Name: dna electroporation, isolation and imaging of myofibers
Uploaded: 2015-12-23T15:00:00.000+00:00
Duration: 7 min 52 s
Description: Watch this Scientific Journal Video about DNA Electroporation, Isolation and Imaging of Myofibers at JoVE.com

This work was supported by National Institutes of Health grants NS047726, NS072027, and AR052646.

The methods in this study were performed in ethical accordance with the Northwestern University Feinberg School of Medicine Institutional Animal Care and Use Committee (IACUC) approved guidelines. All efforts were made to minimize suffering.
1. Experimental Procedure for in vivo Electroporation of the Flexor Digitorum Brevis (FDB) Muscle Bundle in Mouse
<ol>
	<li>Design plasmids for in vivo expression using a promoter known to express in mammalian cells (e.g., cytomegalovirus, CMV) or in muscle (e.g., Muscle Creatine Kinase, MCK) 6,7. Larger plasmids may express at lower levels. The CMV promoter has been used extensively 6,7.</li>
	<li>Purify plasmids from E. coli using endotoxin free conditions per manufacturer&#39;s instructions. Dissolve the plasmid in sterile, endotoxin free TE buffer at 2 - 5 &#181;g plasmid/&#181;l.</li>
	<li>Dilute and aliquot the plasmid to a concentration of 20 &#181;g of DNA in 10 &#181;l of sterile endotoxin free TE (2 &#181;g/&#181;l) buffer per injection into a sterile microcentrifuge tube. Prepare one aliquot per injection.</li>
	<li>Dilute the stock of hyaluronidase to 1x with sterile Phosphate Buffered Saline without calcium and magnesium (PBS-/-) giving a final concentration of 8 units. Aliquot 10 &#181;l of diluted hyaluronidase per injection into a sterile microcentrifuge tube.</li>
	<li>Anesthetize the mouse using 2.5% isoflurane until sedated and unresponsive to tail or toe pinching. Place the animal in a supine position on a heating pad or heating table to maintain normal body temperature.</li>
	<li>Clean the footpad of the mouse with an alternating application of a povidone iodine scrub and alcohol three times using a prep technique originating from the needle site and radiating outwards. This technique maintains the needle site in an aseptic condition, minimizing pathogen introduction.</li>
	<li>Inject the footpad of the mouse with 10 &#181;l of the 1x hyaluronidase solution (2 mg/ml = &#160;8 units) between the skin and the muscle using a sterile 1 ml syringe. Hyaluronidase digests components of the extracellular matrix to permit more efficient entry of plasmid into myofibers. Enter at the base of the heel and advance the needle towards the toes. Slowly release the liquid as the needle is retracted. The injection site under the skin will expand and start to turn pink.</li>
	<li>Repeat steps 1.6 and 1.7 on the contralateral foot.</li>
	<li>Disconnect the anesthesia and place the mouse back in the cage. Allow the mouse to fully recover from anesthesia.</li>
	<li>Wait 2 hr.</li>
	<li>Repeat the anesthetizing and foot sterilization procedures, steps 1.5 and 1.6.</li>
	<li>Inject the diluted plasmid DNA into the footpad in the same fashion as the hyaluronidase. Ensure that the maximum volume is 20 &#181;l or less. For dual plasmid expression inject 20 &#181;g of each plasmid with a maximum volume of 20 &#181;l or less.</li>
	<li>Repeat the plasmid injection on the contralateral footpad or use the contralateral foot for control injections.</li>
	<li>On the foot that was initially injected, lift the skin away from the underlining muscle with fine forceps and place one 27 G needle through the balls of skin near the toes in a position perpendicular to the heal-toe line of the foot. Take a second sterile 27 G needle and place it horizontally through the skin at the heel. Place needles parallel to each other ~1 cm apart.</li>
	<li>Using alligator clamps, clamp electrodes to the parallel needles making sure the needles do not contact one another. The use of modeling clay can secure the clamps in place.</li>
	<li>Connect electrodes to an electrical stimulator.</li>
	<li>Electroporate the muscle by applying 20 pulses, 20 msec in duration/each, at 1 Hz, at 100 V/cm. The toes may twitch with stimulation.</li>
	<li>Repeat the stimulation procedure on the contralateral foot if required.</li>
	<li>Remove needles. Wipe the footpad with povidone iodine scrub.</li>
	<li>Turn off the anesthesia. Return the mouse to the recovery cage and monitor activity. If the procedure was conducted as expected, the animal should regain normal ambulation within 30 min and exhibit no difference in ambulation or activity level from pre-injection behavior. Therefore, no post-procedure analgesics are needed. Upon recovery, return the animal to the vivarium. 
	Note: Protein expression can be assayed as early as 48 hrs after electroporation. However, to allow more complete recovery after electroporation, we prefer isolation and study of myofibers 7 - 14 days after electroporation 10. Expression can be assayed as long as 4 weeks after electroporation.</li>
</ol>
2. Experimental Procedure for Isolation of the Flexor Digitorum Brevis (FDB) Fibers
<ol>
	<li>Thaw collagenase. For each FDB bundle prepare two wells in a 12-well tissue culture plate. In one well add 1 ml of pre-warmed Dulbecco&#39;s Modified Eagles Media plus Bovine Serum Albumin (DMEM + BSA) and in the other well add 1 ml of pre-warmed Ringers Solution. Additionally, add fresh 10 ml of 1x Ringers solution to the 35 mm Sylgard dish.</li>
	<li>Sacrifice the animal through CO2 or anesthetic gas inhalation, followed by cervical dislocation or other method to ensure death.</li>
	<li>Remove the hind feet above the ankle joint using a clean razor blade and submerge in 1x&#160;Ringers solution in the 35 mm Sylgard dish.</li>
	<li>Pin one foot tight, sole facing up, onto the Sylgard dish with 30 G needles through each of the 5 toe tips and at the ankle.&#160;</li>
	<li>Starting at the ankle, snip the skin straight up the side of the foot along the hairline towards the toes, careful not to nick the muscle beneath the skin. Repeat on the other side of foot.</li>
	<li>Holding the skin at the base of the ankle with fine forceps, cut the skin across the heel.</li>
	<li>Lifting the skin flap with forceps, point your scissors toward the skin as to not nick the muscle. Snip the underlying connective tissue, effectively removing the skin.</li>
	<li>To remove the FDB muscle bundle cut the large tendon at the heel to free the tendon from the bone. Holding the tendon with forceps, dissect the FDB bundle from the large white tendon carefully removing any connective tissue attached to the FDB with scissors. If done properly the bundle will easily lift off of the underlying tendons revealing bright white tendons leading to the individual digits. Cut the FDB at the tendons near the toes freeing the bundle from the foot.</li>
	<li>Place the entire muscle bundle in the well containing DMEM + BSA.</li>
	<li>Repeat on the contralateral foot.</li>
	<li>Once all FDB muscles are isolated, add 100 &#181;l of collagenase to each well containing DMEM + BSA and muscle.</li>
	<li>Check the success of electroporation on an inverted fluorescent microscope prior to digestion. If done properly, upwards of 90% transfection efficiency can be achieved.</li>
	<li>Incubate the plate in a humidified, 37 &#176;C incubator at 10% CO2 for approximately 1 hr. The incubation time may vary depending on disease model and background and the lot of collagenase.</li>
	<li>After 1 hr, carefully transfer the FDB bundle to the well containing only Ringers solution.</li>
	<li>Triturate the FDB bundle in the well with the Ringers solution using a 1 ml pipette. Cut the end of the pipette tip with a clean razor blade such that the bundle can easily pass through the pipette tip. Gently triturate the muscle (~15 times) allowing the bundle to pass up and down parallel to the tip. Intact fibers will fall off into the Ringers Solution. When fibers do not easily fall off the bundle, place the muscle back into collagenase solution.</li>
	<li>Plate the isolated fibers on the dish ~ 500 &#181;l per plate.</li>
	<li>Allow the muscle to digest in the well for 15 min.</li>
	<li>Repeat steps 2.15 to 2.17 until the bundle decreases in size and the majority of fibers are dissociated from the bundle (usually a total of 2-3 times). Once dissociated the muscle easily passes through a 1 ml pipette tip.</li>
	<li>Let fibers attach to the dish for a minimum of 15 min. Fresh Ringers solution can be added to the plate to dilute residual collagenase or changed depending on time constraints. 
	Note: Fibers are now ready for imaging.</li>
</ol>
3. Experimental Procedure for Visualization of Laser-induced Membrane Damage using Confocal Microscopy
<ol>
	<li>Load fibers immediately prior to imaging with 300 &#181;l of 2.5 &#181;M FM 1-43 of FM 4-64 dye in Ringers solution. Generally, image fibers 15 min to 2 hr after isolation.&#160;However, fibers can be imaged up to 24 hr post isolation.</li>
	<li>Place the glass bottom dish on a confocal microscope equipped with a UV laser and a 60X / 63X objective.</li>
	<li>Locate a fiber. Ensure the fiber is firmly attached to the dish by tapping the microscope base. Exclude fibers that were damaged in the dissociation process. Damaged fibers may contain membrane blebs, uptake high levels of FM dye, curl or bend intensely.</li>
	<li>To induce membrane damage on the fiber, position the sarcolemma in the center of the imaging window with a field clear of nuclei. Focus on the sarcolemma using the FM dye channel such that the sarcolemma is extremely sharp.</li>
	<li>Place the ROI cross hair on the sarcolemma utilizing the FM 4-64 channel.</li>
	<li>Irradiate a 1 pixel point at 80% power for 3 sec (~ 350 nm x 350 nm area). FM dye will enter the cell at the site of injury. Power and time can be adjusted to increase or decrease the area of insult.</li>
	<li>Capture images of the fiber before damage, at the time of damage, every 2 sec post-damage, then every 10 sec for up to 5 min. Timing can be adjusted depending on need. Individual images can be converted to time-lapse movies in Image J.</li>
	<li>Repeat steps 3.3 through 3.7 on up to 3 fibers per dish.</li>
</ol>

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There are no conflicts of interest.

The use of electroporation to study fluorescently tagged proteins in vivo is ideally suited for the study of muscle given the absence of suitable cell line models that faithfully replicate the entire cell structure of muscle. This protocol describes the utilization of electroporation and high-resolution confocal microscopy to provide detailed imaging of protein localization and translocation after laser wounding. This method can be adapted to study other cellular processes including cytoskeleton reorganization, ...

Individual myofibers are large, highly organized syncytial cells. There are a few cell-culture models for muscle; however, these models have as their major limitation that they do not fully differentiate into mature myofibers. For example, the C2C12 and L6 cell lines are derived from mice and rats, respectively 1-4. Under conditions of serum starvation, the mononuclear &#34;myoblast-like&#34; cells cease proliferation, undergo cell cycle withdrawal, and enter into the myogenic program forming multinucleated cells with sarcomeres, referred to as myotubes. With prolonged culture conditions, myotubes may exhibit contractile properties and &#34;twitch&#34; in culture. Human cell lines have also now been established 5. In addition to these immortalized cell lines, mononuclear myoblasts can be isolated from muscle and under the similar conditions of serum starvation will form myotubes. These cell lines and primary myoblast cultures are highly useful because they can be transfected with plasmids or transduced with viruses and used to study basic cell biological processes. However, these cells, even when induced to form myotubes, lack many of the salient features of mature muscle organization. Specifically, myotubes are much smaller than individual mature myofibers and lack the normal shape of myofibers. Critically, myotubes lack transverse (T-) tubules, the membranous network required for efficient Ca2+ release throughout the myoplasm.
An alternative method to primary myoblast or myogenic cell lines entails using mature myofibers. Transgenesis can be employed to establish expression of tagged proteins, but this method is costly and time consuming. In vivo electroporation of mouse muscle has emerged as a preferred method for its speed and reliability 6-10.
Methods for in vivo electroporation and the efficient isolation of myofibers have been optimized for the mouse flexor digitorum brevis (FDB) muscle 6. The methods can be completed readily and are minimally invasive to induce in vivo expression from plasmids. This approach is now combined with high resolution imaging methods including imaging after laser disruption of the sarcolemma 6,7. The combination of fluorescent dyes and expression of fluorescently labeled proteins can be used to monitor cell biological processes in mature myofibers.

<table><tbody><tr><td>DMEM</td><td>Life Technologies</td><td>11995-073</td><td>Dissection Media&lt;br/&gt; Dilution: 50 ml +100 g BSA</td></tr><tr><td>Collagenase, Type II</td><td>Life Technologies</td><td>17101-015</td><td>Fiber Digestion&lt;br/&gt; Dilution: 160 mg / 4 ml DMEM (stock 40 mg/ml aliquot)</td></tr><tr><td>Falcon 12-well dishes</td><td>Fisher Scientific</td><td>877229</td><td>Fiber Digestion</td></tr><tr><td>Ringers Solution</td><td></td><td></td><td>Fiber Digestion and Imaging&lt;br/&gt; Dilution: 146 mM NaCl&lt;br/&gt; 5 mM&amp;nbsp;&amp;nbsp; KCl&lt;br/&gt; 2 mM&amp;nbsp;&amp;nbsp; CaCl2&lt;br/&gt; 1 mM&amp;nbsp;&amp;nbsp; MCl2&lt;br/&gt; 10 mM HEPES&lt;br/&gt; pH7.4 in H2O</td></tr><tr><td>1 ml Pipettes</td><td>Axygen</td><td>T-1000-C-R</td><td>Digestion</td></tr><tr><td>Razor Blades</td><td>Personna</td><td>94-120-71</td><td>Digestion</td></tr><tr><td>Precision Glide 30 G needle</td><td>BD Biosciences</td><td>305128</td><td>Dissection</td></tr><tr><td>1x PBS (-/-)</td><td>Life Technologies</td><td>14190-250</td><td>Dissection</td></tr><tr><td>Sylgard 184</td><td>Fisher Scientific</td><td>50-366-794</td><td>Dissection</td></tr><tr><td>Forceps</td><td>FST by Dumont</td><td>#5/45</td><td>Dissection</td></tr><tr><td>Forceps</td><td>Roboz</td><td>RS-4913</td><td>Dissection</td></tr><tr><td>Scissors</td><td>World Precision Instruments</td><td>500260</td><td>Dissection</td></tr><tr><td>BSA</td><td>Sigma</td><td>A7906</td><td>Dissection media&lt;br/&gt; Dilution: 100 mg/50 ml DMEM</td></tr><tr><td>Falcon 60 mm dishes</td><td>Fisher Scientific</td><td>08772B</td><td>Dissection- used to hold sylgard</td></tr><tr><td>Clay</td><td></td><td></td><td>Electroporation</td></tr><tr><td>Stimulator&amp;nbsp;</td><td>Grass</td><td>s88x</td><td>Electroporation</td></tr><tr><td>Clamps</td><td>Radio Shack</td><td>270-356</td><td>Electroporation</td></tr><tr><td>Triadine</td><td>Aplicare</td><td>82-220</td><td>Electroporation</td></tr><tr><td>Precision Glide 27 G needle</td><td>BD Biosciences</td><td>305136</td><td>Electroporation</td></tr><tr><td>Simulator</td><td>Grass</td><td>SIU-V</td><td>Electroporation</td></tr><tr><td>Gauze</td><td>Covidien</td><td>2146</td><td>Electroporation</td></tr><tr><td>Hyaluronidase</td><td>Sigma</td><td>H4272</td><td>Injection&lt;br/&gt; Dilution: Add 160 ul PBS (1x) to 40 &amp;micro;l 5x stock</td></tr><tr><td>1 cc U-100 Insulin Syringe (28G1/2)&amp;nbsp;</td><td>BD Biosciences</td><td>329420</td><td>Injection</td></tr><tr><td>Eppendorf Tubes</td><td>Fisher Scientific</td><td>02-682-550</td><td>Injection</td></tr><tr><td>35 mm glass bottom Microwell dish</td><td>MaTek</td><td>P35G-1.5-14-C</td><td>Microscopy</td></tr><tr><td>FM 4-64</td><td>Life Technologies</td><td>T-13320</td><td>Microscopy&lt;br/&gt; Dilution: Final 2.5 ug/ml</td></tr><tr><td>FM 1-43</td><td>Life Technologies</td><td>T-35356</td><td>Microscopy&lt;br/&gt; Dilution: Final 2.5 ug/ml</td></tr><tr><td>Endotoxinfree Plasmid Maxi Kit</td><td>Qiagen</td><td>12362</td><td>Plasmid Purification</td></tr></tbody></table>

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.

Electroporation is a minimally invasive technique that efficiently introduces purified plasmid DNA into the flexor digitorum brevis (FDB) muscle bundle (Figure 1A). Seven days post transfection fluorescently tagged protein is visualized in isolated myofibers (Figure 1B). Isolated fibers are then plated and prepared for imaging on a confocal microscope. Fibers can be subjected to laser induced injury. FM dye can be used to identify the site of membrane dam...

Watch this Scientific Journal Video about DNA Electroporation, Isolation and Imaging of Myofibers at JoVE.com

DNA Electroporation, Isolation and Imaging of Myofibers

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 ...

dna-electroporation-isolation-and-imaging-of-myofibers

Research

JoVE Journal

Biology

12.2K Views.  Northwestern University. 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. 

Video: DNA Electroporation, Isolation and Imaging of Myofibers

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

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant variants) in order to investigate their endogenous distribution and functional relevance. An interesting approach that has been implemented to fulfill this objective in fully differentiated cells is the in vivo transfection of plasmids by various methods into specific tissues such as liver1, skeletal muscle2,3, and even the brain4. We present here a detailed description of the steps that must be followed in order to efficiently transfect genetic material into fibers of the flexor digitorum brevis (FDB) and interosseus (IO) muscles of adult mice using an in vivo electroporation approach. The experimental parameters have been optimized so as to maximize the number of muscle fibers transfected while minimizing tissue damages that may impair the quality and quantity of the proteins expressed in individual fibers. We have verified that the implementation of the methodology described in this paper results in a high yield of soluble proteins, i.e. EGFP and ECFP3, calpain, FKBP12, &beta;2a-DHPR, etc. ; structural proteins, i.e. minidystrophin and &alpha;-actinin; and membrane proteins, i.e. &alpha;1s-DHPR, RyR1, cardiac Na/Ca2+ exchanger , NaV1.4 Na channel, SERCA1, etc., when applied to FDB, IO and other muscles of mice and rats. The efficient expression of some of these proteins has been verified with biochemical3 and functional evidence5. However, by far the most common confirmatory approach used by us are standard fluorescent microscopy and 2-photon laser scanning microscopy (TPLSM), which permit to identify not only the overall expression, but also the detailed intracellular localization, of fluorescently tagged protein constructs. The method could be equally used to transfect plasmids encoding for the expression of proteins of physiological relevance (as shown here), or for interference RNA (siRNA) aiming to suppress the expression of normally expressed proteins (not tested by us yet). It should be noted that the transfection of FDB and IO muscle fibers is particularly relevant for the investigation of mammalian muscle physiology since fibers enzymatically dissociated from these muscles are currently one of the most suitable models to investigate basic mechanisms of excitability and excitation-contraction coupling under current or voltage clamp conditions2,6-8.

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

We describe detailed procedures for the efficient transfection of plasmid DNA into the fibers of foot muscles of live mice using electroporation and the subsequent visualization of protein expression using fluorescence microscopy.

 A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant ...

Minimally Invasive Muscle Embedding (MIME) - A Novel Experimental Technique to Facilitate Donor-Cell-Mediated Myogenesis

Skeletal muscle possesses regenerative capacity due to tissue-resident, muscle-fiber-generating (myogenic) satellite cells (SCs), which can form new muscle fibers under the right conditions. Although SCs can be harvested from muscle tissue and cultured in vitro, the resulting myoblast cells are not very effective in promoting myogenesis when transplanted into host muscle. Surgically exposing the host muscle and grafting segments of donor muscle tissue, or the isolated muscle fibers with their SCs onto host muscle, promotes better myogenesis compared to myoblast transplantation. We have developed a novel technique that we call Minimally Invasive Muscle Embedding (MIME). MIME involves passing a surgical needle through the host muscle, drawing a piece of donor muscle tissue through the needle track, and then leaving the donor tissue embedded in the host muscle so that it may act as a source of SCs for the host muscle. Here we describe in detail the steps involved in performing MIME in an immunodeficient mouse model that expresses a green fluorescent protein (GFP) in all of its cells. Immunodeficiency in the host mouse reduces the risk of immune rejection of the donor tissue, and GFP expression enables easy identification of the host muscle fibers (GFP+) and donor-cell-derived muscle fibers (GFP-). Our pilot data suggest that MIME can be used to implant an extensor digitorum longus (EDL) muscle from a donor mouse into the tibialis anterior (TA) muscle of a host mouse. Our data also suggest that when a myotoxin (barium chloride, BaCl2) is injected into the host muscle after MIME, there is evidence of donor-cell-derived myogenesis in the host muscle, with approximately 5%, 26%, 26% and 43% of the fibers in a single host TA muscle showing no host contribution, minimal host contribution, moderate host contribution, and maximal host contribution, respectively.

Minimally Invasive Muscle Embedding MIME - A Novel Experimental Technique to Facilitate Donor-Cell-Mediated Myogenesis

We describe a novel experimental technique that we call Minimally Invasive Muscle Embedding (MIME), which is based on the evidence that skeletal muscle tissue contains viable myogenic cells that can facilitate donor-cell-mediated myogenesis when implanted into a host muscle.

Skeletal muscle possesses regenerative capacity due to tissue-resident, muscle-fiber-generating (myogenic) satellite cells (SCs), which can form new ...

Medicine

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 ...

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 ...

Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear positioning occurs during skeletal muscle differentiation. Muscle fibers (myofibers) are multinucleated cells formed by the fusion of muscle precursor cells (myoblasts) derived from muscle stem cells (satellite cells) that undergo proliferation and differentiation. Correct nuclear positioning within myofibers is required for the proper muscle regeneration and function. The common procedure to assess myoblast differentiation and myofiber formation relies on fixed cells analyzed by immunofluorescence, which impedes the study of nuclear movement and cell behavior over time. Here, we describe a method for the analysis of myoblast differentiation and myofiber formation by live cell imaging. We provide a software for automated nuclear tracking to obtain a high-throughput quantitative characterization of nuclear dynamics and myoblast behavior (i.e., the trajectory) during differentiation and fusion.

Skeletal muscle differentiation is a highly dynamic process, which particularly relies on nuclear positioning. Here, we describe a method to track nuclei movements by live cell imaging during myoblast differentiation and myotube formation and to perform a quantitative characterization of nuclei dynamics by extracting information from automatic tracking.

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear ...

Isolating Myofibrils from Skeletal Muscle Biopsies and Determining Contractile Function with a Nano-Newton Resolution Force Transducer

Striated muscle cells are indispensable for the activity of humans and animals. Single muscle fibers are comprised of myofibrils, which consist of serially linked sarcomeres, the smallest contractile units in muscle. Sarcomeric dysfunction contributes to muscle weakness in patients with mutations in genes encoding for sarcomeric proteins. The study of myofibril mechanics allows for the assessment of actin-myosin interactions without potential confounding effects of damaged, adjacent myofibrils when measuring the contractility of single muscle fibers. Ultrastructural damage and misalignment of myofibrils might contribute to impaired contractility. If structural damage is present in the myofibrils, they likely break during the isolation procedure or during the experiment. Furthermore, studies in myofibrils provide the assessment of actin-myosin interactions in the presence of the geometrical constraints of the sarcomeres. For instance, measurements in myofibrils can elucidate whether myofibrillar dysfunction is the primary effect of a mutation in a sarcomeric protein. In addition, perfusion with calcium solutions or compounds is almost instant due to the small diameter of the myofibril. This makes myofibrils eminently suitable to measure the rates of activation and relaxation during force production. The protocol described in this paper employs an optical force probe based on the principle of a Fabry-P&#233;rot interferometer capable of measuring forces in the nano-Newton range, coupled to a piezo length motor and a fast-step perfusion system. This setup enables the study of myofibril mechanics with high resolution force measurements.

Presented here is a protocol to assess the contractile properties of striated muscle myofibrils with nano-Newton resolution. The protocol employs a setup with an interferometry-based, optical force probe. This setup generates data with a high signal-to-noise ratio and enables the assessment of the contractile kinetics of myofibrils.

Striated muscle cells are indispensable for the activity of humans and animals. Single muscle fibers are comprised of myofibrils, which consist of ...

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 ...

Enzymatic Isolation and Characterization of Variable Length Mouse Muscle Fibers

Intact Short, Intermediate, and Long Skeletal Muscle Fibers Obtained by Enzymatic Dissociation of Six Hindlimb Muscles of Mice: Beyond Flexor Digitorum Brevis

Skeletal muscle fibers obtained by enzymatic dissociation of mouse muscles are a useful model for physiological experiments. However, most papers deal with the short fibers of the flexor digitorum brevis (FDB), which restrains the scope of results dealing with fiber types, limits the amount of biological material available, and impedes a clear connection between cellular physiological phenomena and previous biochemical and dynamical knowledge obtained in other muscles.
This paper describes how to obtain intact fibers from six muscles with different fiber type profiles and lengths. Using C57BL/6 adult mice, we show the muscle dissection and fiber isolation protocol and demonstrate the suitability of the fibers for Ca2+ transient studies and their morphometric characterization. The fiber type composition of the muscles is also presented. When dissociated, all muscles rendered intact, living fibers that contract briskly for more than 24 h. FDB gave short (&#60;1 mm), peroneus digiti quarti (PDQA) and peroneus longus (PL) gave intermediate (1-3 mm), while extensor digitorum longus (EDL), extensor hallucis longus (EHL), and soleus muscles released long (3-6 mm) fibers.
When recorded with the fast dye Mag-Fluo-4, Ca2+ transients of PDQA, PL, and EHL fibers showed the fast, narrow kinetics reminiscent of the morphology type II (MT-II), known to correspond to type IIX and IIB fibers. This is consistent with the fact that these muscles have over 90% of type II fibers compared with FDB (~80%) and soleus (~65%). Moving beyond FDB, we demonstrate for the first time the dissociation of several muscles, which render fibers spanning a range of lengths between 1 and 6 mm. These fibers are viable and give fast Ca2+ transients, indicating that the MT-II can be generalized to IIX and IIB fast fibers, regardless of their muscle source. These results increase the availability of models for mature skeletal muscle studies.

Author Spotlight: Isolation of Long Muscle Fibers from Mouse Hindlimb Muscles for Studying Excitation-Contraction Coupling Across Fiber Types

We describe a protocol to obtain enzymatically dissociated fibers of different lengths and types from six muscles of adult mice: three of them already described (flexor digitorum brevis, extensor digitorum longus, soleus) and three of them successfully dissociated for the first time (extensor hallucis longus, peroneus longus, peroneus digiti quarti).

Skeletal muscle fibers obtained by enzymatic dissociation of mouse muscles are a useful model for physiological experiments. However, most papers deal ...

DNA Electroporation, Isolation and Imaging of Myofibers

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Chapters in this video

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

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

Minimally Invasive Muscle Embedding (MIME) - A Novel Experimental Technique to Facilitate Donor-Cell-Mediated Myogenesis

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

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

Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion

Isolating Myofibrils from Skeletal Muscle Biopsies and Determining Contractile Function with a Nano-Newton Resolution Force Transducer

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

Intact Short, Intermediate, and Long Skeletal Muscle Fibers Obtained by Enzymatic Dissociation of Six Hindlimb Muscles of Mice: Beyond Flexor Digitorum Brevis