We thank Andrea Bianchi, the Italian Network of Laminopathies and the members of the laboratory for the support and all the constructive comments. We are grateful to Chiara Cordiglieri for the precious help&#160;at confocal microscope. The authors thank Dr. Beatrice Biferali for her help in taking pictures for figures. The work presented in here was supported by My First AIRC Grant n. 18535, AFM-Telethon n. 21030, the Italian Minister of Health n. GR-2013-02355413 and Cariplo 2017-0649 to C.L. C.M. is supported by My First AIRC grant n.18993 and AFM-Telethon n. 22489.

All the experimental procedures were performed under the ethical approval of the Italian Ministry of Health and the Institutional Animal Care and Use Committee (authorization n. 83/2019-PR). The animals were maintained in an authorized facility at San Raffaele Hospital, Milan, Italy (authorization n. N. 127/2012-A).
1. Muscle dissection and myofiber culture
<ol>
	<li>Equipment preparation.
	<ol>
		<li>Before starting, prepare all the necessary solutions as described in Table 1. These solutions need to be freshly prepared.</li>
		<li>Clean all the surfaces and tools that will be used during the procedure with 70% ethanol.</li>
		<li>Before starting with mice sacrifice, perform coating of 100 mm and 35 mm Petri dishes using Horse Serum (HS). Coat all dishes to prevent myofibers from attaching to the plastic. Consider using one 100 mm dish and four 35 mm dishes per mouse.</li>
		<li>After removing the excess of HS, store coated dishes in an incubator at 37 &#176;C for 30 min. Then fill it with washing solution or culture medium (two 35 mm dishes per mouse). 
		NOTE: Alternatively, a solution of 10% HS in Dulbecco&#39;s modified Eagle&#39;s medium (DMEM) can be used to coat dishes. Always use coated dishes. It is possible to use culture dishes of different size, but small dimension Petri dishes are recommended.</li>
		<li>For fibers isolation, prepare sterile Pasteur pipettes as shown in Figure 2. For each&#160;mouse prepare one large hole bore pipette for muscle handling and mechanical disaggregation (Figure 2A) and one small hole pipette for fibers selection (Figure 2B). Cut each glass pipette, possibly using a diamond pen, to the desired length and smooth pipette&#8217;s edges on a flame.</li>
		<li>Coat each pipette by briefly wetting it with HS before use.</li>
	</ol>
	</li>
	<li>Mouse sacrifice and muscle dissection
	<ol>
		<li>Pre-warm the digestion solution at room temperature for 10 min before starting the dissection procedure. Polypropylene FACS round-bottom tubes are the most suitable containers for the purpose.</li>
		<li>Immediately before the beginning of muscle retrieval, sacrifice the mouse according to proper national IACUC recommendation.</li>
		<li>Wet its lower body with 70% ethanol before cutting the skin to make the removal of the hair easier.</li>
		<li>Put the mouse in a prone position on a support of polystyrene covered with aluminum paper and cut the skin starting from the middle of the back longitudinally and in the direction of the legs.</li>
		<li>Carefully remove the skin without touching the muscles and tendons. It is possible to rip off all the skin.</li>
		<li>Cut the two legs of the mouse and rapidly proceed with the dissection. 
		NOTE: If it is more comfortable, it is possible to continue the dissection on the entire mouse but working on the leg allows more mobility and precision in the later cuts.</li>
		<li>Fix the leg on the support at the level of the foot using a pin and start isolating skeletal muscles of interest in this order: TA, EDL, Gastrocnemius and Soleus.</li>
		<li>Lift the lower tendon of TA with a sharp tweezer at ankle height and cut it, then cut with fine scissor all around the TA muscle to the other tendon at the level of the patella (Figure 3A). Transfer into the digestion solution.</li>
		<li>Lift the lower tendon of EDL and separate it from other muscles by gently pulling it upward up to the other tendon. Cut and place it into the digestion solution. 
		NOTE: Since EDL may be extremely small to be cut separately from TA, they can be dissected together (Figure 3B). Then, if the entire muscle is too big, cut it in 2-3 pieces starting from the tendon and following the fibers in a longitudinal direction (Figure 3C).</li>
		<li>Rotate the leg showing the back muscles and fix the foot using the pin. Lift the Achille&#8217;s tendon, Gastrocnemius will automatically separate from other muscles. The upper tendon is up in the back of the patella. Cut it and add the muscle to digestion.</li>
		<li>Lift the external tendon of the leg (with respect to the body) and obtain the Soleus. Gently separate it from the other muscles by scrolling under with the tweezer.</li>
		<li>Do the same for the other leg (steps from 1.2.7. to 1.2.11.).</li>
	</ol>
	</li>
	<li>Muscle digestion
	<ol>
		<li>Incubate the digestion solution containing all the muscles in a water bath at 37 &#176;C for about 45-50 min. During the digestion time, regularly check the muscle to avoid over-digestion. Every 10 min invert the digestion tubes 10x with an energetic movement (avoid vortexing).</li>
		<li>Stop the digestion when muscles start to loosen up and myofibers are visible.</li>
		<li>At the very end of the digestion time shake the samples.</li>
		<li>To stop the digestion, carefully transfer the digestion suspension to a pre-warmed 100 mm Petri dish with 10 mL of washing solution. 
		NOTE: Avoid muscle over-digestion as this will inevitably result in the isolation of hyper-contracted myofibers. Usually with this protocol, muscles of homozygous mutant mice take 45 min to be digested, while muscles of wild type mice take 50-55 min. Digestion time needs to be experimentally validated.</li>
	</ol>
	</li>
	<li>Single myofibers isolation
	<ol>
		<li>First isolate the fibers already dissociated under a dissecting microscope by picking them individually with a coated P200 pipette or small hole Pasteur and transfer them from the 100 mm Petri into a new 35 mm Petri dish with 5 mL of pre-warmed washing solution.</li>
		<li>To release further myofibers, pipette the muscle up and down using a large hole bore glass pipette with warm medium, until fibers are mechanically released. Do not be too persistent as this will result in damaging fibers.</li>
		<li>Continue releasing myofibers from the muscle until the dish contains a desirable amount. If the Petri dish is kept at room temperature for more than 8 min, stop and perform a minimum of 5 min incubation at 37 &#176;C, 5% CO2 to re-equilibrate the medium.</li>
		<li>Before transferring the single myofibers to the culture medium, leave them at 37 &#176;C, 5% CO2 in wash dish for at least 1 h. This helps myofibers to adjust to the in vitro condition. 
		NOTE: Adult myofibers are less susceptible to stress and can be washed ~ 2-3x. However, in this condition (at 19 days post-natal age) it is better to perform only one washing step to prevent myofiber damage. Therefore, always pay attention to keep selected myofibers sufficiently clean by not carrying debris or hypercontracted fibers.</li>
	</ol>
	</li>
	<li>Single myofiber culture
	<ol>
		<li>Transfer individual myofibers to a new pre-warmed dish with the appropriate culture medium (high serum medium to allow satellite cells activation, see Figure 1).</li>
		<li>Change the medium to the isolated myofibers, by transferring them to a new coated dish with new culture medium, only after 48-72 h of culture to avoid any stress that will lead to myofibers hypercontraction and disruption.</li>
	</ol>
	</li>
</ol>
2. Downstream applications: Myofibers crosslinking and immunofluorescence
NOTE: The myofibers-associated satellite cells can be visualized by immunofluorescence (IF) at the time of interest. Since most of the published protocols are optimized to perform IF on adult myofibers, here a detailed protocol is presented to obtain reliable results also on myofibers isolated from post-natal muscles.
<ol>
	<li>Myofiber crosslinking
	<ol>
		<li>Before starting, prepare all the necessary solutions as described in Table 2.</li>
		<li>Precoat with HS as many 1.5 mL microcentrifuge tubes as the number of samples. Be sure to remove all HS before proceeding. Since crosslinked fibers are tougher than the living ones and more difficult to pipette, be sure to crosslink about 200-300 fibers separately per tube.</li>
		<li>Under a dissecting microscope, collect all the fibers that can be considered healthy, transfer them to the microcentrifuge tube and leave the tube vertical for 5 min inside the incubator to allow fibers to settle.</li>
		<li>Remove the supernatant very slowly from the tube.</li>
		<li>Crosslink fibers by adding 1 mL of 4% paraformaldehyde (PFA) at RT to the tube. Do it gently to avoid fibers distress.</li>
		<li>To prevent fibers interweaving during the crosslinking, keep the tube in very gentle agitation for 10 min.</li>
		<li>Keep the tube in a vertical position for 5 min at RT to allow fibers to settle, then discard the supernatant ensuring to remove the majority of PFA volume.</li>
		<li>Add 1 mL of phosphate buffered saline (PBS) and keep the tubes vertical for 5 min at RT to let the fibers to sediment.</li>
		<li>Remove the supernatant and repeat the washing procedure (step 2.1.8.) twice again.</li>
		<li>Keep crosslinked samples at 4 &#176;C. 
		NOTE: It is possible to keep crosslinked myofibers at 4 &#176;C for a week. If the fibers remain more than a week in this condition, this will inevitably result in myofibers intertwinement.</li>
	</ol>
	</li>
	<li>Immunofluorescence
	<ol>
		<li>Keep the tubes containing the fibers standing at RT for at least 5 min to allow for fiber sedimentation.</li>
		<li>Remove the supernatant, leaving just a small volume to be sure not to remove any fiber.</li>
		<li>Add 1 mL of 0.5% Triton X-100 in PBS and incubate for 5 min with gentle agitation.</li>
		<li>Put tubes in vertical position for 5 min, then remove supernatant.</li>
		<li>Add 1.5 mL of PBS and incubate with gentle agitation for 5 min.</li>
		<li>Keep tubes in vertical position for 5 min, then remove the supernatant.</li>
		<li>Add 1 mL of blocking solution and incubate for 1 h at RT with gentle agitation.</li>
		<li>Keep tubes 5 min in vertical position, then remove the supernatant.</li>
		<li>Dilute primary antibodies in blocking solution, add them to the tubes and incubate over-night at 4 &#176;C in gentle agitation (for suggested concentrations see Table of Materials). 
		NOTE: Alternatively, the primary antibody can be incubated for 3 h at RT. However, overnight incubation gave optimal staining. The incubation volume for both primary and secondary antibodies should be of 300 &#181;L when sedimented fibers reach the 100 &#181;L notch of the 1.5 mL microcentrifuge tube. When fibers are less abundant, a volume of 100-200 &#181;L&#160;is recommended.</li>
		<li>Leave the tubes in vertical position for 5 min and then remove the supernatant.</li>
		<li>Perform 3 washes in 1 mL of 0.25% Tween-20 in PBS, incubating for 5 min in gentle agitation and then leaving the tubes standing for 5 min each time. 
		NOTE: From here on, perform all the steps in the dark, to avoid fluorochromes bleaching.</li>
		<li>Dilute secondary antibodies in blocking solution, add them to the tubes&#160;and incubate for 1 h at RT with gentle agitation (for concentrations see Table of Materials).</li>
		<li>Wash twice in 1 mL of 0.1% Tween-20 in PBS, incubating for 5 min with gentle agitation and then leaving the tubes in standing position for 5 min.</li>
		<li>Remove the supernatant, add 1 mL of DAPI solution and incubate for 5 min with gentle agitation.</li>
		<li>Let the tubes stand vertically in a rack for 5 min, then remove the supernatant.</li>
		<li>Wash with 1 mL of PBS, incubating for 5 min with gentle agitation and then leaving the tubes stand in a rack for 5 min.</li>
		<li>Remove the supernatant, leaving a volume of about 50 &#181;L.</li>
	</ol>
	</li>
	<li>Mounting of fluorescently labeled myofibers on microscope slides
	<ol>
		<li>Cut a P200 pipette tip and coat it with blocking solution 
		NOTE:To coat the tip, pipette the solution up and down several times before picking the fibers, this will avoid fibers to stick to the tip wall.</li>
		<li>Collect the fibers from the tube and spread them on a microscope glass slide.</li>
		<li>Under a dissecting microscope, using only the natural light reflected by the mirror, use a new (not cut) P200 pipette tip to spread the fibers and to remove the excess liquid solution.</li>
		<li>Leave the slides to air dry in the dark for about 10-15 min, until very low amount of solution remains.</li>
		<li>Add the mounting medium on the slide (the proper amount of mounting medium should be calibrated on the dimension of the coverslip: for a 24 x 40 mm coverslip, 20 &#181;L is enough) and then slowly lay a cover glass on the area containing the fibers. Be careful not to create bubbles between the glasses.</li>
		<li>Press the coverslip so that the fibers will lay on a single horizontal plan.</li>
		<li>Fix the coverslip using nail polish.</li>
		<li>Store the samples at 4 &#176;C for up to 4 weeks.</li>
		<li>Acquire the desired images of fluorescently labeled myofibers with a confocal microscope.</li>
	</ol>
	</li>
</ol>

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No competing interests.

Isolation of intact single myofibers is an essential method in the field of myogenesis when the main objective is to characterize cell-autonomous regenerative capacities of muscle stem cells within their niche, in healthy and pathological conditions. However, when biochemical or genomic studies are of interest, FACS-isolated satellite cells might be the best option.
Single myofibers isolation allows to follow ex-vivo, but in the most physiological way, the dynamics of all the steps single satellite cells undergo during muscle regeneration, that are: activation, cell division (asymmetric and symmetric), differentiation and return to quiescence by self-renewal. Once myofibers are grown in floating conditions, the single satellite cells activate and expand forming a cluster of cells, all deriving from the same satellite cell. Immunofluorescence analysis for proliferation, differentiation, activation or stemness markers is then optimal to quantify the proportion between cell stages.
The key step in our protocol to obtain viable and intact myofibers can be considered the rapid but gentle muscle dissection, by tendon-to-tendon isolation, to avoid any muscle damage. Our advice is to use only sharp scissors and small sharp tweezers and to limit the entire muscle dissection procedure to ten minutes. When it is difficult to isolate very small muscles (i.e., EDL and TA), it is possible to cut them together and to later divide them by using fine scissors cutting along the longitudinal plan following the fibers. This strategy will eventually give less intact myofibers, but viability will not be compromised. The same must be performed on big muscles like Gastrocnemius to facilitate digestion. Optimization of digestion time, which needs to be empirically validated, and minimal manipulation of isolated fibers are also two crucial aspects for the positive outcome of subsequent analysis.
The advantage of the protocol reported here is that it can be applied on very small mice (in age and dimension), even when their muscles are extremely fragile. Even if not mentioned above, it is possible to follow this protocol of dissection to then culture viable myofibers for longer period using basement membrane-coated dishes18,19. It is important to consider that this situation is completely different from floating condition, where adhesion stimuli and proximity stimuli are absent.

Skeletal muscle is a differentiated tissue with one of the most extended ability to regenerate after exercise or trauma1. This characteristic&#160;is mainly&#160;due to the presence of stem cells, called satellite cells because of their peripheral position between the basal lamina and the plasmalemma of the myofiber2. During post-natal development, satellite cells proliferate and progressively differentiate, thus contributing to skeletal muscle growth. Once in the adulthood, satellite cells enter a reversible quiescent state, and upon physiological or pathological trauma, they activate, proliferate and differentiate in order to repair the damaged muscles3. Defects in the capacity of satellite cells to properly transit through these different regenerative phases and to undergo self-renewal have been firmly linked to muscle wasting, either during physiological aging4,5,6 or in muscle degenerative diseases, such as muscular dystrophies7,8,9,10.
Two main culture approaches exist to study satellite cells ex-vivo: primary myogenic cultures from mononucleated cells, mechanically and chemically dissociated from whole muscle11,12; or culture of isolated myofibers13,14,15,16,17,18,19,20. In the first case, the process of satellite cells isolation involves the trituration of whole muscles extracted from the mouse, a chemical digestion, filtration and fluorescent activated cell sorting (FACS)21. This procedure, although effective in isolating satellite cells from a variety of models, entails several variables that expose satellite cells to stress and disrupts their physiological niche22,23. By contrast, myofiber isolation involves a gentler digestion of muscle tissue with matrix degrading enzymes and a mechanical shredding that causes reduced trauma to stem cells20. This second approach allows a much more efficient retrieval of viable satellite cells, that remain physically attached to their myofiber between the basal lamina and the sarcolemma, thus allowing analysis within their physiological niche19,20.
Many different protocols have been proposed during the past years to properly and efficiently isolate single myofibers from skeletal muscles. Already in 1986 Bischoff proposed a protocol to isolate fibers from the Flexor Digitorum Brevis13 and later, in 1995, Rosenblatt et al. modified the protocol to obtain a more efficient separation of myofibers14. Since then, many other authors proposed adjusted procedures on other muscles, such as Extensor Digitorum Longus (EDL) and Tibialis Anterior (TA)15,16,17,18,19,20, that are longer, even if more fragile, muscles14. Isolated myofibers can then be cultivated both in adhesion, to allow for the expansion of satellite cells-derived myoblasts, or in floating conditions, up to&#160;96 hours, to follow the progeny derived from single satellite cells19 (Figure 1). Variable concentrations of serum within the culture medium are used to trigger satellite cells activation, proliferation and/or differentiation, to study their capacity&#160;to properly transit through these different phases1.
We recently described the epigenetic mechanism behind the exhaustion of the satellite stem cell pool in the mouse model of EDMD, the Lamin &#916;8-11 -/- mouse7. Since these mice usually die between 4-8 weeks of age24, due to severe muscle loss, an attempt was made to capture the molecular defects underlying the early onset of the disease by focusing our analysis on post-natal muscle development. Floating single myofibers were isolated and cultured from wild type and Lamin &#916;8-11 -/- mutant7 19 days-old mice. At this stage, muscle defects are already evident, but mice are still viable. However, since all the above-mentioned protocols for single myofibers extraction were optimized for skeletal muscles of adult mice, we needed to adapt them to our purposes: very small mice in term of age and size, and very fragile myofibers. Thus, we describe here our re-adaptation of the protocol proposed by the Rudnicki laboratory19 to obtain a significant number of single viable myofibers from mice during post-natal development and from severe dystrophic muscles, such as those derived from Lamin &#916;8-11 -/- mice24. The final goal of this approach is to provide a standardized procedure&#160;for the study of myofibers-associated muscle stem cells in any other mouse model when the early stages of post-natal development are of interest, or in the case of mouse models carrying any specific disease that makes myofibers more susceptible to mechanical stress.

<table><tbody><tr><td>4&amp;prime;,6-diamidino-2-phenylindole</td><td>Sigma</td><td>D9542</td><td></td></tr><tr><td>Chicken embryo extract</td><td>Seralab</td><td>CE650-DL</td><td></td></tr><tr><td>Collagenase type I</td><td>SIGMA</td><td>C0130-500MG</td><td></td></tr><tr><td>Donkey anti-Rabbit 488 antibody</td><td>Thermofisher</td><td>R37118</td><td>to be used 1:200</td></tr><tr><td>Dulbecco&#039;s modified Eagle&#039;s medium</td><td>Gibco</td><td>10569-010</td><td></td></tr><tr><td>Fetal bovine serum</td><td>Corning</td><td>35-015-CV</td><td></td></tr><tr><td>Horse serum</td><td>Gibco</td><td>26050-088</td><td></td></tr><tr><td>MyoG antibody</td><td>Millipore</td><td>219998</td><td>to be used 1:100</td></tr><tr><td>Paraformaldehyde</td><td>SIGMA</td><td>P6148</td><td></td></tr><tr><td>Pax7 antibody</td><td>Developmental studies&lt;br/&gt; Hybridoma bank</td><td></td><td>to be used 1:20</td></tr><tr><td>Penicillin and Streptomycin</td><td>Euroclone</td><td>ECB3001D</td><td></td></tr><tr><td>Phosphate saline buffer</td><td>Euroclone</td><td>ECB4004L</td><td></td></tr><tr><td>Prolong gold antifade mountant</td><td>Thermofisher</td><td>P36930</td><td></td></tr><tr><td>Triton X-100</td><td>SIGMA</td><td>T8787</td><td></td></tr><tr><td>Tween-20</td><td>SIGMA</td><td>P1379</td><td></td></tr><tr><td>&lt;strong&gt;Lab equipment&lt;/strong&gt;</td><td>&lt;strong&gt;Manufacturer&lt;/strong&gt;</td><td></td><td></td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>Bunsen burner</td><td></td><td></td><td></td></tr><tr><td>Confocal microscope</td><td>Leica</td><td></td><td></td></tr><tr><td>Diamond pen</td><td>bio-optica</td><td></td><td></td></tr><tr><td>Dissection pins</td><td></td><td></td><td></td></tr><tr><td>FACS polypropylene tubes</td><td>Falcon</td><td></td><td></td></tr><tr><td>Falcon tubes (50 and 15 mL)</td><td>Falcon</td><td></td><td></td></tr><tr><td>Glass coverslips</td><td>Thermofisher</td><td></td><td></td></tr><tr><td>Glass Pasteur pipettes (22cm)</td><td>VWR</td><td></td><td></td></tr><tr><td>Glass slides</td><td>Thermofisher</td><td></td><td></td></tr><tr><td>Micro dissecting scissors</td><td></td><td></td><td></td></tr><tr><td>Micropipette (1 mL and 200 &amp;micro;L)</td><td>Gilson</td><td></td><td></td></tr><tr><td>Micropipette tips</td><td>Corning</td><td></td><td></td></tr><tr><td>Petri dishes (100 and 35 mm diameter)</td><td>Thermofisher</td><td></td><td></td></tr><tr><td>Plastic pipettes (5 and 10 mL)</td><td>VWR</td><td></td><td></td></tr><tr><td>Rubber pipette bulbs</td><td>VWR</td><td></td><td></td></tr><tr><td>Sharp tweezers</td><td></td><td></td><td></td></tr><tr><td>Stereo dissection microscope with transmission illumination</td><td>Leica</td><td></td><td></td></tr><tr><td>Tissue culture hood or lamina flow cabinet</td><td></td><td></td><td></td></tr><tr><td>Tissue culture incubator (humidified, 37&amp;deg;C, 5% CO2)</td><td></td><td></td><td></td></tr><tr><td>Water bath at 37&amp;deg;C</td><td>VWR</td><td></td><td></td></tr></tbody></table>

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.

We typically digest four different muscles (TA, EDL, Soleus and Gastrocnemius) to retrieve a good amount of long and viable fibers that could survive 96 h in growth-factors rich medium (Figure 4A,B). Only the most intact fibers should&#160;be transferred in culture medium, as they will survive; all the others, that are easy to discriminate and select, need to be discarded.
When myofibers are maintained in a growth-factors rich medium satellite cells derived from wild type mice start to activate and proliferate, see Figure 1. Upon 48 h of culture, in healthy condition (Lamin &#916;8-11 +/+), satellite cells upregulate MyoD and undergo their first division. Activated Pax7+/MyoD+ satellite cells then proliferate and by 72 h in culture, they generate cell aggregates bound to the myofiber, that are even more visible at 96 h (Figure 5). During these divisions, some of them can repress MyoD expression, undergoing self-renewal to repopulate the stem cells pool, while those that maintain MyoD become committed to differentiation by downregulating Pax7 expression. After 96 h, satellite cells clusters contain clearly visible MyoG+ committed cells, that can differentiate into new myofibers (Figure 1 and Figure 6). Notably, with this experiment, we described a delayed dynamics of satellite cell differentiation in homozygous mutant Lamin &#916;8-11 mice (-/-) as compared to their wild type counterparts (+/+), see Figure 6.
The final outcome of each single experiment let us think that the protocol developed for single myofibers isolation and culture from this model of severe muscle dystrophy ensures good quality myofibers for all further applications.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61516/61516fig01.jpg" /> 
Figure 1: Graphical representation of satellite cells&#8217; regenerative phases modelled in floating myofibers.&#160;Upon 48 h of culture in growth-factors rich medium, Pax7+ cells get activated and undergo the first division, giving rise to a doublet of Pax7+/MyoD+ cells. MyoD positive cells then proliferate and expand, giving rise, in 72 h of culture, to a cluster of several cells which are the progeny of a single satellite cell. Upon 96 h of culture Pax7+/MyoD+ cells become differentiating Pax7-/MyoG+ cells. During the expansion phase, a subset of Pax7+/MyoD+ cells downregulate MyoD expression undergoing self-renewal into quiescence. <a href="https://www.jove.com/files/ftp_upload/61516/61516fig01large.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/61516/61516fig02.jpg" /> 
Figure 2: Preparation of bore Pasteur pipettes.&#160;(A) Longitudinal and frontal view of how the big hole bore pipette must appear. (B) Longitudinal and frontal view of how the small hole bore pipette must finally appear. <a href="https://www.jove.com/files/ftp_upload/61516/61516fig02large.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/61516/61516fig03.jpg" /> 
Figure 3: Representative pictures of single muscle dissection.&#160;(A) Isolation of the TA muscle. Avoiding the removal of the thin layer covering the muscle protects&#160;the myofibers inside. (B) TA and EDL muscles isolated together still attached to their upper tendon at the level of the patella. (C) Division of TA and EDL after isolation by cutting them along the longitudinal axis. <a href="https://www.jove.com/files/ftp_upload/61516/61516fig03large.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/61516/61516fig04.jpg" /> 
Figure 4: Examples of healthy and viable myofibers.&#160;Representative phase contrast images of viable myofibers in suspension. (A) The red arrow indicates a myofiber with visible sarcomere organization and a satellite cell on its side; the orange arrows indicate some pieces of broken myofibers, and some debris present in the first dish before the final selection for culture. Scale bar 100 &#181;m. (B) More complete view of other myofibers under a smaller magnification. Scale bar 500 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61516/61516fig04large.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/61516/61516fig05.jpg" /> 
Figure 5: Difference&#160;in the dimension of stem cell clusters in wt and mutant mice.&#160;Immunofluorescence staining of myofibers extracted from 19 days Lamin &#916;8-11 mice (+/+ and -/-) after 96 h of culture. Pax7+ satellite cells are shown. The dimension of the cell cluster in most of the cases was significantly bigger in Lamin &#916;8-11 +/+ than in Lamin &#916;8-11 -/- in terms of number of cells. Scale bar 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61516/61516fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61516/61516fig06.jpg" /> 
Figure 6: Representative immunofluorescence experiment.&#160;Immunofluorescence experiment performed, after 96 h of culture, on myofibers extracted from 19 days Lamin &#916;8-11 mice (+/+ and -/-).&#160;Pax7+/MyoG- (red) and Pax7-/MyoG+ (green) cells were observed. Images obtained with a confocal microscope. Scale bar 25 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61516/61516fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Name of the solution in the text</td>
			<td>component</td>
			<td>percentage</td>
			<td>suggested final volume/sample</td>
			<td colspan="2">notes</td>
		</tr>
		<tr>
			<td rowspan="2">washing solution</td>
			<td>DMEM high glucose</td>
			<td>90%</td>
			<td rowspan="2">4 mL</td>
			<td colspan="2" rowspan="2">Keep sterile, keep at 4&#176;C until usage</td>
		</tr>
		<tr>
			<td>Horse serum (HS)</td>
			<td>10%</td>
		</tr>
		<tr>
			<td rowspan="2">digestion solution</td>
			<td>DMEM high glucose</td>
			<td>9.80%</td>
			<td rowspan="2">20 mL</td>
			<td colspan="2" rowspan="2">Extremely harmful powder. Filter solution with 0.22&#181;m filter and then keep sterile. Keep at 4&#176;C until usage</td>
		</tr>
		<tr>
			<td>Collagenase I</td>
			<td>0.20%</td>
		</tr>
		<tr>
			<td rowspan="4">culture medium</td>
			<td>DMEM high glucose</td>
			<td>78%</td>
			<td rowspan="4">10 mL</td>
			<td colspan="2" rowspan="4">Keep sterile, keep at 4&#176;C until usage</td>
		</tr>
		<tr>
			<td>Fetal bovine serum (FBS)</td>
			<td>20%</td>
		</tr>
		<tr>
			<td>Chicken embryo extract (CEE)</td>
			<td>1%</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin (P/S)</td>
			<td>1%</td>
		</tr>
	</tbody>
</table>
Table 1: Recipes for solutions used in section 1.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Name of the solution in the text</td>
			<td>component</td>
			<td>percentage</td>
			<td>suggested final volume/sample</td>
			<td colspan="2">notes</td>
		</tr>
		<tr>
			<td rowspan="2">4% PFA</td>
			<td>Paraformaldehyde (PFA)</td>
			<td>4%</td>
			<td rowspan="2">2-3 mL</td>
			<td colspan="2" rowspan="2">Powder and then solution are extremely harmful</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>96%</td>
		</tr>
		<tr>
			<td rowspan="2">0.5% Triton X-100</td>
			<td>Triton X-100</td>
			<td>0.50%</td>
			<td rowspan="2">2-3 mL</td>
			<td colspan="2" rowspan="2">Triton X-100 is extremely viscous, preferentially cut the tip of the pipette to aliquot it</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>99.50%</td>
		</tr>
		<tr>
			<td rowspan="2">0.25% Tween-20</td>
			<td>Tween-20</td>
			<td>0.25%</td>
			<td rowspan="2">10 mL</td>
			<td colspan="2" rowspan="2">Tween-20 is extremely viscous, preferentially cut the tip of the pipette to aliquot it</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>99.75%</td>
		</tr>
		<tr>
			<td rowspan="2">0.1% Tween-20</td>
			<td>Tween-20</td>
			<td>0.10%</td>
			<td rowspan="2">10 mL</td>
			<td colspan="2" rowspan="2">Tween-20 is extremely viscous, preferentially cut the tip of the pipette to aliquot it</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>99.90%</td>
		</tr>
		<tr>
			<td rowspan="2">blocking solution</td>
			<td>Fetal bovine serum (FBS)</td>
			<td>10%</td>
			<td rowspan="2">10 mL</td>
			<td colspan="2" rowspan="2">Prepare fresh solution and store at 4&#176;C for no longer than 3 weeks (always check clearness before usage)</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>90%</td>
		</tr>
		<tr>
			<td rowspan="2">DAPI</td>
			<td>DAPI</td>
			<td>0.10%</td>
			<td rowspan="2">2-3 mL</td>
			<td colspan="2" rowspan="2">Keep in the dark</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>99.90%</td>
		</tr>
	</tbody>
</table>
Table 2: Recipes for solutions used in section 2.

Watch this Scientific Journal Video about Single Myofiber Isolation and Culture from a Murine Model of Emery-Dreifuss Muscular Dystrophy in Early Post-Natal Development at JoVE.com

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

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 ...

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Research

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Medicine

7.6K Views.  Istituto Nazionale di Genetica Molecolare "Romeo ed Enrica Invernizzi". Here, we propose a method to efficiently obtain single muscle fibers at early post-natal developmental stages from homozygous mutant Lamin Δ8-11 mouse model, a very severe model for Emery-Dreifuss muscular dystrophy (EDMD).

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

In Vitro Differentiation of Mature Myofibers for Live Imaging

Skeletal muscles are composed of myofibers, the biggest cells in the mammalian body and one of the few syncytia. How the complex and evolutionarily conserved structures that compose it are assembled remains under investigation. Their size and physiological features often constrain manipulation and imaging applications. The culture of immortalized cell lines is widely used, but it can only replicate the early steps of differentiation.
Here, we describe a protocol that enables easy genetic manipulation of myofibers originating from primary mouse myoblasts. After one week of differentiation, the myofibers display contractility, aligned sarcomeres and triads, as well as peripheral nuclei. The entire differentiation process can be followed by live imaging or immunofluorescence. This system combines the advantages of the existing ex vivo and in vitro protocols. The possibility of easy and efficient transfection as well as the ease of access to all differentiation stages broadens the potential applications. Myofibers can subsequently be used not only to address relevant developmental and cell biology questions, but also to reproduce muscle disease phenotypes for clinical applications.

In Vitro Differentiation of Mature Myofibers for Live Imaging

Muscle cells are among the most complex eukaryotic cells. We present a protocol for the in vitro differentiation of highly mature myofibers that allows for genetic manipulation and clear imaging during all developmental stages.

Skeletal muscles are composed of myofibers, the biggest cells in the mammalian body and one of the few syncytia. How the complex and evolutionarily ...

Developmental Biology

Direct Reprogramming of Human Fibroblasts into Myoblasts to Investigate Therapies for Neuromuscular Disorders

Investigations into both the pathophysiology and therapeutic targets in muscular dystrophies have been hampered by the limited proliferative capacity of human myoblasts. Several mouse models have been created but they either do not truly represent the human physiopathology of the disease or are not representative of the broad spectrum of mutations found in humans. The immortalization of human primary myoblasts is an alternative to this limitation; however, it is still dependent on muscle biopsies, which are invasive and not easily available. In contrast, skin biopsies are easier to obtain and less invasive to patients. Fibroblasts derived from skin biopsies can be immortalized and transdifferentiated into myoblasts, providing a source of cells with excellent myogenic potential. Here, we describe a fast and direct reprogramming method of fibroblast into a myogenic lineage. Fibroblasts are transduced with two lentiviruses: hTERT to immortalize the primary culture and a tet-inducible MYOD, which upon the addition of doxycycline, induces the conversion of fibroblasts into myoblasts and then mature myotubes, which express late differentiation markers. This quick transdifferentiation protocol represents a powerful tool to investigate pathological mechanisms and to investigate innovative gene-based or pharmacological biotherapies for neuromuscular disorders.

This protocol describes the conversion of skin fibroblasts into myoblasts and their differentiation into myotubes. The cell lines are derived from patients with neuromuscular disorders and can be used to investigate pathological mechanisms and to test therapeutic strategies.

Investigations into both the pathophysiology and therapeutic targets in muscular dystrophies have been hampered by the limited proliferative capacity of ...

Genetics

Isolation and Differentiation of Primary Myoblasts from Mouse Skeletal Muscle Explants

Primary myoblasts are undifferentiated proliferating precursors of skeletal muscle. They can be cultured and studied as muscle precursors or induced to differentiate into later stages of muscle development. The protocol provided here describes a robust method for the isolation and culture of a highly proliferative population of myoblast cells from young adult mouse skeletal muscle explants. These cells are useful for the study of the metabolic properties of skeletal muscle of different mouse models, as well as in other downstream applications such as transfection with exogenous DNA or transduction with viral expression vectors. The level of differentiation and metabolic profile of these cells depends on the length of exposure, and composition of the media used to induce myoblast differentiation. These methods provide a robust system for the study of mouse muscle cell metabolism ex vivo. Importantly, unlike in vivo models, the methods described here provide a cell population that can be expanded and studied with high levels of reproducibility.

Myoblasts are proliferating precursor cells that differentiate to form polynucleated myotubes and eventually skeletal muscle myofibers. Here, we present a protocol for efficient isolation and culture of primary myoblasts from young adult mouse skeletal muscles. The method enables molecular, genetic, and metabolic studies of muscle cells in culture.

Primary myoblasts are undifferentiated proliferating precursors of skeletal muscle. They can be cultured and studied as muscle precursors or induced to ...

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 ...

Biology

Preparation of 3D Decellularized Matrices from Fetal Mouse Skeletal Muscle for Cell Culture

The extracellular matrix (ECM) plays a crucial role in providing structural support for cells and conveying signals that are important for various cellular processes. Two-dimensional (2D) cell culture models oversimplify the complex interactions between cells and the ECM, as the lack of a complete three-dimensional (3D) support can alter cell behavior, making them inadequate for understanding in vivo processes. Deficiencies in ECM composition and cell-ECM interactions are important contributors to a variety of different diseases. 
One example is LAMA2-congenital muscular dystrophy (LAMA2-CMD), where the absence or reduction of functional laminin 211 and 221 can lead to severe hypotony, detectable at or soon after birth. Previous work using a mouse model of the disease suggests that its onset occurs during fetal myogenesis. The present study aimed to develop a 3D in vitro model permitting the study of the interactions between muscle cells and the fetal muscle ECM, mimicking the native microenvironment. This protocol uses deep back muscles dissected from E18.5 mouse fetuses, treated with a hypotonic buffer, an anionic detergent, and DNase. The resultant decellularized matrices (dECMs) retained all ECM proteins tested (laminin &#945;2, total laminins, fibronectin, collagen I, and collagen IV) compared to the native tissue. 
When C2C12 myoblasts were seeded on top of these dECMs, they penetrated and colonized the dECMs, which supported their proliferation and differentiation. Furthermore, the C2C12 cells produced ECM proteins, contributing to the remodeling of their niche within the dECMs. The establishment of this in vitro platform provides a new promising approach to unravel the processes involved in the onset of LAMA2-CMD, and has the potential to be adapted to other skeletal muscle diseases where deficiencies in communication between the ECM and skeletal muscle cells contribute to disease progression.

In this work, a decellularization protocol was optimized to obtain decellularized matrices of fetal mouse skeletal muscle. C2C12 myoblasts can colonize these matrices, proliferate, and differentiate. This in vitro model can be used to study cell behavior in the context of skeletal muscle diseases such as muscular dystrophies.

The extracellular matrix (ECM) plays a crucial role in providing structural support for cells and conveying signals that are important for various ...

Isometric and Eccentric Force Generation Assessment of Skeletal Muscles Isolated from Murine Models of Muscular Dystrophies

Critical to the evaluation of potential therapeutics for muscular disease are sensitive and reproducible physiological assessments of muscle function. Because many pre-clinical trials rely on mouse models for these diseases, isolated muscle function has become one of the standards for Go/NoGo decisions in moving drug candidates forward into patients. We will demonstrate the preparation of the extensor digitorum longus (EDL) and diaphragm muscles for functional testing, which are the predominant muscles utilized for these studies. The EDL muscle geometry is ideal for isolated muscle preparations, with two easily accessible tendons, and a small size that can be supported by superfusion in a bath. The diaphragm exhibits profound progressive pathology in dystrophic animals, and can serve as a platform for evaluating many potential therapies countering fibrosis, and promoting myofiber stability. Protocols for routine testing, including isometric and eccentric contractions, will be shown. Isometric force provides assessment of strength, and eccentric contractions help to evaluate sarcolemma stability, which is disrupted in many types of muscular dystrophies. Comparisons of the expected results between muscles from wildtype and dystrophic muscles will also be provided. These measures can complement morphological and biochemical measurements of tissue homeostasis, as well as whole animal assessments of muscle function.

Muscle function measurements contribute to the evaluation of potential therapeutics for muscle pathology, as well as to the determination of mechanisms underlying physiology of this tissue. We will demonstrate the preparation of the extensor digitorum longus and diaphragm muscles for functional testing. Protocols for isometric and eccentric contractions will be shown, as well as differences in results between dystrophic muscles, representing a pathological state, and wildtype muscles.

Critical to the evaluation of potential  therapeutics for muscular disease are sensitive and reproducible physiological  assessments of muscle function. ...

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 ...

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

The zebrafish has proven to be a valuable model system for exploring skeletal muscle function and for studying human muscle diseases. Despite the many advantages offered by in vivo analysis of skeletal muscle in the zebrafish, visualizing the complex and finely structured protein milieu responsible for muscle function, especially in whole embryos, can be problematic. This hindrance stems from the small size of zebrafish skeletal muscle (60 &mu;m) and the even smaller size of the sarcomere. Here we describe and demonstrate a simple and rapid method for isolating skeletal myofibers from zebrafish embryos and larvae. We also include protocols that illustrate post preparation techniques useful for analyzing muscle structure and function. Specifically, we detail the subsequent immunocytochemical localization of skeletal muscle proteins and the qualitative analysis of stimulated calcium release via live cell calcium imaging. Overall, this video article provides a straight-forward and efficient method for the isolation and characterization of zebrafish skeletal myofibers, a technique which provides a conduit for myriad subsequent studies of muscle structure and function.

Zebrafish are an emerging system for modeling human disorders of the skeletal muscle. We describe a fast and efficient method to isolate skeletal muscle myofibers from embryonic and larval zebrafish. This method yields a high-density myofiber preparation suitable for study of single skeletal muscle fiber morphology, protein subcellular localization, and muscle physiology.

The zebrafish has  proven to be a valuable model system for exploring skeletal muscle function and  for studying human muscle diseases. Despite  the many ...

DNA Electroporation, Isolation and Imaging of Myofibers

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently labeled proteins in the flexor digitorum brevis (FDB) muscle. Conditions have been optimized to provide a large number of high quality myofibers expressing the electroporated plasmid while minimizing muscle damage. The method employs fluorescent tags on various proteins. Combining this expression method with high resolution confocal microscopy permits live cell imaging, including imaging after laser-induced damage. Fluorescent dyes combined with imaging of fluorescently-tagged proteins provides information regarding the basic structure of muscle and its response to stimuli.

This protocol utilizes electroporation to introduce and express fluorescently labeled proteins in mouse muscle fibers. Following recovery after electroporation, fibers are isolated. Individual fibers are then imaged using high resolution confocal microscopy to visualize muscle structure.

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently ...

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.

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

Summary

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Chapters in this video

In Vitro Differentiation of Mature Myofibers for Live Imaging

Direct Reprogramming of Human Fibroblasts into Myoblasts to Investigate Therapies for Neuromuscular Disorders

Isolation and Differentiation of Primary Myoblasts from Mouse Skeletal Muscle Explants

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

Preparation of 3D Decellularized Matrices from Fetal Mouse Skeletal Muscle for Cell Culture

Isometric and Eccentric Force Generation Assessment of Skeletal Muscles Isolated from Murine Models of Muscular Dystrophies

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

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

DNA Electroporation, Isolation and Imaging of Myofibers

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