1. Tissue Collection:
<ol>
<li>Use mice between 7 and 12 weeks old. Usually, at least two animals are used: one is to set up the FACS single-color controls required for compensation and for fluorescence-minus-one isotype control samples. The other is the actual sample for sorting.</li>
<li>Euthanize the mice by CO2 inhalation or according to an ethics protocol of choice. Place mice on a paper towel, facing up, and spray with 70% ethanol thoroughly to prevent contamination with the mouse fur.</li>
<li>Lift the skin in the abdominal region with forceps, then make a small longitudinal cut with sharp scissors.</li>
<li>Peel the two flaps of skin in opposite directions (up and down) to completely expose the hind limb muscles underneath.</li>
<li>Excise the muscle tissue by inserting and extending the scissors along both sides of the tibia.</li>
<li>Repeat the same process for the femur to extract the muscle tissue.</li>
<li>Place all excised muscle tissue in 60mm petri-dishes, using one dish for the tissue from each mouse.</li>
<li>Repeat the above for all mice.</li>
</ol>
2. Dissociate Tissue Into Single Cells by Enzymatic Digestion:
Throughout this section of the protocol, use sterile reagents and tools and work under sterile conditions.
<ol start="9">
<li>Add 2 mL of Collagenase II ( Sigma, cat#6885, 500u/ mL) and 20 &mu;L of 250mM CaCl2 stock to each Petri-dish.</li>
<li>Tear the muscle tissue into small pieces ( about 1 mm3) with two forceps (one serrated, one with 2 teeth, from Fine Science Tools , cat#11051-10, 11154-10), paying attention to remove and discard as much contaminated non-muscle tissue as possible.</li>
<li>Cover the Petri-dishes and incubate them at 37 degrees Celsius for 30 minutes.</li>
<li>Retrieve the Petri-dishes and use a 3 mL syringe plunger to thoroughly mash the tissue pieces. Use one plunger for each sample to avoid contaminations.</li>
<li>Add some (~ 5 mL) sterile cold PBS to each Petri-dish and transfer the tissue sludge into a 50mL Falcon tube (rinse the Petri-dish if necessary to recover all the tissue)</li>
<li>Shake the Falcon tube vigorously, fill the tube with PBS, centrifuge @800rpm x 5 minutes ( Eppendorf bench top Model :5810R), and aspirate the supernatant (repeat 3 times)</li>
<li>Add 1mL mixture of Collagenase D (Roche, Cat# 11088882001, 1.5u/ mL) /Dispase II (Roche, Cat# :04942078001, 2.4u/ mL) and 10&mu;L of CaCl2 (250mM) to each tube, incubate at 37 degrees Celsius with rotation for one hour. </li>
<li>Add 5-10 mL of cold, sterile PBS, vortex and pipette thoroughly to homogenize the tissue.</li>
<li>Fill up tubes with PBS and mix well.</li>
<li>Transfer the liquid to a 40&mu;m cell strainer placed on top of a 50 mL Falcon tube to filter out the remaining large pieces of tissue. Divide each sample evenly into three 50mL Falcon tubes. Fill up each of the tubes with PBS then centrifuge @1600rpm for 5 minutes at 8 degrees Celsius.</li>
<li>Discard the supernatant and collect cells for staining.</li>
</ol>
3. Antibody Staining and Sorting:
Throughout this section of the protocol, use sterile FACS and collection buffers and work under sterile conditions. Commercial antibodies are generally considered sterile if properly handled.
<ol start="20">
<li>For single color and isotype control samples, resuspend the sample in 1mL FACS buffer and divide the suspended cells in nine 1.5mL pre-labelled Eppendorf tubes (100 &mu;L each tube).
<ol>
<li>Set up single color staining mixes according to the following table: 
ATTENTION: The final concentration of isotype control antibodies in the stain should match that of the primary antibody they are the control for. For example, if anti-Sca-1 is used for 1 &mu;g/106 cells, its corresponding isotype must be used at the same concentration. 
<table border="1">
 <tbody>
 <tr>
 <td>Tube #</td>
 <td>Name of tube</td>
 <td>Company</td>
 <td>Cat#</td>
 <td>clone</td> 
 <td>isotype</td>
 <td>dilution</td>
 </tr>
 <tr>
 <td>1</td>
 <td>Non-stain</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>2</td>
 <td>Hoechst33342</td>
 <td>sigma</td>
 <td>B2261</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>2-5ug/ml</td>
 </tr>
 <tr>
 <td>3</td>
 <td>PI</td>
 <td>sigma</td>
 <td>P4864</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>1ug/ml</td>
 </tr>
 <tr>
 <td rowspan="2">4</td> 
 <td rowspan="2">CD31-FITC CD45-FITC</td> 
 <td>ebioscience</td>
 <td>11-0311-85</td>
 <td>390</td>
 <td>Rat IgG2a</td>
 <td>500</td>
 </tr>
 <tr>
 <td>House made</td>
 <td>&nbsp;</td>
 <td>I3/2</td>
 <td>Rat IgG2b</td>
 <td>500</td>
 </tr>
 <tr>
 <td>5</td>
 <td>Sca1-PECY7</td>
 <td>ebioscience</td>
 <td>25-5981-82</td>
 <td>D7</td>
 <td>Rat IgG2a</td>
 <td>5000</td>
 </tr>
 <tr>
 <td>6</td>
 <td>&alpha;-7 APC</td>
 <td>House made</td>
 <td>&nbsp;</td>
 <td>R2F2</td>
 <td>Rat IgG2b</td>
 <td>1000</td>
 </tr>
 </tbody>
</table>
</li>
<li>Set up isotype control staining mixes using the following table: 
 <table border="1">
<tbody>
 <tr>
 <td>Tube #</td>
 <td>Name of tube</td>
 <td>Hoechst</td>
 <td>PI</td>
 <td>CD31- FITC</td>
 <td>CD45- FITC</td>
 <td>Scal- PE-CY7</td>
 <td>a-7- APC</td>
 <td>Rat- IgG2b- APC</td>
 <td>Rat- IgG2a- pecy7</td>
 <td>Rat- IgG2a- FITC</td>
 <td>Rat- IgG2b- FITC</td>
 </tr>
 <tr>
 <td>7</td>
 <td>Iso- CD31 /CD45</td>
 <td>+</td>
 <td>+</td>
 <td>-</td>
 <td>-</td>
 <td>+</td>
 <td>+</td>
 <td>-</td>
 <td>-</td>
 <td>+</td>
 <td>+</td>
 </tr>
 <tr>
 <td>8</td>
 <td>Iso- Scal- pecy7</td>
 <td>+</td>
 <td>+</td>
 <td>+</td>
 <td>+</td>
 <td>-</td>
 <td>+</td>
 <td>-</td>
 <td>+</td>
 <td>-</td>
 <td>-</td>
 </tr>
 <tr>
 <td>9</td>
 <td>Iso- &alpha;-7-APC</td>
 <td>+</td>
 <td>+</td>
 <td>+</td>
 <td>+</td>
 <td>+</td>
 <td>-</td>
 <td>+</td>
 <td>-</td>
 <td>-</td>
 <td>-</td>
 </tr>
 </tbody>
</table>
</li>
<li>Pipet the single color and isotope control staining mixes (we usually adjust the concentration of antibodies so that between 5 and 10 &mu;L of staining mix are added to each sample) into the appropriate Eppendorf tube from Step 20. Mix well and incubate on ice for 1 hour.</li></ol></li>
<li>Stain cells from sample mouse with 800&mu;L of the antibody cocktail (below) for FACS cell sorting. Mix well and incubate on ice for 1 hour. 
Antibody cocktail: Hoechst, PI, CD31-FITC, CD45-FITC, Sca1-PECY7, &alpha;-7 APC 
ATTENTION: The optimal ratio of antibodies to cells must be determined by individually titrating each antibody, as significant differences may exist between batches. The optimal amount of antibody to be used should be expressed in &mu;gs of antibody per 106 target cells and kept constant when using the same batch of antibody. Clearly, if the number of target cells does not change significantly between experiments, or if the staining volume is scaled with the number of cells, a fixed dilution of each antibody can also be used.</li>
<li>Wash: Add approximately 1mL of FACS buffer to each of the 9 control tubes; add approximately 20mL of FACS buffer to the sample tube.</li>
<li>Centrifuge the Eppendorf tubes @ 3000rpm for 5 minutes ( Eppendorf bench top centrifuge, Model:5417C ). Centrifuge the sample tube @1600rpm for 5 minutes (Eppendorf bench top centrifuge, Model:5810R).</li>
<li>Aspirate and discard the supernatant from all the tubes.</li>
<li>Resuspend cells in the FACS buffer (1mL for each of the controls, 4mL for the sorting sample), Pipette cells into "FACS tubes" through the cell strainer cap (BD Falcon cat# 352235, 40&mu;m pore size) to remove remaining clumps that may affect the cytometer.</li>
<li>Set up the FACS Sorter with single color controls and isotype controls.</li>
<li>Collect sorted cells according to the following table in a 3.5 mL collection buffer (95% DMEM, 5%FBS) separately. Normally, when harvesting both hind limbs, a single wild type mouse could yield about 150,000 myogenic progenitor cells or 100,000 adipogenic progenitor cells, with the viability above 95%, as judged by reanalyzing sorted cells by FACS at the end of the procedure (Figure 1) 
Definition of myogenic and adipogenic progenitors based on surface staining 
 <table border="1">
<tbody>
 <tr>
 <td>Antibody/stain </td>
 <td>Myogenic progenitor (MP)</td>
 <td>Adipogenic progenitor (AP)</td>
 </tr>
 <tr>
 <td>Hoechst</td>
 <td>+</td>
 <td>+</td>
 </tr>
 <tr>
 <td>PI</td>
 <td>-</td>
 <td>-</td>
 </tr>
 <tr>
 <td>CD31-FITC</td>
 <td>-</td>
 <td>-</td>
 </tr>
 <tr>
 <td>CD45-FITC</td>
 <td>-</td>
 <td>-</td>
 </tr>
 <tr>
 <td>Sca1-PECY7</td>
 <td>-</td>
 <td>+</td>
 </tr>
 <tr>
 <td>&alpha;-7 APC</td>
 <td>+</td>
 <td>-</td>
 </tr>
 </tbody>
</table>
</li>
</ol>
4. Transplantation:
<ol start="28">
<li>Centrifuge sorted cells @1500rpm for 5 minutes, remove the supernatant and transfer cells to autoclaved Eppendorf tubes (1.5 mL),
rinse cells with 500 &mu;L of sterile PBS (serum free!), then centrifuge @ 3000rpm for 5 minutes. Maintaining all reagents in sterile conditions and working in a tissue culture hood, resuspend myogenic progenitors in PBS, or adipogenic progenitor in Matrigel ( BD cat# 354234) at approximately 106 cells /mL.</li>
<li>Transplant myogenic progenitors or adipogenic progenitor to recipient mice.
<ol>
<li>Anaesthetize the mouse with isoflouran according to your institution policy.</li>
<li>Shave the hair around the injection region, then sterilize the skin by rubbing with 70% ethanol</li>
<li>Inject 20&mu;L of myogenic progenitors or adipogenic progenitors (= 20k cells) using a 3/10 CC insulin syringe ( BD cat# 309300).</li></ol></li>
<li>After an appropriate time (three weeks works well, but myofibers expressing donor-derived transgenic markers are usually evident within one week after transplant), anaesthetize the recipient mice from step 29 by intraperitoneal injection of 400 &mu;L Avertin (Sigma Cat# T04840-2, 25 mg/ mL), then perfuse transcardially with 50mL PBS (contains 10 &mu;M of EDTA) first, followed by 15mL of 4% PFA. Collect target tissue, post-fix with 4% PFA overnight if required, then transfer to 20% sucrose overnight. Embed in OCT, freeze and cryosection.</li>
</ol>
5. Representative Results:
When MP are transplanted into mouse skeletal muscle, they readily fuse with pre-existing myofibers. Therefore any genetic label present in the donor cells will be easily detectable in the fibers that received them. Figure 2 shows an example where the transplanted cells expressed human alkaline phostphatase, revealed histochemically.
 When AP are transplanted the result is highly dependent on the environment of the transplantation site: when transplanted subcutaneously these cells give origins to adipocytes and myofibroblasts (see figure 2). In many other sites, including muscle, they do not survive unless fatty degeneration has been induced3.
 
 <img src="/files/ftp_upload/2476/2476fig1.jpg" alt="Figure 1" /> 
 Figure 1. Sorting strategy for the isolation of progenitor populations from skeletal muscle. (A) Sorting strategy: Viable cells were identified based on forward scatter and side scatter. Hoechst staining was used to exclude anuclear debris and propidium iodide (PI) staining to exclude dead cells. Hematopoietic (CD45) and endothelial (CD31) cells were excluded from the sorting gates. The &alpha;7+ Hoechst+ PI- CD45- CD31- Scal- subset contains all myogenic progenitors (MP). The Scal+ Hoechst+ PI- CD45- CD31- &alpha;7- population contains adipogenic progenitors (AP). (B) Fluorescence-minus-one (FMO) isotype controls confirm the specificity of the stain. (C) Purity checks of MP and AP subsets after sorting.
 <img src="/files/ftp_upload/2476/2476fig2.jpg" alt="Figure 2" /> 
 Figure 2. Representative results following MP and AP transplantation. AP were transplanted subcutaneously (top panel) and MP intramuscularly (bottom panel). In both cases, donor cells originated from a mouse expressing transgenic human alkaline phosphatase, identified by the brown staining.

<ol>
	<li>Buckingham, M., Montarras, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Skeletal+muscle+stem+cells.">Skeletal muscle stem cells.</a> Curr Opin Genet Dev. 18, 330-336 (2008).</li><li>Relaix, F., Marcelle, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Muscle+stem+cells.">Muscle stem cells.</a> Curr Opin Cell Biol. 21, 748-753 (2009).</li><li>Joe, A. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Muscle+injury+activates+resident+fibro/adipogenic+progenitors+that+facilitate+myogenesis.">Muscle injury activates resident fibro/adipogenic progenitors that facilitate myogenesis.</a> Nat Cell Biol. 12, 153-163 (2010).</li><li>Natarajan, A., Lemos, D. R., Rossi, F. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fibro/adipogenic+progenitors:+A+double-edged+sword+in+skeletal+muscle+regeneration.">Fibro/adipogenic progenitors: A double-edged sword in skeletal muscle regeneration.</a> Cell Cycle. 9, (2010).</li><li>Sherwood, R. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+of+adult+mouse+myogenic+progenitors:+functional+heterogeneity+of+cells+within+and+engrafting+skeletal+muscle.">Isolation of adult mouse myogenic progenitors: functional heterogeneity of cells within and engrafting skeletal muscle.</a> Cell. 119, 543-554 (2004).</li><li>Sacco, A., Doyonnas, R., Kraft, P., Vitorovic, S., Blau, H. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Self-renewal+and+expansion+of+single+transplanted+muscle+stem+cells.">Self-renewal and expansion of single transplanted muscle stem cells.</a> Nature. , (2008).</li><li>Montarras, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+isolation+of+satellite+cells+for+skeletal+muscle+regeneration.">Direct isolation of satellite cells for skeletal muscle regeneration.</a> Science. 309, 2064-2067 (2005).</li></ol>

No conflicts of interest declared.

The goal of this protocol is to strike a reasonable balance between high yields and high viability of the purified cells. The most critical step in ensuring that healthy cells are recovered is, predictably, the enzymatic dissociation of the starting tissue. The handling of the tissue should be particularly gentle, which is hard to demonstrate or appreciate even in an audiovisual format. Another key factor is the length of the procedure. The longer it takes to go from the donor to the recipient animal, the lower the viability and therefore, the engraftment efficiency. Should there be any question about viability that is not immediately answered by looking at the frequency of PI positive event in the purified cell samples after sorting, we suggest that a limiting dilution assay to measure clonogenicity is performed3. Typical sorts from undamaged muscle routinely yield about 1 in 15-20 cells capable of initiating myogenic or fibro/adipogenic colonies in vitro. While essentially 100% of the colonies initiated from MPs contain differentiated myotubes, only about one third of colonies obtained from FAPs contain adipocytes in addition to smooth muscle actin positive myofibroblasts.

<table border="1">
<tbody>
 <tr>
 <th>Name of the reagent</th>
 <th>Company</th>
 <th>Catalogue number</th>
 <th>Comments</th>
 </tr>
 <tr>
 <td>Collagenase II</td>
 <td>Sigma-Aldrich</td>
 <td>C-6885,</td>
 <td>500 u/ml</td>
 </tr>
 <tr>
 <td>Collagenase D</td>
 <td>Roche</td>
 <td>11088882001</td>
 <td>1.5 u/ml</td>
 </tr>
 <tr>
 <td>Displace II</td>
 <td>Roche</td>
 <td>04942078001</td>
 <td>2.4 u/ml</td>
 </tr>
 <tr>
 <td>cell strainer</td>
 <td>BD</td>
 <td>352340</td>
 <td>40 &mu;m</td>
 </tr>
 <tr>
 <td>Tube with cell strainer</td>
 <td>BD</td>
 <td>352235</td>
 <td>40 &mu;m</td>
 </tr>
 <tr>
 <td>Hoechst33342</td>
 <td>Sigma-Aldrich</td>
 <td>B2261</td>
 <td>2-5 ug/ml</td>
 </tr>
 <tr>
 <td>CD31-FITC</td>
 <td>Ebiosciences</td>
 <td>11-0311-85</td>
 <td>Clone 390</td>
 </tr>
 <tr>
 <td>CD45-FITC</td>
 <td>Home made</td>
 <td>&nbsp;</td>
 <td>Clone I3/2</td>
 </tr>
 <tr>
 <td>Sca1-PECY7</td>
 <td>Ebiosciences</td>
 <td>25-5981-82</td>
 <td>Clone D7</td>
 </tr>
 <tr>
 <td>&alpha;-7 integrin APC</td>
 <td>Home made</td>
 <td>Inquire with Rossi lab</td>
 <td>R2F2</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Purification of Progenitors from Skeletal Muscle at JoVE.com

Purification of Progenitors from Skeletal Muscle

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 ...

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18.2K Views.  University of British Columbia. Method for the enzymatic dissociation, surface labeling and purification by flow cytometry of fibro/adipogenic and myogenic progenitors from murine skeletal muscle.

Video: Purification of Progenitors from Skeletal Muscle

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 ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections (bupivacaine, cardiotoxin or notexin), muscle transplantations (denervation-devascularization induced regeneration), intensive exercise, but also murine muscular dystrophy models such as the mdx mouse (for a review of these approaches see 1). In zebrafish, genetic approaches include mutants that exhibit muscular dystrophy phenotypes (such as runzel2 or sapje3) and antisense oligonucleotide morpholinos that block the expression of dystrophy-associated genes4. Besides, chemical approaches are also possible, e.g. with Galanthamine, a chemical compound inhibiting acetylcholinesterase, thereby resulting in hypercontraction, which eventually leads to muscular dystrophy5. However, genetic and pharmacological approaches generally affect all muscles within an individual, whereas the extent of physically inflicted injuries are more easily controlled spatially and temporally1. Localized physical injury allows the assessment of contralateral muscle as an internal control. Indeed, we recently used laser-mediated cell ablation to study skeletal muscle regeneration in the zebrafish embryo6, while another group recently reported the use of a two-photon laser (822 nm) to damage very locally the plasma membrane of individual embryonic zebrafish muscle cells7.
Here, we report a method for using the micropoint laser (Andor Technology) for skeletal muscle cell injury in the zebrafish embryo. The micropoint laser is a high energy laser which is suitable for targeted cell ablation at a wavelength of 435 nm. The laser is connected to a microscope (in our setup, an optical microscope from Zeiss) in such a way that the microscope can be used at the same time for focusing the laser light onto the sample and for visualizing the effects of the wounding (brightfield or fluorescence). The parameters for controlling laser pulses include wavelength, intensity, and number of pulses.
Due to its transparency and external embryonic development, the zebrafish embryo is highly amenable for both laser-induced injury and for studying the subsequent recovery. Between 1 and 2 days post-fertilization, somitic skeletal muscle cells progressively undergo maturation from anterior to posterior due to the progression of somitogenesis from the trunk to the tail8, 9. At these stages, embryos spontaneously twitch and initiate swimming. The zebrafish has recently been recognized as an important vertebrate model organism for the study of tissue regeneration, as many types of tissues (cardiac, neuronal, vascular etc.) can be regenerated after injury in the adult zebrafish10, 11.

The method presented here comprises the precise injury of live zebrafish embryos with high-energy laser pulses and the subsequent analysis of these injuries and their recovery with time. We also show how genetically labeled single or groups of skeletal muscle cells can be tracked during and after laser light induced damage.

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections ...

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 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 ...

Identification, Isolation, and Characterization of Fibro-Adipogenic Progenitors (FAPs) and Myogenic Progenitors (MPs) in Skeletal Muscle in the Rat

Fibro-adipogenic Progenitors (FAPs) are resident interstitial cells in skeletal muscle that, together with myogenic progenitors (MPs), play a key role in muscle homeostasis, injury, and repair. Current protocols for FAPs identification and isolation use flow cytometry/fluorescence-activated cell sorting (FACS) and studies evaluating their function in vivo to date have been undertaken exclusively in mice. The larger inherent size of the rat allows for a more comprehensive analysis of FAPs in skeletal muscle injury models, especially in severely atrophic muscle or when investigators require substantial tissue mass to conduct multiple downstream assays. The rat additionally provides a larger selection of muscle functional assays that do not require animal sedation or sacrifice, thus minimizing morbidity and animal use by enabling serial assessments. The flow cytometry/FACS protocols optimized for mice are species specific, notably restricted by the characteristics of commercially available antibodies. They have not been optimized for separating FAPs from rat or highly fibrotic muscle. A flow cytometry/FACS protocol for the identification and isolation of FAPs and MPs from both healthy and denervated rat skeletal muscle was developed, relying on the differential expression of surface markers CD31, CD45, Sca-1, and VCAM-1. As rat-specific, flow cytometry-validated primary antibodies are severely limited, in-house conjugation of the antibody targeting Sca-1 was performed. Using this protocol, successful Sca-1 conjugation was confirmed, and flow cytometric identification of FAPs and MPs was validated by cell culture and immunostaining of FACS-isolated FAPs and MPs. Finally, we report a novel FAPs time-course in a prolonged (14 week) rat denervation model. This method provides the investigators the ability to study FAPs in a novel animal model.

Identification, Isolation, and Characterization of Fibro-Adipogenic Progenitors FAPs and Myogenic Progenitors MPs in Skeletal Muscle in the Rat

This protocol outlines a method to isolate Fibro-adipogenic progenitors (FAPs) and myogenic progenitors (MPs) from rat skeletal muscle. Utilization of the rat in muscle injury models provides increased tissue availability from atrophic muscle for the analysis and a larger repertoire of validated methods to assess muscle strength and gait in free-moving animals.

Fibro-adipogenic Progenitors (FAPs) are resident interstitial cells in skeletal muscle that, together with myogenic progenitors (MPs), play a key role in ...

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 ...

Electroporation of Plasmid DNA into Mouse Skeletal Muscle

Transient gene expression modulation in murine skeletal muscle by plasmid electroporation is a useful tool for assessing normal and pathological physiology. Overexpression or knockdown of target genes enables investigators to manipulate individual molecular events and, thus, better understand the mechanisms that impact muscle mass, muscle metabolism, and contractility. In addition, electroporation of DNA plasmids that encode fluorescent tags allows investigators to measure changes in subcellular localization of proteins in skeletal muscle in vivo. A key functional assessment of skeletal muscle includes the measurement of muscle contractility. In this protocol, we demonstrate that whole muscle contractility studies are still possible after plasmid DNA injection, electroporation, and gene expression modulation. The goal of this instructional procedure is to demonstrate the step-by-step method of DNA plasmid electroporation into mouse skeletal muscle to facilitate uptake and expression in the myofibers of the tibialis anterior and extensor digitorum longus muscles, as well as to demonstrate that skeletal muscle contractility is not compromised by injection and electroporation.

Electroporation of plasmid DNA into skeletal muscle is a viable method to modulate gene expression without compromising muscle contractility in mice.