Name: single myofiber culture assay for the assessment of adult muscle stem cell functionality ex vivo
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Description: Watch this Scientific Journal Video about Single Myofiber Culture Assay for the Assessment of Adult Muscle Stem Cell Functionality Ex Vivo at JoVE.com

We thank Christine Poser and Christina Picker for excellent technical assistance and critical reading of the manuscript. This work was supported by a grant from the Deutsche Forschungsgemeinschaft to JvM (MA-3975/2-1), the Carl Zeiss foundation and the Deutsche Krebshilfe (DKH-JvM-861005).

Sacrificing of animals must be performed in accordance with the national regulations for animal experimentation. The protocol described here was performed in accordance with the guidelines of the Leibniz Institute on Aging - Fritz Lipmann Institute and the European Union (EU) directive 2010/63/EU (license number for organ harvesting: O_JvM_18-20). Essential steps of the protocol are summarized in Figure 1.
1. Preparation of culture plates, media, and Pasteur pipettes
NOTE: All materials and equipment necessary to isolate and culture single myofibers need to be as sterile as possible. Therefore, it is recommended to isolate single myofibers under a semi-sterile dissection hood.
<ol>
	<li>Use sterile HS (horse serum) to coat tissue culture plates. Coating prevents the attachment of single myofibers to the plastic surface. For each mouse, 4 wells of a 12-well plate for the isolation and a dedicated number of wells of a 24-well plate for culturing are required. Incubate the wells of a 12-well plate with 1 mL and the wells of a 24-well plate with 0.5 mL of HS for 5 min at RT (room temperature), then remove the HS and let the plates dry for another 5 min. 
	NOTE: HS can be collected and re-used for coating purposes multiple times, if kept sterile.</li>
	<li>Prepare myofiber isolation medium by supplementing DMEM (Dulbecco&#39;s modified Eagle&#39;s medium with 4.5 g/L glucose and sodium pyruvate) with 20% FBS (fetal bovine serum), filter through 0.22 &#181;m filter. Add medium to the pre-coated isolation plates (12-well plate, 2-4 mL isolation medium per well) approximately 30 min before the isolation and equilibrate in a humidified 37 &#176;C incubator with 5% CO2.</li>
	<li>Prepare myofiber culture medium by supplementing DMEM (Dulbecco&#39;s modified Eagle&#39;s medium with 4.5 g/L glucose and sodium pyruvate) with 20% FBS (fetal bovine serum) and 1% chicken embryo extract, filter through 0.22 &#181;m filter. Add 0.5 mL medium to each well of the pre-coated culture plates (24-well plate) approximately 30 min before the isolation and equilibrate in a humidified 37 &#176;C incubator with 5% CO2.</li>
	<li>Prepare collagenase digestion solution by dissolving 0.2% (w/v) collagenase type 1 (from Clostridium histolyticum) in DMEM (Dulbecco&#39;s modified Eagle&#39;s medium with 4.5 g/L glucose and sodium pyruvate), filter through 0.22 &#181;m filter. For two EDL (extensor digitorum longus) muscles use 2.5 mL collagenase solution in a sterile 15 mL reaction tube. Additionally, pre-heat the solution ~10 min in a circulating water bath at 37 &#176;C before starting the isolation.</li>
	<li>Prepare sterile Pasteur pipettes for trituration of the collagenase-digested muscles. Use a diamond pen to cut the Pasteur pipettes.
	<ol>
		<li>For each mouse, use one large bore pipette with an opening of about 0.3 cm and a length of about 10-12 cm and a second glass pipette with a small opening of about 0.1 cm and a length of approximately 22 cm (Figure 2F).</li>
		<li>Smoothen the edges of both pipettes by heat polishing with the flame of a Bunsen burner. Hold the pipette tip for 5-10 s into the flame with gentle movement to allow equal heat distribution until the sharp glass edges smoothen.</li>
		<li>Immediately before use, coat both types of glass pipettes with sterile HS by filling the whole pipette with ~ 2 mL of HS for 5 min, thereafter, eject the HS and let the pipettes dry for 5 min at RT.</li>
	</ol>
	</li>
</ol>
2. EDL muscle isolation and collagenase digestion
<ol>
	<li>Spray all equipment with 70% ethanol to avoid contamination.</li>
	<li>Sacrifice the mouse in accordance with the national regulations for animal experimentation.</li>
	<li>Spray the hindlimbs of the mouse with 70% ethanol. Use hardened fine curved scissors (24 mm cutting edge) and fine forceps (Dumont 7, curved or straight) to remove the skin and to expose the underlying muscles. Avoid any contact of the underlying muscles with fur (increases the risk of contamination).</li>
	<li>Remove the surrounding fascia with fine curved forceps without damaging the underlying muscles (Figure 2A). Close the forceps to avoid bending.</li>
	<li>Use curved forceps to expose the distal tendons of the TA (tibialis anterior) and EDL muscle. To remove the TA, grab the distal TA tendon with forceps and cut with fine Vannas spring scissors (5 mm cutting edge, 0.35 mm tip diameter). While holding the TA at the tendon, pull it towards the knee and cut the muscle close to the knee (Figure 2B), the EDL muscle is now exposed. 
	NOTE: Make sure that only the TA tendon is grabbed in this step, otherwise the underlying EDL might get damaged. When cutting off the TA muscle make sure that the tendons at the knee can be seen easily afterwards.</li>
	<li>Lift the distal EDL tendon with fine curved forceps and cut with fine Vannas spring scissors (Figure 2C). Expose the proximal EDL tendon by carefully pulling the EDL towards the knee. Cut the proximal tendon with fine Vannas spring scissors. Transfer the EDL muscle to the 37 &#176;C preheated 2.5 mL of collagenase digestion solution in the 15 mL reaction tube from step 1.4. (Figure 2D). 
	NOTE: Make a small incision at the outer knee connective tissue to fully expose the proximal EDL tendon. Make sure to only grab the tendons and not to stretch the EDL too much.</li>
	<li>Repeat steps 2.3. to 2.6. with the second EDL. Add both EDL muscles to the same 15 mL reaction tube filled with 2.5 mL collagenase digestion solution.</li>
	<li>Incubate the EDL muscles in the reaction tube at 37 &#176;C in a circulating water bath. 
	NOTE: Incubation time depends on several factors, such as collagenase activity, age of the mouse and amount of fibrotic tissue. The typical incubation time for EDL muscles of adult mice (2-6 months of age) is 1-1.5 h and for aged mice (18 months of age) 1.5-2 h.</li>
	<li>To avoid excessive digestion of the muscles, check the muscles during the digestion time. Stop the digestion when muscles loosen up and single myofibers are visible (Figure 2E). Transfer muscles carefully with the large bore pipette to the first well of the prewarmed 12-well plate containing myofiber isolation medium (Figure 2G).</li>
</ol>
3. Myofiber dissociation and culture
<ol>
	<li>For the following steps use a stereo binocular microscope, preferentially equipped with a heating plate (37 &#176;C). Use the large bore pipette to flush the muscles with warm isolation medium until single myofibers are released. Dissociate the muscles with the large bore pipette until the desired number of myofibers are floating freely in the solution. 
	NOTE: Avoid excessive trituration to reduce the risk of damaged myofibers. Using a heating plate for myofiber dissociation is strongly advised since temperature drops during the isolation process, which will result in myofiber death.</li>
	<li>Transfer non-contracted myofibers (Figure 2H) with the HS coated small bore glass pipette to the second well filled with isolation medium to wash away debris and contracted (damaged) myofibers (Figure 2I). 
	NOTE: To avoid excessive movement of isolated myofibers, they can be transferred to the second well and then the trituration process can be continued.</li>
	<li>Transfer non-contracted myofibers to the next (3rd) well filled with isolation medium to wash again.</li>
	<li>Use the small-bore glass pipette, coated with HS, to transfer approximately 50-100 non-contracted myofibers to the 24-well plate containing myofiber culture medium.</li>
	<li>Incubate myofibers at 37 &#176;C, 5% CO2 for dedicated time (96 h maximum are recommended). 
	NOTE: MuSCs divide once either planar or apical-basal after 42 h of culture. Additionally, after 72 h of culture MuSCs form multicellular clusters consisting of self-renewing, proliferating or committed (differentiated) MuSCs.</li>
</ol>
4. siRNA transfection
<ol>
	<li>4 h after myofiber isolation, transfect myofiber associated MuSCs (50-100 non-contracted myofibers in one well of the 24-well plate filled with 500 &#181;L culture medium) with lipid based transfection reagent, e.g. RNAiMAX according to the manufacturer&#39;s protocol with a final concentration of 5 pmol siRNA. Therefore, add 25 &#181;L of Opti-MEM with the respective volume of siRNA to 25 &#181;L Opti-MEM containing 1.5 &#181;L of the transfection reagent. Incubate the reaction mix for 5 min and add it then to the 500 &#181;L of culture medium. 
	NOTE: A second transfection after 24 h or 48 h is recommended for longer culture periods, e.g., more than 48 h. A change of medium after transfection is not necessary.</li>
</ol>
5. Fixation and IF staining
<ol>
	<li>For immunostaining use a stereo binocular microscope. All volumes in the following steps are adjusted to one well of a 24-well plate. Perform all following steps using a HS-coated small-bore pipette.</li>
	<li>Carefully discard the myofiber culture medium while leaving some solution in the well (approximately 100 &#181;L per 24-well). Do this for all further steps unless stated otherwise to allow floating of the myofibers. Add 500 &#181;L of 2% PFA to fix the myofibers with their adjacent MuSCs, incubate for 5 min at room temperature (RT).</li>
	<li>Remove the supernatant carefully and wash the myofibers three times with PBS (pH 7.4, 500 &#181;L for 5 min at RT each).</li>
	<li>Add 500 &#181;L permeabilization solution (0.1% Triton X-100, 0.1 M Glycine in PBS, pH 7.4), incubate for 10 min at RT.</li>
	<li>Remove the permeabilization solution and add 500 &#181;L blocking solution (5% HS in PBS, pH 7.4) for 1 h at RT. 
	NOTE: Check the recommended blocking solution for primary antibodies to avoid unspecific binding.</li>
	<li>Remove the blocking solution and add 300 &#181;L of primary antibody dilution (e.g., anti-Pax7 (PAX7, DSHB, undiluted), anti-MyoD (clone G-1, Santa Cruz, 1:200)) per well. Incubate at 4 &#176;C overnight.</li>
	<li>Wash three times with 500 &#181;L of PBS per well (5 min at RT).</li>
	<li>Remove PBS and add 300 &#181;L of secondary antibody dilution (e.g., anti-mouse-IgG1-546 and anti-mouse-IgG2b-488 1:1000) per well. Incubate 1 h at RT protected from light. For the following steps, light reduced conditions are recommended.</li>
	<li>Wash two times with 500 &#181;L of PBS per well (5 min at RT).</li>
	<li>Perform DAPI staining by using 500 &#181;L of the solution per well (final DAPI concentration 10 &#181;g/mL) for 5 min at RT.</li>
	<li>Wash two times with 500 &#181;L of PBS per well (5 min at RT).</li>
	<li>Use a hydrophobic pen to draw a circle on a microscopic glass slide to create a water repellant barrier that will prevent spilling of liquid containing myofibers. Transfer the myofibers in the smallest volume possible to the microscopic glass slide and disperse them on the glass slide. 
	NOTE: Avoid physical pulling of the myofibers over the microscopic glass slide since this might cause friction resulting in detachment of clusters from myofibers.</li>
	<li>Remove the residual liquid with the small-bore Pasteur pipette or a 200 &#181;L pipette.</li>
	<li>Use two drops of aqueous mounting medium and cover the myofibers with a coverslip. Let the slides dry for the time recommended by the manufacturer. Store the slides at 4 &#176;C in the dark.</li>
</ol>

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A. <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+satellite+cells+and+transplantation+into+mice+for+lineage+tracing+in+muscle.">Isolation of satellite cells and transplantation into mice for lineage tracing in muscle.</a> Nature Protocols. 15 (3), 1082-1097 (2020).</li><li>Garry, G. A., Antony, M. L., Garry, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiotoxin+induced+injury+and+skeletal+muscle+regeneration.">Cardiotoxin induced injury and skeletal muscle regeneration.</a> Methods in Molecular Biology. 1460, 61-71 (2016).</li></ol>

The authors declare no competing financial interests.

Here, a method to functionally investigate the role of a specific gene in MuSCs using an in vitro approach is presented. Importantly, in the system described here the MuSCs are cultured in conditions, which resemble the in vivo situation as much as possible preserving most of the interactions of the MuSCs with their niche. This is accomplished by culturing isolated myofibers with their adjacent MuSCs under floating conditions and consecutive siRNA transfection. The procedures of myofiber isolation, siRNA transfection and investigation of MuSC populations during 72 h of culture through immunofluorescence analyses are described. Furthermore, it was demonstrated that around 74% of all MuSCs were transfected with a fluorescently labelled control siRNA after 30 h of culture.
Particular attention should be focused on the careful dissection of the EDL muscle since extensive stretching, pinching, or squeezing will lead to contraction and consecutive death of myofibers. Furthermore, it is important to investigate clusters of MuSCs from at least 20 different myofibers per replicate per&#160;condition. This is necessary since MuSC numbers and properties are varying due to the existence of MuSC subpopulations. When investigating the effect of a specific siRNA on MuSCs using the floating myofiber culture method a comparison of the condition with the targeting siRNA to the non-targeting control should be performed within the same mouse and muscle. This is recommended to avoid mouse-specific differences which might cover or amplify effects of the siRNA. The knockdown efficiency can be determined by immunofluorescence analyses with antibodies directed against the target gene using single myofibers with their adjacent MuSCs. If this is not an option, one can test the siRNA knockdown efficiency in primary myoblasts followed by quantitative RT-PCR or immunoblot analyses. The efficiency of the siRNA should be determined before analyzing the effect of the siRNA on MuSCs on single myofibers. The use of a smart pool consisting of 4 different siRNAs versus a single one increases the knockdown efficiency but also increases the risk of unspecific targeting. A non-targeting siRNA should be used as a control. To directly monitor the transfection efficiency, one can use a fluorescently labelled non-targeting siRNA as performed here. The time point for transfection with the siRNA is around 4 hours after isolation, a time point when the basal lamina surrounding the MuSCs is already permeable for the siRNA. If the effect of a specific siRNA on MuSCs after 72 or 96 h should be investigated, it is recommended to perform a second siRNA transfection after 24 h or 48 h to maintain a high knockdown efficiency.
The myofiber culture assay displays diverse advantages compared to the investigation of MuSCs with conventional cell culture methods. MuSCs stay attached to the myofibers during the whole isolation process, thereby preserving the crucial interaction of the MuSC with its niche19,23,24,25. The preserved interaction of MuSCs with the myofiber is a prerequisite for studying niche dependent effects on MuSC functionality, which cannot be recapitulated in conventional 2D myoblast cultures. For example, during aging MuSCs display impaired myogenic capacity resulting in a reduced efficiency to regenerate muscle tissue after damage20,26. This impairment is at least partially attributable to changes in the MuSC niche, in particular changes in the ECM composition27,28. The myofiber culture protocol allows the study and interference with these aberrant niche changes.
In contrast to the method described here, purification of MuSCs by immunolabeling and -sorting techniques like FACS (fluorescence activated cell sorting) or MACS (magnetic cell sorting) involves the removal of the MuSCs from their niche. Interestingly, 2D cultures of isolated MuSCs from aged muscles loose their extrinsic cues and behave similar to MuSCs isolated from young muscles thereby not recapitulating the in vivo situation appropriately29. Furthermore, the complete dissociation of the muscle tissue and labeling of MuSCs with surface markers results in transcriptomic changes and activation of the cells30,31,32. Another advantage of the myofiber culture system is the possibility to interfere with MuSC functionality at various levels. Manipulation of MuSCs on cultured myofibers can be effectively achieved by siRNA-mediated gene knockdown as described here in detail. Likewise, the application of chemical compounds or the delivery of recombinant proteins is very effective to interfere with stem cell pathways20,28. Furthermore, retro- or lentiviral expression vectors permit the introduction of exogenous genes, i.e., constitutive active mutants33. Additionally, the influence of extrinsic factors on MuSC functionality can be explored in the system described here, e.g., the culture conditions can be supplemented with supernatant from different physiological or pathological sources to model different states like cancer cachexia34,35.
One limitation of the method described here is the fact, that the single myofiber culture system cannot completely recapitulate the impact of all systemic factors or influence of other cell types on MuSCs. Also, the time in which the myofibers can be kept viable in culture is limited and therefore the study of MuSC related processes focuses on early events like activation and myogenic commitment. Furthermore, the investigation of the MuSC interaction with other niche cells such as immune cells or fibro-adipogenic progenitor cells is not possible. To investigate systemic effects on MuSC functionality one can either perform muscle injury experiments followed by the analysis of muscle regeneration in vivo or perform transplantation experiments36,37.
Taken together, the myofiber isolation and culture protocol provides great opportunity for genetic or mechanistic studies on adult MuSCs without the requirement of transgenic mouse models and can potentially reduce animal experimentation.

Skeletal muscle in the adult is a postmitotic tissue mainly composed of multinucleated myofibers, which are the effector cells for voluntary movements. It has a remarkable ability to regenerate, a process that resembles embryonic myogenesis and undergoes impairments in age and disease1. This striking regenerative capacity of skeletal muscle depends on muscle stem cells (MuSCs), which are also termed satellite cells due to their location between the sarcolemma and the basal lamina of myofibers2,3. Under resting conditions MuSCs are quiescent and characterized by the expression of the transcription factor Pax7 and quiescence markers such as Sprouty14,5,6,7,8. Upon activation, e.g., after injury, MuSCs leave the quiescent state and upregulate the myogenic regulatory factor MyoD9. The Pax7/MyoD double positive MuSCs proliferate and differentiate thereby generating myogenic precursor cells, which are also often referred to as myoblasts. Those myoblasts then further differentiate into elongated myocytes, a process coinciding with molecular and morphological changes, e.g., loss of Pax7 and upregulation of Myogenin expression10. Myocytes then eventually fuse to each other or to existing myofibers thereby repairing the damaged tissue. Importantly, a small fraction of muscle stem cells reverts MyoD upregulation and is able to self-renew11. The status of MuSC differentiation and myogenic progression can be readily observed by investigation of myogenic markers such as Pax7, MyoD and Myogenin10.
The culture of single myofibers with their adjacent MuSCs is an excellent method to investigate MuSC functionality in an ex vivo setting since MuSCs are remaining in their endogenous niche12,13. The behavior of MuSCs is regulated by intrinsic signals as well as extrinsic signals provided by the niche, a specialized anatomical location comprising components of the extracellular matrix (ECM) surrounding the MuSCs and the myofiber itself. For instance, one of the extrinsic regulators of MuSC quiescence is Notch signaling. Here, signaling cues are received by MuSCs from both the myofiber and the ECM14,15,16. Moreover, the MuSC niche is important to control the division axis of MuSCs thus regulating the cell fate of MuSC daughter cells17,18. Reasonably, parameters like asymmetric MuSC divisions, myogenic progression and self-renewal can be uniquely assessed in this experimental setup. For instance, a multicellular cluster can form arising from one MuSC after a 72 h culture period, which can be investigated for the occurrence and percentage of distinct myogenic populations such as self-renewing, proliferating and further differentiated MuSCs8,19,20,21. The differentiation state of MuSCs can be determined by investigation of the expression/co-expression of Pax7, MyoD and Myogenin. After 72 h of culture the cells in a cluster can be discriminated by the following parameters: Pax7 only cells are self-renewing MuSCs, while Pax7/MyoD double positive cells are proliferating/activated MuSCs and further differentiated myogenic cells are Myogenin positive22. Furthermore, MuSC numbers or re-entry into the cell cycle/activation can be investigated in addition to myogenic progression, e.g., through immunofluorescence-based analyses based on the parameters described above.
Here, the unique features of the myofiber isolation and culture protocol, e.g., the preservation of the interaction of the MuSC with its niche, are described. Mouse whole EDL (extensor digitorum longus) muscles are carefully dissected, digested by collagenase, and physically triturated to obtain single myofibers with their associated MuSCs for further culture. Moreover, the protocol delineates the steps to transfect MuSCs with siRNA for functional analyses of candidate genes and consecutive immunofluorescence-based analyses without the necessity of transgenic animals.

<table><tbody><tr><td>Alexa Fluor 488 goat anti-rabbit IgG 2b</td><td>ThermoScientific</td><td>A-21141</td><td>use 1:1000 for IF</td></tr><tr><td>Alexa Fluor 546 goat anti-mouse IgG1</td><td>ThermoScientific</td><td>A-21123</td><td>use 1:1000 for IF</td></tr><tr><td>chicken embryo extract</td><td>Seralab</td><td>CE-650-J</td><td>chicken embryo extract containing growth factors etc.</td></tr><tr><td>collagenase type 1</td><td>Sigma</td><td>C0130</td><td></td></tr><tr><td>DMEM (Dulbecco&amp;rsquo;s modified Eagle&amp;rsquo;s medium with 4.5&amp;nbsp;g/l glucose and sodium pyruvate)</td><td>GibCo</td><td>41966029</td><td>cell culture medium</td></tr><tr><td>fetal bovine serum</td><td>Gibco</td><td>10270-106</td><td>fetal bovine serum</td></tr><tr><td>horse serum</td><td>Gibco</td><td>26050-088</td><td></td></tr><tr><td>Lipofectamine RNAiMax</td><td>ThermoScientific</td><td>13778150</td><td>transfection reagent</td></tr><tr><td>MyoD antibody clone G-1</td><td>Santa Cruz</td><td>sc-377460</td><td>dilute 1:200 for IF</td></tr><tr><td>Pax7 antibody</td><td>DSHB</td><td>PAX7</td><td>use undiluted</td></tr><tr><td>siGLO Red Transfection Indicator</td><td>horizon discovery</td><td>D-001630-02-05</td><td>non targeting siRNA</td></tr></tbody></table>

single myofiber culture assay for the assessment of adult muscle stem cell functionality ex vivo

Adult skeletal muscle tissue harbors a stem cell population that is indispensable for its ability to regenerate. Upon muscle damage, muscle stem cells leave their quiescent state and activate the myogenic program ultimately leading to the repair of damaged tissue concomitant with the replenishment of the muscle stem cell pool. Various factors influence muscle stem cell activity, among them intrinsic stimuli but also signals from the direct muscle stem cell environment, the stem cell niche. The isolation and culture of single myofibers with their associated muscle stem cells preserves most of the interaction of the stem cell with its niche and is, therefore, the closest possibility to study muscle stem cell functionality ex vivo. Here, a protocol for the isolation, culture, siRNA transfection and immunostaining of muscle stem cells on their respective myofibers from mouse EDL (extensor digitorum longus) muscles is provided. The experimental conditions outlined here allow the study and manipulation of muscle stem cells ex vivo including investigation of myogenic activity without the inherent need for in vivo animal experiments.

This protocol provides instructions for successful derivation and culture of single myofibers with their associated MuSCs from murine EDL muscles. Essential steps of the protocol are summarized in Figure 1. Careful tendon-to-tendon dissection of the EDL muscles (Figure 2A-C) is critical for a high yield of viable myofibers. Muscle dissociation is achieved first by collagenase digestion (Figure 2D) followed by physical trituration (Figure 2G). Intact myofibers (Figure 2H) are cultured, whereas hypercontracted and dead myofibers (Figure 2I) should be excluded from culture and analysis.
MuSCs remain myofiber-associated during the isolation process. Immunofluorescence staining for the transcription factor Pax7 identifies and distinguishes 7-9 MuSC nuclei from the plethora of myonuclei per myofiber when fixed directly after completion of the isolation process (Figure 3A). Figure 3B shows a magnified area from Figure 3A with the additional brightfield channel, which exposes the subcellular myofibrillary structure of the myotube and demonstrates Pax7 immunofluorescence signal in the nucleus of a MuSC.
Activation and myogenic progression of myofiber associated MuSCs can be analyzed by myogenic marker expression. Associated with freshly isolated myofibers (0 h), mostly quiescent MuSCs can be found, which are characterized by Pax7 expression and absence of MyoD expression, thereby resembling an in vivo homeostatic condition (Figure 4, 0 h). Due to the dissection/dissociation procedure and the myofiber culture media composition, the MuSCs rapidly activate and upregulate the transcription factor MyoD to facilitate proliferation as can be observed at 42 hours, when MuSCs have undergone their first division (Figure 4, 42 h). After 72 hours of culture, MuSCs form clusters of progenies with different myogenic fates that is paralleled by the expression patterns of different myogenic markers (Figure 4, 72 h). Pax7+ only cells resist differentiation and become self-renewing stem cells. Pax7+ and MyoD+ double positive cells are proliferative, whereas MyoD+ only cells have further progressed along the myogenic lineage and will differentiate.
The myofiber culture system allows for efficient interference of MuSC activity by various interventions, which one of them is siRNA transfection as described in detail in this protocol. To monitor the transfection efficiency of myofiber associated MuSCs a fluorescently labeled non-targeting siRNA was transfected. Pax7 positive MuSCs accumulated cytoplasmic siRNA in a granule-like fashion, indicating efficient uptake (Figure 5A). No fluorescent granules were observed in the cytoplasm of myofibers suggesting a natural uptake barrier in myofibers at 4 h after isolation and that siRNA transfection specifically targets MuSCs. Quantification of positively transfected Pax7+ cells per myofiber revealed that more than half of all MuSCs had taken up visible amounts of fluorescently labelled siRNA just before completion of the first round of division at 24 hours. The number of transfected Pax7+ cells increased further up to 74% after 30 hours (Figure 5B). Furthermore, there was no difference in the number of Pax7+ cells per myofiber of transfected or non-transfected conditions at both timepoints, demonstrating no adverse effects on the stem cell numbers due to the transfection procedure (Figure 5C).
In summary, the protocol provides a detailed description of the isolation and culture of EDL single myofibers with their adjacent muscle stem cells. It enables the study of myofiber-associated muscle stem cell activity ex vivo, e.g.,&#160;by immunofluorescence-based analyses. Manipulation of muscle stem cells by siRNA transfection is efficient and provides a methodological basis for functional analyses.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62257/62257fig1v2.jpg" /> 
Figure 1: Schematic summary of essential steps. The essential steps of the isolation and immunostaining procedure are summarized in this figure. Abbreviations: DMEM, Dulbecco&#39;s Modified Eagle&#39;s Medium; FBS, Fetal Bovine Serum; CEE, Chicken Embryo Extract; TA, tibialis anterior; EDL, extensor digitorum longus <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62257/62257fig1v2large.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/62257/62257fig02.jpg" /> 
Figure 2: Mouse EDL dissection and single myofiber isolation. (A) The skin of the mouse hindlimb is removed and the muscle surrounding fascia is pulled away to expose the TA (tibialis anterior) muscle. (B) The TA muscle is removed to expose the EDL (extensor digitorum longus) muscle, marked by the white arrow. (C) The EDL muscle is dissected by cutting its tendons. (D) Two EDL muscles are collagenase digested. (E) Appearance of collagenase digested muscle with visible single myofibers loosening up from the tissue core. (F) Large and small-bore Pasteur pipette with scale. (G) The EDL muscles (marked by black arrows) are physically triturated using a large bore Pasteur pipette. (H) Single intact myofibers are thin and shiny and can be individually collected for culture and analysis. (I) Hypercontracted and dead myofibers are unsuitable for culture and analysis. The microscopic images of 2H and 2I were captured with a microscope using a N-Achroplan 5x objective. Scale bars are 200 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62257/62257fig02large.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/62257/62257fig03.jpg" /> 
Figure 3: Microscopic image of a whole single myofiber with its associated MuSCs. Single myofibers from EDL of young C57BL/6 mice were prepared and PFA-fixed after isolation (0 h). (A) Pax7 and DAPI immunofluorescence staining identifies myofiber associated muscle stem cells (MuSCs, marked by arrows). The microscopic image was captured using the tiles and z-stack function of a microscope equipped with a Plan-Apochromat 40x oil objective. Scale bar is 200 &#181;m. (B) Magnification from (A) showing the myonuclei, one Pax7 positive nucleus and the bright-dark pattern from myofibrillary structures in brightfield. Scale bar is 10 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62257/62257fig03large.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/62257/62257fig04.jpg" /> 
Figure 4: Immunofluorescence images of MuSC activation and myogenic progression during single myofiber culture. Muscle stem cells (MuSCs) of freshly isolated myofibers (0 h) represent a homeostatic state close to in vivo quiescence, characterized by expression of Pax7 and lack of MyoD expression. During single myofiber culture most MuSCs upregulate MyoD and re-enter the cell cycle to divide and proliferate (42 h). After 72 h of culture the MuSC fate can be discriminated based on myogenic marker expression. Cells with Pax7 only expression will self-renew (red arrow) whereas Pax7 and MyoD double positive cells (red/green arrow) continue to proliferate. MyoD only positive cells (green arrow) have committed to myogenic differentiation. Microscopic images were captured using a microscope with a Plan-Apochromat 20x objective (0 h, 42 h) or a LD Plan-Neofluar 40x objective (72 h). Scale bars are 20 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62257/62257fig04large.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/62257/62257fig05.jpg" /> 
Figure 5: Fluorescent siRNA transfection of MuSCs and uptake analysis. EDL single myofibers were prepared and transfected with fluorescently labeled siRNA (siGLO) following the steps provided by this protocol. (A) Cytoplasmatic accumulation of siGLO granules specifically in a Pax7 positive muscle stem cell (MuSC) on a single myofiber at 30 h of culture. The microscopic image was captured using the z-stack and apotome function of a&#160;microscope with a Plan-Apochromat 100x oil objective. Scale bars are 5 &#181;m. (B) Quantification of siRNA uptake by Pax7 positive muscle stem cells at 24 or 30 h of culture. (C) Number of Pax7 positive cells per single myofiber comparing non-transfected and transfected conditions at 24 or 30 hours of culture. Data are&#160;shown as mean with standard deviation of n = 4 mice. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62257/62257fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Single Myofiber Culture Assay for the Assessment of Adult Muscle Stem Cell Functionality Ex Vivo at JoVE.com

Single Myofiber Culture Assay for the Assessment of Adult Muscle Stem Cell Functionality Ex Vivo

In this protocol an in vitro culturing and functional analysis method for muscle stem cells is described, which preserves most of their interactions with their endogenous niche.

Adult skeletal muscle tissue harbors a stem cell population that is indispensable for its ability to regenerate. Upon muscle damage, muscle stem cells ...

single-myofiber-culture-assay-for-assessment-adult-muscle-stem-cell

Nanyang Technological University

Research

JoVE Journal

Biology

4.7K Views.  Leibniz Institute on Aging, Fritz-Lipmann-Institute. In this protocol an in vitro culturing and functional analysis method for muscle stem cells is described, which preserves most of their interactions with their endogenous niche.

Video: Single Myofiber Culture Assay for the Assessment of Adult Muscle Stem Cell Functionality Ex Vivo

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

Preparation and Culture of Myogenic Precursor Cells/Primary Myoblasts from Skeletal Muscle of Adult and Aged Humans

Skeletal muscle homeostasis depends on muscle growth (hypertrophy), atrophy and regeneration. During ageing and in several diseases, muscle wasting occurs. Loss of muscle mass and function is associated with muscle fiber type atrophy, fiber type switching, defective muscle regeneration associated with dysfunction of satellite cells, muscle stem cells, and other pathophysiological processes. These changes are associated with changes in intracellular as well as local and systemic niches. In addition to most commonly used rodent models of muscle ageing, there is a need to study muscle homeostasis and wasting using human models, which due to ethical implications, consist predominantly of in vitro cultures. Despite the wide use of human Myogenic Progenitor Cells (MPCs) and primary myoblasts in myogenesis, there is limited data on using human primary myoblast and myotube cultures to study molecular mechanisms regulating different aspects of age-associated muscle wasting, aiding in the validation of mechanisms of ageing proposed in rodent muscle. The use of human MPCs, primary myoblasts and myotubes isolated from adult and aged people, provides a physiologically relevant model of molecular mechanisms of processes associated with muscle growth, atrophy and regeneration. Here we describe in detail a robust, inexpensive, reproducible and efficient protocol for the isolation and maintenance of human MPCs&#160;and their progeny&#160;&#8212; myoblasts and myotubes from human muscle samples using enzymatic digestion. Furthermore, we have determined the passage number at which primary myoblasts from adult and aged people undergo senescence in an in vitro culture. Finally, we show the ability to transfect these myoblasts and the ability to characterize their proliferative and differentiation capacity and propose their suitability for performing functional studies of molecular mechanisms of myogenesis and muscle wasting in vitro.

This protocol describes a robust, reproducible and simple method of isolation and culture of myoblast progenitor cells from the skeletal muscle of adult and aged people. The muscles used here include foot and leg muscles. This approach enables the isolation of an enriched population of primary myoblasts for functional studies.

Skeletal muscle homeostasis depends on muscle growth (hypertrophy), atrophy and regeneration. During ageing and in several diseases, muscle wasting ...

Isolation of Human Myoblasts, Assessment of Myogenic Differentiation, and Store-operated Calcium Entry Measurement

Satellite cells (SC) are muscle stem cells located between the plasma membrane of muscle fibers and the surrounding basal lamina. They are essential for muscle regeneration. Upon injury, which occurs frequently in skeletal muscles, SCs are activated. They proliferate as myoblasts and differentiate to repair muscle lesions. Among many events that take place during muscle differentiation, cytosolic Ca2+ signals are of great importance. These Ca2+ signals arise from Ca2+ release from internal Ca2+ stores, as well as from Ca2+ entry from the extracellular space, particularly the store-operated Ca2+ entry (SOCE). This paper describes a methodology used to obtain a pure population of human myoblasts from muscle samples collected after orthopedic surgery. The tissue is mechanically and enzymatically digested, and the cells are amplified and then sorted by flow cytometry according to the presence of specific membrane markers. Once obtained, human myoblasts are expanded and committed to differentiate by removing growth factors from the culture medium. The expression levels of specific transcription factors and in vitro immunofluorescence are used to assess the myogenic differentiation process in control conditions and after silencing proteins involved in Ca2+ signaling. Finally, we detail the use of Fura-2 as a ratiometric Ca2+ probe that provides reliable and reproducible measurements of SOCE.

Here, we describe a methodology to obtain a pure population of human myoblasts from adult muscle tissue. These cells are used to study in vitro skeletal muscle differentiation and, in particular, to study proteins involved in Ca2+ signaling.

Satellite cells (SC) are muscle stem cells located between the plasma membrane of muscle fibers and the surrounding basal lamina. They are essential for ...

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 ...

Assessing Functional Metrics of Skeletal Muscle Health in Human Skeletal Muscle Microtissues

Three-dimensional (3D) in vitro models of skeletal muscle are a valuable advancement in biomedical research as they afford the opportunity to study skeletal muscle reformation and function in a scalable format that is amenable to experimental manipulations. 3D muscle culture systems are desirable as they enable scientists to study skeletal muscle ex vivo in the context of human cells. 3D in vitro&#160;models closely mimic aspects of the native tissue structure of adult skeletal muscle. However, their universal application is limited by the availability of platforms that are simple to fabricate, cost and user-friendly, and yield relatively high quantities of human skeletal muscle tissues. Additionally, since skeletal muscle plays an important functional role that is impaired over time in many disease states, an experimental platform for microtissue studies is most practical when minimally invasive calcium transient and contractile force measurements can be conducted directly within the platform itself. In this protocol, the fabrication of a 96-well platform known as &#39;MyoTACTIC&#39;, and en masse production of 3D human skeletal muscle microtissues (hMMTs) is described. In addition, the methods for a minimally invasive application of electrical stimulation that enables repeated measurements of skeletal muscle force and calcium handling of each microtissue over time are reported.

This manuscript describes a detailed protocol to produce arrays of 3D human skeletal muscle microtissues and minimally invasive downstream in situ assays of function, including contractile force and calcium handling analyses.

Three-dimensional (3D) in vitro models of skeletal muscle are a valuable advancement in biomedical research as they afford the opportunity to study ...

Bioengineering

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 ...

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 Quiescent Stem Cell Populations from Individual Skeletal Muscles

Skeletal muscle harbors distinct populations of adult stem cells that contribute to the homeostasis and repair of the tissue. Skeletal muscle stem cells (MuSCs) have the ability to make new muscle, whereas fibro-adipogenic progenitors (FAPs) contribute to stromal supporting tissues and have the ability to make fibroblasts and adipocytes. Both MuSCs and FAPs reside in a state of prolonged reversible cell cycle exit, called quiescence. The quiescent state is key to their function. Quiescent stem cells are commonly purified from multiple muscle tissues pooled together in a single sample. However, recent studies have revealed distinct differences in the molecular profiles and quiescence depth of MuSCs isolated from different muscles. The present protocol describes the isolation and study of MuSCs and FAPs from individual skeletal muscles and presents strategies to perform molecular analysis of stem cell activation. It details how to isolate and digest muscles of different developmental origin, thicknesses, and functions, such as the diaphragm, triceps, gracilis, tibialis anterior (TA), gastrocnemius (GA), soleus, extensor digitorum longus (EDL), and the masseter muscles. MuSCs and FAPs are purified by fluorescence-activated cell sorting (FACS) and analyzed by immunofluorescence staining and 5-ethynyl-2&#180;-deoxyuridine (EdU) incorporation assay.

This protocol describes the isolation of muscle stem cells and fibro-adipogenic progenitors from individual skeletal muscles in mice. The protocol involves single muscle dissection, stem cell isolation by fluorescence activated cell sorting, purity assessment by immunofluorescence staining, and quantitative measurement of S-phase entry by 5-ethynyl-2&#180;-deoxyuridine incorporation assay.

Skeletal muscle harbors distinct populations of adult stem cells that contribute to the homeostasis and repair of the tissue. Skeletal muscle stem cells ...

High-Throughput Contractile Measurements of Hydrogel-Embedded Intact Mouse Muscle Fibers Using an Optics-Based System

In vitro cell culture is a powerful tool to assess cellular processes and test therapeutic strategies. For skeletal muscle, the most common approaches involve either differentiating myogenic progenitor cells into immature myotubes or the short-term ex vivo culture of isolated individual muscle fibers. A key benefit of ex vivo culture over in vitro is the retention of the complex cellular architecture and contractile characteristics. Here, we detail an experimental protocol for the isolation of intact flexor digitorum brevis muscle fibers from mice and their subsequent ex vivo culture. In this protocol, muscle fibers are embedded in a fibrin-based and basement membrane matrix hydrogel to immobilize the fibers and maintain their contractile function. We then describe methods to assess the muscle fiber contractile function using an optics-based, high-throughput contractility system. The embedded muscle fibers are electrically stimulated to induce contractions, after which their functional properties, such as sarcomere shortening and contractile velocity, are assessed using optics-based quantification. Coupling muscle fiber culture with this&#160;system allows for high-throughput testing of the effects of pharmacological agents on contractile function and ex vivo studies of genetic muscle disorders. Finally, this protocol can also be adapted to study dynamic cellular processes in muscle fibers using live-cell microscopy.

Skeletal muscle function can be assessed by quantifying the contractility of isolated muscle fibers, traditionally using laborious, low-throughput approaches. Here, we describe an optics-based, high-throughput method to quantify the contractility of hydrogel-embedded muscle fibers. This approach has applications for drug screening and therapeutic development.

In vitro cell culture is a powerful tool to assess cellular processes and test therapeutic strategies. For skeletal muscle, the most common approaches ...

Harvesting Mouse Skeletal Muscle to Culture Stem Cells and Their Niche

Unfractionated Bulk Culture of Mouse Skeletal Muscle to Recapitulate Niche and Stem Cell Quiescence

Skeletal muscle is the largest tissue of the body and performs multiple functions, from locomotion to body temperature control. Its functionality and recovery from injuries depend on a multitude of cell types and on molecular signals between the core muscle cells (myofibers, muscle stem cells) and their niche. Most experimental settings do not preserve this complex physiological microenvironment, and neither do they allow the ex vivo study of muscle stem cells in quiescence, a cell state that is crucial for them. Here, a protocol is outlined for the ex vivo culture of muscle stem cells with cellular components of their niche. Through the mechanical and enzymatic breakdown of muscles, a mixture of cell types is obtained, which is put in 2D culture. Immunostaining shows that within 1 week, multiple niche cells are present in culture alongside myofibers and, importantly, Pax7-positive cells that display the characteristics of quiescent muscle stem cells. These unique properties make this protocol a powerful tool for cell amplification and the generation of quiescent-like stem cells that can be used to address fundamental and translational questions.

Author Spotlight: Ex Vivo Protocol for Culturing Quiescent Muscle Stem Cells with Niche Components 

Skeletal muscle comprises multiple cell types, including resident stem cells, each with a special contribution to muscle homeostasis and regeneration. Here, the 2D culture of muscle stem cells and the muscle cell niche in an ex vivo setting that preserves many of the physiological, in vivo, and environmental characteristics are described.

Skeletal muscle is the largest tissue of the body and performs multiple functions, from locomotion to body temperature control. Its functionality and ...