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

Summary

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.

Abstract

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–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–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 S100β 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.

Introduction

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 integrated^2,3,4, 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 mitophagy^2,3. Mitochondrial biogenesis is required to maintain and expand healthy mitochondria. Metabolic demand, such as physical exercise, increases expression of the peroxisome proliferator-activated receptor γ co-activator 1α (PGC-1α)⁵. This process occurs via primary endurance signals, specifically the exercise-induced elevation of AMP-activated protein kinase (AMPK), calcium, and reactive oxygen species^6,7. Fusion expands networks, promoting survivability and complementing deficits, while fission mitigates further mitochondrial damage through the preferential exclusion of dysfunctional region^2,3,8. Once removed, excluded regions must undergo mitophagy, preventing the accumulation of dysfunctional organelles and proteins^6,9.

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 quality^3,10. In that regard, during prolonged muscle dysfunction, mitochondrial respiration is reduced, while ROS are increased, and networks are fragmented^3,10. 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 “pinched” off, promoting fragmentation^3,11. 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 muscle^2,12.

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 increase^13,14. Following nerve injury by transection, crush, or stretch, SCs dedifferentiate into a demyelinated progenitor-like state^15,16. 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 degeneration^17,18. 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 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 labeled^5,24. Live-cell imaging typically requires incubating cells for 20–60 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 dispersion^25,26,27, and suitability for concurrent functional measurements. The gluteus maximus (GM) muscle was selected for tSC imaging based on its established injection procedure^28,29, 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.

Protocol

Six-month-old (n=3) adult male DBA/2J [Wild-type (WT)] (RRID: IMSR_JAX:000671) and D2.B10-DMD^D2.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&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 µM

1. Suspend the mitochondrial stain (50 µg) in 322 µL of dimethyl sulfoxide (DMSO) according to the manufacturer’s instructions to prepare a 200 µM stock solution.
2. To prepare 30 mL of 2 µM working solution, transfer 300 µL of 200 µM stock solution into 29.7 mL of sterile saline. Mix thoroughly.
3. Aliquot 250 µL of the 2 µM working solution into microcentrifuge tubes and store at -80 °C until use.
   NOTE: No detectable signal degradation was observed after 3 years of storage at -80 °C.

2. Dye and animal preparation

1. Thaw one frozen aliquot 15 min prior to experimentation. Draw the solution into a 30 G, 1 cc insulin syringe.
2. 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.
3. Clean the shaved injection site with 70% isopropyl alcohol.

3. Tibialis anterior (TA) muscle mitochondrial dye administration and function

1. TA dye administration
   1. Under anesthesia, inject 50 µL of 2 µM mitochondrial stain into the Left or Right TA muscle of anesthetized WT and D2.mdx mice.
   2. Insert the needle at the distal tendon toward the knee and depress the plunger while withdrawing the needle along the injection path.
   3. Incubate the dye in vivo for 1 h prior to dissection.
2. TA muscle function measurements
   1. After 45 min of dye incubation, anesthetize the mouse using isoflurane (2%–3% induction, 1.5%–2% maintenance in oxygen).
   2. Surgically expose the left TA muscle while preserving surrounding tissues. Tie the distal tendon to the load beam of the muscle testing system.
   3. Wrap the exposed muscle with a laboratory wipe and irrigate at 3 mL min⁻¹ with 0.9% sterile saline warmed to 40 °C. Maintain muscle temperature at 32 °C using a heat lamp.
   4. Position platinum/iridium wire electrodes across the muscle belly for direct stimulation.
   5. Attach the load beam to a micrometer and adjust optimal length (L₀) using twitch contractions (T_w) at 1 Hz. Record L₀ with a digital caliper.
   6. 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.
3. TA tissue processing
   1. After incubation, isolate and dissect the TA muscle prior to euthanasia.
   2. 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).
   3. Add cell-permeant nucleic acid stain to a final concentration of 5–10 µg/mL to label nuclei.
   4. Wash samples three times in phosphate-buffered saline (PBS) for 5 min each to remove excess dye.
   5. Dehydrate samples sequentially in 50%, 75%, and 100% ethanol for 5 min each.
   6. Incubate samples sequentially in tissue clearing solutions I and II for 30 min each at room temperature using sufficient volume to fully submerge tissue.
   7. Place cleared TA bundles on a coverslip and gently flatten using a glass block (2.5 cm × 2 cm × 1 cm; mass 7.8 g)²⁶.

4. Imaging of TA muscle samples

1. Image muscle samples with a confocal microscope to obtain high-resolution three–dimensional (3D) optical sections.
2. Acquire Z-stack images at a depth of approximately 100 µm for muscle using a 63x magnification objective lens (N.A. 1.40).
3. Use the imaging system’s confocal software with an excitation wavelength of 646 nm to provide the greatest clarity with the least background.
4. Acquire images using deconvolution mode, employing “System Optimized” parameters with 1 µm 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.

5. GM muscle tSC dye loading and imaging

1. Use S100β mice for specific fluorescent identification of tSCs.
2. Under anesthesia, perform a small skin incision above the GM muscle. Inject 75 μL of a 2 μM far-red fluorescent mitochondrial stain beneath the GM, prior to wound closure²⁵.
3. Close the wound and allow the dye to incubate for 45 minutes to selectively label tSCs without significant labeling of the underlying muscle fibers.
4. Dissect the GM muscles and immediately place them in chilled 0.9% sterile saline. Rapidly trim excess fat and connective tissue.
5. Transfer the dissected GMs onto a coverslip for imaging.
6. 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).
7. 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.
8. Perform confocal microscopy to obtain high-resolution three–dimensional (3D) optical sections.
9. Acquire Z-stack images at a depth of approximately 16 µm 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.
10. 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.
11. Apply 2 µL of general mounting medium onto the dorsal surface of the GM.
12. Gently flatten the tissue using a calibrated glass block (2.5 cm × 2 cm × 1 cm; mass 7.8 g) to ensure a uniform imaging plane.

6. Focused ion beam microscopy

1. Perform focused ion beam scanning electron microscopy (FIB-SEM) according to previously established protocols²⁶.
   1. Apply FIB-SEM specifically to confirm and visualize novel morphological features identified during confocal imaging.

7. Image analysis

1. TA muscle samples
   1. Conduct quantitative analysis of confocal optical sections (1 μm) of TA scans (approximately 75–112 μm thick) using AI-driven image analysis software.
      ​NOTE: Perform multiple supervised training sessions prior to automated segmentation to enhance algorithm adaptation.
   2. Import single optical sections (Z-slices) to teach the software to retain pixelated information representing white mitochondrial dye while excluding the black background.
   3. Develop image masks for investigator review to ensure high-quality analysis and accuracy.
   4. Separate 3D image stacks into single optical sections following algorithm optimization.
   5. Select every fifth Z-slice from each 3D image for analysis, with each data point representing the average of these individual sections per sample.
   6. Quantify mitochondrial morphology to derive spatial distribution and network connectivity metrics.
   7. Standardize intensity thresholds, edge detection settings, and voxel size calibration across all samples to minimize experimental variability.
2. tSCs on GM samples
   1. Assemble 1 μm optical sections into Z-stacks (approximately 6–25 μm thick) and separate images into individual color channels using 2D image analysis software.
   2. Calibrate software using scale bars and apply thresholding to optimize tSC mitochondria visualization with minimal background interference.
   3. Perform binarization and remove background noise to isolate tSC mitochondria or tSCs for the measurement of area and particle counts.
   4. Save binary images to ensure consistent processing and perform all imaging experiments under identical conditions to ensure reproducibility across biological replicates.

8. Immunohistochemistry

1. Embed left TA muscle samples in optimal cutting temperature (OCT) compound to evaluate cross-sectional area (CSA).
   1. Section samples at 10 μm thickness using a cryostat set to -17°C and collect onto microscope slides.
   2. 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).
   3. Mount slides using an appropriate mounting medium prior to imaging.
2. Image slides using a confocal microscope by randomly selecting 2–3 regions of interest (580 x 580 µm) per sample with a 20x (N.A. 0.75) objective.
3. Average values across regions per muscle and analyze approximately 400 myofibers per TA muscle section.
4. Use semi-automatic muscle segmentation software to determine CSA (µm²) by masking the myofibers.

9. Statistical analysis

1. Report summary data as mean ± standard error of the mean (S.E.M.).
2. Perform statistical analysis using a dedicated statistical software, utilizing parametric, unpaired Student’s t-tests to determine significance among group mean differences.
3. 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.
4. Assess Pearson r correlation coefficients to determine the correlation between DMD pathology and mitochondrial pathology. Define statistical significance as P ≤ 0.05.

Results

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 [<100 µm²] (WT, 17.3 ± 3.5%; m...

Discussion

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...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
3-in-1 Whole Animal System	Aurora Scientific, Ontario, Canada	1300A	Muscle funciton system with analysis software
AIVIA	Leica Microsystems, Bellevue, WA, USA	15	AI driven image analysis software
anti-laminin	MilliporeSigma	L9393-.2mL	myofiber boarder label
B6; D2-Tg(S100-EGFP)1Wjt/J Mouse Strain	Jackson Labs, Bar Harbor, ME	JAX:005621	S100β
Cryostar NX50 Cryostat	Epredia, Kalamazoo, MI, USA	https://www.epredia.com/products/histology-instruments/cryotomy/cryostar-nx50	Cryostat
D2.B10-DMDD2.mdx/J Mouse Strain	Jackson Labs, Bar Harbor, ME	JAX:13141	D2.mdx
DBA/2J Mouse Strain	Jackson Labs, Bar Harbor, ME	JAX:00671	WT
Dimethyl Sulfoxide (DMSO)	Fisher Scientific; Hampton, NJ, USA	BP231-100	Mitochondrial stain reconstitution reagent
Hoechst 33342	Thermo Fisher Scientific	H1399	Cell-permeant nucleic acid stain
ImageJ/Fiji	National Institutes of Health, Bethesda, MD, USA	SCR_002285	2D analysis software
Invitrogen ProLong Gold antifade reagent with DAPI	Fisher Scientific, Hampton, NJ, USA	P36941	Mounting medium
KimWipe	Kimberly-Clark, Irving, TX, USA	06-666A	Laboratory wipes
Leica LAS_X software	Leica Biosystems, Wetzlar, Germany	SCR_013673	Confocal software
MitoView Fix 640	Biotium	70075-50ug	Far-red fluorescent mitochondrial stain
NIS-Elements Advanced Research software	Nikon, Melville, NY, USA	SCR_014329	Cross sectional area analysis software
Permount Mounting Medium	Fisher Scientific, Waltham, MA 02454, USA	#SP15-100	General mounting media
Prism 9 software	GraphPad Software, La Jolla, CA, USA	RRID: SCR_002798	Statistical analysis software
Rhodamine (TRITC)	Fisher Scientific; Hampton, NJ, USA	AP132RMI	Secondary antibody
Stellaris 5 White Light Confocal Microscope and corresponding deconvolution software	Leica Microsystems, Wetzlar, Germany	158301313:158201310:158004752: 158301312:158301314:158401100: 158209011:158301130:158204510: 158301120:158401140:158204511: 15500332:15506428:15506517:155 13859:15521522:15525226:155252 32:15525314:15555009:15555017: 15555046:27100022:155933660: 155933666:155933668:158000640 :158002407:158003150:158004141 :158004201:158004421:158004423 :158200681:158202140:158203114: 158203200:158203220:158204201: 158204510:158204709:158301200: 9I_LL_STELLARIS_C:9T-CLSM-APP _CLASSC:9WE_LAS_405:9WE_ LAS_WLL:9WE_LL_STELLARIS_C	Confocal microscope
SZ61	Olympus, Breinigsville, PA, USA	sz61	Stereo microscope
Tissue-Plus O.C.T. Compound	Scigen, Fischer Scientific, Hampton, NJ, USA	23-730-571	Optimal cutting temperature compound
Visikol I	Sigma-Aldrich, St. Louis, MO, USA	H1-30	Tissue clearing solution 1
Visikol II	Sigma-Aldrich, St. Louis, MO, USA	H2-30	Tissue clearing solution 2
α-bungarotoxin 552	Biotrend, Koln, Germany	00014	Acetylcholine receptor dye