Mitochondrial dysfunction is well described in many chronic illnesses including musculoskeletal, neurodegenerative, and cardiovascular diseases. Mitochondrial network morphology has been implicated as a biomarker of disease, correlating increased mitochondrial fragmentation to impaired cellular function. While advancements in imaging techniques further our understanding of mitochondrial dynamics in live cells, easily accessible approaches for accurate quantification of in situ mitochondrial networks in low abundance tissues are lacking. The purpose of this study was to validate a proof-of-concept method capable of quantifying 3D mitochondrial network morphology in whole mount skeletal muscle and then applying it to mitochondrial morphology analysis in cell types otherwise difficult to image within their native environment, terminal Schwann cells (tSCs). Herein, we report that mitochondrial networks were fragmented in dystrophic mouse muscle compared to healthy controls, as observed by others, and correlated with muscle pathology as expected. Using S100β reporter mice to identify Schwann cells we labeled tSC mitochondrial networks in vivo prior to rapid imaging in situ with high-resolution confocal microscopy. Moreover, these methods offer a comprehensive and novel approach enabling the quantification of mitochondria network morphology across multiple cell types (like muscle fibers and tSCs) using standard microscopy available in university core facilities.

The authors thank Drs. Mariappan Muthuchamy, Mendell Rimer, and Peter Nghiem for generously providing the D2.mdx mice used to evaluate tSC mitochondria. This work was supported by the Sydney and J.L. Huffines Institute for Sports Medicine and Human Performance Student Research Grant and by funds from the College of Education and Human Development at Texas A&#38;M University. A.B.M. was also supported by the NIH LRP (NIAMS, 2L40AR077899-02A3).

Six-month-old (n=3) adult male DBA/2J [Wild-type (WT)] (RRID: IMSR_JAX:000671) and D2.B10-DMDD2.mdx/J [D2.mdx (mdx)] (RRID: IMSR_JAX:013141), while three to five-month-old (n=8) adult male and female B6; D2-Tg(S100-EGFP)1Wjt/J (S100B) (RRID: IMSR_JAX:005621) [also pooled with WT given no differences from DBA] were used from an ongoing colony of investigator-maintained mice. The overall experimental workflow for TA and GM injections, tissue processing, and imaging is summarized in Figure 1. An additional D2.mdx mouse was investigator-donated for tSC imaging in mdx (n=1). All animal procedures were approved by the Institutional Animal Care and Use Committee at Texas A&#38;M University (AUP 2022-0215) and were conducted in accordance with the National Research Council Guide for the Care and Use of Laboratory Animals (8th edition)30. Euthanasia was performed by cervical dislocation under an anesthetic overdose of isoflurane.
1. Preparation of fluorescent mitochondrial stain working solution of 2 &#181;M
<ol>
	<li>Suspend the mitochondrial stain (50 &#181;g) in 322 &#181;L of dimethyl sulfoxide (DMSO) according to the manufacturer&#8217;s instructions to prepare a 200 &#181;M stock solution.</li>
	<li>To prepare 30 mL of 2 &#181;M working solution, transfer 300 &#181;L of 200 &#181;M stock solution into 29.7 mL of sterile saline. Mix thoroughly.</li>
	<li>Aliquot 250 &#181;L of the 2 &#181;M working solution into microcentrifuge tubes and store at -80 &#176;C until use. 
	NOTE: No detectable signal degradation was observed after 3 years of storage at -80 &#176;C.</li>
</ol>
2. Dye and animal preparation
<ol>
	<li>Thaw one frozen aliquot 15 min prior to experimentation. Draw the solution into a 30 G, 1 cc insulin syringe.</li>
	<li>Anesthetize the animal with isoflurane. Apply ophthalmic ointment, shave the injection site, and maintain body temperature using a warming lamp with a temperature probe positioned between the animal and the heat source.</li>
	<li>Clean the shaved injection site with 70% isopropyl alcohol.</li>
</ol>
3. Tibialis anterior (TA) muscle mitochondrial dye administration and function
<ol>
	<li>TA dye administration
	<ol>
		<li>Under anesthesia, inject 50 &#181;L of 2 &#181;M mitochondrial stain into the Left or Right TA muscle of anesthetized WT and D2.mdx mice.</li>
		<li>Insert the needle at the distal tendon toward the knee and depress the plunger while withdrawing the needle along the injection path.</li>
		<li>Incubate the dye in vivo for 1 h prior to dissection.</li>
	</ol>
	</li>
	<li>TA muscle function measurements
	<ol>
		<li>After 45 min of dye incubation, anesthetize the mouse using isoflurane (2%&#8211;3% induction, 1.5%&#8211;2% maintenance in oxygen).</li>
		<li>Surgically expose the left TA muscle while preserving surrounding tissues. Tie the distal tendon to the load beam of the muscle testing system.</li>
		<li>Wrap the exposed muscle with a laboratory wipe and irrigate at 3 mL min&#8315;1 with 0.9% sterile saline warmed to 40 &#176;C. Maintain muscle temperature at 32 &#176;C using a heat lamp.</li>
		<li>Position platinum/iridium wire electrodes across the muscle belly for direct stimulation.</li>
		<li>Attach the load beam to a micrometer and adjust optimal length (L0) using twitch contractions (Tw) at 1 Hz. Record L0 with a digital caliper.</li>
		<li>Induce maximum tetanic contractions (P0) three times at 140 Hz with 60 s rest between contractions. Record the highest force value (g). Acquire data using the muscle function system with analysis software.</li>
	</ol>
	</li>
	<li>TA tissue processing
	<ol>
		<li>After incubation, isolate and dissect the TA muscle prior to euthanasia.</li>
		<li>Place isolated muscle fiber bundles in 4% paraformaldehyde at room temperature for 15 min while gently teasing fibers under a stereomicroscope. 
		CAUTION: Paraformaldehyde is toxic and should be handled in a chemical fume hood while wearing appropriate personal protective equipment (lab coat, gloves, and eye protection).</li>
		<li>Add cell-permeant nucleic acid stain to a final concentration of 5&#8211;10 &#181;g/mL to label nuclei.</li>
		<li>Wash samples three times in phosphate-buffered saline (PBS) for 5 min each to remove excess dye.</li>
		<li>Dehydrate samples sequentially in 50%, 75%, and 100% ethanol for 5 min each.</li>
		<li>Incubate samples sequentially in tissue clearing solutions I and II for 30 min each at room temperature using sufficient volume to fully submerge tissue.</li>
		<li>Place cleared TA bundles on a coverslip and gently flatten using a glass block (2.5 cm &#215; 2 cm &#215; 1 cm; mass 7.8 g)26.</li>
	</ol>
	</li>
</ol>
4. Imaging of TA muscle samples
<ol>
	<li>Image muscle samples with a confocal microscope to obtain high-resolution three&#8211;dimensional (3D) optical sections.</li>
	<li>Acquire Z-stack images at a depth of approximately 100 &#181;m for muscle using a 63x magnification objective lens (N.A. 1.40).</li>
	<li>Use the imaging system&#8217;s confocal software with an excitation wavelength of 646 nm to provide the greatest clarity with the least background.</li>
	<li>Acquire images using deconvolution mode, employing &#8220;System Optimized&#8221; parameters with 1 &#181;m step sizes. 
	NOTE: To combat dehydration and photobleaching, transfer the tissues into tissue-clearing solution II and place them on a coverslip. Place two drops of clearing solution 2 directly upon the tissue prior to imaging.</li>
</ol>
5. GM muscle tSC dye loading and imaging
<ol>
	<li>Use S100&#946; mice for specific fluorescent identification of tSCs.</li>
	<li>Under anesthesia, perform a small skin incision above the GM muscle. Inject 75 &#956;L of a 2 &#956;M far-red fluorescent mitochondrial stain beneath the GM, prior to wound closure25.</li>
	<li>Close the wound and allow the dye to incubate for 45 minutes to selectively label tSCs without significant labeling of the underlying muscle fibers.</li>
	<li>Dissect the GM muscles and immediately place them in chilled 0.9% sterile saline. Rapidly trim excess fat and connective tissue.</li>
	<li>Transfer the dissected GMs onto a coverslip for imaging.</li>
	<li>Only if using D2.mdx mice, add fluorescent acetylcholine receptor (AChR) dye at a 1:500 dilution to the microdissection bath. Incubate the tissue for 30 min to visualize the postsynaptic components of the neuromuscular junction (NMJ).</li>
	<li>Use this dual-labeling approach to differentiate postsynaptic mitochondria from tSC-specific mitochondria. 
	CAUTION: Limit dissection and cleaning of the GM to no more than 10 minutes prior to placing onto a coverslip for imaging to minimize morphological changes following dissection. This time limit is critical to minimize post-dissection morphological changes and mitochondrial degradation.</li>
	<li>Perform confocal microscopy to obtain high-resolution three&#8211;dimensional (3D) optical sections.</li>
	<li>Acquire Z-stack images at a depth of approximately 16 &#181;m for tSCs using a 20x magnification objective lens (N.A. 0.75). Acquire high-resolution representative images of individual tSCs using a 63x magnification objective lens.</li>
	<li>Implement mounting procedures to minimize tissue dehydration and photobleaching during the imaging session. 
	CAUTION: Transfer tissues in saline onto a coverslip, ensuring the ventral side faces down for inverted microscopy.</li>
	<li>Apply 2 &#181;L of general mounting medium onto the dorsal surface of the GM.</li>
	<li>Gently flatten the tissue using a calibrated glass block (2.5 cm &#215; 2 cm &#215; 1 cm; mass 7.8 g) to ensure a uniform imaging plane.</li>
</ol>
6. Focused ion beam microscopy
<ol>
	<li>Perform focused ion beam scanning electron microscopy (FIB-SEM) according to previously established protocols26.
	<ol>
		<li>Apply FIB-SEM specifically to confirm and visualize novel morphological features identified during confocal imaging.</li>
	</ol>
	</li>
</ol>
7. Image analysis
<ol>
	<li>TA muscle samples
	<ol>
		<li>Conduct quantitative analysis of confocal optical sections (1 &#956;m) of TA scans (approximately 75&#8211;112 &#956;m thick) using AI-driven image analysis software. 
		&#8203;NOTE: Perform multiple supervised training sessions prior to automated segmentation to enhance algorithm adaptation.</li>
		<li>Import single optical sections (Z-slices) to teach the software to retain pixelated information representing white mitochondrial dye while excluding the black background.</li>
		<li>Develop image masks for investigator review to ensure high-quality analysis and accuracy.</li>
		<li>Separate 3D image stacks into single optical sections following algorithm optimization.</li>
		<li>Select every fifth Z-slice from each 3D image for analysis, with each data point representing the average of these individual sections per sample.</li>
		<li>Quantify mitochondrial morphology to derive spatial distribution and network connectivity metrics.</li>
		<li>Standardize intensity thresholds, edge detection settings, and voxel size calibration across all samples to minimize experimental variability.</li>
	</ol>
	</li>
	<li>tSCs on GM samples
	<ol>
		<li>Assemble 1 &#956;m optical sections into Z-stacks (approximately 6&#8211;25 &#956;m thick) and separate images into individual color channels using 2D image analysis software.</li>
		<li>Calibrate software using scale bars and apply thresholding to optimize tSC mitochondria visualization with minimal background interference.</li>
		<li>Perform binarization and remove background noise to isolate tSC mitochondria or tSCs for the measurement of area and particle counts.</li>
		<li>Save binary images to ensure consistent processing and perform all imaging experiments under identical conditions to ensure reproducibility across biological replicates.</li>
	</ol>
	</li>
</ol>
8. Immunohistochemistry
<ol>
	<li>Embed left TA muscle samples in optimal cutting temperature (OCT) compound to evaluate cross-sectional area (CSA).
	<ol>
		<li>Section samples at 10 &#956;m thickness using a cryostat set to -17&#176;C and collect onto microscope slides.</li>
		<li>Stain muscle cross-sections for laminin to identify myofiber borders using rabbit anti-laminin primary antibody (1:400) and goat anti-rabbit Rhodamine secondary antibody (1:400).</li>
		<li>Mount slides using an appropriate mounting medium prior to imaging.</li>
	</ol>
	</li>
	<li>Image slides using a confocal microscope by randomly selecting 2&#8211;3 regions of interest (580 x 580 &#181;m) per sample with a 20x (N.A. 0.75) objective.</li>
	<li>Average values across regions per muscle and analyze approximately 400 myofibers per TA muscle section.</li>
	<li>Use semi-automatic muscle segmentation software to determine CSA (&#181;m2) by masking the myofibers.</li>
</ol>
9. Statistical analysis
<ol>
	<li>Report summary data as mean &#177; standard error of the mean (S.E.M.).</li>
	<li>Perform statistical analysis using a dedicated statistical software, utilizing parametric, unpaired Student&#8217;s t-tests to determine significance among group mean differences.</li>
	<li>Conduct simple linear regressions on CSA and muscle force from TA muscles, utilizing these as markers of DMD pathology to compare mitochondrial morphology from TA muscle and GM tSCs.</li>
	<li>Assess Pearson r correlation coefficients to determine the correlation between DMD pathology and mitochondrial pathology. Define statistical significance as P &#8804; 0.05.</li>
</ol>

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<li>Pareyson, D., Piscosquito, G., Moroni, I., Salsano, E., Zeviani, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Peripheral+neuropathy+in+mitochondrial+disorders%5BTitle%5D)+AND+%22Lancet+Neurol%22%5BJournal%5D)">Peripheral neuropathy in mitochondrial disorders.</a> Lancet Neurol. 12 (10), 1011-1024 (2013).</li>
<li>Schröder, J. M., Sommer, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mitochondrial+abnormalities+in+human+sural+nerves%3A+Fine+structural+evaluation+of+cases+with+mitochondrial+myopathy%2C+hereditary+and+non-hereditary+neuropathies%2C+and+review+of+the+literature%5BTitle%5D)+AND+%22Acta+Neuropathol%22%5BJournal%5D)">Mitochondrial abnormalities in human sural nerves: Fine structural evaluation of cases with mitochondrial myopathy, hereditary and non-hereditary neuropathies, and review of the literature.</a> Acta Neuropathol. 82 (6), 471-482 (1991).</li>
<li>Schröder, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Neuropathy+associated+with+mitochondrial+disorders%5BTitle%5D)+AND+%22Brain+Pathol%22%5BJournal%5D)">Neuropathy associated with mitochondrial disorders.</a> Brain Pathol. 3 (2), 177-190 (1993).</li>
<li>Santosa, K. B., Keane, A. M., Jablonka-Shariff, A., Vannucci, B., Snyder-Warwick, A. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Clinical+relevance+of+terminal+Schwann+cells%3A+An+overlooked+component+of+the+neuromuscular+junction%5BTitle%5D)+AND+%22J+Neurosci+Res%22%5BJournal%5D)">Clinical relevance of terminal Schwann cells: An overlooked component of the neuromuscular junction.</a> J Neurosci Res. 96 (7), 1125-1135 (2018).</li>
<li>Morton, A. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Inducible+deletion+of+endothelial+cell+Efnb2+delays+capillary+regeneration+and+attenuates+myofibre+reinnervation+following+myotoxin+injury+in+mice%5BTitle%5D)+AND+%22J+Physiol%22%5BJournal%5D)">Inducible deletion of endothelial cell Efnb2 delays capillary regeneration and attenuates myofibre reinnervation following myotoxin injury in mice.</a> J Physiol. 602 (19), 4907-4927 (2024).</li>
<li>Morton, A. B., Norton, C. E., Jacobsen, N. L., Fernando, C. A., Cornelison, D. D. W., Segal, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Barium+chloride+injures+myofibers+through+calcium-induced+proteolysis+with+fragmentation+of+motor+nerves+and+microvessels%5BTitle%5D)+AND+%22Skelet+Muscle%22%5BJournal%5D)">Barium chloride injures myofibers through calcium-induced proteolysis with fragmentation of motor nerves and microvessels.</a> Skelet Muscle. 9 (1), 27(2019).</li>
<li>Sukhorukov, V. M., Dikov, D., Reichert, A. S., Meyer-Hermann, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Emergence+of+the+mitochondrial+reticulum+from+fission+and+fusion+dynamics%5BTitle%5D)+AND+%22PLoS+Comput+Biol%22%5BJournal%5D)">Emergence of the mitochondrial reticulum from fission and fusion dynamics.</a> PLoS Comput Biol. 8 (10), e1002745(2012).</li>
<li>Ferramosca, A., Zara, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mitochondrial+carriers+and+substrate+transport+network%3A+A+lesson+from+Saccharomyces+cerevisiae%5BTitle%5D)+AND+%22Int+J+Mol+Sci%22%5BJournal%5D)">Mitochondrial carriers and substrate transport network: A lesson from Saccharomyces cerevisiae.</a> Int J Mol Sci. 22 (16), (2021).</li>
</ol>

The authors have no competing interests.

The location of mitochondria within cellular structures, as well as their morphology, correlates with mitochondrial health and metabolic demand21. Given the clinical relevance of mitochondrial health, coupled with divergent respiration kinetics and morphologies of different cell types, an easily applied, confocal microscopy-based method for the evaluation of mitochondrial morphologies in tSCs was developed after first evaluating the method in skeletal muscle, providing information on mitochondrial health as indicated by morphology, in a tissue type difficult to maintain in vivo structure upon isolation. In that regard, controlled loading of a fluorescent mitochondrial dye in vivo labels mitochondria in both skeletal muscle and tSCs, enabling quantification of basic morphological metrics in health and disease.
Skeletal muscle is 40% of body mass and is necessary for breathing, myokine secretion, and locomotion3,10. Owing to its high body mass, ease of isolation, and multiplicity of physiological effects, mitochondrial morphology has been investigated in skeletal muscle since the 1960s21. Mitochondrial morphology in skeletal muscle is closely associated with both health and disease2,21,27. Indeed, in genetic muscle diseases such as muscular dystrophies, mitochondria are more fragmented, generating greater amounts of reactive oxygen species (ROS), reducing metabolic efficiency, and contributing to greater overall pathology28. Conversely, focusing on the restoration of healthy mitochondrial networks and morphology in disease reduces markers of myopathy, including improved contractile performance, reduced fibrosis, and lower net ROS production29.
Consistent with reports of skeletal muscle mitochondrial morphology in mouse models of Duchenne Muscular Dystrophy, this method of directly injecting skeletal muscle with a dye attracted to the negative membrane potential of mitochondria revealed fragmented mitochondria in the D2.mdx mouse. Additionally, these markers of mitochondrial pathology were highly correlated with muscle function and cross-sectional area, suggesting the utility of this method for evaluating mitochondrial morphology.
SCs provide insulation for axons, enabling rapid transmission of action potentials31. Given their importance, SC disruption can cause both sensory and motor disorders, such as Charcot-Marie-Tooth disease32. Notably, mitochondrial diseases often result in neuropathic phenotypes, while defects in peripheral nervous system axons often correlate with damage to mitochondrial DNA32. Following investigations of mitochondrial abnormalities using electron microscopy in cross-sections of peripheral nerves, it has been noted that mitochondria appear swollen and malformed, primarily in the enshrouding SCs rather than in the axon33,34. While poorly understood, there is a strong association between peripheral SC mitochondria morphology and neuropathology.
Recent evidence suggests that tSCs are particularly important in diseases such as spinal muscular atrophy and Amyotrophic lateral sclerosis35. Remek SCs or tSCs are unique among peripheral glia, as they remain unmyelinated, enshrouding the neurofilaments hovering above synaptic clefts at the neuromuscular junction (NMJ)20,35. The NMJ is the active site of neuromotor activity. In particular, axonal propagation down the axon promotes a release of acetylcholine to activate myofibers for contraction. Following injury, tSCs aid in the recovery of NMJs20. Concomitant with malformed NMJs in neuromotor pathologies, tSC morphology is equally malformed20,36,37.
Despite the close association of mitochondrial health and morphology with pathology, few studies have examined whole-mount imaging of tSC mitochondria using light microscopy. Given that confocal laser scanning microscopy is readily available at most university research cores, it was decided to develop a method to image and quantify mitochondrial morphology in tSCs in situ. Notably, the most striking characteristics of this analysis of healthy tSC mitochondria are the loops found at each bifurcation of the recognizable pretzel-like configuration (Figure 4C,D and Figure 5A&#8211;D). While stand-alone loops and individual mitochondrial globules are associated with mitochondrial fission, interconnected segments and loops are indicators of fusion events38, culminating in healthy mitochondrial interconnectivity and participation in substrate shuttling39. In agreement, the current study revealed mitochondrial fragmentation in tSCs from D2.mdx mice, suggesting reduced mitochondrial function.
Limitations of the current study included the ability to study only n=3 DBA and D2.mdx mice to establish proof of concept. In that regard, the power of the correlations between mitochondrial network connectivity and DMD pathology is low; however, previous studies have already established that D2.mdx mice exhibit deficits in CSA, muscle function, and mitochondrial connectivity, thereby increasing the applicability of these findings. The tSC images were acquired in sufficient numbers from 8 animals. Tissue thickness, as well as the location of tSCs (superficial or deep), dictates the ease of labeling and imaging tSC mitochondria with reduced background from surrounding muscles, currently restraining the tSC labeling technique to thin muscles with superficial NMJs and requiring optimization of dye incubation prior to experimentation.
Mitochondrial morphology has been used to infer mitochondrial health and the locations of metabolic activity. Taken together, injecting mitochondrial dyes into soft tissues in vivo provides unique advantages for understanding mitochondrial morphology in tissues otherwise difficult to obtain, such as tSCs. The present findings demonstrate a highly interconnected mitochondrial network within each tissue type. Disconnected mitochondria correlate with markers of dysfunction, consistent with other reports, demonstrating the validity of this approach. This technique is useful for analyzing tSC mitochondria within their reticulum, which are typically destroyed during dissection and may serve as a foundation for application to other tissues difficult to evaluate in vitro.

Mitochondria are remarkably plastic organelles that act as the lungs of the cell, consuming oxygen during cellular respiration to support adenosine triphosphate (ATP) production in response to metabolic demand1. While once thought of as discrete organelles, it is now understood that mitochondria are highly integrated2,3,4, sharing energy substrate throughout their network in accord with local requirements. Integration provides a distinct advantage over isolation: connected mitochondria support energy deficits, maintaining electrochemical gradients for oxidative phosphorylation during elevated metabolic demand. Indeed, the distribution, volume, and morphology of mitochondria suit the energy needs in their vicinity5. In health, regular turnover of mitochondrial networks is maintained through mitochondrial biogenesis, fusion and fission events, and mitophagy2,3. Mitochondrial biogenesis is required to maintain and expand healthy mitochondria. Metabolic demand, such as physical exercise, increases expression of the peroxisome proliferator-activated receptor &#947; co-activator 1&#945; (PGC-1&#945;)5. This process occurs via primary endurance signals, specifically the exercise-induced elevation of AMP-activated protein kinase (AMPK), calcium, and reactive oxygen species6,7. Fusion expands networks, promoting survivability and complementing deficits, while fission mitigates further mitochondrial damage through the preferential exclusion of dysfunctional region2,3,8. Once removed, excluded regions must undergo mitophagy, preventing the accumulation of dysfunctional organelles and proteins6,9.
Skeletal muscle comprises about 40% of body mass and relies on mitochondria for aerobic metabolism and cellular signaling to maintain homeostasis10. Collectively, in conditions of skeletal muscle dysfunction, mitochondrial quality control mechanisms are impaired10. Investigations of muscle dysfunction typically measure mitochondrial respiration, volume, morphology, and reactive oxygen species (ROS) production to assess mitochondrial quality3,10. In that regard, during prolonged muscle dysfunction, mitochondrial respiration is reduced, while ROS are increased, and networks are fragmented3,10. Perturbations in mitochondrial morphology arise largely from activation of mitochondrial fission proteins, dynamin-related protein 1 (DRP1) and mitochondrial fission 1 (FIS1)2,8,11. In response to muscle atrophy, FIS1, residing on the surface of the outer mitochondrial membrane, recruits DRP1 to oligomerize with multiple DRP1 dimers to create a filament around the mitochondria, decreasing in size until a given section of mitochondria is &#8220;pinched&#8221; off, promoting fragmentation3,11. Together, overactivity of mitochondrial fission, reductions in fusion of newly formed mitochondria, and dysfunctional mitophagy are responsible for the fragmented appearance of dysfunctional mitochondrial networks, like those described in inactive or diseased muscle2,12.
The neuromuscular junction (NMJ) is the site of muscular excitation. Action potentials initiated in the soma propagate along axons, ensheathed by myelinating Schwann cells (SCs), and terminate at NMJs, which are encapsulated by terminal Schwann cells (tSC). NMJs can regenerate following injury. However, the likelihood and speed of regeneration are diminished as the distance between the site of injury and the NMJ increase13,14. Following nerve injury by transection, crush, or stretch, SCs dedifferentiate into a demyelinated progenitor-like state15,16. Demyelinated SCs proliferate and work in concert with macrophages to remodel the local environment for regeneration by clearing debris between the site of injury and the proximal axon stump through Wallerian degeneration17,18. Migrating SCs signal an inflammatory response that remodels the environment for successful nerve regeneration18. Nerve regeneration occurs inside the neural sheath formed by the progenitor-like SCs, which form bands of Bunger, through which the axons regrow19. The axon stump then extends filopodia to initiate reinnervation14. Following reinnervation, the presynapse can fully regenerate, provided tSCs reinvest at the NMJ20.
Mitochondrial morphology varies markedly between cell types, conforming to tissue function21. Reports from liver tissues suggest that mitochondria are more compact and spherical, whereas those in the white matter in the brain are more elongated and tubular22. Upon challenge, such as in conditions of aging and oxidative stress, mitochondria in these tissues exhibit a doughnut-like phenotype. However, in osteoblasts, mitochondrial doughnut formation and fragmentation were linked with increased secretion of mitochondria and mitochondrial-derived vesicles, and with osteoblast maturation23. While skeletal muscle mitochondrial networks have been extensively examined across multiple species, limitations remain in imaging mitochondrial networks from tSCs of the peripheral nervous system. These pose difficult&#160;to capture within their native environments, as cell ultrastructure is significantly altered following isolation, and these tissues are difficult to target with dye in vivo without also loading accompanying skeletal muscle.
Imaging techniques used to capture mitochondrial morphology in living systems can be categorized as either light or electron microscopy5. With traditional light capture microscopy, dyes can label live cells, fixed cells can be immunolabeled, and mitochondria can be genetically labeled5,24. Live-cell imaging typically requires incubating cells for 20&#8211;60 min prior to fluorescence analysis. Dyes are directed to mitochondria via membrane potential and, therefore, may be altered under conditions of altered membrane potential5. Capturing mitochondrial morphology of tSCs within skeletal muscle via dye loading is theoretically possible; however, it would require imaging of whole tissues and detailed labeling techniques. In particular, it captures mitochondria within intact structures, without interference from skeletal muscle labeling. To overcome these limitations, the mitochondrial fluorescent stain was used to label living skeletal muscle in healthy and dystrophic mice, followed by labeling tSC mitochondria for in vivo and in situ imaging. The tibialis anterior (TA) muscle was selected to optimize skeletal muscle imaging due to its superficial anatomical location, well-characterized injection volume and myofascial dispersion25,26,27, and suitability for concurrent functional measurements. The gluteus maximus (GM) muscle was selected for tSC imaging based on its established injection procedure28,29, its thin architecture, which minimizes background fluorescence, and the superficial ventral positioning of neuromuscular junctions between the GM and gluteus medius, enabling preferential labeling of tSCs prior to skeletal muscle fibers.

<table><tbody><tr><td>3-in-1 Whole Animal System</td><td>Aurora Scientific, Ontario, Canada</td><td>1300A</td><td>Muscle funciton system with analysis software</td></tr><tr><td>AIVIA&amp;nbsp;</td><td>Leica Microsystems, Bellevue, WA, USA</td><td>15</td><td>AI driven image analysis software</td></tr><tr><td>anti-laminin&amp;nbsp;</td><td>MilliporeSigma</td><td>L9393-.2mL</td><td>myofiber boarder label</td></tr><tr><td>B6; D2-Tg(S100-EGFP)&lt;sup&gt;1Wjt&lt;/sup&gt;/J Mouse Strain</td><td>Jackson Labs, Bar Harbor, ME</td><td>JAX:005621</td><td>S100&amp;beta;</td></tr><tr><td>Cryostar NX50 Cryostat</td><td>Epredia, Kalamazoo, MI, USA</td><td>https://www.epredia.com/products/histology-instruments/cryotomy/cryostar-nx50</td><td>Cryostat</td></tr><tr><td>D2.B10-DMD&lt;sup&gt;D2.mdx&lt;/sup&gt;/J Mouse Strain</td><td>Jackson Labs, Bar Harbor, ME</td><td>JAX:13141</td><td>D2.mdx</td></tr><tr><td>DBA/2J Mouse Strain</td><td>Jackson Labs, Bar Harbor, ME</td><td>JAX:00671</td><td>WT</td></tr><tr><td>Dimethyl Sulfoxide (DMSO)</td><td>Fisher Scientific; Hampton, NJ, USA</td><td>BP231-100</td><td>Mitochondrial stain reconstitution reagent</td></tr><tr><td>Hoechst 33342</td><td>Thermo Fisher Scientific</td><td>H1399</td><td>Cell-permeant nucleic acid stain</td></tr><tr><td>ImageJ/Fiji&amp;nbsp;</td><td>National Institutes of Health, Bethesda, MD, USA</td><td>SCR_002285</td><td>2D analysis software</td></tr><tr><td>Invitrogen ProLong Gold antifade reagent with DAPI&amp;nbsp;</td><td>Fisher Scientific, Hampton, NJ, USA</td><td>P36941</td><td>Mounting medium</td></tr><tr><td>KimWipe</td><td>Kimberly-Clark, Irving, TX, USA</td><td>06-666A</td><td>Laboratory wipes</td></tr><tr><td>Leica LAS_X software&amp;nbsp;</td><td>Leica Biosystems, Wetzlar, Germany</td><td>SCR_013673</td><td>Confocal software</td></tr><tr><td>MitoView Fix 640</td><td>Biotium</td><td>70075-50ug</td><td>Far-red fluorescent mitochondrial stain</td></tr><tr><td>NIS-Elements Advanced Research software&amp;nbsp;</td><td>Nikon, Melville, NY, USA</td><td>SCR_014329</td><td>Cross sectional area analysis software</td></tr><tr><td>Permount Mounting Medium&amp;nbsp;</td><td>Fisher Scientific, Waltham, MA 02454, USA</td><td>#SP15-100</td><td>General mounting media</td></tr><tr><td>Prism 9 software&amp;nbsp;</td><td>GraphPad Software, La Jolla, CA, USA</td><td>RRID: SCR_002798</td><td>Statistical analysis software</td></tr><tr><td>Rhodamine (TRITC)</td><td>Fisher Scientific; Hampton, NJ, USA</td><td>AP132RMI</td><td>Secondary antibody</td></tr><tr><td>Stellaris 5 White Light Confocal Microscope and corresponding deconvolution software</td><td>Leica Microsystems, Wetzlar, Germany</td><td>158301313:158201310:158004752:&lt;br/&gt; 158301312:158301314:158401100:&lt;br/&gt; 158209011:158301130:158204510:&lt;br/&gt; 158301120:158401140:158204511:&lt;br/&gt; 15500332:15506428:15506517:155&lt;br/&gt; 13859:15521522:15525226:155252&lt;br/&gt; 32:15525314:15555009:15555017:&lt;br/&gt; 15555046:27100022:155933660:&lt;br/&gt; 155933666:155933668:158000640&lt;br/&gt; :158002407:158003150:158004141&lt;br/&gt; :158004201:158004421:158004423&lt;br/&gt; :158200681:158202140:158203114:&lt;br/&gt; 158203200:158203220:158204201:&lt;br/&gt; 158204510:158204709:158301200:&lt;br/&gt; 9I_LL_STELLARIS_C:9T-CLSM-APP&lt;br/&gt; _CLASSC:9WE_LAS_405:9WE_&lt;br/&gt; LAS_WLL:9WE_LL_STELLARIS_C</td><td>Confocal microscope</td></tr><tr><td>SZ61</td><td>Olympus, Breinigsville, PA, USA</td><td>sz61</td><td>Stereo microscope</td></tr><tr><td>Tissue-Plus O.C.T. Compound&amp;nbsp;</td><td>Scigen, Fischer Scientific, Hampton, NJ, USA</td><td>23-730-571</td><td>Optimal cutting temperature compound</td></tr><tr><td>Visikol I</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>H1-30</td><td>Tissue clearing solution 1</td></tr><tr><td>Visikol II</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>H2-30</td><td>Tissue clearing solution 2</td></tr><tr><td>&amp;alpha;-bungarotoxin 552</td><td>Biotrend, Koln, Germany</td><td>00014</td><td>Acetylcholine receptor dye</td></tr></tbody></table>

Mitochondrial network morphology is widely used as an indicator of cellular health; however, quantifying mitochondrial architecture within intact tissues remains technically challenging. Terminal Schwann cells (tSCs), which are essential for the maintenance and regeneration of neuromuscular junctions, are particularly difficult to analyze in situ due to their anatomical location and sensitivity to tissue disruption. This protocol describes a reproducible approach for labeling and quantifying three&#8211;dimensional mitochondrial network morphology in whole-mount skeletal muscle and in tSCs in mice using standard confocal microscopy. The method employs in vivo delivery of a membrane potential&#8211;sensitive mitochondrial dye followed by rapid tissue processing and high-resolution confocal imaging. Image stacks are analyzed to quantify mitochondrial network connectivity, area, and fragmentation. The protocol is first validated in dystrophic and healthy skeletal muscle to confirm expected differences in mitochondrial morphology and is subsequently adapted to visualize and quantify mitochondrial networks in tSCs identified using S100&#946; reporter mice. This approach enables the analysis of mitochondrial morphology within intact neuromuscular tissues without requiring transgenic mitochondrial reporters or specialized imaging platforms. The protocol requires only equipment commonly available in university core facilities and can be adapted to other thin or superficially accessible tissues.

Mitochondrial networks displayed expected reductions in connectivity 
Representative 3D fluorescent images of TA muscle confocal scans in two groups, WT and mdx, are presented (Figure 2). In comparison with the WT group, mitochondrial connectivity appears reduced in mdx mice compared to the WT (Figure 2A–D). Greater numbers of small mitochondrial fragments were observed in mdx mice [&lt;100 µm2] (WT, 17.3 ± 3.5%; mdx, 36.3 ± 4.9%; P = 0.015) and reduced numbers of large interconnected mitochondrial networks [1000 µm2] compared to WT mice (WT, 23.5 ± 0.26%; mdx, 16.8 ± 1.02%; P = 0.001) (Figure 3A,B).
tSC imaging in situ 
Fluorescent confocal microscopy revealed mitochondrial morphology of tSCs in situ. Both tSC and mitochondrial morphologies are preserved following labeling, rapid removal, sample preparation, and imaging (Figure 4A–C). Mitochondria of tSCs were observed to have interconnected loops (Figure 4C,D) near bifurcations. In contrast to the interconnected nature of tSC mitochondria from WT mice, mdx tSC mitochondria were fragmented, lacking loops near bifurcations (Figure 4E,F).
Further validation of tSC mitochondrial morphology was performed qualitatively comparing fluorescent confocal imaging to focused ion beam scanning electron microscopy, demonstrating similar doughnut shapes in tSC mitochondria (Figure 5A–D). Representative imaging of green tSCs with red mitochondria from WT mice was included for reference (Figure 5E). Analysis of post-synaptic components revealed subsarcolemmal mitochondria, void of the characteristic pretzel-like configuration, but clustered around AChRs embedded upon the surface of muscle fibers (Figure 5F–H).
Image analysis revealed that (green) tSCs comprised about 367 ± 27.0 μm2 and (red) mitochondria make up ~31 ± 2.49% of tSCs (Figure 6A,B). Comparing tSC mitochondria from WT and mdx, tSC mitochondria from mdx muscle occupied a greater area than WT [WT 111 ± 10.1 μm2; mdx 271 ± 25.9 μm2; P &lt; 0.001] (Figure 6C). mdx tSC mitochondria were also greatly fragmented compared to WT [WT 207 ± 18.4; mdx 518 ± 77.1; P &lt; 0.001] (Figure 6D). However, the relative size of the particles did not vary between groups [WT 0.53 ± 0.03; mdx 0.54 ± 0.05; P = 0.836] (Figure 6E).
D2.mdx mouse functional deficits for comparison 
To evaluate muscle pathology in diseased mice, muscle force was measured via direct electrical stimulation. Consistent with previous reports26, muscle twitch (Tw) (g) and peak muscle force (P0) (g) produced by TA muscles of mdx mice were significantly less than the WT group, with greater variability observed in mdx Tw (means ± S.E.M: Tw, WT 38 ± 4.426 g; mdx, 23 ± 15.33 g; P = 0.035; P0, WT 137.0 ± 4.43 g; mdx, 78.59 ± 15.33 g; P = 0.006) (Figure 7A,B). Myofiber CSA was binned into 400 μm2 increments to create a relative frequency histogram plot for WT and mdx groups. Force production across increasing stimulation frequencies was also significantly reduced in mdx TA muscles compared to WT (Figure 7C). Consistent with reports, the mdx phenotype shifts the CSA relative frequency curve to the left of WT with an almost 60% reduction in average fiber size (Figure 7D–F).
Muscle function and histology correlate with mitochondrial area and connectivity 
Linear regression analysis was performed to determine whether markers of muscle pathology were associated with mitochondrial morphology. No significant correlation was observed between TA muscle CSA and the frequency of mitochondrial networks smaller than 100 µm2 or approximately 500 µm2, although slight trends were noted (Figure 8A,B). In contrast, mitochondrial networks with an area ≥1000 µm2 showed a significant positive correlation with TA muscle CSA (Figure 8C,D). Similarly, TA muscle force showed only weak or non-significant relationships with mitochondrial networks &lt;100 µm2 and ~500 µm2 (Figure 8E,F). However, mitochondrial networks ≥1000 µm2 were significantly correlated with TA muscle force (Figure 8G,H). Analysis of tSC mitochondrial morphology in the GM muscle revealed that tSC mitochondrial area was inversely correlated with TA muscle force (Figure 8I), while tSC mitochondrial fragmentation also correlated with TA muscle force (Figure 8J). Furthermore, tSC mitochondrial area correlated with TA muscle cross-sectional area (CSA) (Figure 8K), whereas tSC mitochondrial fragment area did not show a significant relationship with CSA (Figure 8L).
<img alt="Mouse surgical experiment diagram with anesthesia, injection, dissection, imaging process timeline." class="xfigimg" src="/files/ftp_upload/69732/69732fig1v2.jpg" data-altupdatedat="1785816036000" data-alt="Figure 1"> 
Figure 1: Laboratory workflow for TA injections. (A) Left TA muscle was injected with Coomassie blue dye to demonstrate dye spread. Injections began with syringe insertion at the distal tendon of the TA and continued insertion pointed at the TA origin. Injections were made while the syringe was withdrawn along the injection path (yellow bar). (B) Timeline of TA injection, dissection, and imaging. Laboratory workflow for GM injections. (C) Timeline of GM injection, dissection, and imaging. (D) GM shaving and animal preparation. (E) Inhaled anesthesia with a temperature probe between the mouse and the heat lamp. A 3 mm incision along the lumbar spine. (F) GM injection made directly beneath the GM. (G) 3 mm incision closed with 6-0 nylon suture during the 45 min incubation period. <a href="https://www.jove.com/files/ftp_upload/69732/69732fig1largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Confocal microscopy image showing mitochondria and nuclei in muscle fibers, WT vs mdx comparison." class="xfigimg" src="/files/ftp_upload/69732/69732fig2v2.jpg" data-altupdatedat="1785816036000" data-alt="Figure 2"> 
Figure 2: Representative images of TA muscle confocal scans from WT and mdx mice. (A,B) 3D mitochondrial scans (red) with nuclei (blue) from WT and mdx mouse muscle. (C,D) One micrometer-thick optical sections of mitochondria (white) from healthy and dystrophic muscle. Scale bar = 25 µm. <a href="https://www.jove.com/files/ftp_upload/69732/69732fig2largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Mitochondrial area comparison, Bar graphs, WT vs mdx, Cellular biology, Statistical analysis." class="xfigimg" src="/files/ftp_upload/69732/69732fig3v2.jpg" data-altupdatedat="1785816036000" data-alt="Figure 3"> 
Figure 3: Quantification of mitochondrial morphological parameters in WT and mdx mouse TA muscle. (A) Area of mitochondria measured throughout a Z-stack. Statistics: Student's t-test. (B) Relative frequency of occurrence of discrete mitochondria with areas binned by 500 µm2. Statistics: Student's t-test. <a href="https://www.jove.com/files/ftp_upload/69732/69732fig3largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Fluorescence microscopy image of mitochondria and tSC in muscle cells, showing structural differences." class="xfigimg" src="/files/ftp_upload/69732/69732fig4v2.jpg" data-altupdatedat="1785816036000" data-alt="Figure 4"> 
Figure 4: tSC mitochondrial morphology from GM muscle. (A–C) tSC in green and mitochondria in red. Scale bar = 20 µm. (D) tSC mitochondria in red from D2.mdx mouse GM. Scale bar = 2.5 µm. Close-up images of mitochondrial bifurcations; yellow arrows depict interconnected loops of mitochondria at each bifurcation. Scale bar = 20 µm. (E) tSC mitochondria from D2.mdx mice. (F) Close image of D2.mdx tSC mitochondria, yellow arrows highlight fragmented mitochondria. Scale bar = 2.5 µm. <a href="https://www.jove.com/files/ftp_upload/69732/69732fig4largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Mitochondria analysis: microscopy images, Schwann cells, loop structure, nuclei labeling, cellular study." class="xfigimg" src="/files/ftp_upload/69732/69732fig5v2.jpg" data-altupdatedat="1785816036000" data-alt="Figure 5"> 
Figure 5: Validation of tSC mitochondrial imaging in GM muscle. (A) Zoomed in view of mitochondrial doughnut (yellow arrow) with confocal microscopy. Scale bar = 1 µm. (B–D) Images of a tSC mitochondrial doughnut at multiple angles using focused ion beam scanning electron microscopy. Scale bar = 0.5 µm. (E) Image of green tSC with red mitochondria and adjacent red mitochondria in a neighboring muscle fiber. Scale bar = 50 µm. (F) 3D image of AChR (teal) and nuclei (blue), (G) Sub-sarcolemma mitochondria (red), and nuclei, (H) merged. Scale bar = 20 µm. <a href="https://www.jove.com/files/ftp_upload/69732/69732fig5largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Bar chart analysis of tSC metrics in WT vs. mdx for mitochondrial research results." class="xfigimg" src="/files/ftp_upload/69732/69732fig6v2.jpg" data-altupdatedat="1785816036000" data-alt="Figure 6"> 
Figure 6: tSC mitochondrial morphology parameters quantified. Individual dots represent individual tSCs in GM muscles. (A) Size of tSCs in WT mice reported in μm2. (B) Percent of tSC area occupied by mitochondria. (C) tSC mitochondrial area in WT and mdx mice, indicating greater area occupied by tSCs in mdx. (D) tSC number of mitochondrial fragments compared between WT and mdx, indicating greater fragmentation in mdx. (E) Average fragment size is unchanged in mdx tSC compared with WT. Statistics: Student's t-test. <a href="https://www.jove.com/files/ftp_upload/69732/69732fig6largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Static equilibrium analysis; bar graphs, muscle fiber images, and size distribution chart; WT vs. mdx." class="xfigimg" src="/files/ftp_upload/69732/69732fig7v2.jpg" data-altupdatedat="1785816036000" data-alt="Figure 7"> 
Figure 7: Comparisons of pathological markers between WT and mdx mouse TA muscles. (A) TA muscle twitch (Tw). (B) Peak Muscle Force (P0), and (C) force frequency curve. Muscle functional parameters were reduced in the mdx TA muscle compared to the WT. (D) Representative images of TA muscle cross sections with laminin (white) and myofiber interiors (red). Scale bar = 100 µm. (E,F) Fiber size is significantly reduced, and the fiber size frequency of occurrence was shifted to the left in mdx mice compared to WT. Statistics: Student's t-test. <a href="https://www.jove.com/files/ftp_upload/69732/69732fig7largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Mitochondrial area vs muscle CSA and peak force graphs; correlation analysis; biological data evaluation." class="xfigimg" src="/files/ftp_upload/69732/69732fig8v2.jpg" data-altupdatedat="1785816036000" data-alt="Figure 8"> 
Figure 8: Relationships between mitochondrial morphology and markers of mdx muscle pathology. (A–D) Linear regression analysis of TA mitochondrial network size distribution relative to TA muscle cross-sectional area (CSA). Mitochondrial areas &lt;100 µm2 (A) and ~500 µm2 (B) showed no correlation with CSA, whereas networks ≥1000 µm2 (C,D) were positively correlated with CSA. (E–H) Linear regression analysis of mitochondrial network size distribution relative to TA muscle force. Mitochondrial areas &lt;100 µm2 (E) and ~500 µm2 (F) showed weak or no correlation with force, whereas networks ≥1000 µm2 correlated with TA muscle force (G,H). (I–L) Relationships between mitochondrial morphology in tSCs from the GM and TA muscle pathology. tSC mitochondrial area correlated with TA muscle force (I) and CSA (K), while mitochondrial fragmentation correlated with TA muscle force (J) but not CSA (L). Statistics: Linear regression with Pearson’s r correlation. <a href="https://www.jove.com/files/ftp_upload/69732/69732fig8largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Local injections of a mitochondrial dye label terminal Schwann cell mitochondria in vivo for confocal microscopy. This method utilizes proof of concept demonstrated in skeletal muscle tissue from mice with healthy or diseased muscle to provide high-resolution mitochondrial visualization.

methods-for-situ-quantification-mitochondrial-morphology-muscle

Methods for In situ Quantification of Mitochondrial Morphology in Muscle and Terminal Schwann Cells of Mice

Mitochondrial network morphology is widely used as an indicator of cellular health; however, quantifying mitochondrial architecture within intact tissues ...

Our research investigates mitochondrial morphology of terminal Schwann cells at the neuromuscular junction for the first time. Our approach enables quantification of mitochondrial morphology in otherwise hard-to-obtain tissues, such as capillary and lymphatic beds. To begin, obtain a wild type and a D2-mdx mouse.After confirming the anesthetic depth by toe-pinch reflex, insert the needle at the distal tendon toward the right knee. Inject 50 microliters of two-micromolar far red fluorescent mitochondrial stain on the tibialis interior, or the TA, muscle while withdrawing the needle along the injection path, and incubate the dye in vivo for one hour. While maintaining the animal under anesthesia, use scissors and forceps to dissect and isolate the TA muscle prior to euthanasia.Place the isolated TA muscle in 4%paraformaldehyde at room temperature for 15 minutes. Then, under a stereo microscope, gently separate the fibers by teasing. Add cell-permeant nucleic acid stain to the 4%paraformaldehyde to obtain a final concentration of five to 10 micrograms per milliliter for nuclear labeling.Then, wash the TA muscle three times in PBS for five minutes each to remove excess dye. Dehydrate the muscle sequentially in 50%75%and 100%ethanol for five minutes each. Then, incubate the sample sequentially in tissue-clearing solutions one and two for 30 minutes each, ensuring they are fully submerged.Place the muscle on a coverslip. Use a 2.5-by-2-by-1-centimeter glass block weighing 7.8 grams to gently flatten it. Obtain an S100 beta mouse.After confirming the anesthetic depth by toe-pinch reflex, make a small skin incision above the gluteus maximus, or GM, muscle. Inject 75 microliters of two-micromolar far red fluorescent mitochondrial stain beneath the GM muscle. Close the wound and let the dye incubate for 45 minutes to selectively label terminal Schwann cells.While maintaining the animal under anesthesia, dissect the GM muscle within 10 minutes, and immediately place it in chilled 0.9%sterile saline. Only if using D2-mdx mice, add fluorescent acetylcholine receptor dye at a one-to-500 dilution to the microdissection bath, and incubate the tissue for 30 minutes. Trim the excess fat and connective tissue, and transfer the GM muscle onto a coverslip with the ventral side facing down for inverted microscopy.Add two microliters of general mounting medium onto the dorsal surface of the GM, and flatten the tissue using glass block before imaging. In the software, set the excitation wavelength to 646 nanometers. After adding two drops of clearing solution two, select the 63X objective lens to image the TA muscle, and set the Z stack depth close to 100 micrometers.Then, select deconvolution mode, and apply system optimized parameters. Set the step size to one micrometer and acquire images. To acquire terminal Schwann cells'images, select the 20 times magnification objective lens, set the Z stack depth to approximately 17 micrometers, and acquire images.Then, select the 63 times magnification objective lens and acquire images of individual terminal Schwann cells. Import individual Z slices to train the software to recognize the mitochondrial signal. And conduct quantitative analysis using AI-driven software.Develop image masks for investigator review. Separate 3D image stacks into individual optical sections following algorithm optimization. Select every fifth Z slice from each 3D image, and calculate the average per sample.Standardize imaging parameters across all samples, and quantify mitochondrial morphology. Perform immunohistochemistry and statistical analysis to assess group differences and correlations. Mitochondrial connectivity was reduced in mdx muscle compared to the wild type mice.A greater number of small mitochondrial fragments and a reduced number of large interconnected mitochondrial networks were observed in mdx mice. In wild type mice, mitochondria in terminal Schwann cells exhibited interconnected loop structures near bifurcations. In contrast, in mdx mice, mitochondria appeared fragmented and lacked loop formations near bifurcations.Confocal and electron microscopy confirmed similar donut-shaped mitochondrial structures in terminal Schwann cells. Postsynaptic analysis showed sub-sarcolemmal mitochondria clustered around acetylcholine receptors, lacking the characteristic pretzel-like structure. Image analysis revealed that terminal Schwann cell mitochondria in mdx muscle showed increased area and fragmentation compared to wild type muscle, with no difference in relative particle size.Muscle force measurements showed reduced twitch with greater variability and decreased peak force in mdx muscle. Force production across increasing stimulation frequencies was also significantly reduced in mdx muscles. The mdx muscle showed a leftward shift in fiber size, indicating approximately 60%smaller myofibers compared to wild type.This protocol allows researchers to study and quantify mitochondrial morphology in skeletal muscle and terminal Schwann cells. Future studies can adapt this approach to investigate mitochondrial morphology in micro-circulation and lymphatic vessels.

Methods for In situ Quantification of Mitochondrial Morphology In Muscle and Terminal Schwann Cells 

Kleiton A. S. Silva

Sarcopenia is a muscle disease marked by the gradual, progressive, generalized, and inherent loss of muscle mass and function due to aging. This condition increases the risk of falls, fractures, frailty, prolonged hospital stays, and mortality. Currently, there are no FDA-approved drugs to treat this degenerative muscle condition, and regular exercise training remains the only recommended therapy. Although sarcopenia is a significant clinical issue, experts in skeletal muscle continue to seek ways to evaluate and treat the condition.This video-based collection of methods will focus on approaches that assess the overall physical and motor function of animals. It will cover exercise prescriptions for sarcopenic animals, both in vivo and ex vivo muscle function, the characterization of aged myofibers, and the ultrastructure and functionality of mitochondria.

JoVE Medicine serves as a connection between basic research and clinical applications. Articles may present medical procedures, case studies or clinical trial methodologies. Included studies are often modeled towards predictive, preventative, personalized, and practical approaches to treatment of disease. The goal of research methodologies in this section is to improve overall patient care and prognosis.

Medicine

Mighty Muscle: Traditional and Contemporary Methods to Identify and Understand Sarcopenia

Sarcopenia is a muscle disease marked by the gradual, progressive, generalized, and inherent loss of muscle mass and function due to aging. Curated by Kleiton A. S. Silva. Explore Medicine techniques on JoVE.

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Methods for In situ Quantification of Mitochondrial Morphology in Muscle and Terminal Schwann Cells of Mice