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.
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Demonbreun, A. R., McNally, E. M. DNA Electroporation, Isolation and Imaging of Myofibers. J. Vis. Exp. (106), e53551, doi:10.3791/53551 (2015).
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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.
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 "myoblast-like" 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 "twitch" 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.
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
- 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.
- Purify plasmids from E. coli using endotoxin free conditions per manufacturer's instructions. Dissolve the plasmid in sterile, endotoxin free TE buffer at 2 - 5 µg plasmid/µl.
- Dilute and aliquot the plasmid to a concentration of 20 µg of DNA in 10 µl of sterile endotoxin free TE (2 µg/µl) buffer per injection into a sterile microcentrifuge tube. Prepare one aliquot per injection.
- 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 µl of diluted hyaluronidase per injection into a sterile microcentrifuge tube.
- 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.
- 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.
- Inject the footpad of the mouse with 10 µl of the 1x hyaluronidase solution (2 mg/ml = 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.
- Repeat steps 1.6 and 1.7 on the contralateral foot.
- Disconnect the anesthesia and place the mouse back in the cage. Allow the mouse to fully recover from anesthesia.
- Wait 2 hr.
- Repeat the anesthetizing and foot sterilization procedures, steps 1.5 and 1.6.
- Inject the diluted plasmid DNA into the footpad in the same fashion as the hyaluronidase. Ensure that the maximum volume is 20 µl or less. For dual plasmid expression inject 20 µg of each plasmid with a maximum volume of 20 µl or less.
- Repeat the plasmid injection on the contralateral footpad or use the contralateral foot for control injections.
- 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.
- 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.
- Connect electrodes to an electrical stimulator.
- 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.
- Repeat the stimulation procedure on the contralateral foot if required.
- Remove needles. Wipe the footpad with povidone iodine scrub.
- 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.
2. Experimental Procedure for Isolation of the Flexor Digitorum Brevis (FDB) Fibers
- 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'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.
- Sacrifice the animal through CO2 or anesthetic gas inhalation, followed by cervical dislocation or other method to ensure death.
- Remove the hind feet above the ankle joint using a clean razor blade and submerge in 1x Ringers solution in the 35 mm Sylgard dish.
- 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.
- 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.
- Holding the skin at the base of the ankle with fine forceps, cut the skin across the heel.
- 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.
- 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.
- Place the entire muscle bundle in the well containing DMEM + BSA.
- Repeat on the contralateral foot.
- Once all FDB muscles are isolated, add 100 µl of collagenase to each well containing DMEM + BSA and muscle.
- Check the success of electroporation on an inverted fluorescent microscope prior to digestion. If done properly, upwards of 90% transfection efficiency can be achieved.
- Incubate the plate in a humidified, 37 °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.
- After 1 hr, carefully transfer the FDB bundle to the well containing only Ringers solution.
- 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.
- Plate the isolated fibers on the dish ~ 500 µl per plate.
- Allow the muscle to digest in the well for 15 min.
- 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.
- 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.
3. Experimental Procedure for Visualization of Laser-induced Membrane Damage using Confocal Microscopy
- Load fibers immediately prior to imaging with 300 µl of 2.5 µM FM 1-43 of FM 4-64 dye in Ringers solution. Generally, image fibers 15 min to 2 hr after isolation. However, fibers can be imaged up to 24 hr post isolation.
- Place the glass bottom dish on a confocal microscope equipped with a UV laser and a 60X / 63X objective.
- 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.
- 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.
- Place the ROI cross hair on the sarcolemma utilizing the FM 4-64 channel.
- 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.
- 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.
- Repeat steps 3.3 through 3.7 on up to 3 fibers per dish.
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 damage (Figure 1C)
During the digestion process, a small number of myofibers may be injured (white arrow) (Figure 2A). Damaged myofibers can be identified by membrane blebbing present at the sarcolemma (black arrow). The damaged fiber is not suitable for further experimentation and should not be used. Representative confocal images showing the expression of a fluorescent fusion protein within the isolated myofibers Figure 2B. Utilization of the CMV promoter drives optimal protein expression in myofibers. DIC imaging shows an optimal fiber. FM 4-64 is present at the membrane at low levels prior to damage (Figure 2C).
Representative image of a myofiber loaded with FM 4-64 imaged on a Leica SP 5 confocal microscope. The sarcolemma is centered in the field of view (white boxes) and ROI crosshair (white x) is aligned on the crisp, fluorescent edge of the sarcolemma in an area devoid of myonuclei (Figure 3, middle). Nuclei are avoided as they can influence FM dye analysis post imaging. Upon membrane damage, FM dye minimally enters normal myofibers (Figure 3, right). Muscle fibers defective in membrane repair will display increased levels of FM dye uptake upon laser-induced damage.
Figure 1. Schematic of in vivo Electroporation and Laser Damage Protocol. (A) Hyaluronidase is injected into the footpad of an anesthetized mouse. Plasmid DNA is then injected and voltage is applied. (B) Seven days post injection, the flexor digitorum brevis (FDB) bundle (white arrow middle panel) is isolated from the footpad leaving behind exposed tendons (black arrow). (C) Fibers are plated and laser-induced damage is performed on live myofibers. Left panel,scale 50 µm. Right panel, scale 5 µm. Please click here to view a larger version of this figure.
Figure 2. Isolated Myofibers Expressing Fluorescently Tagged Proteins. Seven days post electroporation, protein expression is detected throughout the myofiber. (A) The isolation procedure results in a small number of unusable myofibers with distinct membrane blebbing (black arrow) and membrane disruptions (white arrow). Scale 5 µm. (B) Representative image of a healthy myofiber. Evenly spaced sarcomeres are visualized in DIC imaging (left). A myofiber expressing annexin A6 tagged with GFP (center panel). There is minimal FM dye signal prior to damage (right panel). Scale 5 µm. Please click here to view a larger version of this figure.
Figure 3. Alignment of Laser Crosshairs on the FM-positive Sarcolemma. The region of interest on the sarcolemma is centered within the field of view. The sarcolemma is magnified (white dotted box) and ROI crosshairs aligned on the sharp FM-positive edge of the sarcolemma in an area devoid of nuclei (center panel). Upon damage, FM dye enters at the site of injury (right panel, white arrow). Scale 5 µm. Please click here to view a larger version of this figure.
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, trafficking, and fusion.
Using this protocol, up to 90% of myofibers within the FDB muscle bundle express plasmid, and this can be seen readily by fluorescence microscopy both before and after myofiber dissociation. There is typically a gradient of expression with higher expression observed nearer to the injection site. Because of the gradient of expression, the effect of varying protein expression can be monitored from a single experiment. As with all transfection methods, this approach is limited by issues related to overexpression. Notably, with high level expression, protein aggregation can occur. In our experience, the mCherry fluorescent tag demonstrates more aggregation than the enhanced green fluorescent protein (eGFP) tag reducing resolution and sharpness of protein domains (for example see Figure 4 in 7). Additionally, mTurquise2 and Venus fluorescent tags both demonstrate sufficient fluorescence expression. However, both proteins are less bright than eGFP.
This method can be used to image living myofibers under a variety of conditions including testing pharmaceutical agents and cell wounding. We used laser injury to disrupt the sarcolemma, but other wounding processes could be adapted to this method. To this end, formation of membrane blebs created by osmotic shock as well as membrane injury induced by rolling glass beads can also be assessed utilizing electroporated myofibers 11,12. For laser wounding, the type and power of the laser, as well as the objective will influence the length of time needed to create a membrane lesion 13,14. Furthermore, external buffers, specifically the concentration of calcium and FM dye within the buffer will influence the repair process 13,15. This methodology has been tested on both Nikon and Leica confocal imaging systems and both are capable of performing the technique. FM 4-64 is well suited for experimentation with green fluorescent protein 7. Others have used FM 1-43, another lipophilic dye that binds negatively charged phospholipids with an excitation peak at 470 nm and emission peak at 610 nm, which limits the application of this dye 13,15,16. Photobleaching and phototoxicity are both possible outcomes due to over-imaging. Determining the proper balance between exposure time, image frequency and number of channels imaged are parameters easily manipulated to reduce these effects.
There are no conflicts of interest.
This work was supported by National Institutes of Health grants NS047726, NS072027, and AR052646.
|DMEM||Life Technologies||11995-073||Dissection Media
Dilution: 50 ml +100 g BSA
|Collagenase, Type II||Life Technologies||17101-015||Fiber Digestion
Dilution: 160 mg / 4 ml DMEM (stock 40 mg/ml aliquot)
|Falcon 12-well dishes||Fisher Scientific||877229||Fiber Digestion|
|Ringers Solution||Fiber Digestion and Imaging
Dilution: 146 mM NaCl
5 mM KCl
2 mM CaCl2
1 mM MCl2
10 mM HEPES
pH7.4 in H2O
|1 ml Pipettes||Axygen||T-1000-C-R||Digestion|
|Precision Glide 30 G needle||BD Biosciences||305128||Dissection|
|1x PBS (-/-)||Life Technologies||14190-250||Dissection|
|Sylgard 184||Fisher Scientific||50-366-794||Dissection|
|Forceps||FST by Dumont||#5/45||Dissection|
|Scissors||World Precision Instruments||500260||Dissection|
Dilution: 100 mg/50 ml DMEM
|Falcon 60 mm dishes||Fisher Scientific||08772B||Dissection- used to hold sylgard|
|Precision Glide 27 G needle||BD Biosciences||305136||Electroporation|
Dilution: Add 160 ul PBS (1x) to 40 µl 5x stock
|1 cc U-100 Insulin Syringe (28G1/2)||BD Biosciences||329420||Injection|
|Eppendorf Tubes||Fisher Scientific||02-682-550||Injection|
|35 mm glass bottom Microwell dish||MaTek||P35G-1.5-14-C||Microscopy|
|FM 4-64||Life Technologies||T-13320||Microscopy
Dilution: Final 2.5 ug/ml
|FM 1-43||Life Technologies||T-35356||Microscopy
Dilution: Final 2.5 ug/ml
|Endotoxinfree Plasmid Maxi Kit||Qiagen||12362||Plasmid Purification|
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