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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L., Levitt, D. E., Molina, P. E., Simon, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+expression+of+adipocyte+and+myotube+extracellular+vesicle+miRNA+cargo+in+chronic+binge+alcohol-administered+siv-infected+male+macaques.">Differential expression of adipocyte and myotube extracellular vesicle miRNA cargo in chronic binge alcohol-administered siv-infected male macaques.</a> Alcohol. 108, 1-9 (2023).</li><li>Kargl, C. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Angiogenic+potential+of+skeletal+muscle+derived+extracellular+vesicles+differs+between+oxidative+and+glycolytic+muscle+tissue+in+mice.">Angiogenic potential of skeletal muscle derived extracellular vesicles differs between oxidative and glycolytic muscle tissue in mice.</a> Sci Rep. 13 (1), 18943 (2023).</li><li>Vann, C. 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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.
...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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)&#160;</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&#160;</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&#39;s Modified Eagle&#39;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>
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			<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&#181;L F 768G/8</td>
			<td>Rainin</td>
			<td>30808038</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette tips ER LTS 20&#181;L F 960G/10</td>
			<td>Rainin</td>
			<td>30807966</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette tips ER LTS 200&#181;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>
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		<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>
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			<td>Scissors</td>
			<td>SPI, USA</td>
			<td></td>
			<td></td>
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		<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 &#181;m pore size</td>
		</tr>
		<tr>
			<td>Wild-type C57BL/6J mice</td>
			<td>Jackson Laboratory</td>
			<td>stock #000664</td>
			<td></td>
		</tr>
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</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 &#17...

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

Research

JoVE Journal

Biology

683 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

Adult and Embryonic Skeletal Muscle Microexplant Culture and Isolation of Skeletal Muscle Stem Cells

Cultured embryonic and adult skeletal muscle cells have a number of different uses. The micro-dissected explants technique described in this chapter is a robust and reliable method for isolating relatively large numbers of proliferative skeletal muscle cells from juvenile, adult or embryonic muscles as a source of skeletal muscle stem cells. The authors have used micro-dissected explant cultures to analyse the growth characteristics of skeletal muscle cells in wild-type and dystrophic muscles. Each of the components of tissue growth, namely cell survival, proliferation, senescence and differentiation can be analysed separately using the methods described here. The net effect of all components of growth can be established by means of measuring explant outgrowth rates. The micro-explant method can be used to establish primary cultures from a wide range of different muscle types and ages and, as described here, has been adapted by the authors to enable the isolation of embryonic skeletal muscle precursors.
Uniquely, micro-explant cultures have been used to derive clonal (single cell origin) skeletal muscle stem cell (SMSc) lines which can be expanded and used for in vivo transplantation. In vivo transplanted SMSc behave as functional, tissue-specific, satellite cells which contribute to skeletal muscle fibre regeneration but which are also retained (in the satellite cell niche) as a small pool of undifferentiated stem cells which can be re-isolated into culture using the micro-explant method.

The micro-dissected explants technique is a robust and reliable method for isolating proliferative skeletal muscle cells from juvenile, adult or embryonic muscles as a source of skeletal muscle stem cells. Uniquely, these cells have been clonally derived to produce skeletal muscle stem cell lines used for in vivo transplantation.

Cultured embryonic and adult skeletal muscle cells have a number of different uses. The micro-dissected explants technique described in this chapter is a ...

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 ...

Purification of Progenitors from Skeletal Muscle

Skeletal muscle contains multiple progenitor populations of distinct embryonic origins and developmental potential. Myogenic progenitors, usually residing in a "satellite cell position" between the myofiber plasma membrane and the laminin-rich basement membrane that ensheaths it, are self-renewing cells that are solely committed to the myogenic lineage1,2. We have recently described a second class of vessel associated progenitors that can generate myofibroblasts and white adipocytes, which responds to damage by efficiently entering proliferation and provides trophic support to myogenic cells during tissue regeneration3,4. One of the most trusted assays to determine the developmental and regenerative potential of a given cell population relies on their isolation and transplantation5-7. To this end we have optimized protocols for their purification by flow cytometry from enzymatically dissociated muscle, which we will outline in this article. The populations obtained using this method will contain either myogenic or fibro/adipogenic colony forming cells: no other cell types are capable of expanding in vitro or surviving in vivo delivery. However, when these populations are used immediately after the sort for molecular analysis (e.g qRT-PCR) one must keep in mind that the freshly sorted subsets may contain other contaminant cells that lack the ability of forming colonies or engrafting recipients.

Method for the enzymatic dissociation, surface labeling and purification by flow cytometry of fibro/adipogenic and myogenic progenitors from murine skeletal muscle.

Skeletal muscle contains multiple progenitor populations of distinct embryonic origins and developmental potential. Myogenic progenitors, usually residing ...

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 ...

Isolation of Mitochondria from Minimal Quantities of Mouse Skeletal Muscle for High Throughput Microplate Respiratory Measurements

Dysfunctional skeletal muscle mitochondria play a role in altered metabolism observed with aging, obesity and Type II diabetes. Mitochondrial respirometric assays from isolated mitochondrial preparations allow for the assessment of mitochondrial function, as well as determination of the mechanism(s) of action of drugs and proteins that modulate metabolism. Current isolation procedures often require large quantities of tissue to yield high quality mitochondria necessary for respirometric assays. The methods presented herein describe how high quality purified mitochondria (~ 450 &#181;g) can be isolated from minimal quantities (~75-100 mg) of mouse skeletal muscle for use in high throughput respiratory measurements. We determined that our isolation method yields 92.5&#177; 2.0% intact mitochondria by measuring citrate synthase activity spectrophotometrically. In addition, Western blot analysis in isolated mitochondria resulted in the faint expression of the cytosolic protein, GAPDH, and the robust expression of the mitochondrial protein, COXIV. The absence of a prominent GAPDH band in the isolated mitochondria is indicative of little contamination from non-mitochondrial sources during the isolation procedure. Most importantly, the measurement of O2 consumption rate with micro-plate based technology and determining the respiratory control ratio (RCR) for coupled respirometric assays shows highly coupled (RCR; &#62;6 for all assays) and functional mitochondria. In conclusion, the addition of a separate mincing step and significantly reducing motor driven homogenization speed of a previously reported method has allowed the isolation of high quality and purified mitochondria from smaller quantities of mouse skeletal muscle that results in highly coupled mitochondria that respire with high function during microplate based respirometirc assays.

Here, we present a modification of a previously reported method that allows for the isolation of high quality and purified mitochondria from smaller quantities of mouse skeletal muscle. This procedure results in highly coupled mitochondria that respire with high function during microplate based respirometirc assays.

Dysfunctional skeletal muscle mitochondria play a role in altered metabolism observed with aging, obesity and Type II diabetes. Mitochondrial ...

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 ...

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules derived from the parental cells. EVs function as intra- and inter-species communication vectors while also contributing to the interaction between bacteria and host organisms in the context of infection and colonization. Given the multitude of functions attributed to EVs in health and disease, there is a growing interest in isolating EVs for in vitro and in vivo studies. It was hypothesized that the separation of EVs based on physical properties, namely size, would facilitate the isolation of vesicles from diverse bacterial cultures. 
The isolation workflow consists of centrifugation, filtration, ultrafiltration, and size-exclusion chromatography (SEC) for the isolation of EVs from bacterial cultures. A pump-driven tangential flow filtration (TFF) step was incorporated to enhance scalability, enabling the isolation of material from liters of starting cell culture. Escherichia coli was used as a model system expressing EV-associated nanoluciferase and non-EV-associated mCherry as reporter proteins. The nanoluciferase was targeted to the EVs by fusing its N-terminus with cytolysin A. Early chromatography fractions containing 20-100 nm EVs with associated cytolysin A - nanoLuc were distinct from the later fractions containing the free proteins. The presence of EV-associated nanoluciferase was confirmed by immunogold labeling and transmission electron microscopy. This EV isolation workflow is applicable to other human gut-associated gram-negative and gram-positive bacterial species. In conclusion, combining centrifugation, filtration, ultrafiltration/TFF, and SEC enables scalable isolation of EVs from diverse bacterial species. Employing a standardized isolation workflow will facilitate comparative studies of microbial EVs across species.

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Bacteria secrete nanometer-sized extracellular vesicles (EVs) carrying bioactive biological molecules. EV research focuses on understanding their biogenesis, role in microbe-microbe and host-microbe interactions and disease, as well as their potential therapeutic applications. A workflow for scalable isolation of EVs from various bacteria is presented to facilitate standardization of EV research.

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules ...

Isolation of Mitochondria from Mouse Skeletal Muscle for Respirometric Assays

Most of the cell&#39;s energy is obtained through the degradation of glucose, fatty acids, and amino acids by different pathways that converge on the mitochondrial oxidative phosphorylation (OXPHOS) system, which is regulated in response to cellular demands. The lipid molecule Coenzyme Q (CoQ) is essential in this process by transferring electrons to complex III in the electron transport chain (ETC) through constant oxidation/reduction cycles. Mitochondria status and, ultimately, cellular health can be assessed by measuring ETC oxygen consumption using respirometric assays. These studies are typically performed in established or primary cell lines that have been cultured for several days. In both cases, the respiration parameters obtained may have deviated from normal physiological conditions in&#160;any given organ or tissue.
Additionally, the intrinsic characteristics of cultured single fibers isolated from skeletal muscle impede this type of analysis. This paper presents an updated and detailed protocol for the analysis of respiration in freshly isolated mitochondria from mouse skeletal muscle. We also provide solutions to potential problems that could arise at any step of the process. The method presented here could be applied to compare oxygen consumption rates in diverse transgenic mouse models and study the mitochondrial response to drug treatments or other factors such as aging or sex. This is a feasible method to respond to crucial questions about mitochondrial bioenergetics metabolism and regulation.

Here, we describe a detailed method for mitochondria isolation from mouse skeletal muscle and the subsequent analysis of respiration by Oxygen Consumption Rate (OCR) using microplate-based respirometric assays. This pipeline can be applied to study the effects of multiple environmental or genetic interventions on mitochondrial metabolism.

Most of the cell&#39;s energy is obtained through the degradation of glucose, fatty acids, and amino acids by different pathways that converge on the ...

Isolation and Characterization of Cyanobacterial Extracellular Vesicles

Cyanobacteria are a diverse group of photosynthetic, Gram-negative bacteria that play critical roles in global ecosystems and serve as essential biotechnology models. Recent work has demonstrated that both marine and freshwater cyanobacteria produce extracellular vesicles -&#160;small membrane-bound structures released from the outer surface of the microbes. While vesicles likely contribute to diverse biological processes, their specific functional roles in cyanobacterial biology remain largely unknown. To encourage and advance research in this area, a detailed protocol is presented for isolating, concentrating, and purifying cyanobacterial extracellular vesicles. The current work discusses methodologies that have successfully isolated vesicles from large cultures of Prochlorococcus, Synechococcus, and Synechocystis. Methods for quantifying and characterizing vesicle samples from these strains are presented. Approaches for isolating vesicles from aquatic field samples are also described. Finally, typical challenges encountered with cyanobacterial vesicle purification, methodological considerations for different downstream applications, and the trade-offs between approaches are also discussed.

The present protocol providesdetailed descriptions for isolation, concentration,and characterization of extracellular vesicles from cyanobacterial cultures. Approaches for purifying vesicles from cultures at different scales, trade-offs among methodologies, and considerations for working with field samples are also discussed.