This work was supported by Department of Defense W81XWH2210261 (to Yao Yao).

This protocol isolates extracellular vesicles (EVs) from skeletal muscle tissue using mechanical detachment, enzymatic dissociation, filtration, and differential ultracentrifugation. It allows for the isolation of EVs from individual or combined rodent limb muscle groups, including the tibialis anterior (TA), extensor digitorum longus (EDL), soleus (SOL), gastrocnemius (GAS), and quadriceps (QUA). Animal use was approved by the Institutional Animal Care and Use Committee at the University of Georgia, and the study complied with the National Institutes of Health Guidelines for the care and use of animals in research. The animals were housed in the animal facilities within the Department of Animal and Dairy Science at the University of Georgia. Wild-type (C57BL/6J) adult male mice were used in this study, with tissue harvested from mice sacrificed at 3 months of age. Details of the reagents and equipment used are provided in the Table of Materials.
1. Mechanical detachment of muscle groups from the mouse
<ol>
	<li>Euthanize the mice with CO2, followed by cervical dislocation (following institutionally approved protocols). Shave both hindlimbs.</li>
	<li>Secure the mouse leg in a sterile plate with the anterior side of the leg facing upwards (Figure 1A).</li>
	<li>Make a small incision (5 mm) in the skin on the lateral side of the mouse ankle using dissection scissors.</li>
	<li>Insert one blade of the scissors carefully into the incision, positioning it beneath the skin but not in the underlying muscle tissue. Then, cut the skin upwards towards the knee to extend the incision length.</li>
	<li>Grab the skin along the incision using rough-ended tweezers and expand the opening until the muscle is visible.</li>
	<li>Locate the TA muscle, which runs from the knee to the ankle on the lateral side of the tibia bone11, and use sharp-end tweezers to grasp and peel off the thin layer of connective tissue covering the muscle (Figure 1B).</li>
	<li>Near the ankle, identify the two tendons at the end of the TA, one directly connected to the TA muscle and the other, the EDL tendon, which is smaller and located next to the TA tendon on the lateral side. Sever both tendons using dissection scissors.</li>
	<li>Grab both tendons with rough-ended tweezers. Then, use another pair of rough or sharp-ended tweezers to separate the tendons, thus separating the TA from the EDL (Figure 1C).</li>
	<li>Lift the TA by the freshly severed tendon with rough-ended tweezers. Then, cut the TA tendon near the knee using dissection scissors, thus removing the muscle. Repeat this process to remove the EDL, and store both muscles in PBS on ice.</li>
	<li>Release the leg and flip the mouse over so that the posterior face is facing up.</li>
	<li>With the skin on the leg now partially removed, cut the skin near the ankle using dissection scissors. Then, use rough-ended tweezers to grasp and peel the skin upwards towards the back of the knee, exposing the GAS.</li>
	<li>Sever the GAS tendon near the ankle using dissection scissors (Figure 1D).</li>
	<li>Grasp the GAS tendon with rough-ended tweezers and lift it upwards to expose the underside of the GAS.</li>
	<li>Locate the small SOL tendon on the underside of the GAS close to the knee (where the GAS is still connected). Sever the SOL tendon using dissection scissors, causing the SOL to contract upwards (Figure 1E).</li>
	<li>Grasp the severed SOL tendon with sharp-ended tweezers and lift it gently, thus detaching the SOL from the underside of the GAS.</li>
	<li>Sever the end of the SOL where it attaches to the severed end of the GAS using dissection scissors, thus separating the two muscles. Preserve the SOL in PBS on ice.</li>
	<li>Cut the end of the GAS near the knee and place it in PBS on ice.</li>
	<li>After removing the four listed muscles below the knee, flip the mouse over so that the anterior face is upwards.</li>
	<li>If needed, peel the skin further with rough-ended tweezers to expose the QUA. Position the mouse leg at a 90&#176; angle to force the QUA to flex. Sever the tendon closest to the knee using dissection scissors.
	<ol>
		<li>Separate the QUA from the femur by cutting between the muscle and bone until the QUA is only connected by the proximal tendon (Figure 1E). Then, sever the proximal tendon and remove the QUA. Place the QUA in PBS on ice.</li>
	</ol>
	</li>
	<li>Ensure that after muscle detachments, the tibia and fibula are clearly exposed (Figure 1F).</li>
	<li>Rinse the muscles several times with PBS to remove contaminants such as fur and blood (Figure 1G). Then, dry the muscles gently with paper towels. Use an analytical balance to measure the muscle weight, which will be used later to calculate the EV collected per gram of muscle.</li>
</ol>
2. Enzymatic dissociation of detached muscle chunks
<ol>
	<li>Gently remove muscle tendons from the muscle sample using sharp-end tweezers, splinter tweezers, scalpels, or scissors (Figure 2A).</li>
	<li>After tendon removal, cut the muscle longitudinally along the muscle fibers (from tendon to tendon) into 1 mm-thick muscle chunks using scissors or a scalpel to increase the surface area-to-volume ratio (Figure 2B,C). 
	NOTE: Try to limit myofiber damage during cutting. For larger muscles such as the TA, GAS, and QUA, make additional cuts compared to smaller muscles like the EDL and SOL to ensure a roughly equal surface area-to-volume ratio across all muscle samples (Figure 2D).</li>
	<li>Prepare 10 mL of fresh digestive enzyme buffer (DEB) for each mouse and keep it on ice until use.
	<ol>
		<li>To prepare DEB, mix 2 mg of collagenase II enzyme per 1 mL of Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM) with 1% penicillin-streptomycin solution. Filter the solution through a 0.2 &#181;m filter after mixing.</li>
		<li>Ensure the DEB has an enzymatic activity of over 250 units. Vortex the DEB until the solution is thoroughly mixed and the collagenase II enzyme powder is no longer visible.</li>
	</ol>
	</li>
	<li>Separate the muscle tissue into aliquots, each containing 20 mg of muscle tissue. Then, add 1 mL of DEB to each aliquot for enzymatic dissociation.</li>
	<li>Incubate the DEB and muscle mixture on a shaker in the incubator at 37 &#176;C for 12 h at a shaking speed of 100 rotations per minute (rpm) (Figure 2E).</li>
</ol>
3. Filtration and low-speed centrifugation to enrich SkM-EV
NOTE: Set the refrigerated centrifuge to 4 &#176;C and run it until it reaches the set temperature. Keep all needed tubes and samples on ice until ready for use.
<ol>
	<li>Following the enzymatic digestion procedures, ensure no discernible muscle chunks are visible (Figure 3A). Before refrigerated centrifugation, extract 10 &#181;L of solution from the DEB-muscle mixture and transfer it into a 0.2 mL tube for cell viability testing if needed.</li>
	<li>Prepare and label new empty 1.5 mL microcentrifuge tubes. Set up a 0.45 &#181;m filter and a 5 mL syringe by removing the plunger (do not discard) and screwing the hub into the top end of the 0.45 &#181;m filter. Position the filter so the bottom end is placed into a newly labeled empty tube (Figure 3D). Filter the 1x PBS through a 0.2 &#181;m filter.</li>
	<li>Centrifuge the mixture for 10 min at 1,000 x g and 4 &#176;C to pellet any debris (Figure 3B). Transfer the supernatant (around 1 mL) into the syringe using a P200 micro-pipette (Figure 3C,E).</li>
	<li>Reinsert the syringe plunger and slowly push the supernatant through the filter, allowing it to pass through and collect in the newly labeled 1.5 mL microcentrifuge tube (Figure 3F).</li>
	<li>Remove the plunger again and pipette roughly 200 &#181;L of 1x PBS into the syringe.</li>
	<li>Reinsert the plunger and slowly push the 200 &#181;L of 1x PBS through the filter (Figure 3G) to remove any remaining supernatant from the filter and into the labeled 1.5 mL microcentrifuge tube.</li>
	<li>Repeat the filtration process for other tubes, using a new filter for each tissue sample.</li>
	<li>After filtration, ensure the solution is transparent and devoid of residual muscle chunks (Figure 3H).</li>
</ol>
4. Differential ultracentrifugation for SkM-EV purification
NOTE: Set the refrigerated centrifuge to 4 &#176;C and run it until it reaches the set temperature. Keep all needed tubes and samples on ice and ready for use.
<ol>
	<li>Transfer the filtered solution to new 1.5 mL microtubes (suitable for up to 200,000 x g). Balance all microtubes with an additional 1x PBS to ensure equal volume before placing them into the micro-ultracentrifuge.</li>
	<li>After ensuring the volume of all tubes is equal, close the tubes and place them into the rotor of the micro-ultracentrifuge to be balanced (Figure 4A).</li>
	<li>Turn on the micro-ultracentrifuge and set it to run for 1 h at 100,000 x g and 4 &#176;C (Figure 4B).</li>
	<li>Place the rotor loaded with the tubes into the micro-ultracentrifuge, close the lid, and turn on the vacuum (Figure 4C).</li>
	<li>Once the vacuum reaches level 2, press the start button and wait until the centrifugation process is complete (Figure 4D).</li>
	<li>After the micro-ultracentrifuge cycle is complete, click on the vacuum button to repressurize the chamber before removing the rotor and tubes. Keep the tubes on ice and position them upright to avoid disturbing the pellet.</li>
	<li>Carefully remove the supernatant with a P200 micropipette, avoiding disturbance of the pellet (Figure 4E).</li>
	<li>Eject the tip, then add 50 &#181;L of 1x PBS into the tube after removing the supernatant using a P200 micropipette. For GAS and QUA samples, add up to 500 &#181;L of 1x PBS to aid dissolution (Figure 4F).</li>
	<li>Resuspend the pellet by gently pipetting up and down using a P20 micropipette (or P200 micropipette if 500 &#181;L was added) until it is fully broken up and no longer visible.</li>
	<li>Close the tube and store it at 4 &#176;C for short-term storage (less than one week), -80 &#176;C for long-term storage, or place the tube back on ice if the sample will be immediately characterized using nano-flow cytometry, Western blot, or other assays.</li>
</ol>

Optimized Isolation of Skeletal Muscle-Derived EVs for Research and Therapeutics

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The authors declare no conflicts of interest.

The protocol begins with dissecting hindlimb muscle groups from mice or acquiring postmortem human muscle tissue based on available protocols16. The following skeletal muscle extracellular vesicles (SkM-EVs) isolation procedures comprise three primary sections: mild enzymatic digestion of the tissue to liberate interstitial SkM-EVs, filtration and low-speed centrifugation, and ultracentrifugation. Finally, various techniques are applied to profile the isolated SkM-EVs.
Considering the significant differences among limb muscles in terms of size, myofiber type, and resistance to external stimuli17, this protocol provides detailed procedures for isolating SkM-EVs from each individual muscle, including the tibialis anterior (TA), extensor digitorum longus (EDL), soleus (SOL), gastrocnemius (GAS), and quadriceps (QUA). The techniques employed to detach muscle groups from mice and dissociate cells from these muscle tissues are adopted from those utilized for isolating intact myofibers and stem cells from adult rodent muscles11,18.
Subsequently, the tissue is enzymatically digested with collagenase to loosen the extracellular matrix (ECM), facilitating the release of cells while minimizing damage to cellular plasma membranes19. Specifically, collagenase II is often used in muscle tissue digestion to prepare a single-cell suspension20,21. However, enzymatic digestion with collagenase may result in the degradation of proteins on the surface of EVs, potentially compromising downstream analyses, such as proteomic profiling. To ensure a high EV yield while mitigating off-target protein degradation, this protocol includes several modifications to the collagenase-based tissue digestion compared to previously established protocols for muscle cell dissociation. First, a lower enzyme concentration is employed than what has been used for single-cell suspension preparation for flow cytometry. Second, the digestion duration is optimized to 12 h instead of 24 h to maximize extraction efficiency and minimize cell death22. Furthermore, DMEM is used to dissolve collagenase, providing a gentle environment for tissue digestion. These optimizations aim to reduce stimulus, control digestive damage, and preserve cell viability and the integrity of SkM-EVs. The cell viability immediately following enzymatic dissociation was evaluated. As expected, it was around 50%-60%, comparable to other studies using similar strategies to dissociate muscle tissue into single cells for various applications, such as flow cytometry-based cell sorting. 
Nano-flow cytometry-based analysis was applied to characterize the SkM-EVs isolated using this protocol. The median and mean size of SkM-EVs was around 70-80 nm, with a general distribution from 45-150 nm, which aligns with the range reported by other studies9,23. MemGlow-488 probes, a plasma membrane-targeting dye, confirmed that most particles (around 95%) of the SkM-EVs sample possess a membrane structure. Furthermore, Western blots were performed to verify the expression of EV-positive markers (CD8114 and Alix15) and negative markers (Tubulin) in isolated SkM-EVs.
The protocol described in this manuscript has been successfully applied to five different skeletal muscles: the TA, EDL, SOL, GAS, and QUA. These five skeletal muscles differ in size and myofiber type. Yet, the Skm-EVs isolated from these skeletal muscles exhibited consistent size distribution, particle count, and MemGlow-488 staining percentage (Figure 5), indicating the protocol's reliability across different muscle types. Notably, the muscles used to test this protocol were dissected from healthy WT mice. However, when isolating SkM-EVs from disease models, specific steps may need to be optimized. For instance, in chronic muscle injuries, increased fibrotic tissue may compromise digestion efficiency, requiring additional adjustments such as modifying enzyme concentration or digest time to ensure optimal SkM-EV yield.
Given the intrinsic nature of EVs in transferring bioactive cargos between cells nearby or at long distances, EVs, including SkM-EVs, are widely involved in critical physiological and pathologic processes. A reliable approach to isolate and characterize SkM-EVs will facilitate the exploration of EV-mediated intercellular communications in muscle development and related disease pathogenesis. In addition, collecting EVs from specific tissues with high quantity and purity holds the potential of developing EVs as diagnostic biomarkers for particular diseases. Moreover, an established strategy to isolate and purify tissue-specific EVs will enable the application of EVs for drug delivery due to their low immunogenicity and tissue-targeting specificity. Overall, further improvements for EV isolation, such as immunoaffinity capture-based purification, are needed to elucidate their function and activities in developmental and disease conditions and fulfill their biomedical applications.

Extracellular vesicles (EVs) are nano-sized, membrane-bound particles released by cells into the extracellular environment. These EVs transfer nucleic acids, proteins, and lipids to modulate multiple signaling pathways upon uptake by recipient cells. Specifically, EVs isolated from skeletal muscle (SkM-EVs) have been implicated in muscle degeneration, regeneration, and growth by transferring myogenic regulatory microRNAs or transcription factors1,2,3,4. As our understanding of SkM-EV subpopulations and characteristics continues to evolve, these vesicles show promise as biomarkers for early diagnostics5,6 and as therapeutic targets for neuromuscular disorders. This expanding field offers exciting opportunities for both pathological investigation and clinical applications.
Current research on SkM-EVs predominantly focuses on those obtained from cell culture supernatants following ex vivo culture2,7,8, raising concerns regarding the fidelity of target cell lines after successive passages and the susceptibility of EV characteristics to alteration in artificial environments. This highlights the need to isolate SkM-EVs directly from muscle tissues to avoid ex vivo culture influences and better represent in vivo conditions. Although several approaches for isolating EVs from muscle tissues have been developed9,10, studies still vary significantly in protocol details, impacting consistency and comparability. A standardized protocol for SkM-EV isolation and characterization is urgently needed to ensure reliability across related studies.
Considering the physiological characteristics of skeletal muscle, we developed a protocol to isolate and purify SkM-EVs from rodent skeletal muscle samples using mechanical detachment, enzymatic dissociation, filtration, and differential ultracentrifugation. Through comprehensive analysis using nano-flow cytometry, BCA assay, and Western blot assay, we found that this protocol consistently yields SkM-EVs of high purity and quantity within a limited timeframe. It can be applied to skeletal muscle samples in various conditions, including acute and chronic muscle injuries. Furthermore, with appropriate adjustments, this method can be tailored for muscle tissues from human patients or other animal models. Standardizing the SkM-EV isolation process is essential for ensuring consistency in SkM-EV quality, facilitating comparisons of EV characteristics -- such as yield, size, cargo composition, and function -- and advancing their potential applications as diagnostic biomarkers, as well as enhancing our understanding of their roles in various physiological and pathological processes.

<table><tbody><tr><td>0.6 mL microcentrifuge tubes</td><td>ThermoFisher Scientific</td><td>05-408-120</td><td></td></tr><tr><td>1.5 mL micro tube (C)&amp;nbsp;</td><td>Eppendorf</td><td>5720411009</td><td></td></tr><tr><td>1.5 mL microcentrifuge tubes</td><td>ThermoFisher Scientific</td><td>3448</td><td>Clear, graduated, nonsterile</td></tr><tr><td>BD 5 mL Syringe</td><td>Becton Dickinson</td><td>14-827-52&amp;nbsp;</td><td>luer-lok tip</td></tr><tr><td>Cell Counting Slides, Dual Chamber for Cell Counter</td><td>Bio-Rad</td><td>145-0011</td><td></td></tr><tr><td>Centrifuge 5430R</td><td>Eppendorf</td><td>22620667</td><td></td></tr><tr><td>Cleaning solution</td><td>NanoFCM Inc.</td><td></td><td></td></tr><tr><td>Collagenase II</td><td>Worthington Biochemical Corporation</td><td>LS004176</td><td></td></tr><tr><td>DPBS (10x) Liquid (-calcium,-magnesium)</td><td>Cytiva</td><td>SH30378.02</td><td>Final used dilution is 1x</td></tr><tr><td>DPBS/Modified (1x)</td><td>Cytiva</td><td>SH30264.02</td><td></td></tr><tr><td>Dulbecco&#039;s Modified Eagle&#039;s Medium (DMEM)</td><td>Genesee Scientific</td><td>25-500</td><td></td></tr><tr><td>E-POD Pure Water Remote Dispenser</td><td>Millipore Sigma</td><td>ZRXSP0D01</td><td></td></tr><tr><td>FA-45-30-11 Microcentrifuge Rotor</td><td>Eppendorf</td><td>05-401-503</td><td></td></tr><tr><td>Fisher Vortex Genie 2</td><td>ThermoFisher Scientific</td><td>12-812</td><td></td></tr><tr><td>Flow NanoAnalyzer</td><td>NanoFCM Inc.</td><td></td><td></td></tr><tr><td>Heratherm OGS60 General Protocol Oven</td><td>ThermoFisher Scientific</td><td>51028112</td><td>Gravity convection, 2.3 cu.ft. 120 V</td></tr><tr><td>MemGlow-488 Probe</td><td>Cytoskeleton Inc.</td><td>MG01-02</td><td></td></tr><tr><td>Millipak Special Millipore</td><td>Millipore Sigma</td><td>TANKMPK02</td><td></td></tr><tr><td>Penicillin-Streptomycin</td><td>Gibco</td><td>15070063</td><td></td></tr><tr><td>Pipet-Lite LTS Pipette L-1000XLS+</td><td>Rainin</td><td>17014382</td><td></td></tr><tr><td>Pipet-Lite LTS Pipette L-200XLS+</td><td>Rainin</td><td>17014391</td><td></td></tr><tr><td>Pipet-Lite LTS Pipette L-20XLS+</td><td>Rainin</td><td>17014392</td><td></td></tr><tr><td>Pipette tips ER LTS 1000&amp;micro;L F 768G/8</td><td>Rainin</td><td>30808038</td><td></td></tr><tr><td>Pipette tips ER LTS 20&amp;micro;L F 960G/10</td><td>Rainin</td><td>30807966</td><td></td></tr><tr><td>Pipette tips ER LTS 200&amp;micro;L F 960G/10</td><td>Rainin</td><td>30807967</td><td></td></tr><tr><td>Quality Control beads</td><td>NanoFCM Inc.</td><td></td><td></td></tr><tr><td>S110-AT Fixed Angle Rotor</td><td>ThermoFisher Scientific</td><td>45539</td><td>1,10,000 rpm</td></tr><tr><td>S16 exo 65-155nm beads</td><td>NanoFCM Inc.</td><td></td><td></td></tr><tr><td>Scissors</td><td>SPI, USA</td><td></td><td></td></tr><tr><td>Sorvall mX 150+ Micro-Ultracentrifuge</td><td>ThermoFisher Scientific</td><td>50135641</td><td></td></tr><tr><td>TC 20 automated cell counter</td><td>Bio-Rad</td><td>1450102</td><td></td></tr><tr><td>Trypan Blue Solution</td><td>Corning</td><td>25-900-CI</td><td></td></tr><tr><td>Tweezers</td><td>SPI, USA</td><td></td><td></td></tr><tr><td>VWR 15mL centrifuge tubes</td><td>Avantor</td><td>89039-668</td><td></td></tr><tr><td>Whatman Puradisc 13 Filters</td><td>Cytiva</td><td>6782-1304</td><td>0.45 &amp;micro;m pore size</td></tr><tr><td>Wild-type C57BL/6J mice</td><td>Jackson Laboratory</td><td>stock #000664</td><td></td></tr></tbody></table>

enzymatic isolation of skeletal muscle interstitial extracellular vesicles

Extracellular vesicles (EVs) are lipid bilayer-enclosed nanoparticles released by cells to transport bioactive cargo, such as proteins, RNAs, and DNAs, for intercellular communication. Investigating EV-mediated crosstalk among cells in muscle homeostasis and diseases offers significant potential to enhance our understanding of muscle development, regeneration, and atrophy. However, current protocols for isolating skeletal muscle-derived EVs (SkM-EVs) face challenges in achieving high purity and yield, primarily due to difficulties in releasing EVs from muscle tissues without compromising cellular membranes. This article presents an efficient protocol for SkM-EV isolation, comprising mechanical detachment, enzymatic dissociation, filtration, and ultracentrifugation. These steps are optimized to enhance EV release from muscle tissues, yielding high-purity SkM-EVs. Subsequently, nano-flow cytometry, BCA assay, and Western blot assay are performed to characterize the quantity and quality of the isolated SkM-EVs. This protocol holds promise for establishing a reliable platform to obtain tissue-derived EVs for advancing basic research, disease diagnosis, and drug delivery.

Following this protocol, skeletal muscle extracellular vesicles (SkM-EVs) were successfully extracted from various muscle groups, including the tibialis anterior (TA), extensor digitorum longus (EDL), soleus (SOL), gastrocnemius (GAS), and quadriceps (QUA) from wild-type mice. Subsequent characterization of these SkM-EVs was conducted through nano-flow cytometry, elucidating their distinct features as summarized in Table 1. Remarkably, from a mere 1 g of muscle tissue, an impressive yield of 1.18e14 &#177; 4.86e13 SkM-EVs was obtained, with a mean particle size of 73.93 nm &#177; 2.70 nm and a median particle size of 72.55 nm &#177; 2.42 nm. The size distribution showed a variance of 13.28 nm &#177; 1.71 nm (Figure 5), aligning precisely with the recognized definition of EVs12,13.
To ensure the integrity of the isolated SkM-EVs, MemGlow-488, a fluorogenic membrane probe, was employed to label the lipid structure within the SkM-EV samples (Figure 5), revealing an astounding 96.40% &#177; 2.42% preservation of lipid-wrapped membranes across the isolated population. Additionally, the purity of the SkM-EVs was rigorously assessed utilizing the Bicinchoninic acid (BCA) protein assay, which revealed a 4.82 &#181;g &#177; 2.28 &#181;g of proteins present per trillion EVs. Western blot analysis demonstrated the expression of EV-positive markers (CD8114 and Alix15) in isolated SkM-EVs, confirming their purity and absence of cellular Tubulin contaminants (Figure 6).
Next, cell viability assays using trypan blue dye were conducted to determine the viability of muscle resident cells during the isolation process. These tests confirmed that the protocol did not significantly impair the muscle resident cells, recording a cell viability of around 55%. Further analysis and characterization can be applied to these SkM-EVs to explore and understand their role as vital mediators of intercellular communication in muscle development and physiology.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67439/67439fig01.jpg" /> 
Figure 1:&#160;Procedure for mechanical detachment of mouse muscle groups. (A) The mouse leg is secured in a sterile plate with the anterior side facing upwards. (B) The TA is located, and sharp-end tweezers are used to grab and peel off the thin layer of connective tissue covering the muscle. (C) The EDL, adjacent to the TA, is identified and carefully separated from the TA. (D) The GAS and SOL tendons are located and severed. (E) The QUA muscle is identified and carefully separated from the femur. (F) Following muscle detachments, the tibia and fibula are clearly exposed. (G) The detached muscles are rinsed several times with PBS to remove any contaminants, such as fur and blood. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/67439/67439fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67439/67439fig02.jpg" /> 
Figure 2: Enzymatic dissociation of detached muscle chunks. (A) Muscle tendons are gently removed from the muscle sample. (B-D) Scissors or a scalpel are used to cut the muscle longitudinally along the muscle fibers (from tendon to tendon) into 1 mm-thick muscle chunks, ensuring uniformity in size to facilitate even enzymatic dissociation. (E) The DEB and muscle mixture is incubated on the shaker within the incubator for 12 h at 37 &#176;C with a shaking speed of 100 rpm. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/67439/67439fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67439/67439fig03.jpg" /> 
Figure 3: Filtration and low-speed centrifugation to enrich SkM-EVs. (A) After enzymatic digestion, no discernible muscle chunks should be visible in the mixture. (B) The mixture is centrifuged for 10 min at 1,000 x g and 4 &#176;C to pellet any remaining debris. (C-E) Using a 200 &#181;L pipette, the supernatant is carefully transferred 200 &#181;L at a time into a syringe. (F) The syringe plunger is reinserted, and the supernatant is slowly pushed through the filter to remove any remaining debris. (G) The plunger is reinserted, and 200 &#181;L of 1x PBS is slowly pushed through the filter to wash it. (H) After filtration, the solution should be transparent and devoid of any residual muscle chunks. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/67439/67439fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/67439/67439fig04.jpg" /> 
Figure 4: Differential ultracentrifugation for SkM-EV purification. (A) After ensuring the volume of all tubes is equal, the tubes are closed and placed in the micro-ultracentrifuge rotor, ensuring proper balance. (B) The micro-ultracentrifuge is turned on and set to 100,000 g for 1 h at 4 &#176;C. (C) The balanced rotor with the tubes is loaded into the micro-ultracentrifuge. The lid is closed, and the vacuum is turned on. (D) Once the vacuum reaches level 2, the start button is pressed to allow the ultracentrifuge to run until completion. (E) The supernatant is carefully removed without disturbing the pellet (indicated by the arrow). (F) After removing the supernatant, 50 &#181;L of 1x PBS is added into the tube using a 200 &#181;L or 100 &#181;L micropipette. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/67439/67439fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/67439/67439fig05.jpg" /> 
Figure 5: Size distribution pattern and MemGlow-488 staining. Size distribution pattern and MemGlow-488 staining of isolated SkM-EVs from different muscle groups, including TA (A,B), EDL (C,D), SOL (E,F), GAS (G,H), and QUA (I,J). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/67439/67439fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/67439/67439fig06.jpg" /> 
Figure 6: Western blot analysis. Western blot analysis of Tubulin, CD81, and Alix in SkM-EVs and skeletal muscle lysis. Equal amounts of protein (5 &#181;g per lane) are loaded onto the gel for each sample. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/67439/67439fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Parameter</td>
			<td>Mean &#177; Standard deviation (SD)</td>
		</tr>
		<tr>
			<td>SkM-EV yield (particles/gram of muscle)</td>
			<td>1.18e14 &#177; 4.86e13</td>
		</tr>
		<tr>
			<td>Mean particle size (nm)</td>
			<td>73.93 &#177; 2.70</td>
		</tr>
		<tr>
			<td>Median particle size (nm)</td>
			<td>72.55 &#177; 2.42</td>
		</tr>
		<tr>
			<td>Particle size distribution variance (nm)</td>
			<td>13.28 &#177; 1.71</td>
		</tr>
		<tr>
			<td>MemGlow-488-positive EV percentage (%)</td>
			<td>96.54 &#177; 1.85</td>
		</tr>
		<tr>
			<td>Protein concentration (&#181;g protein/ trillion EV)</td>
			<td>4.82 &#177; 2.28</td>
		</tr>
	</tbody>
</table>
Table 1: Characterization of isolated SkM-EVs. Size distribution and protein content are measured by nano-flow cytometry and the BCA protein assay, respectively.

Enzymatic Isolation of Skeletal Muscle Interstitial Extracellular Vesicles

This protocol aims to isolate and purify skeletal muscle interstitial extracellular vesicles (SkM-EVs) from rodent muscle tissues through mechanical detachment, enzymatic dissociation, filtration, and differential ultracentrifugation. It demonstrates consistency and reliability. The isolated SkM-EVs provide insights into muscle homeostasis and diseases, with potential applications as diagnostic biomarkers or therapeutic vehicles.

Extracellular vesicles (EVs) are lipid bilayer-enclosed nanoparticles released by cells to transport bioactive cargo, such as proteins, RNAs, and DNAs, ...

enzymatic-isolation-skeletal-muscle-interstitial-extracellular

Universidad San Sebastian

Research

JoVE Journal

Biology

788 Views.  University of Georgia. This protocol aims to isolate and purify skeletal muscle interstitial extracellular vesicles (SkM-EVs) from rodent muscle tissues through mechanical detachment, enzymatic dissociation, filtration, and differential ultracentrifugation. It demonstrates consistency and reliability. The isolated SkM-EVs provide insights into muscle homeostasis and diseases, with potential applications as diagnostic biomarkers or therapeutic vehicles.

Video: Enzymatic Isolation of Skeletal Muscle Interstitial Extracellular Vesicles

Isolation and Quantitative Immunocytochemical Characterization of Primary Myogenic Cells and Fibroblasts from Human Skeletal Muscle

The repair and regeneration of skeletal muscle requires the action of satellite cells, which are the resident muscle stem cells. These can be isolated from human muscle biopsy samples using enzymatic digestion and their myogenic properties studied in culture. Quantitatively, the two main adherent cell types obtained from enzymatic digestion are: (i) the satellite cells (termed myogenic cells or muscle precursor cells), identified initially as CD56+ and later as CD56+/desmin+ cells and (ii) muscle-derived fibroblasts, identified as CD56&#8211; and TE-7+. Fibroblasts proliferate very efficiently in culture and in mixed cell populations these cells may overrun myogenic cells to dominate the culture. The isolation and purification of different cell types from human muscle is thus an important methodological consideration when trying to investigate the innate behavior of either cell type in culture. Here we describe a system of sorting based on the gentle enzymatic digestion of cells using collagenase and dispase followed by magnetic activated cell sorting (MACS) which gives both a high purity (&#62;95% myogenic cells) and good yield (~2.8 x 106 &#177; 8.87 x 105 cells/g tissue after 7 days in vitro) for experiments in culture. This approach is based on incubating the mixed muscle-derived cell population with magnetic microbeads beads conjugated to an antibody against CD56 and then passing cells though a magnetic field. CD56+ cells bound to microbeads are retained by the field whereas CD56&#8211; cells pass unimpeded through the column. Cell suspensions from any stage of the sorting process can be plated and cultured. Following a given intervention, cell morphology, and the expression and localization of proteins including nuclear transcription factors can be quantified using immunofluorescent labeling with specific antibodies and an image processing and analysis package.

The main adherent cell types derived from human muscle are myogenic cells and fibroblasts. Here, cell populations are enriched using magnetic-activated cell sorting based on the CD56 antigen. Subsequent immunolabelling with specific antibodies and use of image analysis techniques allows quantification of cytoplasmic and nuclear characteristics in individual cells.

The repair and regeneration of skeletal muscle requires the action of satellite cells, which are the resident muscle stem cells. These can be isolated ...

Developmental Biology

Extraction of Extracellular Vesicles from Whole Tissue

Circulating and interstitial small membrane-bound extracellular vesicles (EVs) represent promising targets for the development of novel diagnostic or prognostic biomarker assays, and likely serve as important players in the progression of a vast spectrum of diseases. Current research is focused on the characterization of vesicles secreted from multiple cell and tissue types in order to better understand the role of EVs in the pathogenesis of conditions including neurodegeneration, inflammation, and cancer. However, globally consistent and reproducible techniques to isolate and purify vesicles remain in progress. Moreover, methods for extraction of EVs from solid tissue ex vivo are scarcely described. Here, we provide a detailed protocol for extracting small EVs of interest from whole fresh or frozen tissues, including brain and tumor specimens, for further characterization. We demonstrate the adaptability of this method for multiple downstream analyses, including electron microscopy and immunophenotypic characterization of vesicles, as well as quantitative mass spectrometry of EV proteins.

Here, we provide a detailed protocol to isolate small extracellular vesicles (EVs) from whole tissues, including brain and tumor specimens. This method offers a reproducible technique to extract EVs from solid tissue for further downstream analyses.

Circulating and interstitial small membrane-bound extracellular vesicles (EVs) represent promising targets for the development of novel diagnostic or ...

Cancer Research

An Enrichment Method for Small Extracellular Vesicles Derived from Liver Cancer Tissue

Small extracellular vesicles (sEVs) derived from tissue can reflect the functional status of the source cells and the characteristics of the tissue's interstitial space. The efficient enrichment of these sEVs is an important prerequisite to the study of their biological function and a key to the development of clinical detection techniques and therapeutic carrier technology. It is difficult to isolate sEVs from tissue because they are usually heavily contaminated. This study provides a method for the rapid enrichment of high-quality sEVs from liver cancer tissue. The method involves a four-step process: the incubation of digestive enzymes (collagenase D and DNase &#921;) with tissue, filtration through a 70 &#181;m cell strainer, differential ultracentrifugation, and filtration through a 0.22 &#181;m membrane filter. Owing to the optimization of the differential ultracentrifugation step and the addition of a filtration step, the purity of the sEVs obtained by this method is higher than that achieved by classic differential ultracentrifugation. It provides an important methodology and supporting data for the study of tissue-derived sEVs.

Here we describe the enrichment of small extracellular vesicles derived from liver cancer tissue through an optimized differential ultracentrifugation method.

Small extracellular vesicles (sEVs) derived from tissue can reflect the functional status of the source cells and the characteristics of the tissue's ...

Isolating Small Extracellular Vesicles and Extracting Their Peptidome

Identification of Peptides of Small Extracellular Vesicles from Bone Marrow-Derived Macrophages

Small extracellular vesicles (sEVs) are typically secreted by the exocytosis of multivesicular bodies (MVBs). These nanovesicles with a diameter of &lt;200 nm are present in various body fluids. These sEVs regulate various biological processes such as gene transcription and translation, cell proliferation and survival, immunity and inflammation through their cargos, such as proteins, DNA, RNA, and metabolites. Currently, various techniques have been developed for sEVs isolation. Among them, the ultracentrifugation-based method is considered the gold standard and is widely used for sEVs isolation. The peptides are naturally biomacromolecules with less than 50 amino acids in length. These peptides participate in a variety of biological processes with biological activity, such as hormones, neurotransmitters, and cell growth factors. The peptidome is intended to systematically analyze endogenous peptides in specific biological samples by liquid chromatography-tandem mass spectrometry (LC-MS/MS). Here, we introduced a protocol to isolate sEVs by differential ultracentrifugation and extracted peptidome for identification by LC-MS/MS. This method identified hundreds of sEVs-derived peptides from bone marrow-derived macrophages.

Author Spotlight: Peptidome Extraction from Small Extracellular Vesicles Isolated from Bone Marrow-Derived Macrophages 

This protocol describes a procedure to isolate small extracellular vesicles from macrophages by differential ultracentrifugation and extract the peptidome for identification by mass spectrometry.

Small extracellular vesicles (sEVs) are typically secreted by the exocytosis of multivesicular bodies (MVBs). These nanovesicles with a diameter of ...

Biochemistry

Mitochondrial Isolation from Skeletal Muscle

Mitochondria are organelles controlling the life and death of the cell. They participate in key metabolic reactions, synthesize most of the ATP, and regulate a number of signaling cascades2,3. Past and current researchers have isolated mitochondria from rat and mice tissues such as liver, brain and heart4,5. In recent years, many researchers have focused on studying mitochondrial function from skeletal muscles.
Here, we describe a method that we have used successfully for the isolation of mitochondria from skeletal muscles 6. Our procedure requires that all buffers and reagents are made fresh and need about 250-500 mg of skeletal muscle. We studied mitochondria isolated from rat and mouse gastrocnemius and diaphragm, and rat extraocular muscles. Mitochondrial protein concentration is measured with the Bradford assay. It is important that mitochondrial samples be kept ice-cold during preparation and that functional studies be performed within a relatively short time (~1 hr). Mitochondrial respiration is measured using polarography with a Clark-type electrode (Oxygraph system) at 37&deg;C7. Calibration of the oxygen electrode is a key step in this protocol and it must be performed daily. Isolated mitochondria (150 &mu;g) are added to 0.5 ml of experimental buffer (EB). State 2 respiration starts with addition of glutamate (5mM) and malate (2.5 mM). Then, adenosine diphosphate (ADP) (150 &mu;M) is added to start state 3. Oligomycin (1 &mu;M), an ATPase synthase blocker, is used to estimate state 4. Lastly, carbonyl cyanide p-[trifluoromethoxy]-phenyl-hydrazone (FCCP, 0.2 &mu;M) is added to measurestate 5, or uncoupled respiration 6. The respiratory control ratio (RCR), the ratio of state 3 to state 4, is calculated after each experiment. An RCR &ge;4 is considered as evidence of a viable mitochondria preparation.
In summary, we present a method for the isolation of viable mitochondria from skeletal muscles that can be used in biochemical (e.g., enzyme activity, immunodetection, proteomics) and functional studies (mitochondrial respiration).

This protocol describes a procedure to study the respiration of mitochondria isolated from skeletal muscles. This method was adapted from Scorrano et al. (2007). The mitochondrial isolation procedure requires about 2 hours. The mitochondrial respiration can be completed in about 1 hour.

Mitochondria are organelles controlling the life and death of the cell. They participate in key metabolic reactions, synthesize most of the ATP, and ...

Isolation and Culture of Individual Myofibers and their Satellite Cells from Adult Skeletal Muscle

Muscle regeneration in the adult is performed by resident stem cells called satellite cells. Satellite cells are defined by their position between the basal lamina and the sarcolemma of each myofiber. Current knowledge of their behavior heavily relies on the use of the single myofiber isolation protocol. In 1985, Bischoff described a protocol to isolate single live fibers from the Flexor Digitorum Brevis (FDB) of adult rats with the goal to create an in vitro system in which the physical association between the myofiber and its stem cells is preserved 1. In 1995, Rosenblattmodified the Bischoff protocol such that myofibers are singly picked and handled separately after collagenase digestion instead of being isolated by gravity sedimentation 2, 3. The Rosenblatt or Bischoff protocol has since been adapted to different muscles, age or conditions 3-6. The single myofiber isolation technique is an indispensable tool due its unique advantages. First, in the single myofiber protocol, satellite cells are maintained beneath the basal lamina. This is a unique feature of the protocol as other techniques such as Fluorescence Activated Cell Sorting require chemical and mechanical tissue dissociation 7. Although the myofiber culture system cannot substitute for in vivo studies, it does offer an excellent platform to address relevant biological properties of muscle stem cells. Single myofibers can be cultured in standard plating conditions or in floating conditions. Satellite cells on floating myofibers are subjected to virtually no other influence than the myofiber environment. Substrate stiffness and coating have been shown to influence satellite cells' ability to regenerate muscles 8, 9 so being able to control each of these factors independently allows discrimination between niche-dependent and -independent responses. Different concentrations of serum have also been shown to have an effect on the transition from quiescence to activation. To preserve the quiescence state of its associated satellite cells, fibers should be kept in low serum medium 1-3. This is particularly useful when studying genes involved in the quiescence state. In serum rich medium, satellite cells quickly activate, proliferate, migrate and differentiate, thus mimicking the in vivo regenerative process 1-3. The system can be used to perform a variety of assays such as the testing of chemical inhibitors; ectopic expression of genes by virus delivery; oligonucleotide based gene knock-down or live imaging. This video article describes the protocol currently used in our laboratory to isolate single myofibers from the Extensor Digitorum Longus (EDL) muscle of adult mice (6-8 weeks old).

Isolation and culture of myofibers is the gold standard in vitro system to study the transition of satellite cells through quiescence, activation and differentiation. Importantly, the single myofiber culture system preserves the myofiber/stem cell association, which is an essential component of the muscle stem cell niche.

Muscle  regeneration in the adult is performed by resident stem cells called satellite  cells. Satellite cells are defined by  their position between the ...

Isolation of Blood-vessel-derived Multipotent Precursors from Human Skeletal Muscle

Since the discovery of mesenchymal stem/stromal cells (MSCs), the native identity and localization&#160;of MSCs have been obscured by their retrospective isolation in culture. Recently, using fluorescence-activated cell sorting (FACS), we and other researchers prospectively identified and purified three subpopulations of multipotent precursor cells associated with the vasculature of human skeletal muscle. These three cell populations: myogenic endothelial cells (MECs), pericytes (PCs), and adventitial cells (ACs), are localized respectively to the three structural layers of blood vessels: intima, media, and adventitia. All of these human blood-vessel-derived stem cell (hBVSC) populations not only express classic MSC markers but also possess mesodermal developmental potentials similar to typical MSCs. Previously, MECs, PCs, and ACs have been isolated through distinct protocols and subsequently characterized in separate studies. The current isolation protocol, through modifications to the isolation process and adjustments in the selective cell surface markers, allows us to simultaneously purify all three hBVSC subpopulations by FACS from a single human muscle biopsy. This new method will not only streamline the isolation of multiple BVSC subpopulations but also facilitate future clinical applications of hBVSCs for distinct therapeutic purposes.

Blood vessels within human skeletal muscle harbor several multi-lineage precursor populations that are ideal for regenerative applications. This isolation method allows simultaneous purification of three multipotent precursor cell populations respectively from three structural layers of blood vessels: myogenic endothelial cells from intima, pericytes from media, and adventitial cells from adventitia.

Since the discovery of mesenchymal stem/stromal cells (MSCs), the native identity and localization&#160;of MSCs have been obscured by their retrospective ...

Single Myofiber Isolation and Culture from a Murine Model of Emery-Dreifuss Muscular Dystrophy in Early Post-Natal Development

Autosomal dominant Emery-Dreifuss muscular dystrophy (EDMD) is caused by mutations in the LMNA gene, which encodes the A-type nuclear lamins, intermediate filament proteins that sustain the nuclear envelope and the components of the nucleoplasm. We recently reported that muscle wasting in EDMD can be ascribed to intrinsic epigenetic dysfunctions affecting muscle (satellite) stem cells regenerative capacity. Isolation and culture of single myofibers is one of the most physiological ex-vivo approaches to monitor satellite cells behavior within their niche, as they remain between the basal lamina surrounding the fiber and the sarcolemma. Therefore, it represents an invaluable experimental paradigm to study satellite cells from a variety of murine models. Here, we describe a re-adapted method to isolate intact and viable single myofibers from post-natal hindlimb muscles (Tibialis Anterior, Extensor Digitorum Longus, Gastrocnemius and Soleus). Following this protocol, we were able to study satellite cells from Lamin &#916;8-11 -/- mice, a severe EDMD murine model, at only 19 days after birth.
We detail the isolation procedure, as well as the culture conditions for obtaining a good amount of myofibers and their associated satellite-cells-derived progeny. When cultured in growth-factors rich medium, satellite cells derived from wild type mice activate, proliferate, and eventually differentiate or undergo self-renewal. In homozygous Lamin &#916;8-11 -/- mutant mice these capabilities are severely impaired.
This technique, if strictly followed, allows to study all processes linked to the myofiber-associated satellite cell even in early post-natal developmental stages and in fragile muscles.

Here, we propose a method to efficiently obtain single muscle fibers at early post-natal developmental stages from homozygous mutant Lamin &#916;8-11 mouse model, a very severe model for Emery-Dreifuss muscular dystrophy (EDMD).

Autosomal dominant Emery-Dreifuss muscular dystrophy (EDMD) is caused by mutations in the LMNA gene, which encodes the A-type nuclear lamins, intermediate ...

Medicine

Isolation of Intact Mitochondria from Skeletal Muscle by Differential Centrifugation for High-resolution Respirometry Measurements

Mitochondria are involved in cellular energy metabolism and use oxygen to produce energy in the form of adenosine triphosphate (ATP). Differential centrifugation at low- and high-speed is commonly used to isolate mitochondria from tissues and cultured cells. Crude mitochondrial fractions obtained by differential centrifugation are used for respirometry measurements. The differential centrifugation technique is based on the separation of organelles according to their size and sedimentation velocity. The isolation of mitochondria is performed immediately after tissue harvesting. The tissue is immersed in an ice-cold homogenization medium, minced using scissors and homogenized in a glass homogenizer with a loose-fitting pestle. The differential centrifugation technique is efficient, fast and inexpensive and the mitochondria obtained by differential centrifugation are pure enough for respirometry assays. Some of the limitations and disadvantages of isolated mitochondria, based on differential centrifugation, are that the mitochondria can be damaged during the homogenization and isolation procedure and that large amounts of the tissue biopsy or cultured cells are required for the mitochondrial isolation.

Here, a quadriceps muscle specimen is taken from an anaesthetized pig and mitochondria are isolated by differential centrifugation. Then, the respiratory rates of mitochondrial respiratory chain complexes I, II and IV are determined using high-resolution respirometry.

Mitochondria are involved in cellular energy metabolism and use oxygen to produce energy in the form of adenosine triphosphate (ATP). Differential ...

Isolation and Characterization of Exosomes from Skeletal Muscle Fibroblasts

Exosomes are small extracellular vesicles released by virtually all cells and secreted in all biological fluids. Many methods have been developed for the isolation of these vesicles, including ultracentrifugation, ultrafiltration, and size exclusion chromatography. However, not all are suitable for large scale exosome purification and characterization. Outlined here is a protocol for establishing cultures of primary fibroblasts isolated from adult mouse skeletal muscles, followed by purification and characterization of exosomes from the culture media of these cells. The method is based on the use of sequential centrifugation steps followed by sucrose density gradients. Purity of the exosomal preparations is then validated by western blot analyses using a battery of canonical markers (i.e., Alix, CD9, and CD81). The protocol describes how to isolate and concentrate bioactive exosomes for electron microscopy, mass spectrometry, and uptake experiments for functional studies. It can easily be scaled up or down and adapted for exosome isolation from different cell types, tissues, and biological fluids.

This protocol illustrates the 1) the isolation and culture of primary fibroblasts from the adult mouse gastrocnemius muscle as well as 2) purification and characterization of exosomes using a differential ultracentrifugation method combined with sucrose density gradients followed by western blot analyses.

Exosomes are small extracellular vesicles released by virtually all cells and secreted in all biological fluids. Many methods have been developed for the ...