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Method Article

Isolation of Small Extracellular Vesicles from Murine Skeletal Muscle and Bone Marrow by Size-Exclusion Chromatography

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DOI:

10.3791/70894

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June 23rd, 2026

 ,  ,  , 
1Integrated Program in Neuroscience, McGill University, 2McGill Group for Suicide Studies, Douglas Mental Health University Institute, 3Center for Applied Nanomedicine, Research Institute of the McGill University Health Center, 4Department of Psychiatry, McGill University

In This Article

Summary

This protocol describes the isolation of small extracellular vesicles from murine skeletal muscle and bone marrow using enzymatic digestion and size-exclusion chromatography, enabling reproducible recovery of vesicle-enriched fractions from structurally complex, low-yield tissues for downstream molecular analysis.

Abstract

Small extracellular vesicles (sEVs) are nanosized, lipid-bound particles that mediate intercellular communication through the transfer of proteins, lipids, and ribonucleic acids (RNAs). Isolation of sEVs from solid tissues such as skeletal muscle (SkM) and bone marrow (BM) remains challenging due to low yield and contamination from non-vesicular components. This protocol describes a reproducible workflow for isolating sEVs from mouse SkM and BM. SkM is enzymatically digested, whereas BM is processed directly and subsequently subjected to differential centrifugation and size-exclusion chromatography (SEC). For SkM, the void volume (~2.5 mL) is discarded, and the subsequent ~1.6 mL is collected as EV-enriched fractions, typically subdivided into 400 µL sequential fractions (F1–F4). For BM, a 700 µL void volume is discarded, followed by collection of an ~850 µL EV-enriched fraction, which can be subdivided into ~170 µL sequential fractions (F1–F5) and pooled based on EV marker enrichment. Using this approach, sEVs can be isolated from small tissue volumes (approximately 500 µL from a single quadriceps and 150 µL from pooled femur and tibia BM), yielding ~108 particles·mL−1·mg−1 tissue with particle diameters predominantly <200 nm. Vesicle integrity and purity are validated using transmission electron microscopy, nanoparticle tracking analysis, and Western blotting for canonical EV markers, with additional characterization by single-particle interferometric detection to quantify tetraspanin-defined vesicle subpopulations. This method enables reproducible isolation of high-purity sEVs from structurally complex tissues using minimal input material, supporting downstream molecular and functional analyses of tissue-derived vesicles.

Introduction

Small extracellular vesicles (sEVs) are nanosized, lipid-bound particles that mediate intercellular communication through the transfer of proteins, lipids, and nucleic acids1. They are released by all cell types and are present in virtually all biological fluids, where they function as key carriers of bioactive cargo, including peptides and micro ribonucleic acids (RNAs)2. Their cargo is selectively sorted, enabling precise delivery of specific signals3, and their vesicular structure allows them to bypass the typical gradient and dilution constraints of freely secreted factors by delivering concentrated molecular payloads over long distances4. Although EVs from biofluids (e.g., plasma, serum, and saliva) provide a convenient snapshot of systemic signaling, they represent a highly mixed population originating from many tissues5. This makes it challenging to determine which organ is producing a specific biological signal or how EV communication is regulated at the tissue level. Because circulating EVs are pooled together, tissue-specific changes can be diluted or masked, limiting the ability to identify the true source of signals6.

Skeletal muscle (SkM) and bone marrow (BM) are major metabolically and functionally active tissues with distinct cellular compositions and signaling roles7,8. Both tissues may release EV populations enriched with unique, tissue-specific cargo; however, these signals can be easily overshadowed in plasma by EVs from high-turnover tissues such as immune cells8,9. Directly isolating EVs from muscle or BM increases sensitivity and enables detection of subtle, tissue-specific adaptations that may not be observable in circulation. Tissue-derived EVs therefore provide essential complementary information, revealing when a specific organ releases EVs, how long those signals persist, and which adaptations are acute versus chronic. Despite increasing interest in EVs, most established isolation methods have been developed for biofluids (e.g., plasma and serum) or cell culture systems. In these contexts, EVs can be obtained with relatively high yield and minimal structural disruption. However, isolating EVs directly from solid tissues such as SkM and BM presents notable technical challenges. These tissues are structurally complex and heterogeneous, requiring mechanical or enzymatic dissociation steps that can introduce artifacts, including high levels of soluble proteins, organelle fragments, and non-vesicular nanoparticles, making it difficult to achieve high-purity EV preparations.

Another major challenge in tissue-derived EV isolation is the lack of standardized workflows for fraction definition and separation10. Differential centrifugation alone often results in co-isolation of protein aggregates and large vesicles, while ultracentrifugation-based methods, although widely used, can lead to vesicle aggregation, deformation, and variable recovery depending on rotor type and experimental conditions. Density gradient approaches improve purity but are time-intensive, low-throughput, and difficult to standardize across laboratories11. Consequently, there is currently no widely adopted, reproducible method optimized for isolating small EVs from solid tissues with high purity and consistent yield10.

Size-exclusion chromatography (SEC) has emerged as a promising alternative for EV isolation because it separates particles based on size without subjecting vesicles to high centrifugal forces. SEC enables efficient separation of small EVs from soluble proteins and low-molecular-weight contaminants while preserving vesicle integrity and biological function. Importantly, SEC allows for clear definition of elution fractions, enabling reproducible identification of EV-enriched fractions and improved consistency across experiments12. However, SEC workflows require careful optimization for tissue-derived samples, particularly with respect to upstream tissue processing, debris removal, and fraction collection. To address these limitations, we developed a reproducible and standardized protocol for isolating sEVs from SkM and BM. This method integrates controlled enzymatic digestion for fibrous tissues, differential centrifugation for debris removal, and SEC-based purification to isolate high-purity vesicles. The workflow is specifically optimized to minimize contamination from intracellular components while preserving vesicle integrity and enabling consistent recovery from low-yield tissues.

This protocol fills a critical methodological gap by providing a practical and scalable approach for isolating tissue-derived sEVs with defined fractionation and validated purity, achieved through nanoparticle tracking analysis (NTA), Western blotting of canonical EV markers, electron microscopy for morphological assessment, and single-particle EV analysis using antibody capture, interferometric sizing, and multichannel fluorescence imaging. It is particularly well suited for studies requiring tissue-specific EV analysis, where biofluid-derived vesicles are insufficient to resolve the cellular origin of signals. Researchers should use this method when the goal is to obtain high-purity sEVs from solid tissues with minimal contamination and preserved vesicle integrity. In contrast, ultracentrifugation-based methods may be preferred for high-yield applications where purity is less critical, and density gradient approaches may be selected when isolating specific EV subpopulations despite lower throughput. Together, this protocol provides a standardized framework for the isolation and characterization of tissue-derived sEVs, enabling reproducible downstream molecular, biochemical, and functional analyses.

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Protocol

All animal procedures were approved by the McGill University Animal Care Committee (DOUG10029) and conducted in accordance with the guidelines of the Canadian Council on Animal Care.

1. SkM-derived sEV (SkM-sEV) Isolation Protocol

  1. Sample Collection and Tissue Dissociation
    1. Tissue harvest and preservation
      1. Snap-freeze SkM tissue immediately after excision by immersing it in prechilled methylbutane.
        NOTE:  To prepare prechilled methylbutane, place a container of methylbutane in a dry ice bath and gradually add small pieces of dry ice directly into the liquid. Allow the solution to cool, the temperature is typically within the range of −70°C to −80°C. Maintain the methylbutane in the dry ice bath during use to preserve temperature. Immerse the tissue for approximately 3–4 s until fully frozen, as indicated by complete tissue hardening and loss of flexibility.
      2. Transfer the tissue to cryovials.
      3. Place the samples on dry ice.
        NOTE: Store the samples at −80°C until further processing. Avoid repeated freeze–thaw cycles. Rapid snap-freezing effectively preserves EVs when samples are sealed properly. Dehydration or vesicle damage may occur if tissues are left exposed or undergo multiple freeze–thaw cycles13.
    2. Tissue dissociation
      ​NOTE: Perform all steps in a biosafety cabinet using sterile instruments and gloves to prevent contamination, which may otherwise compromise EV purity.
      1. Weigh tissue
        1. Sterilize tweezers with 70% ethanol and allow them to air-dry.
        2. Transfer the frozen tissue onto a pre-weighed suspension culture dish.
        3. Weigh the tissue using a milligram balance (typically ~80–120 mg per sample, depending on mouse age and sex).
          NOTE: The protocol is validated for input tissue masses of ~80–120 mg. For samples within this range, use the reagent volumes specified in Steps 1.2.3.1, 1.3.1.3, and 1.3.2.1 without modification. For samples outside this range, scale all reagent volumes proportionally to tissue mass while maintaining the validated ratio of RPMI-1640, collagenase D, and DNase I (1000:40:4 µL per 100 mg tissue). This corresponds to approximately 10 µL RPMI-1640, 0.4 µL collagenase D (100 mg/mL stock), and 0.04 µL DNase I (20,000 U/mL stock) per mg of tissue. Maintain these proportions to preserve the final in-reaction enzyme concentrations. Digestion efficiency and EV yield should be empirically validated when working substantially outside the 80–120 mg range.
        4. Return the sample to a pre-chilled tube and keep it on ice.
      2. Prepare enzymes
        1. Thaw aliquots of collagenase D (100 mg/mL stock) and DNase I (20,000 U/mL stock) on ice (see Supplementary File 1, Step 1.4 for enzyme stock preparation). These enzymes are added in Step 1.3.2.1 to achieve final in-reaction concentrations of 3.8 mg/mL collagenase D and 77 U/mL DNase I. The final digestion volume is 1.044mL.
        2. Mix DNase I gently by inversion. Do not vortex.
      3. Tissue preparation for digestion
        1. Add 800 µL RPMI-1640 to a sterile suspension culture dish placed on ice.
        2. Transfer the tissue to the dish.
        3. Cut the tissue into 6–8 pieces of roughly uniform size (2–3 mm per side) using a sterile disposable scalpel. Stabilize the tissue with sterile forceps during cutting to ensure even fragmentation. Complete this step within 1 min/sample to maintain consistency across samples.
        4. Keep the dish on ice throughout the procedure to minimize degradation.
    3. Tissue transfer and enzymatic digestion
      1. Tissue transfer
        1. Trim the end of a sterile P1000 pipette tip using a sterile blade to produce an opening of 2–3 mm in diameter, sufficient to allow free passage of the tissue fragments generated in Step 1.2.3.3.
          NOTE: Alternatively, use commercially available wide-bore (wide-orifice) 1 mL pipette tips, which provide a standardized opening (typically ~2–3 mm internal diameter) and eliminate variability associated with manual trimming.
        2. Transfer the tissue fragments and RPMI-1640 medium into pre-labeled 1.5 mL tubes using a P1000 pipette fitted with the modified tip. Perform 2–3 successive aspirations to recover residual fragments and medium. Visually inspect the dish and pipette tip after transfer and use sterile forceps to transfer any remaining fragments to ensure no visible tissue remains.
        3. Add 200 µL of RPMI-1640 to each tube.
      2. Enzymatic digestion
        1. Add 40 µL of collagenase D stock (100 mg/mL) and 4 µL of DNase I stock (20,000 U/mL) to each tube from aliquots thawed on ice. This yields final in-reaction concentrations of 3.8 mg/mL collagenase D and 77 U/mL DNase I. For samples outside the 80–120 mg tissue range, adjust enzyme volumes proportionally as described in the NOTE following Step 1.2.1.3.
          NOTE: These volumes are calibrated for tissue masses within the typical 80–120 mg range. For tissue masses outside this range, scale all reagent volumes proportionally as described in the NOTE following Step 1.2.1.3 (0.4 µL collagenase D stock and 0.04 µL DNase I stock per mg of tissue), which preserves the same final in-reaction enzyme concentrations regardless of tissue mass.
        2. Add the enzymes to all samples simultaneously when processing multiple samples.
        3. Keep the tubes on ice during enzyme addition.
        4. Gently invert the tubes to mix. Do not vortex.
        5. Incubate the tubes at 37°C for 30 min on a nutating mixer (24 rpm, 20° tilt angle; see Table of Materials). Ensure even mixing during incubation.
          NOTE: Digest each sample in a total volume of 1.044 mL (1000 µL RPMI-1640, 40 µL collagenase D, and 4 µL DNase I per sample; equivalent to ~25:1 buffer-to-collagenase ratio).
    4. Removal of debris
      NOTE: Initiate SEC column equilibration at the start of the first centrifugation (Step 1.4.1.1). The SEC column requires 20–25 min for flushing and equilibration. Starting equilibration at this point ensures that the column is ready for sample loading immediately after completion of the filtration step (Step 1.4.3).
      1. First centrifugation
        1. Centrifuge the samples at 300 × g for 10 min at 4°C to pellet undigested debris, intact cells, and remaining tissue fragments.
        2. Carefully transfer the supernatant to new sterile 1.5 mL microcentrifuge tubes without disturbing the pellet.
      2. Second centrifugation
        1. Centrifuge the samples at 2,000 × g for 15 min at 4°C to pellet large debris, dead cells, apoptotic bodies, and large vesicles.
        2. Carefully transfer the supernatant to new sterile 1.5 mL microcentrifuge tubes without disturbing the pellet.
          ​NOTE: The second centrifugation is optional and should be included when additional debris removal is required. Perform this step if the supernatant appears visibly turbid or contains residual particulate material after the first centrifugation. This step is also recommended if prior preparations have shown contamination by intracellular markers (e.g., Calnexin or BiP) in EV-enriched SEC fractions or the presence of non-vesicular particles by transmission electron microscopy. The need for this step should be validated during initial protocol optimization for each tissue type.
      3. Filtration
        1. Pre-wet sterile 30 µm cell strainers by rinsing them with RPMI-1640 to equilibrate the membrane and minimize sample loss. Discard the rinse.
        2. Filter each sample through the pre-wetted 30 µm cell strainer into sterile low-protein-binding 1.5 mL microcentrifuge tubes to remove remaining cellular debris and large particles before SEC.
        3. Allow the samples to pass through the filter by gravity.
        4. Allow 10–15 min per sample for filtration.
          NOTE: Filtration is complete when no visible liquid remains above the filter membrane and the filtrate volume in the collection tube has stabilized. If filtration is slower or stalls, this may indicate clogging due to residual debris; in such cases, consider repeating the clarification step or reducing sample load.
        5. Collect 550–600 µL of filtered supernatant for subsequent loading onto the SEC column. For consistency across samples, normalize the loading volume to 500 µL per column (see Step 2.2.1), regardless of the total volume recovered.
  2. Size-exclusion chromatography (SEC)
    1. Equilibrate SEC column
      NOTE: Perform this step during centrifugation if possible.
      1. Flush the SEC column with 17 mL of 1× PBS (0.22 µm-filtered).
      2. Allow the entire volume to pass through the column by gravity at an expected flow rate of 0.75 mL/min (20–25 min total for the 17 mL flush volume). Faster or slower flow rates may indicate improper column equilibration or blockage.
      3. Maintain the column and PBS at room temperature (RT, 21°C ± 2°C) during equilibration to ensure consistent flow rate and optimal column performance.
      4. Prepare low-protein-binding 1.5 mL microcentrifuge tubes for collection of all fractions.
      5. Prepare an additional tube for collection of the void volume if required.
    2. Sample loading and elution
      1. Load 500 µL of filtered supernatant onto the SEC column.
      2. Allow the sample to fully enter the resin. The endpoint is reached when the liquid meniscus reaches the surface of the resin bed, with no visible liquid remaining above the resin while the resin remains visibly wet.
      3. Do not allow the meniscus to drop below the resin surface or the column to run dry, as this introduces air into the resin bed and disrupts elution. When the liquid meniscus reaches the resin surface (as defined in Step 2.2.2), immediately proceed to Step 2.2.4 without delay.
      4. Immediately add 2 mL PBS to initiate elution.
      5. Collect the void volume (~2.5 mL).
      6. Discard the void volume or retain it as a non-EV fraction.
      7. Begin collecting the EV-enriched fraction immediately after the 2.5 mL void volume has eluted.
      8. Collect a total of 1.6 mL of EV-enriched eluate following the void volume, either as a single continuous fraction in one tube or as sequential fractions that are subsequently pooled.
      9. Alternatively, collect sequential fractions of 0.4 mL to optimize purity versus yield. Smaller fraction volumes are recommended when higher purity is required (e.g., for proteomic analyses or sensitive downstream applications), whereas larger fractions may be used to maximize overall EV yield.
      10. Assess fractions for EV enrichment using characterization methods (e.g., Western blot for EV markers and contaminants, and NTA), and pool EV-enriched fractions as defined in the characterization section.
        NOTE: Keep the column flowing continuously. As one fraction is finishing, add PBS to maintain a constant liquid above the resin bed and prevent the column from running dry. Use low-protein-binding, nuclease-free tubes for collection.
    3. Concentration
      1. Concentrate EV samples using centrifugal filters (100 kDa molecular weight cutoff [MWCO]) for downstream applications (e.g., RNA sequencing, proteomics, or Western blot).
      2. Pre-rinse the filters with PBS before use.
      3. Centrifuge the samples at 4,000 × g at 4°C, checking every 2–3 min until the desired final volume (typically 50–100 µL) is reached. Use lower final volumes (e.g., ~50 µL) for applications requiring higher concentration and higher volumes (e.g., ~100 µL) when maximizing recovery is preferred.
        CAUTION: Avoid over-concentration by stopping centrifugation once the target volume is achieved.
      4. Recover the retentate by gentle pipetting.
      5. Record pre- and post-concentration sample volumes to estimate recovery, if required. NTA may be used to assess particle concentration before and after concentration, if available.
    4. Storage
      1. Keep EV-enriched fractions on ice or at 4°C for immediate assays (e.g., NTA and transmission electron microscopy [TEM]) and process within 24 h for TEM and within 48 h for NTA.
      2. Aliquot EV samples into low-protein-binding tubes for storage.
      3. Store samples at −80°C for long-term use.
      4. Avoid repeated freeze–thaw cycles.

2. BM-derived sEV (BM-sEV) Isolation Protocol

  1. Sample collection and tissue dissociation
    1. Tissue harvest and preservation
      1. Do not snap-freeze bones immediately after excision, as this hardens the marrow and complicates extraction using the poked-tube method. Instead, keep bones on ice or at 4°C during short handling periods prior to processing or freezing as described in Step 3.1.1.4 (placement on dry ice prior to −80°C storage).
      2. Remove surrounding muscle and connective tissue from dissected bones.
      3. Transfer the bones to cryovials.
      4. Place the samples on dry ice.
        ​NOTE: Store the samples at −80°C until further processing.
    2. BM processing
      ​NOTE: Perform all steps in a biosafety cabinet using sterile instruments and gloves to prevent contamination, which may otherwise compromise EV purity.
      1. Pre-weigh and label 1.5 mL centrifuge tubes before adding any material. Calculate BM mass by comparing tube weights before and after extraction.
      2. Prepare the poked-tube setup by inserting a 0.2 mL PCR tube (cap intact) into a 1.5 mL centrifuge tube with a snug fit. Puncture the bottom of the 0.2 mL tube once with a sterile 16 G needle held perpendicular to the base, producing a single uniform perforation of ~1.6 mm diameter. Prepare one setup per leg per mouse.
      3. Place the leg, cleaned of all muscles and surrounding tissue, in a sterile Petri dish.
      4. Separate the femur and tibia by cutting through the joint space.
      5. Cut both ends of the femur and tibia to expose the red marrow. Place the bones into the punctured PCR tube positioned inside the 1.5 mL tube and seal the assembly with parafilm to ensure a secure setup during centrifugation.
      6. Centrifuge the samples for 10 s in a benchtop microcentrifuge at approximately 8,000 × g, or the closest achievable setting on the instrument used, at room temperature.
      7. Verify successful marrow extraction by three criteria: (i) the bones appear visibly white with no residual red marrow, (ii) the pellet in the lower tube appears dark red and compact, typically filling the conical tip of the 1.5 mL tube, and (iii) the measured marrow mass (Step 3.1.2.9) falls within the expected range of ~15–30 mg per mouse.
      8. Reposition or recut the bones and repeat the brief spin if visible red marrow remains within the bone.
      9. Weigh the tubes again to determine the mass of extracted marrow (typically ~15–30 mg per sample, depending on mouse age, sex, and extraction efficiency).
        NOTE: The protocol is validated for marrow yields of ~15–30 mg per sample. Because the RPMI-1640 resuspension volume is held constant at 250 µL regardless of marrow mass (Step 3.1.2.10), EV yield is normalized to the measured starting marrow mass during downstream analysis (particles·mL−1·mg−1 tissue). For samples substantially below 15 mg, pool marrow from matched mice (same age, sex, and bone) prior to resuspension to maintain sufficient input for downstream characterization. For samples substantially above 30 mg, either split the marrow across two separate resuspensions of 250 µL each or scale the RPMI-1640 volume proportionally (~12.5 µL per mg of marrow) to prevent incomplete resuspension; in either case, apply the same per-mass normalization during analysis.
      10. Add 250 µL RPMI-1640 to each tube. Maintain a constant volume across samples regardless of marrow mass to allow normalization of EV yield to starting marrow mass. Keep the samples on ice.
      11. Resuspend the marrow by gently pipetting up and down 10–15 times using a P1000 pipette until the suspension appears visually homogeneous, with no visible clumps or intact tissue fragments remaining. Avoid forceful pipetting or bubble formation, which may disrupt vesicle integrity.
  2. Removal of debris
    1. First centrifugation
      1. Centrifuge the samples at 300 × g for 10 min at 4°C to pellet undigested debris, intact cells, and remaining tissue fragments.
      2. Carefully transfer the supernatant to new sterile 1.5 mL microcentrifuge tubes without disturbing the pellet.
    2. Second centrifugation
      1. Centrifuge the samples at 2,000 × g for 15 min at 4°C to remove large debris.
      2. Carefully transfer the supernatant to new sterile 1.5 mL microcentrifuge tubes without disturbing the pellet.
    3. Third centrifugation
      1. Centrifuge the samples at 10,000 × g for 30 min at 4°C to remove apoptotic bodies and large vesicles.
      2. Carefully transfer the supernatant to new sterile 1.5 mL microcentrifuge tubes without disturbing the pellet.
      3. Ensure a final volume of 200 µL supernatant is obtained.
  3. Size-exclusion chromatography (SEC)
    1. Column preparation
      1. Equilibrate the SEC column by flushing with 6 mL of 1× PBS (0.22 µm-filtered).
        NOTE: The equilibration volume is column-specific and should match the column type listed in the Table of Materials. This column requires a 6 mL equilibration volume, which differs from the column used for SkM samples.
      2. Allow the entire volume to pass through by gravity.
      3. Maintain the column and PBS at RT during equilibration to ensure optimal flow rate and column performance.
    2. Sample loading and elution
      1. Load 150 µL of filtered supernatant onto the SEC column.
      2. Allow the sample to fully enter the resin. The endpoint is reached when the liquid meniscus reaches the surface of the resin bed with no visible liquid remaining above the resin while the resin remains wet (as described in Step 2.2.2).
      3. Do not allow the meniscus to drop below the resin surface or the column to run dry (see endpoint defined in Step 3.3.2.2), as this introduces air into the resin bed and disrupts elution. When the liquid meniscus reaches the resin surface, immediately proceed to Step 3.3.2.4 without delay.
      4. Immediately add 550 µL PBS to initiate elution.
      5. Collect the void volume (700 µL).
      6. Discard the void volume or retain it as a non-EV fraction.
      7. Begin collecting the EV-enriched fraction immediately after the 700 µL void volume has eluted.
      8. Collect 850 µL of EV-enriched eluate following the 700 µL void volume.
      9. Alternatively, collect sequential fractions of approximately 170 µL to optimize purity versus yield.
        NOTE: Smaller fraction volumes are recommended when higher purity is required (e.g., for proteomic analyses or sensitive downstream applications), whereas larger fractions may be used to maximize overall EV yield.
      10. Assess fractions for EV markers and pool EV-enriched fractions as required.
        NOTE: Keep the column flowing continuously. As one fraction is finishing, add PBS to maintain a constant liquid above the resin bed and prevent the column from running dry.

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Results

To assess the effectiveness of this protocol for isolating tissue-derived sEVs, representative results from SkM and BM samples are presented. Successful isolation was defined by (i) enrichment of canonical EV markers (TSG101, CD63, CD81, and CD9) in early SEC fractions, (ii) minimal detection of intracellular contaminants (e.g., Calnexin and BiP), (iii) a unimodal particle size distribution with the majority of particles <200 nm, (iv) visualization of intact, membrane-bound vesicles by TEM, and (v) higher particle con...

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Discussion

This protocol describes a robust and reproducible approach for isolating sEV-enriched preparations directly from SkM and BM, two low-yield and highly heterogeneous tissues. While EV isolation is well established for biofluids and cell culture systems, standardized methods for tissue-derived EVs remain limited due to challenges associated with tissue dissociation, contamination from intracellular material, and variability in yield. Here, we outline critical steps, troubleshooting strategies, and limitations that influence...

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Disclosures

The authors declare no conflicts of interest.

Acknowledgements

We thank Jennie Yang, Dariusz Żurawek, and Volodymyr Yerko for technical assistance and helpful discussions. We are grateful to the Center for Applied Nanomedicine platform, particularly Nadim Tawil and Laura Montermini, for support with extracellular vesicle characterization and data acquisition. We also thank the animal facility staff, Guylaine Gadoury and Tara Thomson, for their assistance with mouse handling.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
1.5 mL centrifuge tubeSarstedt72.690.300General microcentrifuge tube used throughout protocol
1.5 mL low-protein-binding tubeSarstedt72.706.600Used for EV handling, SEC fraction collection, and storage
0.2 mL PCR tubeFisher Scientific14-222-262Used for poked-tube bone marrow extraction setup
16 G needleBD305197Used to puncture the 0.2 mL PCR tube
30 µm cell strainerMiltenyi Biotec 130-098-458Used for skeletal muscle sample filtration
70% ethanolSigma-Aldrich1.11727Used to sterilize tweezers
Biosafety cabinetNANAUsed for sterile tissue handling
Bone cutting instrumentFisher Scientific10-001-976Used to separate femur and tibia and expose marrow
Bovine serum albumin (BSA)BioShopALB001Used for Western blot blocking and antibody dilution
Calcium chloride dihydrate (CaCl2·2H2O)BioShopCCL302Used for HEPES buffer preparation
Carbon-coated copper grid, 200-meshSigma-Aldrich930342Used for TEM sample preparation
Centrifugal filter, 100 kDa MWCOMilliporeSigmaUFC810096Used for EV concentration
Collagenase DRoche11088858001Used for skeletal muscle enzymatic digestion
CryovialSarstedt72.690.300Used for tissue and bone storage
Deionized water (ultrapure)NANAUsed for buffer preparation and TEM washing steps
DNase IRoche11284932001Used for skeletal muscle enzymatic digestion
Dry iceNANAUsed for sample cooling and transport
Enhanced chemiluminescence substrateBiorad1705061Used for Western blot detection
Filter unit, 0.22 µmSarstedt83.3941.501Used to filter PBS where applicable
Forceps / tweezersMilliporeSigmaF3892Used for tissue handling
GlutaraldehydeNANAUsed for TEM fixation
GlycineNANAUsed in TEM grid washing steps
Hank’s Balanced Salt Solution (HBSS)Gibco14175-095Used for DNase I stock preparation
HEPESBioShopHEP001.500Used for HEPES-NaOH buffer preparation
Horseradish peroxidase-conjugated streptavidinThermo FisherS911Used for Western blot detection
Imaging system for chemiluminescenceBiorad12003153Used to image Western blot membranes
KClBioShopPOC308Used for HEPES buffer preparation
Magnesium chloride (MgCl2)BioShopMAG510.500Used for HEPES buffer preparation
2-MethylbutaneThermo Fisher78-78-4Used for skeletal muscle snap-freezing
Milligram balanceMilliporeSigmaZ741359Used for tissue and marrow weighing
NaClBioShopSOD004.1Used for HEPES buffer preparation
Nano-particle tracking analysis instrumentMalvern PanalyticalNanoSight NS300NanoSight NS300 used for particle sizing and counting
NaOHBioShopSHY500.500Used for pH adjustment
Nitrocellulose membraneBioRad1704271Used for Western blot transfer
Nutating mixerLabnet S0500Used during enzymatic digestion
P1000 pipetteMilliporeSigmaFA10006M-1EAUsed for tissue transfer
Pipette tip, 1 mLSarstedt70.3050.275Tip is trimmed to widen opening for tissue fragment transfer
Phosphate-buffered saline (PBS)WisentINC 811.012-FLUsed throughout protocol; also used for SEC equilibration and elution
Phosphatase inhibitorMilliporeSigma4906845001Used for Western blot lysate preparation
Petri dishMilliporeSigmaCLS351146Used for bone marrow preparation
Polyacrylamide gel, 4–20% gradient stain-freeBioRad4561094Used for Western blot separation
Protease inhibitorMilliporeSigma11697498001Used for Western blot lysate preparation
RPMI-1640 mediumSigma-AldrichR7388-500MLUnsupplemented; no FBS or antibiotics
RIPA buffer, 10×Merck20-188Used for protein extraction for Western blotting
Scalpel, sterile disposableUltident Paragon02-90000-10Used to mince skeletal muscle tissue
SEC column, original 70 nmIZONICO70PUsed for skeletal muscle SEC
SEC column, single 70 nmIZONICS70PUsed for bone marrow SEC
Sodium azideSigma-AldrichS2002Added to PBS for washing the column
Sodium cacodylate bufferNANAUsed for TEM fixation buffer
Luer Slip Single Use Syringe without Needle MedlineSYR101020Used during NTA acquisition if applicable
Transmission electron microscopeFEINATecnai G2 Spirit Twin 120 kV Cryo-TEM used for TEM imaging
Tween-20BioShopTWN508.1Used in PBS-T for Western blotting
Uranyl acetateNANAUsed for TEM negative staining
Whatman filter paperMilliporeSigmaWHA1003055Used to wick excess TEM stain

References

  1. Whitham, M., et al. Extracellular vesicles provide a means for tissue crosstalk during exercise. Cell Metab. 27 (1), 237-251.e4 (2018).
  2. Kumar, M. A., et al. Extracellular vesicles as tools and targets in therapy for diseases. Signal Transduct Target Ther. 9 (1), 27 (2024).
  3. Fonseka, P., Mathivanan, S. Extracellular vesicles biogenesis, cargo sorting and implications in disease conditions. Cells. 12 (2), (2023).
  4. Taraboletti, G., et al. Bioavailability of VEGF in tumor-shed vesicles depends on vesicle burst induced by acidic pH. Neoplasia. 8 (2), 96-103 (2006).
  5. Dakup, P. P., et al. Challenges and opportunities in state-of-the-art proteomics analysis for biomarker development from plasma extracellular vesicles. Proteomics. , (2025).
  6. Li, Y., et al. EV-origin: Enumerating the tissue-cellular origin of circulating extracellular vesicles using exLR profile. Comput Struct Biotechnol J. 18, 2851-2859 (2020).
  7. Shero, J. A., Lindholm, M. E., Sandri, M., Stanford, K. I. Skeletal muscle as a mediator of interorgan crosstalk during exercise: Implications for aging and obesity. Circ Res. 136 (11), 1407-1432 (2025).
  8. Vanhie, J. J., et al. The role of exercise- and high-fat diet-induced bone marrow extracellular vesicles in stress hematopoiesis. Front Physiol. 13, 1054463 (2022).
  9. Pinto, A. C., et al. Exploiting the therapeutic potential of contracting skeletal muscle-released extracellular vesicles in cancer: Current insights and future directions. J Mol Med (Berl). 102 (5), 617-628 (2024).
  10. Welsh, J. A., et al. Minimal information for studies of extracellular vesicles (MISEV2023): From basic to advanced approaches. J Extracell Vesicles. 13 (2), e12404 (2024).
  11. De Sousa, K. P., et al. Isolation and characterization of extracellular vesicles and future directions in diagnosis and therapy. Wiley Interdiscip Rev Nanomed Nanobiotechnol. 15 (1), e1835 (2023).
  12. Guo, J., et al. Establishment of a simplified dichotomic size-exclusion chromatography for isolating extracellular vesicles toward clinical applications. J Extracell Vesicles. 10 (11), e12145 (2021).
  13. Ahmadian, S., et al. Different storage and freezing protocols for extracellular vesicles: A systematic review. Stem Cell Res Ther. 15 (1), 453 (2024).
  14. Watanabe, S., et al. Skeletal muscle releases extracellular vesicles with distinct protein and microRNA signatures that function in the muscle microenvironment. PNAS Nexus. 1 (4), pgac173 (2022).
  15. Magalhaes-Gama, F., et al. Exploring cell-derived extracellular vesicles in peripheral blood and bone marrow of B-cell acute lymphoblastic leukemia pediatric patients: Proof-of-concept study. Front Immunol. 15, 1421036 (2024).
  16. Potter, D. R., et al. Mesenchymal stem cell-derived extracellular vesicles attenuate pulmonary vascular permeability and lung injury induced by hemorrhagic shock and trauma. J Trauma Acute Care Surg. 84 (2), 245-256 (2018).
  17. Osteikoetxea, X., et al. Differential detergent sensitivity of extracellular vesicle subpopulations. Org Biomol Chem. 13 (38), 9775-9782 (2015).
  18. Kim, T., Hong, J. W., Lee, L. P. Efficient methods of isolation and purification of extracellular vesicles. Nano Converg. 12 (1), 45 (2025).
  19. Bracht, J. W. P., et al. Choice of size-exclusion chromatography column affects recovery, purity, and miRNA cargo analysis of extracellular vesicles from human plasma. Extracell Vesicles Circ Nucl Acids. 5 (3), 497-508 (2024).
  20. Mo, W., et al. Extracellular vesicle-mediated bidirectional communication between the liver and other organs: Mechanistic exploration and prospects for clinical applications. J Nanobiotechnology. 23 (1), 190 (2025).

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