) The ratio of total mitochondrial branch length per unit of image area was not different between old and young mice indicative of similar mitochondrial contents. (C) The reduced average length of each mitochondrial branch indicated possible mitochondrial fragmentation (D) The Mean Form Factor (MFF) calculated as P²/4πarea analyzing the shape of the mitochondrial branches with lower values being categorized as round and larger values equating to elongated features was also reduced in older mice. (F) Displays the average diameter of all the mitochondrial branches in the image showing reductions in older mice which be interpreted to mean reduced mitochondrial fusion. (G) Represents the absolute number of mitochondria that are not tethered to one another. Data are presented as mean ± SD. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction for multiple comparisons. Significance was determined at p < 0.05. (*p < 0.05, **p < 0.01, ***p < 0.001). The scale bar is 5 μm. The units of length and area used to analyze the images are reported in micrometers (μm). Images were collected utilizing Zeiss CLSM 880 with the same laser power and gain settings. Quantification analyses were conducted in ImageJ utilizing the Mitochondrial Analyzer plugin to optimize image analysis. = Males = Females.

Table 2.

Longitudinal Sections (2D) Measurements.

(2D Longitudinal)	YMM (n=5)	YFM (n=5)	OMM (n=5)	OFM (n=5)
Branch Length·Area−1	2.20 ± 0.22	2.07 ± 0.23	2.11 ± 0.27	2.15 ± 0.34
MFF ( *# )	3.57 ± 0.63	3.13 ± 0.58	2.49 ± 0.12	2.63 ± 0.24
Average Branch Diameter ( *# )	0.36 ± 0.01	0.36 ± 0.01	0.34 ± 0.02	0.35 ± 0.02
Mito Branches·Area−1 ( *# )	2.71 ± 0.30	2.56 ± 0.32	2.04 ± 0.12	2.22 ± 0.26
Average Branch Length ( *# )	1.10 ± 0.15	0.98 ± 0.18	0.71 ± 0.09	0.82 ± 0.12
Mitochondrial Count ( *# )	374 ± 108	350 ± 140	404 ± 202	535 ± 356

Quantitative measurements of mitochondrial and cellular structure parameters from n=5 mice in each of the 4 groups of skeletal muscle from young and old, male and female mice. Data were grouped by sex (Male and Female) and age (Young and Old). Data are reported as mean ± SD and reporting changes that are associated with aging. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction for multiple comparisons. Significance was determined at p < 0.05, (* significantly different for males, # significantly different for females, *# significantly different for both groups).

Figure 9 a-g.-. Transverse Sections of mouse skeletal muscle indicating mitochondrial fragmentation in mouse skeletal muscle.

(A) Transverse muscle sections (100x magnification) stained with COx and separately conjugated to (AF 488 nm Green) for mitochondria in young (Y) (n = 20 fibers, 10 male, M and 10 female, F) and old (O) (n = 20 fibers, 10M and 10F) n=5 mice in each of the 4 groups. (B) The ratio of total mitochondrial branch length per unit of image area was also similar between old and young mice. (C) The reduced average length of each mitochondrial branch showed similar decreases with age (D) The Mean Form Factor (MFF) significantly lower in female mice whereas the opposite was found for (F) Average branch diameter (G) Represents the absolute count of mitochondria was also found to be the same. Data are presented as mean ± SD. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction for multiple comparisons. Significance was determined at p < 0.05, (*p < 0.05, **p < 0.01, ***p < 0.001). The scale bar is 5 μm. The units of length and area used to analyze the images are reported in μm. Images were collected utilizing Zeiss CLSM 880 with the same laser power and gain settings. Quantification analyses were conducted in ImageJ utilizing the Mitochondrial Analyzer plugin to optimize image analysis. Image processing explained in the text. = Males = Females.

Table 3.

Transverse Sections (2D) Measurements.

(2D Transverse)	YMM (n=5)	YFM (n=5)	OMM (n=5)	OFM (n=5)
Branch Length·Area− 1	2.04 ± 0.22	2.37 ± 0.31	1.90 ± 0.18	2.26 ± 0.17
MFF ( # )	2.43 ± 0.41	3.20 ± 0.87	2.05 ± 0.21	2.29 ± 0.32
Average Branch Diameter	0.43 ± 0.02	0.38 ± 0.02	0.40 ± 0.03	0.36 ± 0.01
Mito Branches·Area− 1	1.75 ± 0.17	2.20 ± 0.29	2.15 ± 0.25	2.67 ± 0.50
Average Branch Length ( *# )	1.13 ± 0.16	1.10 ± 0.17	0.93 ± 0.13	0.87 ± 0.12
Mitochondrial Count	374 ± 108	350 ± 140	404 ± 202	535 ± 356

Quantitative measurements of mitochondrial and cellular structure parameters from n=5 mice in each of the 4 groups of skeletal muscle from young and old, male and female mice. The data is grouped by sex (Male and Female) and age (Young and Old). Data are reported as mean ± SD and reporting changes that are associated with aging. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction for multiple comparisons. Significance was determined at p < 0.05, (* significantly different for males, # significantly different for females, *# significantly different for both groups).

From a three-dimensional (3D) perspective, reconstructions of mitochondrial reticula in mouse muscles also revealed minor age-related changes to mitochondrial morphology. In the longitudinal plane specifically, thinner branches (F (1,8) = 20.40, p = 0.0020) (Table 4) and increased branch endpoints relative to volume (F (1,8) = 5.504, p =0.0470) (Table 4). In 3D transverse measurements, increased sphericity (F (1,16) = 6.222, p = 0.0239) (Table 5) and smaller branch diameters (F (1,16) = 17.59, p = 0.0007) (Table 5) mirrored the findings in 2D analyses related to the thinning of the mitochondrial network with age.

Table 4.

Longitudinal Sections (3D) Measurements.

3D (Longitudinal)	YMM (n=3)	YFM (n=3)	OMM (n=3)	OFM (n=3)
Average Branch Diameter ( *# )	0.30 ± 0.01	0.33 ± 0.02	0.26 ± 0.01	0.28 ± 0.02
Branch Endpoints·Volume−1	0.66 ± 0.10	0.81 ± 0.46	1.34 ± 0.11	1.15 ± 0.58
Branches·Volume−1	4.88 ± 0.60	4.93 ± 0.58	5.02 ± 0.69	4.86 ± 0.65
Mean Branch Length ( * )	1.66 ± 0.02	1.63 ± 0.04	1.47 ± 0.02	1.62 ± 0.12
Total Branch Length·Volume−1	0.07 ± 0.02	0.08 ± 0.03	0.10 ± 0.03	0.09 ± 0.03
Sphericity	0.02 ± 0.01	0.01 ± 0.02	0.02 ± 0.02	0.01 ± 0.01

Quantitative measurements of mitochondrial and cellular structure parameters from n=3 mice in each of the 4 groups of skeletal muscle. The data is grouped by sex (Male and Female) and age (Young and Old). Data are reported as mean ± SD and reporting changes that are associated with aging. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction for multiple comparisons. Significance was determined at p < 0.05, (* significantly different for males, # significantly different for females, *# significantly different for both groups).

Table 5.

Transverse Sections (3D) Measurements.

3D (Transverse)	YMM (n=3)	YFM (n=3)	OMM (n=3)	OFM (n=3)
Average Branch Diameter ( *# )	0.35 ± 0.02	0.35 ± 0.03	0.30 ± 0.02	0.30 ± 0.03
Branch Endpoints·Volume−1	0.81 ± 0.12	0.94 ± 0.15	0.84 ± 0.24	0.95 ± 0.33
Branches·Volume−1	1.43 ± 0.30	1.42 ± 0.24	1.62 ± 0.24	1.65 ± 0.41
Mean Branch Length	1.75 ± 0.10	1.73 ± 0.11	1.71 ± 0.21	1.76 ± 0.09
Total Branch Length·Volume−1	2.50 ± 0.54	2.36 ± 0.49	2.66 ± 0.36	2.22 ± 0.77
Sphericity ( *# )	0.06 ± 0.02	0.07 ± 0.02	0.10 ± 0.06	0.09 ± 0.04

Quantitative measurements of mitochondrial and cellular structure parameters from n=3 mice in each of the 4 groups of skeletal muscle. The data is grouped by sex (Male and Female) and age (Young and Old). Data are reported as mean ± SD and reporting changes that are associated with aging. Statistical significance was assessed using an ordinary two-way ANOVA with biological replicates, with Bonferroni correction for multiple comparisons. Significance was determined at p < 0.05, (* significantly different for males, # significantly different for females, *# significantly different for both groups).

Together, these findings provide a static yet comprehensive view of the mitochondrial network and how it may become somewhat fragmented with age. Despite these structural changes, mitochondrial function, as assessed by phosphorylation coupling efficiency (ADP:O) and performance in healthy humans in vivo (50), is preserved in aging skeletal muscle.

DISCUSSION

Given inconsistencies in the literature on muscle mitochondrial functioning in aging, we sought to assess the issue of inconsistency utilizing two approaches. In the current investigation, we utilized a multifaceted reductionist approach to evaluate the effects of aging on skeletal muscle mitochondrial morphology and function in a mouse model. Separately, we quantified muscular efficiencies in healthy young and older men and women (50). We found that muscular efficiency during graded exercise testing was preserved in healthy older men and women (50), but still numerous metabolic age-related postprandial metabolic adjustments were seen in older men and women (58–60). By comparing muscle and mitochondrial isolates from young and older male and female mice, we gained insight into how skeletal muscle mitochondrial function is preserved in aging. Again, we identified numerous subtle but significant changes in mitochondrial morphology, and to a much lesser extent, functionality related to energy-substrate partitioning that occurs in aging. These findings are discussed subsequently.

Muscle Mitochondrial Reticulum Energy Coupling Efficiency is Preserved in Aging

It is understood that a well-coupled mitochondrial reticulum serves as a conductive highway for intracellular energy distribution (8, 10, 11, 49, 61). In mouse skeletal muscle, representative data indicate that mitochondrial efficiency, as defined by the coupling between ADP phosphorylation and O₂ consumption (ADP:O) was unaffected by age. Our results are inconsistent with those of investigators who assessed mitochondrial ADP:O by titrating supraphysiological concentrations of ADP into mitochondrial preparations (28, 34, 62, 63). In contrast, we employed physiological ADP concentrations and obtained consistent and expected adenosine phosphorylation coupling efficiencies, regardless of whether pyruvate or palmitoyl-carnitine was used as the energy substrate. Hence, the results we obtained on muscle mitochondrial preparations from young and older mouse muscles were consistent with what we determined on muscular efficiencies in healthy older men and women during exercise.

In discussing ADP-linked respiration, we observed small, but significantly lower rates of O₂ consumption in mitochondrial preparations from older mice. We attributed those results to limitations in substrate uptake and oxidation, specifically to reduced expression of mitochondrial monocarboxylate transporters (mPC and mMCT) and β-oxidation enzymes (HADH and CPT1b). Moreover, because there were either minimal or no age-associated differences in the maximal catalytic efficiencies of enzymes in mitochondrial preparations, likely, the observed changes were again due to limitations in transportation and enzyme abundances. Although phosphorylation efficiency was consistent between young and older mitochondrial preparations, we found reductions to the RCR, an indicator of membrane integrity in those preparations.

Reduced Mitochondrial RCR Indirectly Characterizes Membrane Fragility

The literature reports mixed findings regarding RCR in mitochondrial preparations from older individuals (7, 26, 64). Therefore, when making cross-study comparisons, the analytical procedures used must be considered. These include the tissue sampled, isolation technique, substrate titration, timing, instrumentation, and environmental conditions (65, 66). The lower RCR in mitochondrial preparations from older mice was likely due to elevated rates of ADP-limited (S4) O₂ consumption. Indeed, the increase in S4 respiration has been traditionally associated with proton leak, which suggests mitochondrial membrane damage in isolation or in vivo. Techniques to isolate mitochondria are known to cause membrane damage due to chemical, shear, and centrifugal forces applied when compartmentalizing the cell, which has been shown to exaggerate the functional age effect of mitochondrial fragments (67). For that reason, we used relatively gentle muscle homogenization in the presence of a protease inhibitor to produce mitochondrial membrane fragments. Because we used the same isolation procedures to obtain mitochondrial preparations in skeletal muscle from both young and older mice, it can be speculated that the mitochondrial reticulum is more prone to membrane damage during isolation than to aging per se.

As a cautionary note, we are concerned that the observed decline in RCR of mitochondrial preparations from older muscle is more an isolation artifact than a physiological difference. An increase in S4 respiration and a decrease in RCR of the mitochondrial reticulum in aging would manifest as an increase in basal metabolic rate (BMR) in aging. However, to the contrary, it is well established that BMR declines in aging (68). And, as already mentioned, leg cycle ergometer studies on healthy young and older men and women (50) did not find age-related decreases in muscular efficiency that might have occurred had there been physiologically significant decreases in mitochondrial respiratory control or OxPhos coupling.

Reduced Markers of Mitochondrial Fusion and Fission in Older Mouse Skeletal Muscle

The mechanism coordinating mitochondrial membrane fusion includes MFN1, which specifically maintains outer mitochondrial membrane integrity, and MFN2, which plays a much smaller role in membrane fusion but is primarily responsible for maintaining cellular metabolism and apoptosis (40). Considering the former, it becomes apparent why the RCR data, an indirect but often utilized marker of membrane integrity obtained from older mice, was lower than that of the younger cohort. Examination DRP1 and FIS1 protein yields in older animals resulted in lower abundances. Those results were consistent with previous evidence indicating lower levels of fission proteins or their functions in aging muscle (41). There is evidence that DRP1 phosphorylation at the Ser 616 residue leads to increased mitochondrial membrane fission while phosphorylation of the Ser 637 residue has been shown to inhibit fission (69–73). In our experiments we focused on the abundance of phosphorylated Ser 637 residue that was not significantly lower in older mouse muscle. Consequently, when studying aging skeletal muscle future investigations planning to utilize pDRP1 as a proxy for mitochondrial membrane fission are encouraged to quantify the Ser616 to Ser637 ratio. Nevertheless, age-related changes in levels of the proteins of mitochondrial dynamics were apparent. However, it is important to note that the diminished enzyme abundances reflect static amounts and not turnover rates, making it difficult to infer the state of its activity in vivo. Investigations regarding mitochondrial protein turnover comparing young and old skeletal muscle have reported both slower synthetic rates following short-term labeling (74), while others have observed no differences in turnover rates utilizing long-term tracer labeling schemes (75). These changes, together with visual data on mitochondrial networks obtained using confocal microscopy, suggest a pattern consistent with increased mitochondrial protein degradation in aging.

Older Skeletal Muscle is Characterized by Fragile Mitochondrial Membranes and a Disorganized Reticulum

Our findings demonstrated significant reductions in the average mitochondrial branch diameter in both longitudinal and transverse muscle histological sections, observed in both 2-dimensional (2D) and 3-dimensional (3D) analyses of old and young mice. Notably, previous investigations used 3D reconstruction techniques to explore the relationship between mitochondrial structure and dynamics. One investigation utilized skeletal muscle biopsies from young and older humans to study the effects of circadian rhythms on mitochondrial morphology (76). However, while biopsies obtained from the younger group in that study were categorized as having normal height and stature, the older group was classified as obese, thus raising the issue of comorbidities affecting the results and their interpretation. Similarly, another investigation on cardiac mitochondrial morphology in older adults with or without heart failure identified changes in reticulum structure and membrane damage as they relate to disease (77). The authors speculated there were likely no structural decrements in mitochondrial morphology in older healthy individuals, but again, without a healthy group from which to compare, their conclusions are not definitive.

Our results showing mitochondrial disorganization in muscles of healthy aging mice are supported by results of studies on a variety of different animal models. For instance, changes in mitochondrial branching and morphology have been observed when 2D immuno-histological sections of skeletal muscle from mice were stained with fluorescent mitochondrial probes (78, 79). Even so, results of those studies reinforce the need to consider the 3D and dynamic nature of the mitochondrial reticulum on metabolism (61).

To our knowledge, we are the first to observe a decline in the diameter of mitochondrial reticulum branches in muscles of an aging mammal. If true, loss of reticulum integrity in aging may make it more susceptible to stress. Our observations of mitochondrial reticulum branch thinning, along with reduced RCR can be interpreted to indicate a loss in mitochondrial membrane integrity in aging. Additionally, we observed a reduction in average mitochondrial branch length, which may be linked to decreased abundance of mitochondrial dynamic proteins. Hopefully, data on turnover rates of mitochondrial proteins in aging are forthcoming. Moreover, while others may equate shorter branch lengths to lower mitochondrial content in our 3D reconstructions, the ratio of mitochondrial reticulum branches normalized to the total muscle area was indistinguishable between groups, consistent with our enzymatic measurements used to estimate mitochondrial content. Finally, it is important to note that changes in mitochondrial morphology and enzyme abundances occurred without reductions in mitochondrial phosphorylation efficiency, as determined by the ADP:O ratio. Regardless, if aging results in decreases in diameters of mitochondrial reticulum branches it may be that they are more susceptible to damage by external stressors as well as isolation procedures.

Above we cited the report of Duong et al on working muscle coupling efficiency during mild-to moderate exercise in young and aged humans (50). No age-related muscular efficiency effect was observed. However, should mitochondrial membrane integrity be more susceptible to exercise stress, then we might find an elevation in post-exercise oxygen consumption (EPOC). Above we also cited some of the rich literature on exercise training on muscle mitochondrial protein abundances. However, if mitochondrial membranes were more susceptible to stress in aging, then recovery oxygen consumption should be elevated in older individuals postexercise. However, in a literature search we were unable to find a body of literature on recovery O₂ consumption (EPOC) in aging. Conversely, if fitness was considered, heart rate recovered faster in older fit than in older unfit men, or younger fit or unfit men (80).

In summary, despite preservation of muscular exercise efficiency in older men and women (50), the null hypothesis of unchanged mitochondrial structure and function in muscles of aging mice is rejected. Given the considerable number of mitochondrial parameters examined, some mitochondrial decrements in aging may be due to random chance. However, despite the overall robustness and resilience of the mitochondrial reticulum in a mouse model of aging, we found numerous deficits and no age-related improvements in mitochondrial abundance, structure, or function. Those mitochondrial deficits are not apparent in energy coupling efficiency, but rather may translate to changes in energy substrate partitioning in human aging (58–60).

DATA AVAILABILITY

Data derived from public domain information: Source data for this study were derived from the following resources at Figshare: [10.6084/m9.figshare.29669213].