Isolation, Purification, and Characterization of Functional Mitochondria Derived from Mouse Skeletal Muscle for Mitochondrial Transplantation

Summary

A standardized protocol combining trypsin digestion and differential centrifugation was established for the isolation, purification, and functional characterization of mitochondria from mouse skeletal muscle, as well as for evaluating their application in mitochondrial transplantation.

Abstract

Mitochondria are essential organelles that regulate energy metabolism, signal transduction, and cellular homeostasis in eukaryotic cells. Mitochondrial dysfunction contributes to the pathogenesis of numerous diseases and has prompted the development of mitochondrial transplantation as a regenerative therapeutic strategy. The successful application of mitochondrial transplantation depends on the availability of highly purified and functionally intact mitochondria. Skeletal muscle is a suitable donor source due to its high mitochondrial content, metabolic activity, and accessibility. This study established a standardized, reproducible protocol for the isolation, purification, and characterization of functional mitochondria from mouse skeletal muscle and evaluated their use in mitochondrial transplantation. The procedure consisted of two major stages. First, mitochondria were isolated from the skeletal muscle of C57BL/6 mice using trypsin digestion followed by differential centrifugation. Second, the isolated mitochondria were characterized to evaluate purity, ultrastructure, and functional activity. Mitochondrial purity was assessed by bicinchoninic acid (BCA) protein quantification and Western blot analysis. Ultrastructural integrity was examined by transmission electron microscopy. Functional activity was evaluated using JC-1 and mitochondrial fluorescent labeling together with measurements of oxygen consumption, ATP production capacity, and respiratory control ratio using a high-resolution respirometry system. The isolated mitochondria exhibited preserved membrane potential, intact ultrastructure, and stable respiratory activity, indicating suitability for downstream functional studies and mitochondrial transplantation applications.

Introduction

Mitochondria are the primary sites of adenosine triphosphate (ATP) production and play central roles in apoptosis, calcium homeostasis, and the metabolism of reactive oxygen species (ROS). The functional integrity of mitochondria is therefore critical for maintaining cellular homeostasis and determining cell fate^1,2. Mitochondrial dysfunction, including mitochondrial DNA (mtDNA) mutations and electron transport chain (ETC) abnormalities, leads to impaired energy metabolism, oxidative stress, and dysregulated apoptosis^3,4. These abnormalities are associated with a broad range of diseases, including inherited mitochondrial disorders, neurodegenerative diseases, and cardiovascular diseases^5,6. Conventional therapeutic strategies for mitochondrial dysfunction, such as antioxidants and metabolic cofactors, generally provide limited efficacy because they alleviate symptoms without directly restoring mitochondrial function^7,8. Consequently, mitochondrial transplantation has emerged as a promising therapeutic approach in regenerative medicine. This strategy involves the delivery of intact exogenous mitochondria into damaged or dysfunctional cells to restore mitochondrial activity and cellular energy metabolism^9,10. Previous studies have demonstrated that exogenous mitochondria can be internalized by recipient cells and incorporated into the endogenous mitochondrial network, resulting in improved bioenergetic function and enhanced cell survival^11,12,13.

Skeletal muscle is considered a suitable donor for mitochondrial isolation because of its high mitochondrial content, metabolic activity, and accessibility^14,15,16. Mitochondrial transplantation requires several critical steps, including mitochondrial isolation, purification, functional characterization, and efficient delivery into recipient cells. However, substantial methodological variability remains among the protocols reported to date, particularly in procedures involving tissue homogenization, enzymatic digestion, temperature control, and purification. Such variability can affect mitochondrial yield, purity, and functional integrity, thereby limiting experimental reproducibility.

To improve methodological consistency and reproducibility, this study presents a standardized protocol for isolating and characterizing functional mitochondria from mouse skeletal muscle using trypsin digestion and differential centrifugation. The isolated mitochondria were evaluated for purity, ultrastructural integrity, and functional activity using bicinchoninic acid (BCA) protein quantification, Western blotting, transmission electron microscopy, fluorescence-based membrane potential assays, and high-resolution respirometry. In addition, the uptake of isolated mitochondria by recipient cells was assessed to evaluate their suitability for mitochondrial transplantation applications. This protocol provides a reproducible workflow for obtaining functional skeletal muscle-derived mitochondria for downstream experimental studies.

Protocol

All animal experiments were performed in a specific pathogen-free (SPF) facility and approved by the Animal Ethics and Use Committee of the Third Affiliated Hospital of Air Force Medical University (Approval No.: IACUC-2024kq-065).

Male C57BL/6 mice (6-8 weeks old, 18-20 g) were purchased from the Laboratory Animal Center of Air Force Medical University (Production License: SCXK (Shaanxi) 2024-002). The mice were housed in individually ventilated cages (IVCs) under positive pressure and clean conditions, with a temperature of 20-26 °C, relative humidity 40%-70%, a 12 h light-dark cycle, and at least 15 air changes per hour. Sterilized diet, drinking water, and environmental enrichment were provided.

The reagents, chemicals, and experimental instruments used in the protocol are listed in the Table of Materials.

1. Animal sacrifice and tissue collection

1. Place the mouse into an airtight euthanasia chamber and euthanize via gradual carbon dioxide (CO₂) inhalation, 30%-70% chamber volume per minute.
2. Confirm death by cessation of spontaneous breathing and heartbeat, loss of pupillary reflexes, and no response to mechanical stimulation.
3. Immerse the euthanized C57BL/6 mice in 75% ethanol for 5 min for disinfection.
   NOTE: To avoid alcohol-induced tissue fixation and cellular damage, limit the immersion time to 10 min. Use fresh tissue throughout the protocol because freezing compromises mitochondrial viability and functional integrity.
4. Place the mouse into a sterile Petri dish within a laminar flow hood with the abdomen facing upward to fully expose the abdominal and hindlimb regions.
5. Lift the skin of the lower abdomen near the genital region using forceps and make a small incision. Perform blunt dissection to separate the skin from the underlying tissue. Extend the incision superiorly along the midline to the ankle and laterally toward both hindlimbs to expose the hindlimb muscle groups completely.
6. Identify the quadriceps femoris muscle on the anterior aspect of the thigh and carefully separate the muscle along the direction of the muscle fibers using blunt dissection. Sever the distal patellar ligament and proximal tendon to isolate the muscle tissue completely.
   NOTE: Wash the tissue three times in pre-chilled phosphate-buffered saline (PBS) supplemented with 2% penicillin-streptomycin using a fresh Petri dish for each wash to minimize contamination.
7. Place the isolated muscle tissue immediately into pre-chilled PBS and rinse thoroughly to remove hair and blood from the tissue surface.
8. Gently blot excess moisture from the tissue using sterile gauze. Divide the tissue into 1.5 mL microcentrifuge tubes and weigh the samples. Use approximately 100 mg of tissue per sample to maintain consistency between preparations.

2. Tissue mincing and homogenization

1. Place the microcentrifuge tube containing the tissue from Step 1.8 on ice. Mince the tissue into small fragments approximately 1-2 mm³ in size using sterile scissors.
2. Add 8 volumes of pre-chilled 0.25% trypsin and digest the tissue fragments on ice for 15 min.
   NOTE: Calculate the required trypsin volume according to tissue mass (1 mg ≈ 1 µL). For example, add 640 µL of 0.25% trypsin to 80 mg of tissue.
3. Centrifuge the mixture at 1,000 × g for 5 min at 4 °C. Discard the supernatant and retain the tissue pellet. Resuspend the pellet in 1 mL pre-chilled mitochondrial isolation buffer containing D-mannitol (215 mM), sucrose (75 mM), HEPES (20 mM), EGTA (1 mM), and fatty acid-free bovine serum albumin (0.5% w/v).
4. Transfer the suspension to a glass homogenizer maintained on ice. Homogenize the tissue using 10-12 gentle up-and-down strokes until the suspension becomes turbid.
   NOTE: Avoid rotating the glass pestle laterally at the bottom of the homogenizer because excessive mechanical force may damage mitochondrial membranes and impair mitochondrial function.

3. Differential centrifugation

1. Transfer the homogenate into a 1.5 mL microcentrifuge tube and centrifuge at 1,000 × g for 5 min at 4 °C.
2. Transfer the supernatant to a fresh 1.5 mL microcentrifuge tube and centrifuge at 12,000 × g for 5 min at 4 °C to obtain the crude mitochondrial pellet.
   NOTE: Carefully collect the supernatant using a 1 mL syringe while avoiding disturbance of the pellet and residual tissue debris.

4. Mitochondrial purification and storage

1. Resuspend the crude mitochondrial pellet gently in pre-chilled wash buffer containing sucrose (250 mM), HEPES (10 mM, pH 7.4), EGTA (1 mM), and bovine serum albumin (0.15% w/v). Centrifuge the suspension at 1,000 × g for 5 min at 4 °C.
2. Transfer the supernatant to a fresh centrifuge tube and centrifuge at 12,000 × g for 10 min at 4 °C to obtain the purified mitochondrial pellet.
   NOTE: Use freshly isolated mitochondria immediately whenever possible. Short-term storage at 4 °C or long-term storage at −80 °C in mitochondrial storage buffer containing sucrose (0.25 M), Tris-MOPS (10 mM, pH 7.4), and EGTA (0.1 mM) is feasible. However, freezing may impair mitochondrial enzymatic activity and respiratory function. Use frozen samples primarily for protein analysis.

5. Mitochondrial and cytosolic protein extraction and BCA quantification

1. Mitochondrial protein extraction
   1. Resuspend the purified mitochondrial pellet in 50 µL of RIPA lysis buffer per 100 mg of starting tissue and incubate the suspension on ice for 15 min.
   2. Centrifuge the lysate at approximately 14,000 × g for 15 min at 4 °C. Transfer the supernatant to a fresh 1.5 mL microcentrifuge tube to obtain the mitochondrial protein fraction.
2. Cytosolic protein extraction
   1. Centrifuge the cytosolic supernatant obtained in Step 4.2 at 12,000 × g for 15 min at 4 °C. Discard the supernatant and retain the pellet as the enriched cytosolic fraction.
   2. Resuspend the cytosolic pellet in 50 µL of RIPA lysis buffer and process the sample as described in Steps 5.1.1-5.1.2.
      NOTE: Prepare the lysis buffer freshly with protease inhibitors and perform all procedures on ice to minimize protein degradation. Mitochondria used in this step can be either fresh or frozen.
3. BCA protein quantification
   1. Prepare the BCA working solution by mixing reagent A and reagent B at a ratio of 50:1 (v/v). Add 200 µL of working solution to each well of a 96-well plate.
      NOTE: Prepare the BCA working solution immediately before use and use it within 1 h at room temperature.
   2. Dilute the bovine serum albumin (BSA) stock solution (2 mg/mL) to 0.5 mg/mL and keep the solution on ice until use.
   3. Prepare the standard curve by adding 20, 19, 18, 16, 12, 8, 4, or 0 µL of BSA solution to separate wells. Adjust the final volume to 20 µL using NaCl solution.
   4. Add 4 µL of mitochondrial protein and 16 µL of NaCl solution to one set of wells. Add 4 µL of cytosolic protein and 16 µL of NaCl solution to another set of wells.
   5. Add 200 µL of BCA working solution to each well and incubate the plate at 37 °C in the dark for 30 min. Measure absorbance at 562 nm using a microplate reader.
   6. Construct a protein standard curve with a correlation coefficient (R²) ≥ 0.999. Calculate protein concentrations (µg/µL) by interpolating absorbance values from the standard curve.

6. Purity assessment of skeletal muscle-derived mitochondria

1. Dilute mitochondrial and cytosolic protein samples to a final concentration of 1.5 µg/µL. Prepare 20 µL reaction mixtures containing 30 µg of protein and 4 × loading buffer. Adjust the remaining volume with NaCl solution. Boil the samples for 10 min and store them at −80 °C until use.
2. Separate protein samples using 4%-20% gradient precast gels. Perform electrophoresis at 140-150 V for 50 min.
3. Activate the PVDF membrane (0.22 µm) in methanol for 1 min. Transfer proteins using the rapid transfer buffer at 400 mA for 30 min.
4. Remove the PVDF membrane from the transfer sandwich and wash it in TBST for 5 min. Block the membrane with 5% skim milk at room temperature for 1 h with gentle shaking.
5. Dilute primary antibodies (COX IV, cytochrome c, β-actin) 1:1,000 in antibody dilution buffer. Incubate the membrane with primary antibodies overnight at 4 °C.
6. Wash the membrane three times with TBST for 10 min per wash.
7. Incubate the membrane with secondary antibody diluted 1:10,000 at room temperature for 1 h.
8. Wash the membrane three times with TBST for 10 min each. Develop the blot and capture the images.

7. MitoTracker staining of skeletal muscle-derived mitochondria

1. Dilute the staining working solution to a final concentration of 200 nM in pre-warmed medium immediately before use.
2. Resuspend the mitochondrial pellet in 100 µL of the working solution and incubate at 37 °C in the dark for 20 min.
3. Add five volumes of pre-chilled storage buffer after incubation. Centrifuge the suspension at 12,000 × g for 10 min at 4 °C and discard the supernatant containing unbound dye.
4. Transfer an aliquot of the stained mitochondrial suspension onto a glass slide and cover the sample with a coverslip.
5. Capture fluorescence images using a laser scanning confocal microscope with appropriate excitation and emission settings for red fluorescent mitochondrial dye detection.

8. Assessment of mitochondrial membrane potential using JC-1 staining

1. CCCP (carbonyl cyanide m-chlorophenyl hydrazone) treatment group
   1. Treat the mitochondrial suspension with CCCP at a final concentration of 10 µM and incubate for 20 min to dissipate mitochondrial membrane potential.
   2. Incubate the CCCP-treated mitochondrial suspension with 250 µL of JC-1 staining solution at 37 °C for 20 min with gentle agitation.
2. JC-1 treatment group
   1. Resuspend the mitochondrial pellet in 100 µL of JC-1 working solution and incubate at 37 °C for 20 min protected from light.
   2. Prepare 1× JC-1 staining buffer by diluting the 5× stock solution with distilled water at a ratio of 2:8 (v/v) and maintain the buffer on ice.
   3. Remove the staining solution and wash the mitochondrial pellet twice with pre-chilled 1× JC-1 staining buffer.
   4. Resuspend the stained mitochondria in PBS and image them using a laser scanning confocal microscope. Use appropriate excitation and emission settings for the detection of green fluorescent JC-1 monomers and red fluorescent JC-1 aggregates.

9. Ultrastructural analysis of isolated skeletal muscle mitochondria by transmission electron microscopy

1. Fix freshly isolated mitochondria in 2.5% glutaraldehyde prepared in 0.2 M phosphate buffer (pH 7.4) and incubate the samples overnight at 4 °C.
2. Wash the samples with 0.1 M phosphate-buffered saline (PBS, pH 7.4) and post-fix the samples in 1% osmium tetroxide for 1 h at room temperature.
3. Dehydrate the samples through a graded ethanol series followed by infiltration, embedding, and polymerization in epoxy resin.
4. Section the embedded samples at a thickness of 50-70 nm using an ultramicrotome.
5. Double-stain the sections with uranyl acetate and lead citrate. Observe and image the samples using a transmission electron microscope.

10. Cellular uptake of skeletal muscle-derived mitochondria

1. Culture NCTC clone 929 (L929; RRID: CVCL_0462) cells obtained from a commercial supplier in complete medium containing MEM Alpha supplemented with 10% fetal bovine serum and 1% penicillin-streptomycin.
2. When the cells reach 80%-90% confluence, aspirate the medium and wash the cells twice with pre-warmed PBS. Add trypsin-EDTA and incubate until the cells detach.
3. Add complete medium to terminate digestion and generate a single-cell suspension by gentle pipetting. Centrifuge the suspension at 1,000 × g for 5 min and discard the supernatant.
4. Resuspend the cell pellet in fresh complete medium and seed the cells into new culture dishes at an appropriate density. Incubate the cells at 37 °C in a humidified atmosphere containing 5% CO₂.
5. Seed passage 5 (P5) cells into 35 mm glass-bottom dishes and culture the cells overnight.
6. Label isolated mitochondria with a mitochondria-specific red fluorescent dye at 37 °C in the dark for 20 min and wash the mitochondria with PBS to remove unbound dye.
7. Add red fluorescent dye-labeled mitochondria (20 µg mitochondrial protein per dish) to the cells and incubate for 24 h at 37 °C in a humidified atmosphere containing 5% CO₂.
8. Stain the cells with a fluorescent actin probe for 30 min to visualize the actin cytoskeleton and counterstain nuclei with a fluorescent nuclear dye.
   NOTE: Carefully control serum concentration during co-incubation because excessive serum may reduce mitochondrial uptake efficiency. The mitochondria used in this step are fresh.

11. Measurement of mitochondrial respiratory function using high-resolution respirometry

1. Instrument preparation
   1. Clean the respiratory chambers, electrode compartments, and stoppers.
   2. Place a magnetic stirring bar into each chamber and verify the integrity of the oxygen electrode membrane.
2. Instrument calibration
   1. Add 2.1 mL of mitochondrial respiration buffer (MiR05, pH 7.10) into each respiratory chamber and ensure that no air bubbles are present.
   2. Equilibrate the high-resolution respirometry system at 37 °C for 30 min and perform daily air calibration after stabilization of the oxygen baseline.
      NOTE: Avoid introducing air bubbles into the respiratory chamber, as they interfere with oxygen electrode signals and compromise data quality. The mitochondria used in this step are fresh.
3. Sample loading and equilibration
   1. Add freshly isolated mitochondria containing approximately 30 µg of protein into the respiratory chamber and seal the chamber.
   2. Equilibrate the chamber for 5-8 min and record basal oxygen consumption.
      NOTE: Maintain mitochondria on ice and perform respiration measurements immediately after isolation to preserve activity.
4. SUIT protocol
   1. Add 5 µL pyruvate (2 M), 10 µL glutamate (2 M), and 10 µL malate (400 mM) sequentially. Stabilize the reaction for 5 min and record CI Leak respiration.
   2. Add 10 µL ADP (500 mM) to induce oxidative phosphorylation and record CI OXPHOS respiration. Add 10 µL magnesium chloride (0.3 M).
   3. Add 20 µL succinate (1 M) and record CI+CII OXPHOS respiration after stabilization.
   4. Add 2 µL ATP synthase inhibitor (0.01 mM) and record proton leak respiration.
   5. Add CCCP (1 mM) incrementally in three 2 µL additions at 2 min intervals until oxygen consumption reaches a plateau. Record maximal electron transfer system capacity.
   6. Add 1 µL rotenone (1 mM) to inhibit complex I activity.
   7. Add 1 µL antimycin A (5 mM) to inhibit complex III activity and record residual oxygen consumption.
   8. Add 5 µL ascorbate (800 mM) and 5 µL N,N,N',N'-tetramethyl-p-phenylenediamine (200 mM). Record maximal complex IV respiratory activity after stabilization.
      NOTE: Aliquot all reagents before freezing to avoid repeated freeze-thaw cycles. Add CCCP gradually, as excessive uncoupling inhibits mitochondrial respiration.
5. Data analysis
   1. Obtain mitochondrial respiration parameters, including CI Leak, CI OXPHOS, CI+CII OXPHOS, CI+CII Leak, CI+CII ET, CII ET, and CIV, using the data analysis software.
   2. Calculate the respiratory control ratio (RCR; State 3 respiration/State 4 respiration) and ATP production capacity. Retain all values to three significant figures for statistical analysis. The software used for data analysis was Oroboros Datlab 8 (Oroboros Instruments).

Results

A schematic overview of the mitochondrial isolation and purification workflow is shown in Figure 1. All procedures were performed at 4 °C to preserve mitochondrial integrity and functional activity. Freshly isolated skeletal muscle tissue was minced and digested with trypsin to facilitate tissue dissociation, then homogenized in isotonic isolation buffer using a glass homogenizer. Differential centrifugation was subsequently performed to separate mitochondria from tissue debris and cytosolic contaminants. Initial centrifugation at 1,000 × g removed nuclei and large debris, whereas subsequent centrifugation at 12,000 × g yielded a crude mitochondrial pellet. Additional washing and centrifugation steps further reduced cytosolic contamination, yielding purified mitochondria suitable for downstream structural and functional analyses (Figure 1).

Figure 1: Schematic diagram of mitochondrial isolation and purification. Mitochondria of high purity were isolated through a combined approach of controlled trypsin digestion and differential centrifugation. All procedures were performed at 4 °C to preserve mitochondrial structural and functional integrity. Freshly isolated skeletal muscle tissue was subjected to enzymatic digestion, homogenization, and sequential centrifugation steps to obtain purified mitochondria suitable for downstream functional and structural analyses. Please click here to view a larger version of this figure.

The functional activity of isolated mitochondria was evaluated using JC-1 staining and fluorescent mitochondrial labeling (Figure 2). JC-1 staining demonstrated strong red fluorescence in freshly isolated mitochondria under basal conditions, indicating preservation of mitochondrial membrane potential. Following treatment with carbonyl cyanide m-chlorophenyl hydrazone (CCCP), red fluorescence decreased, and green fluorescence increased, consistent with mitochondrial membrane depolarization. These findings confirmed that the isolated mitochondria retained membrane potential responsiveness and remained functionally active after isolation. Fluorescent labeling further demonstrated that mitochondria appeared as discrete, uniformly distributed punctate structures throughout the field, without obvious aggregation or dye precipitation. The low background fluorescence and uniform staining pattern suggested preservation of mitochondrial integrity and membrane potential following isolation.

Figure 2: Characterization of functional activity of isolated mitochondria. (A) Assessment of mitochondrial membrane potential by JC-1 staining. Purified mitochondria were stained with JC-1 and observed under fluorescence microscopy. Distinct red fluorescent aggregates were observed in the control mitochondria group, indicating preserved mitochondrial membrane potential. Following treatment with the uncoupler CCCP, fluorescence shifted predominantly toward green JC-1 monomers, indicating membrane depolarization and collapse of membrane potential (scale bar = 50 µm). (B) Mitochondrial fluorescent labeling of purified mitochondria. Fluorescently labeled mitochondria appeared as scattered, punctate structures without obvious aggregation and with minimal background fluorescence, indicating preserved mitochondrial integrity and a stable membrane potential (scale bar = 100 µm). (C) Quantitative analysis of JC-1 fluorescence intensity ratios in different groups after JC-1 staining. Data are presented as mean ± SEM (n = 3). Statistical significance was analyzed using an unpaired t-test (****P < 0.0001). Please click here to view a larger version of this figure.

The ultrastructural integrity of isolated mitochondria was examined using transmission electron microscopy (Figure 3). Low-magnification imaging revealed purified mitochondria with predominantly oval morphology, whereas high-magnification imaging demonstrated intact outer membranes and clearly defined cristae structures. No obvious swelling, rupture, or structural disruption was observed. For quantitative assessment, five randomly selected fields containing approximately 40-60 mitochondria per field were analyzed. These observations demonstrated that the isolation procedure effectively preserved mitochondrial ultrastructure and generated mitochondria suitable for downstream functional studies.

Figure 3: Ultrastructural integrity of isolated skeletal muscle mitochondria assessed by transmission electron microscopy. (Left) Representative TEM image at low magnification (4,000×; scale bar = 2 µm) showing purified mitochondria with predominantly oval morphology. Please click here to view a larger version of this figure.

(Right) High-magnification TEM image (40,000×; scale bar = 200 nm) demonstrating smooth and continuous outer mitochondrial membranes and clearly defined cristae structures. Yellow arrows indicate intact outer mitochondrial membranes, whereas white arrows indicate well-organized mitochondrial cristae. No evidence of swelling, rupture, or structural disruption was observed. Five randomly selected fields containing approximately 40-60 mitochondria per field were analyzed. These findings demonstrate that the isolation protocol preserved mitochondrial ultrastructural integrity and provided a reliable structural basis for downstream functional assays.

Mitochondrial yield and purity were subsequently evaluated by protein quantification and immunoblot analysis (Figure 4). Comparison between the trypsin digestion (TD) and normal extraction (NE) methods showed that the TD method yielded approximately 2.3-2.4 µg mitochondrial protein per mg tissue, whereas the NE method yielded approximately 1.7-1.8 µg/mg tissue. Immunoblot analysis demonstrated strong enrichment of mitochondrial markers, cytochrome c oxidase and COX IV, in mitochondrial fractions, with minimal detection in cytosolic fractions. In contrast, the cytosolic marker β-actin exhibited only weak signals in mitochondrial fractions, indicating limited cytosolic contamination. Comparison between the TD and NE groups suggested that trypsin digestion improved mitochondrial yield and purity. Collectively, these findings demonstrated that the protocol generated mitochondria suitable for downstream molecular and functional analyses.

Figure 4: Yield and purity comparison of purified mitochondria. (A) Comparison of mitochondrial protein yield per unit tissue mass obtained using the trypsin digestion (TD) and normal extraction (NE) methods. The TD group yielded approximately 2.3-2.4 µg/mg tissue, whereas the NE group yielded approximately 1.7-1.8 µg/mg tissue. Data are presented as mean ± SEM (n = 3). Statistical significance was analyzed using an unpaired t-test (*P < 0.05). (B) Immunoblot analysis of mitochondrial and cytosolic marker proteins in mitochondrial fractions isolated using the TD and NE methods. (C-D) Quantitative analysis of mitochondrial marker proteins cytochrome c oxidase and COX IV. Mitochondrial markers were strongly enriched in mitochondrial fractions but showed minimal expression in cytosolic fractions. The cytosolic marker β-actin exhibited only weak expression in mitochondrial fractions, indicating efficient removal of cytosolic contaminants. Compared with the NE group, the TD group demonstrated improved mitochondrial purity. Data are presented as mean ± SEM (n = 3). Statistical significance was analyzed using an unpaired t-test (*P < 0.05; **P < 0.01). Please click here to view a larger version of this figure.

To further evaluate mitochondrial respiratory function, oxygen consumption was measured using high-resolution respirometry (Figure 5). Mitochondria isolated using the TD method exhibited respiratory control ratio (RCR) values of approximately 9-10, whereas mitochondria isolated using the NE method exhibited RCR values of approximately 8-9. Because RCR values greater than 4 are generally considered indicative of preserved mitochondrial respiratory activity, these results demonstrated that mitochondria isolated using both methods retained functional integrity. The higher RCR values observed in the TD group suggested better preservation of mitochondrial respiratory function than in the NE group. Biochemical coupling efficiency was also assessed using the equation 1 − (leak respiration rate/electron transfer respiration rate), with values approaching 1 indicating tighter respiratory coupling and more efficient ATP production. Both groups exhibited coupling efficiencies approaching 1, indicating that mitochondrial bioenergetic function was preserved following isolation.

Figure 5: Assay of respiratory function in purified mitochondria. (A) Comparison of mitochondrial respiratory control ratio (RCR) between the trypsin digestion (TD) and normal extraction (NE) groups. RCR values in both groups were greater than 4, indicating preservation of mitochondrial respiratory activity following isolation. The TD group exhibited slightly higher RCR values than the NE group, suggesting better preservation of respiratory function. Data are presented as mean ± SEM (n = 3). Statistical significance was analyzed using an unpaired t-test (**P < 0.01). (B) Comparison of mitochondrial biochemical coupling efficiency and ATP synthesis capacity between the TD and NE groups. Coupling efficiency values approached 1 in both groups, indicating highly coupled respiration and efficient ATP production. Data are presented as mean ± SEM (n = 3). Statistical significance was analyzed using an unpaired t-test (*P < 0.05). Please click here to view a larger version of this figure.

The uptake of isolated mitochondria by recipient cells was evaluated using L929 cells (Figure 6). Cells were co-incubated with red fluorescent dye-labeled skeletal muscle-derived mitochondria, and the actin cytoskeleton was visualized using a fluorescent actin probe. Confocal microscopy revealed punctate red fluorescent structures distributed throughout the cytoplasm, indicating that L929 cells had taken up exogenous mitochondria. Quantitative analysis demonstrated a time-dependent increase in mitochondrial uptake at 12, 24, and 48 h. However, only a modest increase was observed between 24 h and 48 h, suggesting that uptake efficiency approached saturation after 24 h. In addition, recipient L929 cells exhibited increased mitochondrial membrane potential following mitochondrial uptake, indicating improved mitochondrial function. These findings demonstrated that skeletal muscle-derived mitochondria isolated using this protocol could be effectively internalized by recipient cells while maintaining functional activity.

Figure 6: Cellular uptake of skeletal muscle-derived mitochondria by L929 cells. (A) Uptake of purified mitochondria by L929 cells at different incubation times (12 h, 24 h, and 48 h). Mitochondria-fluorescent dye-labeled mitochondria (red) were detected within the cytoplasm of recipient cells, while phalloidin staining (green) was used to visualize the actin cytoskeleton. Nuclear staining was performed using DAPI (blue). Confocal microscopy confirmed the internalization of exogenous mitochondria by L929 cells (scale bar = 50 µm). (B) Quantitative analysis of mitochondrial uptake efficiency by L929 cells at different time points. (C) JC-1 staining analysis of mitochondrial membrane potential in L929 cells following uptake of skeletal muscle-derived mitochondria. (D) Quantitative analysis of JC-1 aggregate-to-monomer fluorescence ratios in recipient L929 cells after mitochondrial uptake. Mitochondrial membrane potential was significantly increased following internalization of active skeletal muscle-derived mitochondria. Data are presented as mean ± SEM (n = 3). Statistical significance was analyzed using one-way ANOVA (*P < 0.05; **P < 0.01; ***P < 0.001). Please click here to view a larger version of this figure.

Discussion

Mitochondrial transplantation is an emerging strategy in cell therapy and regenerative medicine that restores physiological function in cells with impaired energy metabolism through the delivery of functionally intact mitochondria. In addition to regulating cellular energy metabolism, this technique can modulate key biological processes such as apoptosis and oxidative stress, thereby providing novel therapeutic opportunities for a variety of diseases^4,17,18,19. However, the clinical translation of mitochondrial transplantation still faces several challenges, among them the lack of standardized, efficient, and reproducible mitochondrial isolation protocols, which remains a major bottleneck.

To address this issue, the present study established a standardized and visualized experimental protocol based on differential centrifugation for the efficient isolation of functional free mitochondria from C57BL/6 mice skeletal muscle. By optimizing the purification procedure and incorporating trypsin pre-digestion, washing, and purification steps, this method effectively improves tissue dissociation, preserves mitochondrial structural integrity, and reduces contamination from non-target cellular components.

The proposed protocol achieves a favorable balance between mitochondrial purity and yield. Compared with conventional grinding-based extraction methods, the present method is faster, more practical, and more cost-effective, making it suitable for routine laboratory applications and donor mitochondrial preparation. Existing mitochondrial isolation approaches each possess certain limitations. For example, density-gradient centrifugation and immunoaffinity purification can achieve higher purity but are time-consuming and costly, whereas nanoscale probe-based approaches remain insufficiently developed for routine use^20,21. For dense tissues such as skeletal muscle, the optimized trypsin pre-digestion step (0.25% trypsin at 37 °C for 10-15 min), combined with gentle homogenization, better preserves mitochondrial integrity than conventional differential centrifugation alone.

In this study, a commercially available differential centrifugation kit was used under low-temperature conditions throughout the procedure and following a standardized workflow, enabling rapid acquisition of structurally intact mitochondria. To address potential problems encountered during the extraction process, including low yield, protein contamination, and abnormal membrane potential, several troubleshooting strategies are proposed.

Low mitochondrial yield is commonly caused by insufficient homogenization, inadequate tissue quantity, or centrifugal loss. These issues can be minimized by ensuring adequate pre-digestion, performing gentle homogenization (10-15 passes using a tight-fitting homogenizer), using at least 100-150 mg of fresh skeletal muscle tissue, strictly controlling centrifugation conditions (1,000 × g for debris removal and 10,000-12,000 × g for mitochondrial pelleting), and resuspending samples using wide-bore pipette tips. Protein contamination mainly results from incomplete debris removal or lipid interference; therefore, repeating the 1,000 × g centrifugation step, incorporating two additional 12,000 × g washing steps, and carefully removing adipose tissue are recommended. Abnormal mitochondrial membrane potential is often associated with mechanical damage or temperature fluctuations. Excessive homogenization should therefore be avoided, the entire procedure should be performed on ice and completed within 60-90 min, and isolation buffers should be supplemented with EGTA and 0.5% BSA. Incomplete homogenization can be improved by ensuring adequate enzymatic digestion, selecting an appropriate homogenizer, and performing post-homogenization filtration.

Although the present protocol significantly improves the efficiency and purity of mitochondrial isolation, several limitations remain. First, the procedure is highly dependent on tissue freshness; ideally, tissue samples should be processed within 1 h postmortem. Second, residual trypsin may degrade mitochondrial surface proteins, including translocase-related proteins, if not adequately removed during washing, potentially affecting subsequent cellular uptake efficiency. Third, mitochondrial storage time remains limited, as prolonged storage at 4 °C or cryopreservation substantially reduces mitochondrial activity.

At the translational level, additional challenges remain, including improving in vivo delivery efficiency, understanding long-term mechanisms of mitochondrial integration, and clarifying potential interactions between transplanted mitochondria and the host genome. With continued advances in mitochondrial isolation and purification technologies, further methodological innovations are expected to accelerate the clinical translation of mitochondrial transplantation therapy.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Anti-Cytochrome C antibody	Abcam	EPR1327	Mitochondrial Characterization
Anti-fade mounting medium	PlantChemMed	PC-90016	Reduce photobleaching of fluorescent dyes
BCA Protein Assay Kit	Zhonghui Hecai Biotechnology Co., Ltd.	PQ003	Protein concentration assay
Beta-actin antibody	Cell Signaling technology	3700s	Beta-actin antibody was used to detect the purity of isolated mitochondria
BSA	Sigma	A6003	During isolation, it binds free fatty acids to protect mitochondrial activity.
Centrifuge	Eppendorf	5418R	4 °C Centrifugation
CCCP	Solarbio	C6700	to uncouple mitochondrial oxidative phosphorylation and induce maximal proton leakage
Confocal Laser Scanning Microscope	Olympus Corporation	FV4000	Immunofluorescence Staining Observation
COX IV polyclonal antibody	Proteintech	11242-1-AP	Mitochondrial Characterization
DAPI	Plant Chem Medicine	PC-90004	For nuclear counterstaining
D-mannitol	Solarbio	IM00402	maintain osmotic balance and prevent mitochondrial swelling during isolation
EGTA	Sigma	E4378	Commonly used in mitochondrial isolation buffers to protect mitochondrial function by specifically binding Ca2+
Fetal bovine serum	Every Green	11011-08611	Culture of L929 cells
Forceps	RWD Life Science Co., Ltd.	F11003-11	Used to grasp, hold and lift tissues
Glass homogenizer	Beyotime	FGH005	Used for tissue homogenization and isolation of mitochondria
Glutaraldehyde	LEAGENE	DF0151	It rapidly fixes and stabilizes cellular ultrastructure by cross-linking proteins, preventing autolysis to preserve the natural morphology of organelles.
HEPES	Thermo Fisher	J67485	Provides a stable pH environment.
Incubator	Thermo Fisher	51032874	Culture of L929 cells
JC-1	Solarbio	M8650	Detecting changes in the membrane potential of purified mitochondria.
L929 cell line	Procell	CL-0137	For mitochondrial endocytosis assay
Lead citrate	Sigma	15326	to stain ultrathin sections for fine subcellular structure observation.
MEM Alpha basic (1x)	Gibco	6125510	Culture of L929 cells
Methanol	Fuyu Chemical (China)	67-56-1	As a component of the transfer buffer, it is used to remove SDS and facilitate the transfer of proteins from the gel to the solid-phase membrane.
Microplate Reader	Thermo Fisher	Varioskan LUX	Measure the absorbance of JC-1-stained mitochondria
Mitochondrial Membrane Potential Detection Kit	Solarbio	M8650	Detection of Mitochondrial Membrane Potential in Skeletal Muscle
Mitochondrial isolation Buffer	Solarbio	SM0020	Kits for Extraction Based on Differential Centrifugation
Mitochondria Storage Buffer	Solarbio	SM0020	For mitochondrial preservation
Mito Tracker  Red CMXRos	Yeasen Biotechnology	40741ES50	Mitochondrial Characterization
MOPS	MedChemExpress	HY-D0859	AS a buffer in solution preparation
Multi-rAb HRP-Goat Anti-Mouse Recombinant Secondary Antibody (H+L)	Proteintech	RGAM001	RGAM001 targets Mouse IgG (H+L) in Western blot
Multi-rAb HRP-Goat Anti-Rabbit Recombinant Secondary Antibody (H+L)	Proteintech	RGAR001	RGAR001 targets to Rabbit IgG (H+L) in ELISA and Western blot
Non-fat dry milk	NCM Biotech	WB6503	It is mainly used as a blocking agent to reduce background signal by occupying non-specific binding sites on the membrane.
O2k-FluoRespirometer	Oroboros Instruments	O2K	Determine mitochondrial respiratory function and oxygen metabolism.
Osmium tetroxide	Sigma	75633	or postfixation to preserve lipids and enhance membrane contrast in TEM samples.
PBS (1x)	Plant Chem Medicine	PC-00003	Clean Muscle Tissue
Penicillin-Streptomycin	Procell	PB180120	Culture of L929 cells
Phalloidin-488	Beyotime	C1035	Label cytoskeletal F-actin for fluorescence microscopic imaging
Polyvinylidene fluoride (PVDF) membrane	Millipore (Merck)	IPVH00010	Western blotting
PowerPac Basic	BIO-RAD	164-5050	For protein separation by SDS-PAGE electrophoresis
Precast gradient polyacrylamide gel (4–12%)	ACE Bioscience	ET12412LGel	Used for SDS-PAGE separation of mitochondrial and cellular total proteins
Protease inhibitor	Shaanxi Zhonghui Hecai Biomedical Technology	PL032	Inhibits the activity of endogenous and exogenous proteases, preventing the degradation of target proteins during sample processing.
RIPA lysis buffer	Zhonghui Hecai Biotechnology Co., Ltd.	PL001-2A	Used for total protein extraction of animal and cellular tissues
Sodium Chloride Injection (0.9%)	Xi’an Jingshuang Shuanghe Pharmaceutical Co., Ltd.	Approval No. H61020014	Cell/tissue experimental buffering, rinsing and osmotic pressure maintenance
Sterile gloves	Jiangsu Yangzi Lide Medical Devices Co., Ltd.	YMZ000-1	Isolate contamination and protect operators and experimental samples
Sucrose	Sucrose	S818048	Provides osmotic support and maintains mitochondrial morphology and functional integrity.
TBST	NCM Biotech	WB20500	Washing to reduce background, serving as a diluent for antibodies and blocking solutions, while protecting the stability of antigen-antibody binding by maintaining a stable pH and ionic environment.
Tissue Homogenate Assay Kit(contains Adenosine diphosphate (ADP) ,Succinate,Rotenone,Antimycin A ,Oligomycin,Pyruvate,Malate, Glutamate,Ascorbate,N,N,N',N'- tetramethyl-p-phenylenediamine (TMPD))	Beijing Huawei Zhongyi Technology Co., Ltd.	023012	Detection of mitochondrial OCR in tissue homogenate adapted for Oroboros O2k respirometer
Tissue scissors	RWD Life Science Co., Ltd.	S12003-09	For cutting and dissecting soft tissues, separating organs and fascia in animal dissection
Transmission electron microscope	HITACHI	HT7800	Analyze the ultrastructural morphology of mitochondria
Trypsin	Plant Chem Medicine	PC-90001	Organ pre-digestion
Universal Antibody Diluent	NCM Biotech	WB100D	Antibody Diluent
Ultramicrotome	Leica	EMUC7	Used to prepare ultrathin sections for transmission electron microscopy (TEM)
Uranyl acetate	Electron Microscopy Sciences	22400	to stain ultrathin sections and improve intracellular structural contrast.
Vertical electrophoresis cell	BIO-RAD	165-8001	For protein separation by SDS-PAGE electrophoresis
Wash Buffer	Solarbio	SM0020	Wash and purify isolated mitochondria, maintaining their structural and functional integrity
75% ethanol	Shanghai Macklin Biochemical Co., Ltd.	E885996	For surface disinfection of laboratory apparatus and experimental materials