The authors declare no competing financial interests.

All animal procedures were approved by the Animal Welfare Department of the Th&#252;ringer Landesamt f&#252;r Lebensmittelsicherheit und Verbraucherschutz (03-048/16; TLV; Bad Langensalza, Germany).
1. Cardiotoxin-induced Muscle Injury
NOTE: Perform all experiments involving living mice in accordance with the national Animal Welfare Act and under appropriate aseptic conditions. Also, sacrificing of animals must be performed in accordance with the national Animal Welfare Act.
<ol>
	<li>Disinfect the inhalation box used for inhalation anesthesia and the surgery area with 70% ethanol. Place a sterile surgical cloth on the heating pad where the surgery will take place.</li>
	<li>Turn on the heating pad at 37 &#176;C.</li>
	<li>Administer analgesics (e.g., 1 mg/kg body weight meloxicam subcutaneously) 15 min prior to start of the surgery. 
	NOTE: For a typical experiment, use mice which are at least 6 weeks old, sex-matched and in a good general physical condition.</li>
	<li>Prepare the cardiotoxin solution (20 &#181;M in 0.9 % NaCl), store the solution at -20 &#176;C.</li>
	<li>Allow the cardiotoxin solution to reach room temperature (RT).</li>
	<li>Transfer the mouse into the inhalation box and induce anesthesia with isoflurane (initiation 3.5 &#8211; 5 % in pure oxygen). Check the depth of anesthesia with a lack of response to a toe pinch.</li>
	<li>Place the mouse on a clean paper towel and maintain anesthesia with a nose mask (1.5 &#8211; 3 % isoflurane in pure oxygen). Shave the injection area of the cranial lower leg (from knee to paw, Figure 1A). Remove all excessive loose hair. 
	NOTE: Physically separating the shaving and injection area in the room will provide more sterile conditions for the injections.</li>
	<li>Place the mouse in the lateral recumbent&#160;position on the heating pad covered with a sterile surgical cloth and maintain anesthesia with a nose mask (1.5 &#8211; 3% isoflurane in pure oxygen).</li>
	<li>Disinfect the injection area of the cranial lower leg (from knee to paw) with 70% ethanol.</li>
	<li>Perform a toe pinch before starting the intramuscular injection to ensure an adequate depth of anesthesia.</li>
	<li>Inject 50 &#181;L cardiotoxin (20 &#181;M in 0.9% NaCl) into the tibialis anterior muscle using an insulin syringe with a 29-gauge needle. First, pierce the skin just distal of the knee.</li>
	<li>Insert the needle fully into the muscle (in myofiber orientation/parallel to the tibia bone) and inject the cardiotoxin slowly (10-20 s) along the full length of the muscle while moving the needle back and forth to allow an even distribution of the cardiotoxin thereby injuring the whole tibialis anterior muscle (Figure 1B,C). 
	NOTE: Injure the tibialis anterior muscle from only one leg, the contralateral tibialis anterior muscle can serve as an internal control to determine that skeletal muscles were not pathologically affected before cardiotoxin injury.</li>
	<li>Transfer the mouse back to its cage placed on a heating pad and monitor the recovery process until the animal is conscious and becomes ambulatory. 
	NOTE: The mice will only show a slight limp. If mice are limping heavily and do not put weight on the leg at all, sacrifice the mouse.</li>
	<li>Administer analgesics during the following 2 days (e.g., 1 mg/kg body weight meloxicam subcutaneously, every 24 h) and monitor them on a weekly basis.</li>
</ol>
2. Injection of Self-delivering siRNA into the Regenerating Tibialis Anterior Muscle (At Day 3 Post Cardiotoxin Injury)
<ol>
	<li>Prepare the siRNA solution by resuspending the siRNA (e.g., siRNA smart pool) in 0.9% NaCl (final concentration of 2 &#181;g/&#181;L) according to the manufacturer&#8217;s instruction. 
	NOTE: The siRNA is chemically modified to facilitate the passive uptake by cells and protect it from nuclease degradation. No transfection reagents are needed, thereby reducing toxicity. Using a smart pool of four independent siRNA sequences targeting the same gene increases knockdown efficiency.</li>
	<li>Store the siRNA at -20 &#176;C and transfer it on ice to the surgery room.</li>
	<li>Disinfect the inhalation box used for inhalation anesthesia and the surgery area with 70% ethanol. Place a sterile surgical cloth on the heating pad where the surgery will take place.</li>
	<li>Turn on the heating pad at 37 &#176;C.</li>
	<li>Transfer the mouse into the inhalation box and induce anesthesia with isoflurane (initiation 3.5 &#8211; 5% in pure oxygen). Check the depth of anesthesia with a lack of response to a toe pinch.</li>
	<li>Place the mouse in the lateral recumbent&#160;position on the heating pad covered with a sterile surgical cloth and maintain anesthesia with a nose mask (1.5 &#8211; 3% isoflurane in pure oxygen).</li>
	<li>Disinfect the injection area of the cranial lower leg (from knee to paw).</li>
	<li>Perform a toe pinch before starting the intramuscular injection to ensure an adequate depth of anesthesia.</li>
	<li>Inject up to 50 &#181;L siRNA solution (up to 100 &#181;g total in 0.9% NaCl; directed against the target gene; use scrambled siRNA as control) into the injured tibialis anterior muscle using an insulin syringe with a 29 G needle. First, pierce the skin just distal of the knee, then insert the needle into the tibialis anterior muscle.</li>
	<li>Insert the needle fully into the muscle (in myofiber orientation/parallel to the tibia bone) and inject the siRNA solution slowly (10 &#8211; 20 s) along the full length of the muscle while moving the needle back and forth to allow an even distribution of the siRNA along the whole tibialis anterior muscle.</li>
	<li>Transfer the mouse back to its cage placed on a heating pad and monitor the recovery process until the animal is conscious and becomes ambulatory. 
	NOTE: Analgesia is not necessarily following the injection of siRNA.</li>
</ol>
3. Dissection of Tibialis Anterior Muscle
<ol>
	<li>Prepare the freezing solution (2 parts OCT compound and 1 part 30% sucrose in deionized water) at least 12 h before usage to avoid air bubbles.</li>
	<li>Prepare freezing molds by wrapping an aluminum foil around a pencil and seal it with tape. It is important that the bottom of the mold provides an even, closed surface. 
	NOTE: Smaller freezing molds allow the faster freezing of the muscle avoiding freezing artifacts.</li>
	<li>Sacrifice the mouse, e.g., by CO2 inhalation, at the respective time point of regeneration and spray the whole mouse and dissection tools with 70% ethanol. 
	NOTE: Cervical dislocation can be applied, in addition, to avoid excessive bleeding at the leg when isolating the tibialis anterior muscle.</li>
	<li>Remove the fur and skin from the injured hindlimb by cutting the skin at the ankle with extra fine sharp scissors (cutting edge: 13&#160;mm) and pulling up the skin towards the knee using forceps.</li>
	<li>To expose the tibialis anterior tendon at the ankle, pull the remaining skin towards the foot or cut with sharp scissors. 
	NOTE: The mouse can be pinned to a support board to allow better fixation.</li>
	<li>Before harvesting the tibialis anterior muscle, remove the fascia using fine forceps (Dumont 5 or 7, straight or curved). Pinch the closed forceps through the fascia next to the tibia bone at the ankle of the injured leg (Figure 2A). Move the forceps towards the knee thereby tearing the fascia and exposing the tibialis anterior muscle (Figure 2B).</li>
	<li>To isolate the tibialis anterior muscle, expose the distal tendon and grab it with fine forceps (Dumont 7, curved). Cut the tendon using spring scissors (cutting edge: 5&#160;mm, tip diameter: 0.35&#160;mm) and pull the muscle (holding it at the tendon) towards the knee.</li>
	<li>To harvest the tibialis anterior muscle, cut it as close to the knee as possible using sharp scissors.</li>
	<li>Prior to freezing, cut the tibialis anterior muscle at the mid belly region of the muscle with straight scissors into two halves of equal size to allow the analysis of the mid belly region (Figure 2C,D).</li>
	<li>Fill the freezing mold halfway with the freezing solution.</li>
	<li>Insert the two halves of the tibialis anterior muscle into the freezing mold with the mid belly region facing the bottom of the mold (Figure 2E). Make sure the tibialis anterior halves are inserted in an upright position leaning against the wall of the freezing mold to avoid tipping of the muscle. 
	NOTE: The more freezing solution is in the mold, the longer the freezing will take, thus increasing the risk of cryo-artifacts.</li>
	<li>Hold the freezing mold using forceps and transfer it halfway into liquid nitrogen (Figure 2F). Make sure that no liquid nitrogen is entering the freezing mold during the freezing process.</li>
	<li>Observe the freezing process, the freezing medium changes color from transparent to white and becomes solid. Submerse the freezing mold in liquid nitrogen for a few seconds and transfer the freezing mold to a -80 &#176;C freezer or to dry ice for future processing.</li>
	<li>Store the frozen muscles in the freezing molds at -80 &#176;C until further usage. 
	NOTE: The freezing molds fit well in 24-well plates or 1.5 mL reaction tubes allowing labeling and organized storage.</li>
	<li>Handle the muscle in the freezing molds for further usage always on dry ice.</li>
</ol>
4. Cryosectioning of Regenerating Tibialis Anterior Muscle
<ol>
	<li>Before starting sectioning, set the chamber temperature of the cryostat to -21 &#176;C, the object temperature to -20 &#176;C and the cutting thickness to 10, 12 or 14 &#181;m (depending on the future applications) (Figure 3A).</li>
	<li>Transfer the muscle sample in the freezing mold into the cryostat and let it adjust to the temperature for several minutes. During this time, label the microscope slides.</li>
	<li>Remove the foil around the embedded muscle using precooled forceps inside the cryostat. Avoid touching of the sample as the cryomedium starts to thaw easily.</li>
	<li>Mount the sample with the mid belly region of the tibialis anterior muscle directed upwards onto the metal sample holder of the cryostat using cryomedium. This ensures that the cutting site of the muscle (bottom of the mold) is facing the experimenter.</li>
	<li>Insert the sample holder with the mounted sample into the sectioning mechanism. 
	NOTE: To ensure proper sectioning of the sample, use a new blade before starting.</li>
	<li>First, trim the sample block (30 &#181;m) to get an even sectioning plane and to reach the respective region of the muscle (Figure 3B).</li>
	<li>After trimming, cut consecutive sections of the respective thickness (e.g., 14 &#181;m).</li>
	<li>Collect the sections on microscope glass slides by holding the slide face down over the section. The section is attaching to the slide (Figure 3C).</li>
	<li>Store sections at -80 &#176;C or -20 &#176;C until further use or directly use for immunofluorescence or immunohistochemistry.</li>
</ol>
5. Immunostaining for Markers of Regeneration
<ol>
	<li>Perform all further steps in a humidified chamber.</li>
	<li>Briefly, equilibrate the sections at room temperature.</li>
	<li>Fix sections with 1 mL of 2% PFA (in PBS, pH 7.4) per slide for 5 min at RT.</li>
	<li>Remove the 2% PFA solution by pouring it into the respective waste container.</li>
	<li>Wash 3 times with 1 mL of PBS (pH 7.4) for 5 min at RT.</li>
	<li>Remove the PBS by pouring it into a waste container.</li>
	<li>Per slide, add 1 mL of the permeabilization solution (0.1 % Triton X-100, 0.1 M Glycine in PBS pH 7.4) for 10 min.</li>
	<li>Remove the permeabilization solution by pouring it into a waste container.</li>
	<li>Add 150 &#181;L of the blocking solution (M.O.M. 1:40 in PBS pH 7.4) per slide and cover with the coverslip. Incubate for 1 h at RT.</li>
	<li>Remove the coverslip and the blocking solution, apply 100 &#181;L of primary antibody solution [PAX7, DSHB, mouse IgG1, undiluted or devMHC, DSHB, undiluted, mouse IgG1and laminin (rabbit, 1:1,000)] per slide. Cover with coverslip. Incubate O/N (overnight) at 4 &#176;C. 
	NOTE: Perform a control staining by omitting the primary antibodies, incubate the section with blocking solution instead.</li>
	<li>Wash 3 times with 1 mL of PBS (pH 7.4) for 5 min at RT.</li>
	<li>Remove PBS by pouring it into a waste container.</li>
	<li>Add 100 &#181;L of the secondary antibody solution (Alexa Fluor 546 goat anti-mouse IgG1 and Alexa Fluor 488 donkey anti-rabbit IgG in blocking solution, 1: 1,000) per slide and cover with the coverslip. Incubate for 1 h at RT.</li>
	<li>Incubate the slides in the dark, e.g., use an aluminum foil to cover or use a black humidified chamber. 
	NOTE: From now on all steps should be performed in reduced light conditions since some secondary antibodies are light sensitive.</li>
	<li>Remove the coverslip and the secondary antibody solution.</li>
	<li>Wash 3 times with 1 mL of PBS (pH 7.4) for 5 min at RT.</li>
	<li>Remove the PBS by pouring it into a waste container.</li>
	<li>Perform DAPI staining by adding 1 mL of the solution per slide to a final concentration of 10 &#181;g/mL for 5 min at RT.</li>
	<li>Remove the DAPI staining solution by pouring it into a waste container.</li>
	<li>Wash 3 times with 1 mL PBS (pH 7.4) for 5 min at RT.</li>
	<li>Completely remove the PBS used for washing.</li>
	<li>Apply 2 &#8211; 3 drops of aqueous mounting medium and directly cover the slide with a new coverslip.</li>
	<li>Let the slides dry for 1 h at RT in the dark.</li>
	<li>Store the stained sections at 4 &#176;C in the dark until further analysis using a fluorescence microscope.</li>
</ol>
6. Hematoxylin and Eosin Staining
<ol>
	<li>Perform all further steps in a humidified chamber.</li>
	<li>Fix sections with 1 mL of 2% PFA (in PBS, pH 7.4) per slide for 5 min at RT.</li>
	<li>Remove the 2% PFA solution by pouring it into the respective waste container.</li>
	<li>Transfer the slides into a Coplin jar.</li>
	<li>Gently rinse the slides with tap water. 
	NOTE: Do not turn on the faucet too high since that might damage the sections.</li>
	<li>Transfer the slides to the humidified chamber. Remove the liquid.</li>
	<li>Apply 1 mL of Hematoxylin staining solution (Gill No 3) for 2 min.</li>
	<li>Transfer the slides into a Coplin jar.</li>
	<li>Gently rinse the slides with the tap water until the nuclei turn blue. 
	NOTE: Do not turn on the faucet too high since that might damage your sections.</li>
	<li>Transfer the slides to the humidified chamber. Remove the liquid.</li>
	<li>Apply 1 mL of Eosin Y solution and incubate for 2 min.</li>
	<li>Transfer the slides into a Coplin jar.</li>
	<li>Gently rinse the slides with tap water until the tap water coming off the slides is clear.</li>
	<li>Remove all liquid and let the slides dry for 2 min.</li>
	<li>Mount in mounting medium suitable for immunohistochemistry.</li>
</ol>

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We would like to thank Christina Picker and Christine Poser for excellent technical assistance and Saskia Steiner for help with the determination of siRNA transfection efficiency. We thank the core service histology for great technical support, especially Sabine Landmann and Linda Rothenburger. We also thank the animal facility mouse at the FLI for excellent support. This work was supported by grants from the Deutsche Forschungsgemeinschaft (MA-3975/2-1) and the Deutsche Krebshilfe (DKH-JvM-861005) to J.v.M.

Here we present a method to investigate the function of a specific gene during regeneration of skeletal muscle without the need of transgenic animals. This is accomplished by the combination of cardiotoxin induced muscle injury with the injection of a self-delivering siRNA into the regenerating skeletal muscle at day 3 after injury. We have described in detail the procedures of muscle injury by cardiotoxin, the injection of the self-delivering siRNA and processing of the harvested muscles to analyze the progress of regeneration. We demonstrate that injection of the snake venom cardiotoxin into skeletal muscle effectively injures the whole muscle and that self-delivering siRNAs are found in around 75% of all satellite cells among other cell types two days after their injection into the regenerating skeletal muscle (Figure 4, Figure 5).
Particular attention should be focused on a homogeneous injury of the tibialis anterior muscle since varying degrees of injury influence the regeneration outcome and thereby also the effect of the siRNA might be affected. Furthermore, it is critically important that the whole regenerating area is injected with the self-delivering siRNA. For the analysis of the regeneration process, it is recommended to always compare similar areas of the regenerating skeletal muscle, therefore, the tibialis anterior muscle should be cut in half to always compare the mid-belly region of the muscle. When analyzing the muscle, the whole cryosection needs to be analyzed since myofiber composition differs in the tibialis anterior muscle and might therefore regenerate differently.
The function of satellite cells can be investigated by various experimental procedures including their culture on the adjacent isolated single myofibers17, by transplantation and by analyzing regeneration of skeletal muscle after induced injury18,19. Investigating the function of satellite cells by using an in vivo induced injury model, e.g., injection of cardiotoxin, provides the ability to analyze satellite cell function also in terms of their interaction with other cell types such as macrophages and investigation of the influence of systemic factors2. Injury of skeletal muscle can be accomplished by various means, e.g., eccentric exercise, freeze injury, injection of BaCl2 or injection of snake venoms such as cardiotoxin or notexin18. While eccentric exercise is probably the most physiological injury method, the injury to one particular muscle is only limited20. Freeze injuries can be applied when the migration of satellite cells towards the site of injury is the aim of the study or only a specific part of the muscle should be injured. Experimentally the disadvantage of freeze injuries is the open surgery which needs to be performed to apply the precooled metal probe. Injection of BaCl2 or snake venoms is the most dramatic method of injury, thereby challenging satellite cell function the most. Furthermore, the injection is minimally invasive, surgery time, in general, is less than five minutes and does not involve suturing, etc. thereby minimizing the risk of infections.
Muscle injury is mostly used to investigate the functional consequences of loss of gene functions, e.g., loss of Pax77,21. Especially if aged mice are the focus of the scientific question, the generation or use of transgenic mice is often not feasible. Injection of self-delivering siRNAs targeting a specific gene is a viable alternative in those cases and has been used successfully22. In brief, the tibialis anterior muscle of mice was injured by injection of cardiotoxin and self-delivering siRNAs directed against fibronectin (FN) were injected at day 3 after injury. Muscles were analyzed 10 days after injury and a significant decrease in satellite cell numbers was observed in siFN versus scrambled siRNA control conditions. The knockdown efficiency was determined in whole muscle lysates by quantitative Real Time PCR 2 days after siRNA injection, a reduction in expression levels of 58% was achieved suggesting that delivery and knockdown efficiency are sufficient for functional analyses22. Alternatives for testing the knockdown efficiency are either immunoblot or immunofluorescence analyses with antibodies directed against the target gene. The efficiency and specificity of the siRNA used for in vivo injections should be determined before injection into mice, e.g., by testing the efficiency in isolated satellite cells or primary myoblasts. The use of a smart pool consisting of 4 different siRNAs versus a single siRNA increases the efficiency of knockdown but also increases the risk of unspecific targeting. The specificity of all siRNA sequences used should be tested in cell culture to avoid off-target effects. As a control, a non-targeting scrambled siRNA should be used since the injection of a siRNA per se might influence the regeneration process due to the injection and, thereby, additional damage of the muscle. The time point of injecting the siRNA is depending on the scientific question and on the expression profile of the target gene. Generally, an injection of self-delivering siRNA at day 3 after cardiotoxin injury targets most genes important for satellite cell proliferation since satellite cell proliferation peaks around day 3 after injury. The time point for the first siRNA injection should not be less than 48 h after cardiotoxin injury since the injection volume of cardiotoxin is quite high and reabsorption of the liquid should have taken place before injecting additional solutions into the muscle. In general, multiple injections of siRNAs or a combination of different siRNAs is possible although one must consider that each injection into the regenerating muscle is causing additional damage.
One limitation of the described method is the fact, that the effect observed is not necessarily only depending on knockdown of the target gene in satellite cells but might be attributed to other cell types such as immune cells or fibro-adipogenic progenitor cells. Therefore, it is necessary to combine those experiments with experiments investigating a pure population of satellite cells. One can either perform experiments using floating myofiber cultures, where satellite cells are cultured on their adjacent myofibers or perform transplantation experiments using isolated satellite cells17.
An alternative to the injection of self-delivering siRNAs is the injection of small molecule inhibitors or recombinant proteins which can be performed, depending on the scientific question. For instance, the injection of the extracellular matrix protein fibronectin or small molecule inhibitors of JAK/STAT signaling have been successfully performed in aged mice15,16. The analysis of a particular gene function in one specific cell type during regeneration of skeletal muscle, e.g., in satellite cells, is only possible through the use of an inducible genetic mouse model. Injections of self-delivering siRNAs, recombinant proteins or small molecule inhibitors might affect multiple cell types in regenerating skeletal muscle.

Skeletal muscle is the body&#8217;s largest tissue representing approximately 40% of total body weight enabling voluntary locomotion. The tissue architecture of skeletal muscle is mainly composed of postmitotic, terminally differentiated, multinucleated myofibers as well as various other cells from the peripheral nervous system, vascular system, and interstitial cells1. Importantly, skeletal muscle has a tremendous capacity to regenerate and restore function upon injury or damage2. This process depends on the tissue resident muscle stem cells also called satellite cells3,4. Satellite cells are located between the myofiber and the basal lamina and characterized by the expression of the transcription factor Pax75,6,7. Under homeostatic conditions, satellite cells are quiescent but get activated due to traumatic injury, e.g., through eccentric exercise or experimentally through injection of the snake venom cardiotoxin6,8. Once activated, Pax7 positive satellite cells co-express MyoD and Myf5, which commits the stem cells to myogenic differentiation. Satellite cells that resist the upregulation of commitment factors will retain their stemness potential and return to quiescence to replenish the stem cell pool for future demands. After the expansion of the myogenic progenitor pool, transcriptional networks are activated by differentiation factors like Myogenin to initiate the cell cycle exit and terminal differentiation. These myoprogenitors then fuse to each other or to existing myofibers contributing myonuclei to maintain the size of the myonuclear domain. The myofibers express terminal muscle differentiation genes like Myosin Heavy Chain. Finally, the newly formed myofibers grow and mature to build the functional units of skeletal muscle9,10.
The regeneration of skeletal muscle can be affected by various conditions including muscle diseases or aging11,12, ranging from mild impairments to life threatening conditions, e.g., in Duchenne muscular dystrophy13,14. Therefore, regenerative medicine aims to restore damaged or malfunctioning skeletal muscle tissue using its inherent regenerative power by targeting satellite cell function15,16. To use its full potential a comprehensive understanding of satellite cells in their endogenous niche during regeneration of skeletal muscle is required. Although experimental approaches exist to isolate satellite cells adjacent to their myofibers17, the full complexity of cellular and systemic interactions of satellite cells with their environment can only be recapitulated in vivo. In that regard, great knowledge about skeletal muscle regeneration has been acquired using mouse injury models2,18.
Here we introduce a specific experimental mouse injury model to study the stem cell mediated regeneration of cardiotoxin-induced damage of the tibialis anterior muscle in vivo. Cardiotoxin, a snake derived cytolytic toxin that causes myofiber depolarization and necrosis, is injected into the tibialis anterior muscle, which in turn will trigger tissue degeneration followed by regeneration. To analyze the function of genes during acute regeneration, self-delivering siRNAs are injected at day 3 after injury, at the peak of the satellite cell expansion. Experimental animals are sacrificed at various timepoints and the tibialis anterior muscles are collected. The dissected muscles are frozen and processed for further cryo-sectioning. Immunofluorescence microscopy is then used to analyze markers of regeneration. This method allows for the investigation of the function of a single gene during satellite cell driven regeneration of skeletal muscle using wild type mice.

<table>
	<tbody>
		<tr>
			<td>isoflurane</td>
			<td>Henry Schein</td>
			<td>Isothesia</td>
			<td>inhalation narcotics</td>
		</tr>
		<tr>
			<td>Hot Plate 062</td>
			<td>Labotect</td>
			<td>13854</td>
			<td></td>
		</tr>
		<tr>
			<td>Anesthesia System (Tec 7) with inhalation box + nose masks</td>
			<td>Tem Sega</td>
			<td>Minihub</td>
			<td></td>
		</tr>
		<tr>
			<td>shaver for rodents</td>
			<td>isis</td>
			<td>GT420</td>
			<td></td>
		</tr>
		<tr>
			<td>cardiotoxin</td>
			<td>Latoxan</td>
			<td>L8102</td>
			<td>snake venom needed for muscle injury</td>
		</tr>
		<tr>
			<td>Accell siRNA smart pool</td>
			<td>Dharmacon</td>
			<td>depending on your target gene</td>
			<td>self delivering siRNA</td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 donkey anti-rabbit IgG</td>
			<td>Thermo Fisher</td>
			<td>A-21206</td>
			<td>secondary antibody</td>
		</tr>
		<tr>
			<td>Alexa Fluor 546 goat anti-mouse IgG1</td>
			<td>Thermo Fisher</td>
			<td>A-21123</td>
			<td>secondary antibody</td>
		</tr>
		<tr>
			<td>Coverslips</td>
			<td>VWR</td>
			<td>631-1574</td>
			<td></td>
		</tr>
		<tr>
			<td>CV Mount</td>
			<td>Leica</td>
			<td>14046430011</td>
			<td>mounting medium for immunohistochemistry</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma Aldrich</td>
			<td>D9542</td>
			<td>nuclear staining</td>
		</tr>
		<tr>
			<td>devMHC antibody</td>
			<td>DHSB</td>
			<td>F.1652</td>
			<td></td>
		</tr>
		<tr>
			<td>Eosin Y</td>
			<td>Thermo Fisher</td>
			<td>73104</td>
			<td></td>
		</tr>
		<tr>
			<td>Haematoxylin Gill No3</td>
			<td>Sigma-Aldrich</td>
			<td>GHS316-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>insulin syringe (29g)</td>
			<td>Terumo</td>
			<td>3SS05M2813</td>
			<td>syringue used for muscle injections</td>
		</tr>
		<tr>
			<td>laminin antibody</td>
			<td>Sigma Aldrich</td>
			<td>L9393</td>
			<td></td>
		</tr>
		<tr>
			<td>M.O.M blocking reagent</td>
			<td>Vector labs</td>
			<td>MKB-2213</td>
			<td>blocking for immunofluorescent staining</td>
		</tr>
		<tr>
			<td>Meloxicam</td>
			<td>Boehringer Ingelheim</td>
			<td>Metacam</td>
			<td>analgesics</td>
		</tr>
		<tr>
			<td>OCT</td>
			<td>Thermo Fisher</td>
			<td>6502</td>
			<td>tissue embedding</td>
		</tr>
		<tr>
			<td>Pax7 antibody</td>
			<td>DSHB</td>
			<td>PAX7</td>
			<td>satellite cell specific antibody</td>
		</tr>
		<tr>
			<td>ProLong Gold Antifade Mountant</td>
			<td>Thermo Fisher</td>
			<td>P36934</td>
			<td>aequos mounting medium</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Carl Roth</td>
			<td>4621.1</td>
			<td>tissue embedding</td>
		</tr>
		<tr>
			<td>Superfrost plus</td>
			<td>Thermo Scientific</td>
			<td>J1830AMNZ</td>
			<td>microscope slides</td>
		</tr>
		<tr>
			<td>TritonX-100</td>
			<td>Amresco</td>
			<td>0694-1L</td>
			<td>permeabilization reagent</td>
		</tr>
		<tr>
			<td colspan="4"></td>
		</tr>
		<tr>
			<td colspan="4">Dissection tools</td>
		</tr>
		<tr>
			<td>Dumont 5, straight</td>
			<td>Fine Science Tools</td>
			<td>11295-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont 7, curved</td>
			<td>Fine Science Tools</td>
			<td>11272-40</td>
			<td></td>
		</tr>
		<tr>
			<td>Extra fine Bonn scissors (cutting edge: 13&#160;mm)</td>
			<td>Fine Science Tools</td>
			<td>14084-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Narrow pattern forceps</td>
			<td>Fine Science Tools</td>
			<td>11002-16</td>
			<td></td>
		</tr>
		<tr>
			<td>Spring scissors (cutting edge: 5&#160;mm, tip diameter: 0.35&#160;mm)</td>
			<td>Fine Science Tools</td>
			<td>91500-09</td>
			<td></td>
		</tr>
	</tbody>
</table>

analyzing satellite cell function during skeletal muscle regeneration by cardiotoxin injury and injection of self-delivering sirna in vivo

Skeletal muscle possesses an enormous capacity to regenerate after injury. This process is mainly driven by muscle stem cells, also termed satellite cells. Satellite cells are characterized by the expression of the transcription factor Pax7 and their location underneath the basal lamina in the resting skeletal muscle. Upon injury, satellite cells get activated, undergo self-renewal or differentiation to either form new myofibers or to fuse with damaged ones. The functionality of satellite cells in vivo can be investigated using a cardiotoxin based injury model of skeletal muscle. To study the function of one gene during the regeneration of skeletal muscle, transgenic mouse models are mostly used. Here, we present an alternative method to transgenic mice, to investigate the gene function in satellite cells during regeneration, e.g., in cases where transgenic mice are not available. We combine the cardiotoxin mediated injury of a specific skeletal muscle with the injection of a self-delivering siRNA into the regenerating muscle which is then taken up by satellite cells among other cells. Thereby, we provide a method to analyze gene function in satellite cells during regeneration under physiological conditions without the need for transgenic mice.

A typical result of a hematoxylin and eosin (H&#38;E) staining of a tibialis anterior muscle before and after cardiotoxin mediated injury is presented in Figure 4A,B. In the control muscles, the architecture of the muscle is intact as seen by the localization of the nuclei in the periphery of the myofibers and the lack of accumulation of mononucleated cells in the interstitial space (Figure 4A). 7 days after cardiotoxin mediated injury new myofibers are formed marked by centrally located nuclei (Figure 4B). Furthermore, an accumulation of mononucleated cells can be observed, consisting mostly of satellite cells but also non-myogenic cells like immune cells. The whole muscle should be injured characterized by the central location of all myonuclei and the accumulation of mononucleated cells on the whole muscle section.
To characterize the progression and success of the regeneration process, several immunofluorescent staining using markers of regeneration can be performed. The number of satellite cells can be evaluated by staining for Pax7, the canonical marker for satellite cells (Figure 4C,D). Three days after injury the number of satellite cells increases (Figure 4D), satellite cells are not located under the basal lamina anymore. To further analyze the regeneration process, newly formed myofibers can be stained with antibodies directed to developmental myosin (Figure 4E). Newly formed myofibers display centrally located nuclei and a strong expression of developmental myosin. As the myofibers mature, expression of developmental myosin decreases, the diameter of the myofiber increases while nuclei are migrating to the periphery of the myofibers.
The influence of a single gene on the process of regeneration can be investigated using transgenic mice or as shown here by injection of a self-delivering siRNA. Fluorescently labeled self-delivering scrambled control siRNA was injected into the regenerating tibialis anterior muscle at day 3 after injury, the time point when the proliferation of satellite cells peaks. Two days after injection of the siRNA satellite cells were analyzed for the presence of the fluorescently labeled siRNA (Figure 4F). We analyzed how many satellite cells had taken up the fluorescently labeled siRNA two days after injection into the regenerating muscle (Figure 5). About 75% of all satellite cells in the regenerating muscle were positive for the fluorescently labeled siRNA. Furthermore, we determined that about 74% of all regenerating myofibers were positive for the fluorescently labeled siRNA suggesting that either 74% of the regenerating myofibers had taken up the siRNA or that siRNA-positive satellite cells either had fused with each other to form new myofibers or that fusion with already existing regenerating myofibers occurred (Figure 5). This suggests that satellite cells take up the self-delivering siRNA and that the siRNA persists in the satellite cells for at least two days.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60194/60194fig01.jpg" /> 
Figure 1: Injection of cardiotoxin into the tibialis anterior muscle.&#160;(A) Mice are anesthetized by inhalation of isoflurane and the lower limb is shaved. (B) A 29 G needle is injected into the tibialis anterior muscle (C) and moved along the tibia bone during injection of the cardiotoxin solution. <a href="https://www.jove.com/files/ftp_upload/60194/60194fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60194/60194fig02.jpg" /> 
Figure 2: Steps involved in dissection and freezing of the injured tibialis anterior muscle.&#160;(A) The hind limb muscles are exposed (B) and the fascia surrounding the tibialis anterior muscle is ripped to expose the muscle. (C) After removing the muscle, it is cut at the mid belly region into two halves of similar size using sharp straight scissors (D). (E) The halves of the dissected muscle are embedded in an aluminum foil mold filled with freezing solution and frozen in liquid nitrogen (F). <a href="https://www.jove.com/files/ftp_upload/60194/60194fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60194/60194fig03.jpg" /> 
Figure 3: Equipment and steps involved in cryosectioning of tibialis anterior muscles.&#160;(A) The chamber temperature of the cryostat is set to -21 &#176;C. (B) The tibialis anterior muscle in the freezing solution is mounted on the sample holder. (C) Sections of 14 &#181;m thickness are attached to glass microscope slides. <a href="https://www.jove.com/files/ftp_upload/60194/60194fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60194/60194fig04.jpg" /> 
Figure 4: Representative images of regenerating tibialis anterior muscles.&#160;H&#38;E staining of tibialis anterior muscles before (A) and 7 days after cardiotoxin injury (B). (C) In uninjured muscle Pax7 positive (in red) satellite cells are located underneath the basal lamina marked by laminin (in green). Nuclei are counterstained with DAPI (in blue). Pax7 positive cells are marked by an arrow. (D) After 10 days of regeneration, myofibers are characterized by centrally located nuclei (in blue), Pax7 is shown in red, laminin in green. Pax7 positive cells are marked by an arrow. (E) Newly formed myofibers express developmental myosin (in red), laminin in green, nuclei are counterstained with DAPI (in blue). (F) Fluorescently labeled self-delivering siRNA (siRED, depicted in red) is still found in satellite cells (Pax7 positive, in green, marked by an arrow in the inset) 2 days after injection of the self-delivering siRNA into regenerating tibialis anterior muscle (injection at day 3 after cardiotoxin mediated injury). Laminin is shown in white, nuclei are counterstained with DAPI (in blue). Scale bar = 100 &#181;m (A and B),&#160;50 &#181;m (C-E), 25 &#181;m (F). <a href="https://www.jove.com/files/ftp_upload/60194/60194fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60194/60194fig05.jpg" /> 
Figure 5: Quantification of uptake of fluorescently labeled siRNA.&#160;(A) Quantification of siRNA uptake by satellite cells (Pax7+ cells) and regenerating myofibers 2 days after injection into regenerating skeletal muscle. Injection was performed at day 3 after cardiotoxin mediated injury. n = 3, error bars show SEM. (B) Quantification underlying the graph depicted in (A). <a href="https://www.jove.com/files/ftp_upload/60194/60194fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Analyzing Satellite Cell Function During Skeletal Muscle Regeneration by Cardiotoxin Injury and Injection of Self-delivering siRNA In Vivo at JoVE.com

Analyzing Satellite Cell Function During Skeletal Muscle Regeneration by Cardiotoxin Injury and Injection of Self-delivering siRNA In Vivo

We describe an in vivo method to investigate the functional loss of a specific gene in satellite cells by using a combination of cardiotoxin mediated injury of skeletal muscle and the injection of a self-delivering siRNA.

Skeletal muscle possesses an enormous capacity to regenerate after injury. This process is mainly driven by muscle stem cells, also termed satellite ...

analyzing-satellite-cell-function-during-skeletal-muscle-regeneration

Research

JoVE Journal

Genetics

14.7K Views.  Leibniz-Institute on Aging - Fritz-Lipmann-Institute. We describe an in vivo method to investigate the functional loss of a specific gene in satellite cells by using a combination of cardiotoxin mediated injury of skeletal muscle and the injection of a self-delivering siRNA.

Video: Analyzing Satellite Cell Function During Skeletal Muscle Regeneration by Cardiotoxin Injury and Injection of Self-delivering siRNA In Vivo

siRNA Transfection and EMSA Analyses on Freshly Isolated Human Villous Cytotrophoblasts

Human primary villous cytotrophoblasts are a very useful source of primary cells to study placental functions and regulatory mechanisms, and to comprehend diseases related to pregnancy. In this protocol, human primary villous cytotrophoblasts freshly isolated from placentas through a standard DNase/trypsin protocol are microporated with small interfering RNA (siRNA). This approach provided greater efficiency for siRNA transfection when compared to a lipofection-based method. Transfected cells can subsequently be analyzed by standard Western blot within a time frame of 3-4 days post-transfection. In addition, using cultured primary villous cytotrophoblasts, Electrophoretic Mobility Shift Assay (EMSA) analysis was optimized and performed on extracts from days 1 to 4. The use of these cultured primary cells and the protocol described allow for an evaluation of the implication of specific genes and transcription factors in the process of villous cytotrophoblast differentiation into a syncytiotrophoblast-like cell layer. However, the limited time span allowable in culture precludes the use of methods requiring more time, such as generation of a stable cell population. Therefore testing of this cell population requires highly optimized gene transfer protocols.

This protocol describes a method for efficiently transfecting siRNA in freshly isolated human villous cytotrophoblasts using microporation and identifying DNA-protein complexes in these cells. Transfected cells can be monitored by Western blot and EMSA analyses during the 4-day culture time.

Human primary villous cytotrophoblasts are a very useful source of primary cells to study placental functions and regulatory mechanisms, and to comprehend ...

RNA Interference-based Investigation of the Function of Heat Shock Protein 27 during Corneal Epithelial Wound Healing

Small interfering RNA (siRNA) is among the most widely used RNA interference methods for the short-term silencing of protein-coding genes. siRNA is a synthetic RNA duplex created to specifically target a mRNA transcript to induce its degradation and it has been used to identify novel pathways in various cellular processes. Few reports exist regarding the role of phosphorylated heat shock protein 27 (HSP27) in corneal epithelial wound healing. Herein, cultured human corneal epithelial cells were divided into a scrambled control-siRNA transfected group and a HSP27-specific siRNA-transfected group. Scratch-induced directional wounding assays, and western blotting, and flow cytometry were then performed. We conclude that HSP27 has roles in corneal epithelial wound healing that may involve epithelial cell apoptosis and migration. Here, step-by-step descriptions of sample preparation and the study protocol are provided.

Herein, we present a protocol to use heat shock protein 27 (HSP27)-specific small interfering RNA to assess the function of HSP27 during corneal epithelial wound healing. RNA interference is the best method for effectively knocking-down gene expression to investigate protein function in various cell types.

Small interfering RNA (siRNA) is among the most widely used RNA interference methods for the short-term silencing of protein-coding genes. siRNA is a ...

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative without tools to study gene function. The development and sharing of detailed protocols for the establishment of new model systems in laboratories, and for tools to carry out functional studies, is thus crucial for leveraging the power of genomics. Coleoptera (beetles) are the largest clade of insects and occupy virtually all types of habitats on the planet. In addition to providing ideal models for fundamental research, studies of beetles can have impacts on pest control as they are often pests of households, agriculture, and food industries. Detailed protocols for rearing and maintenance of D. maculatus laboratory colonies and for carrying out dsRNA-mediated interference in D. maculatus are presented. Both embryonic and parental RNAi procedures-including apparatus set up, preparation, injection, and post-injection recovery-are described. Methods are also presented for analyzing embryonic phenotypes, including viability, patterning defects in hatched larvae, and cuticle preparations for unhatched larvae. These assays, together with in situ hybridization and immunostaining for molecular markers, make D. maculatus an accessible model system for basic and applied research. They further provide useful information for establishing procedures in other emerging insect model systems.

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Here, we present protocols for rearing an intermediate-germ beetle, Dermestes maculatus (D. maculatus) in the lab. We also share protocols for embryonic and parental RNAi and methods for analyzing embryonic phenotypes to study gene function in this species.

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative ...

Retroviral Scanning: Mapping MLV Integration Sites to Define Cell-specific Regulatory Regions

Moloney murine leukemia (MLV) virus-based retroviral vectors integrate predominantly in acetylated enhancers and promoters. For this reason, mLV integration sites can be used as functional markers of active regulatory elements. Here, we present a retroviral scanning tool, which allows the genome-wide identification of cell-specific enhancers and promoters. Briefly, the target cell population is transduced with an mLV-derived vector and genomic DNA is digested with a frequently cutting restriction enzyme. After ligation of genomic fragments with a compatible DNA linker, linker-mediated polymerase chain reaction (LM-PCR) allows the amplification of the virus-host genome junctions. Massive sequencing of the amplicons is used to define the mLV integration profile genome-wide. Finally, clusters of recurrent integrations are defined to identify cell-specific regulatory regions, responsible for the activation of cell-type specific transcriptional programs.
The retroviral scanning tool allows the genome-wide identification of cell-specific promoters and enhancers in prospectively isolated target cell populations. Notably, retroviral scanning represents an instrumental technique for the retrospective identification of rare populations (e.g. somatic stem cells) that lack robust markers for prospective isolation.

Here, we describe a protocol for genome-wide mapping of the integration sites of Moloney murine leukemia virus-based retroviral vectors in human cells.

Moloney murine leukemia (MLV) virus-based retroviral vectors integrate predominantly in acetylated enhancers and promoters. For this reason, mLV ...

High Fat Diet Feeding and High Throughput Triacylglyceride Assay in Drosophila Melanogaster

Heart disease is the number one cause of human death worldwide. Numerous studies have shown strong connections between obesity and cardiac malfunction in humans, but more tools and research efforts are needed to better elucidate the mechanisms involved. For over a century, the genetically highly tractable model of Drosophila has been instrumental in the discovery of key genes and molecular pathways that proved to be highly conserved across species. Many biological processes and disease mechanisms are functionally conserved in the fly, such as development (e.g., body plan, heart), cancer, and neurodegenerative disease. Recently, the study of obesity and secondary pathologies, such as heart disease in model organisms, has played a highly critical role in the identification of key regulators involved in metabolic syndrome in humans.
Here, we propose to use this model organism as an efficient tool to induce obesity, i.e., excessive fat accumulation, and develop an efficient protocol to monitor fat content in the form of TAGs accumulation. In addition to the highly conserved, but less complex genome, the fly also has a short lifespan for rapid experimentation, combined with cost-effectiveness. This paper provides a detailed protocol for High Fat Diet (HFD) feeding in Drosophila to induce obesity and a high throughput triacylglyceride (TAG) assay for measuring the associated increase in fat content, with the aim to be highly reproducible and efficient for large-scale genetic or chemical screening. These protocols offer new opportunities to efficiently investigate regulatory mechanisms involved in obesity, as well as provide a standardized platform for drug discovery research for rapid testing of the effect of drug candidates on the development or prevention of obesity, diabetes and related metabolic diseases.

High Fat Diet Feeding and High Throughput Triacylglyceride Assay in Drosophila Melanogaster

This is a high fat diet feeding protocol to induce obesity in Drosophila, a model for understanding fundamental molecular mechanisms implicated in lipotoxicity. It also provides a high throughput triacylglyceride assay for measuring fat accumulation in Drosophila and potentially other (insect) models under various dietary, environmental, genetic or physiological conditions.

Heart disease is the number one cause of human death worldwide. Numerous studies have shown strong connections between obesity and cardiac malfunction in ...

Constitutive and Inducible Systems for Genetic In Vivo Modification of Mouse Hepatocytes Using Hydrodynamic Tail Vein Injection

In research models of liver cancer, regeneration, inflammation, and fibrosis, flexible systems for in vivo gene expression and silencing are highly useful. Hydrodynamic tail vein injection of transposon-based constructs is an efficient method for genetic manipulation of hepatocytes in adult mice. In addition to constitutive transgene expression, this system can be used for more advanced applications, such as shRNA-mediated gene knock-down, implication of the CRISPR/Cas9 system to induce gene mutations, or inducible systems. Here, the combination of constitutive CreER expression together with inducible expression of a transgene or miR-shRNA of choice is presented as an example of this technique. We cover the multi-step procedure starting from the preparation of sleeping beauty-transposon constructs, to the injection and treatment of mice, and the preparation of liver tissue for analysis by immunostaining. The system presented is a reliable and efficient approach to achieve complex genetic manipulations in hepatocytes. It is specifically useful in combination with Cre/loxP-based mouse strains and can be applied to a variety of models in the research of liver disease.

Constitutive and Inducible Systems for Genetic In Vivo Modification of Mouse Hepatocytes Using Hydrodynamic Tail Vein Injection

Hydrodynamic tail vein injection of transposon-based integration vectors enables stable transfection of murine hepatocytes in vivo. Here, we present a practical protocol for transfection systems that enables the long-term constitutive expression of a single transgene or combined constitutive and doxycycline-inducible expression of a transgene or miR-shRNA in the liver.

In research models of liver cancer, regeneration, inflammation, and fibrosis, flexible systems for in vivo gene expression and silencing are highly ...

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA mutation rates. Changes in replication timing occur during development and in cancer, but the role replication timing plays in development and disease is not known. Zebrafish were recently established as an in vivo model system to study replication timing. Here is detailed the protocols for using the zebrafish to determine DNA replication timing. After sorting cells from embryos and adult zebrafish, high-resolution genome-wide DNA replication timing patterns can be constructed by determining changes in DNA copy number through analysis of next generation sequencing data. The zebrafish model system allows for evaluation of the replication timing changes that occur in vivo throughout development, and can also be used to assess changes in individual cell types, disease models, or mutant lines. These methods will enable studies investigating the mechanisms and determinants of replication timing establishment and maintenance during development, the role replication timing plays in mutations and tumorigenesis, and the effects of perturbing replication timing on development and disease.

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Zebrafish were recently used as an in vivo model system to study DNA replication timing during development. Here is detailed the protocols for using zebrafish embryos to profile replication timing. This protocol can be easily adapted to study replication timing in mutants, individual cell types, disease models, and other species.

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA ...

Using Mouse Oocytes to Assess Human Gene Function During Meiosis I

Embryonic aneuploidy is the major genetic cause of infertility in humans. Most of these events originate during female meiosis, and albeit positively correlated with maternal age, age alone is not always predictive of the risk of generating an aneuploid embryo. Therefore, gene variants might account for incorrect chromosome segregation during oogenesis. Given that access to human oocytes is limited for research purposes, a series of assays were developed to study human gene function during meiosis I using mouse oocytes. First, messenger RNA (mRNA) of the gene and gene variant of interest are microinjected into prophase I-arrested mouse oocytes. After allowing time for expression, oocytes are synchronously released into meiotic maturation to complete meiosis I. By tagging the mRNA with a sequence of a fluorescent reporter, such as green fluorescent protein (Gfp), the localization of the human protein can be assessed in addition to the phenotypic alterations. For example, gain or loss of function can be investigated by establishing experimental conditions that challenge the gene product to fix meiotic errors. Although this system is advantageous in investigating human protein function during oogenesis, adequate interpretation of results should be undertaken given that protein expression is not at endogenous levels and, unless controlled for (i.e. knocked out or down), murine homologs are also present in the system.

As the genetic variants associated with human disease begin to become uncovered, it is becoming increasingly important to develop systems with which to rapidly evaluate the biological significance of those identified variants. This protocol describes methods for evaluating human gene function during female meiosis I using mouse oocytes.

Embryonic aneuploidy is the major genetic cause of infertility in humans. Most of these events originate during female meiosis, and albeit positively ...

Dissection of Enhancer Function Using Multiplex CRISPR-based Enhancer Interference in Cell Lines

Multiple enhancers often regulate a given gene, yet for most genes, it remains unclear which enhancers are necessary for gene expression, and how these enhancers combine to produce a transcriptional response. As millions of enhancers have been identified, high-throughput tools are needed to determine enhancer function on a genome-wide scale. Current methods for studying enhancer function include making genetic deletions using nuclease-proficient Cas9, but it is difficult to study the combinatorial effects of multiple enhancers using this technique, as multiple successive clonal cell lines must be generated. Here, we present Enhancer-i, a CRISPR interference-based method that allows for functional interrogation of multiple enhancers simultaneously at their endogenous loci. Enhancer-i makes use of two repressive domains fused to nuclease-deficient Cas9, SID and KRAB, to achieve enhancer deactivation via histone deacetylation at targeted loci. This protocol utilizes transient transfection of guide RNAs to enable transient inactivation of targeted regions and is particularly effective at blocking inducible transcriptional responses to stimuli in tissue culture settings. Enhancer-i is highly specific both in its genomic targeting and its effects on global gene expression. Results obtained from this protocol help to understand whether an enhancer is contributing to gene expression, the magnitude of the contribution, and how the contribution is affected by other nearby enhancers.

This protocol describes the steps needed to design and perform multiplexed targeting of enhancers with the deactivating fusion protein SID4X-dCas9-KRAB, also known as enhancer interference (Enhancer-i). This protocol enables the identification of enhancers that regulate gene expression and facilitates the dissection of relationships between enhancers regulating a common target gene.

Multiple enhancers often regulate a given gene, yet for most genes, it remains unclear which enhancers are necessary for gene expression, and how these ...

Purification and Transplantation of Myogenic Progenitor Cell Derived Exosomes to Improve Cardiac Function in Duchenne Muscular Dystrophic Mice

Duchene Muscular Dystrophy (DMD) is an X-linked recessive genetic disease caused by a lack of functional dystrophin protein. The disease cannot be cured, and as the disease progresses, the patient develops symptoms of dilated cardiomyopathy, arrhythmia, and congestive heart failure. The DMDMDX mutant mice do not express dystrophin, and are commonly used as a mouse model of DMD. In our recent study, we observed that intramyocardial injection of wide type (WT)-myogenic progenitor cells-derived exosomes (MPC-Exo) transiently restored the expression of dystrophin in the myocardium of DMDMDX mutant mice, which was associated with a transient improvement in cardiac function suggesting that WT-MPC-Exo may provide an option to relieve the cardiac symptoms of DMD. This article describes the technique of MPC-Exo purification and transplantation into hearts of DMDMDX mutant mice.

Here, we present a protocol to transiently improve cardiac function in Duchenne muscular dystrophy mice by transplanting exosomes derived from normal myogenic progenitor cells.

Duchene Muscular Dystrophy (DMD) is an X-linked recessive genetic disease caused by a lack of functional dystrophin protein. The disease cannot be cured, ...

Analyzing Satellite Cell Function During Skeletal Muscle Regeneration by Cardiotoxin Injury and Injection of Self-delivering siRNA In Vivo

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Constitutive and Inducible Systems for Genetic In Vivo Modification of Mouse Hepatocytes Using Hydrodynamic Tail Vein Injection

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Using Mouse Oocytes to Assess Human Gene Function During Meiosis I

Dissection of Enhancer Function Using Multiplex CRISPR-based Enhancer Interference in Cell Lines

Purification and Transplantation of Myogenic Progenitor Cell Derived Exosomes to Improve Cardiac Function in Duchenne Muscular Dystrophic Mice