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+A2:D32D42A2:D31AA2:D29</td>
			<td>Life Technologies</td>
			<td>11995-073</td>
			<td>Dissection Media 
			Dilution: 50ml +100g BSA</td>
		</tr>
		<tr>
			<td>Collagenase, Type II</td>
			<td>Life Technologies</td>
			<td>17101-015</td>
			<td>Fiber Digestion 
			Dilution: 160mg / 4ml DMEM (stock 40mg/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 
			Dilution: 146 mM NaCl 
			5 mM&#160;&#160; KCl 
			2 mM&#160;&#160; CaCl2 
			1 mM&#160;&#160; MCl2 
			10mM HEPES 
			pH7.4 in H2O</td>
		</tr>
		<tr>
			<td>1ml Pipettes</td>
			<td>Axygen</td>
			<td>T-1000-C-R</td>
			<td>Digestion</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>Razor Blades</td>
			<td>Personna</td>
			<td>94-120-71</td>
			<td>Digestion</td>
		</tr>
		<tr>
			<td>Precision Glide 30G needle</td>
			<td>BD Biosciences</td>
			<td>305128</td>
			<td>Dissection</td>
		</tr>
		<tr>
		</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 
			Dilution: 100mg / 50ml DMEM</td>
		</tr>
		<tr>
			<td>Falcon 60mm dishes</td>
			<td>Fisher Scientific</td>
			<td>08772B</td>
			<td>Dissection- used to hold sylgard</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>Clay</td>
			<td></td>
			<td></td>
			<td>Electroporation</td>
		</tr>
		<tr>
			<td>Stimulator&#160;</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>
		</tr>
		<tr>
			<td>Precision Glide 27G needle</td>
			<td>BD Biosciences</td>
			<td>305136</td>
			<td>Electroporation</td>
		</tr>
		<tr>
		</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 
			Dilution: Add 160ul PBS (1X) to 40&#181;l 5x stock</td>
		</tr>
		<tr>
			<td>1cc U-100 Insulin Syringe (28G1/2)&#160;</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>35mm glass bottom Microwell dish</td>
			<td>MaTek</td>
			<td>P35G-1.5-14-C</td>
			<td>Microscopy</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>FM 4-64</td>
			<td>Life Technologies</td>
			<td>T-13320</td>
			<td>Microscopy 
			Dilution: Final 2.5 ug/ml</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>FM 1-43</td>
			<td>Life Technologies</td>
			<td>T-35356</td>
			<td>Microscopy 
			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>
		<tr>
		</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

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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

Isolation of Genomic DNA from Mouse Tails

Electroporation of Mycobacteria

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly caused by the thick and waxy cell wall, but is compounded by the fact that most molecular techniques have been developed for distantly-related species such as Escherichia coli and Bacillus subtilis. In spite of these obstacles, mycobacterial plasmids have been identified and DNA transformation of many mycobacterial species have now been described. The most successful method for introducing DNA into mycobacteria is electroporation. Many parameters contribute to successful transformation; these include the species/strain, the nature of the transforming DNA, the selectable marker used, the growth medium, and the  conditions for the electroporation pulse. Optimized methods for the transformation of both slow- and fast-grower are detailed here. Transformation efficiencies for different mycobacterial species and with various selectable markers are reported.

Mycobacterial pathogenic strategies remain poorly understood. The slow growth rate of most species, the impenetrable nature of the cell-wall, and the hazards of working with pathogens make mycobacteria difficult to study and are largely responsible for our poor understanding of these organisms. In this video we will demonstrate the technique of electroporation, which involves subjecting cells to a brief high electrical impulse to allow the entry of DNA. It is the most widely used method for introducing DNA into mycobacterial cells.

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly ...

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 ...

Electroporation of Craniofacial Mesenchyme

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. For example, electroporation has been used with great success to study neural and retinal development in Xenopus, chicken and mouse 1-10. However, it is important to note that in all of these studies, investigators were not targeting soft tissues. Because we are interested in craniofacial development, we adapted a method to target facial mesenchyme.
When we searched the literature, we found, to our surprise, very few reports of successful gene transfer into cartilaginous tissue. The majority of these studies were gene therapy studies, such as siRNA or protein delivery into chondrogenic cell lines, or, animal models of arthritis 11-13. In other systems, such as chicken or mouse, electroporation of facial mesenchyme has been challenging (personal communications, Dept of Craniofacial Development, KCL). We hypothesized that electroporation into procartilaginous and cartilaginous tissues in Xenopus might work better. In our studies, we show that gene transfer into the facial cartilages occurs efficiently at early stages (28), when the facial primordium is still comprised of soft tissue prior to cartilage differentiation.
Xenopus is a very accessible vertebrate system for analysis of craniofacial development. Craniofacial structures are more readily visible in Xenopus than in any other vertebrate model, primarily because Xenopus embryos are fertilized externally, allowing analyses of the earliest stages, and facilitating live imaging at single cell resolution, as well as reuse of the mothers 14. Among vertebrate models developing externally, Xenopus is more useful for craniofacial analysis than zebrafish, as Xenopus larvae are larger and easier to dissect, and the developing facial region is more accessible to imaging than the equivalent region in fish. In addition, Xenopus is evolutionarily closer to humans than zebrafish (&tilde;100 million years closer) 15. Finally, at these stages, Xenopus tadpoles are transparent, and concurrent expression of fluorescent proteins or molecules will allow easy visualization of the developing cartilages. We anticipate that this approach will allow us to rapidly and efficiently test candidate molecules in an in vivo model system.

Craniofacial cartilages develop in close contact with other tissues and are difficult to manipulate in live animals. We are using electroporation to deliver molecular tools during growth of the craniofacial skeleton while bypassing early embryonic effects. This approach will allow us to efficiently test candidate molecules in vivo.

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

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 ...

Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method offers the opportunity of investigating interactions occurring in solution (thus avoiding problems due to closeby surfaces as in other single molecule methods), controlling the DNA extension and tracking interaction dynamics as a function of both mechanical parameters and DNA sequence. The methods for establishing successful optical trapping and nanometer localization of single molecules are illustrated. We illustrate the experimental conditions allowing the study of interaction of lactose repressor (lacI), labeled with Atto532, with a DNA molecule containing specific target sequences (operators) for LacI binding. The method allows the observation of specific interactions at the operators, as well as one-dimensional diffusion of the protein during the process of target search. The method is broadly applicable to the study of protein-DNA interactions but also to molecular motors, where control of the tension applied to the partner track polymer (for example actin or microtubules) is desirable.

Here we describe the instrumentation and methods for detecting single fluorescently-labeled protein molecules interacting with a single DNA molecule suspended between two optically trapped microspheres.

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

Reduced-gravity Environment Hardware Demonstrations of a Prototype Miniaturized Flow Cytometer and Companion Microfluidic Mixing Technology

Until recently, astronaut blood samples were collected in-flight, transported to earth on the Space Shuttle, and analyzed in terrestrial laboratories. If humans are to travel beyond low Earth orbit, a transition towards space-ready, point-of-care (POC) testing is required. Such testing needs to be comprehensive, easy to perform in a reduced-gravity environment, and unaffected by the stresses of launch and spaceflight. Countless POC devices have been developed to mimic laboratory scale counterparts, but most have narrow applications and few have demonstrable use in an in-flight, reduced-gravity environment. In fact, demonstrations of biomedical diagnostics in reduced gravity are limited altogether, making component choice and certain logistical challenges difficult to approach when seeking to test new technology. To help fill the void, we are presenting a modular method for the construction and operation of a prototype blood diagnostic device and its associated parabolic flight test rig that meet the standards for flight-testing onboard a parabolic flight, reduced-gravity aircraft. The method first focuses on rig assembly for in-flight, reduced-gravity testing of a flow cytometer and a companion microfluidic mixing chip. Components are adaptable to other designs and some custom components, such as a microvolume sample loader and the micromixer may be of particular interest. The method then shifts focus to flight preparation, by offering guidelines and suggestions to prepare for a successful flight test with regard to user training, development of a standard operating procedure (SOP), and other issues. Finally, in-flight experimental procedures specific to our demonstrations are described.

Spaceflight blood diagnostics need innovation. Few demonstrations have been published illustrating in-flight, reduced-gravity health diagnostic technology. Here we present a method for construction and operation of a parabolic flight test rig for a prototype point-of-care flow-cytometry design, with components and preparation strategies adaptable to other setups.

Until recently, astronaut blood samples were collected in-flight, transported to earth on the Space Shuttle, and analyzed in terrestrial laboratories. If ...