Name: using cleavage under targets and tagmentation (cut&tag) assay in mouse myoblast research
Uploaded: 2024-03-01T13:50:00.000+00:00
Duration: 7 min 9 s
Description: Watch this Scientific Journal Video about Author Spotlight: Navigating Challenges and Innovations in Muscle Stem Cell Studies at JoVE.com

This work was supported by the Strategic Priority Research Program of the Chinese Academy of Science (XDA16020400 to PH); the National Natural Science Foundation of China (32170804 to PH).

The methods presented in this manuscript are all approved by the Institutional Animal Care and Use Committee of Guangzhou Laboratory. Mice used to generate this manuscript&#39;s representative results were housed and maintained in accordance with the guidelines of the Institutional Animal Care and Use Committee of Guangzhou Laboratory.
1. Myoblast isolation from mouse hindlimb muscles (Example of using 1 mouse)
<ol>
	<li>Dilute glacial acetic acid with ddH2O to 0.02 M and autoclave it.</li>
	<li>Dilute collagen (from rat tail) to 0.1 mg/mL with 0.02 M acetic acid.</li>
	<li>Add this collagen solution into culture dishes to coat the dish bottom in a 37&#160;&#176;C cell incubator for overnight.</li>
	<li>Before use, remove the collagen solution and wash the dish twice with 1x PBS. Collagen solution can be reused for 8-10 times.</li>
	<li>Sacrifice an 8-12-week-old mouse. (Old mice have poor yields of MuSCs and myoblasts.)</li>
	<li>Isolate and place all hindlimb muscles from the mouse into a 10-cm dish.</li>
	<li>Use 1x PBS (with penicillin/streptomycin) to rinse the muscles 4 times. After each rinse, always transfer the muscles into a new plate.</li>
	<li>After the final rinse, remove PBS and mince up the muscles with scissors.</li>
	<li>Add 6 mL of 1x PBS containing dispase II (1.1 U/mL) and collagenase II (830 U/mL) to digest the minced muscles, and the digestion should last for ~1.5 h.</li>
	<li>To perform digestion, transfer the minced muscles into a tube in a 37 &#176;C water bath or directly leave the minced muscles within the dish and place it in a 37 &#176;C CO2 cell incubator.</li>
	<li>If using a cell incubator to digest, rock the dish every 15-30 min to mix the digestion well. 
	NOTE: Dispase II should be first made as 11 U/mL stock with DMEM media (no serum), and collagenase II should be made as 10,000 U/mL stock with 1x PBS.</li>
	<li>When the digestion is complete, use a 10-mL plastic pipet to flush the digested tissues up and down several times to homogenize the digestion mixture thoroughly.</li>
	<li>Add 10 mL of MuSC/Myoblast growth media to slow down the digestion. Details on the preparation of MuSC/Myoblast growth media can be found in Section 2.</li>
	<li>Run the mixture through a 70-&#181;m strainer. Use 1x PBS to rinse the digestion plate and run this PBS through the strainer as well, in order to collect all the cells in the digestion plate.</li>
	<li>Centrifuge at 350 x g for 10 min and discard the supernatant with vacuum.</li>
	<li>Resuspend the cell pellet with 20 mL of 1x PBS and run the cells through a 40-&#181;m strainer.</li>
	<li>Centrifuge at 350 x g for 5 min and remove the supernatant with vacuum.</li>
	<li>Use 20 mL of MuSC/Myoblast growth media to resuspend the cells and seed the cells in a 10-cm normal plastic plate. MuSCs cannot attach to the plate bottom during this time and will stay floating mostly in the media. 
	NOTE: This is the first pre-plating on plastic plates.</li>
	<li>After 1.5 h, briefly rock the plate and transfer the liquid into two collagen-coated 10-cm dishes, each dish containing 10 mL of culture. Now, the MuSCs will attach and grow on the collagen-coated plates. Change the media after 2 days. Collagen-coated plates are prepared as in steps 1-4 of this section.</li>
	<li>After another day, passage the cells: Remove the media, wash the cells once with 1x PBS, and trypsinize the cells.</li>
	<li>Use MuSC/Myoblast growth media to flush the cells off the plates, move the cells onto a new plastic plate, and incubate in the cell incubator for 40 min. 
	NOTE: This is the second pre-plating on plastic plates.</li>
	<li>Pour the culture into a new plastic plate and incubate in the cell incubator for another 20 min. 
	NOTE: This is the third pre-plating on plastic plates.</li>
	<li>Then split the supernatant in the plate into 8 collagen-coated 10-cm dishes to subculture the cells. The passage ratio here is 2: 8 (1: 4). After three times of pre-plating described above, the myoblasts can reach over 90% pure. Therefore, after this point, there is no need to pre-plate when further passaging the myoblasts.</li>
</ol>
2. Preparation of MuSC/Myoblast growth media (Example of preparing from 5 mice)
<ol>
	<li>Sacrifice 5 young and healthy wild-type mice, isolate the spleens, and rinse the spleens once with 1x PBS (with penicillin/streptomycin).</li>
	<li>Use scissors to remove the dark parts/spots on the spleens, if any. Discard the spleen if most of it has turned black.</li>
	<li>Place all 5 spleens into a 40-&#181;m strainer on the top of a 50 mL tube.</li>
	<li>Open a sterile 1 mL syringe package, pull out the plunger, and discard the barrel.</li>
	<li>In the 40-&#181;m strainer, hold the rubber stopper and use the thumb rest of the plunger to grind and disassociate the spleen tissues into single cells. Be careful not to touch and contaminate the thumb rest of the plunger when tearing open the syringe package.</li>
	<li>While grinding the spleens, occasionally use 1x PBS to wash off the dissociated spleen cells into the 50 mL tube underneath the strainer.</li>
	<li>After all the spleen cells have been flushed into the 50 mL tube through the strainer, centrifuge the tube at 500 x g for 10 min.</li>
	<li>Use 1 mL of 1x Red blood cell lysis buffer to resuspend the pellet. Lyse the red blood cells for ~1 min. 
	NOTE: 10x Red blood cell lysis buffer contains 155 mM NH4Cl, 10 mM NaHCO3, and 1 mM EDTA disodium salt in ddH2O. Filter to sterilize, and do not autoclave. Dilute with ddH2O to 1x before use.</li>
	<li>Fill up the 50 mL tube with 1x PBS to cease the lysing process of red blood cells.</li>
	<li>Strain the cell suspension again with a 40-&#181;m strainer, and then centrifuge at 500 x g for 10 min.</li>
	<li>Resuspend the cells with 2 mL of RPMI-1640 full media (RPMI-1640 media with 10% FBS and penicillin/streptomycin) with very gentle pipetting.</li>
	<li>Add more RPMI-1640 full media into the tube and mix well. Distribute the cell suspension into ten T75 culture flasks. The final culture volume in each T75 culture flask should be 20 mL.</li>
	<li>Add 10 &#181;L of Concanavalin-A stock into each T75 culture flask and mix well. Concanavalin-A activates the proliferation of T-cells. 
	NOTE: Concanavalin-A stock is 4 mg/mL and prepared by dissolving Concanavalin-A in 1x PBS. The Concanavalin-A itself is used here, and do not confuse it with the Concanavalin-A beads used in CUT&#38;Tag assays later on.</li>
	<li>After 48 h, supplement each T75 flask with another 20 mL of RPMI-1640 full media so the total volume will add up to 40 mL.</li>
	<li>Another 24 h later, collect all the culture in each T75 flask and centrifuge at 600 x g for 3 min.</li>
	<li>The supernatant is T-cell conditional media and can be stored at -20 &#176;C to -80 &#176;C for up to 2 months. Discard the spleen cell pellets. Before use, thaw the T-cell conditional media and further sterilize the media using a 0.22 &#181;m filter.</li>
	<li>To make Ham&#39;s F10 full media, add 100 mL of FBS, 300 &#181;L of bFGF stock, and 6 mL of penicillin/streptomycin into 500 mL of Ham&#39;s F10 media. 
	NOTE: bFGF stock is 5 &#181;g/mL and prepared by dissolving bFGF in Ham&#39;s F10 media.</li>
	<li>Mix the Ham&#39;s F10 full media with filtered T-cell conditional media described above at 1:1 ratio to make MuSC/Myoblast growth media.</li>
</ol>
3. CUT&#38;Tag with myoblasts (Example of 3 CUT&#38;Tag reactions)
<ol>
	<li>Preparation of key reagents
	<ol>
		<li>Purify the pA/G-Tn5 transposase in-house from bacterial culture using published method14. This enzyme only reaches its functional form till it is mounted with library-making DNA adaptors. Most importantly, pA/G-Tn5 is commercially available from many sources.</li>
		<li>The oligos to make adaptors are shown in Table 1. Specifically, anneal Oligo-Rev with Oligo-A and Oligo-B, respectively, to make double-stranded Adaptor-A and Adaptor-B accordingly. The details for annealing to make the adaptors can be found elsewhere14.</li>
		<li>Incubate Adaptor-A and Adaptor-B with pA/G-Tn5 transposase at room temperature (RT) for 2 h. Then, this transposase will take its functional form. In the following sections of this manuscript, the functional form of pA/G-Tn5 will be termed pA/G-Tn5a. 
		NOTE: If the pA/G-Tn5 is purchased instead of in-house made, make sure to clarify whether the bought item is pA/G-Tn5 mounted with adaptors (in other words, ready to use) or it is just the pA/G-Tn5 enzyme itself. If it is just the enzyme, prepare the two adaptors described above and mount them onto the pA/G-Tn5 before finally using this enzyme for CUT&#38;Tag.</li>
		<li>For the recipes of the Binding buffer, Wash buffer, Dig-wash buffer, 300-Dig buffer, and Tagmentation buffer, readers can also refer to Kaya-Okur et al.1.</li>
		<li>Prepare the Binding buffer, which contains 20 mM HEPES pH 7.5, 10 mM KCl, 1 mM CaCl2,&#160;and 1 mM MnCl2.</li>
		<li>Prepare the Wash buffer, which contains 20 mM HEPES pH 7.5, 150 mM NaCl, 0.5 mM Spermidine, and 1x Protease inhibitor cocktail.</li>
		<li>Make digitonin as a concentrated stock in DMSO, and then prepare Dig-wash buffer by adding digitonin into Wash buffer to a concentration of 0.5 mg/mL.</li>
		<li>Prepare Antibody buffer by adding EDTA to a concentration of 2 mM and BSA to a concentration of 0.1%, into the Dig-wash buffer.</li>
		<li>Prepare 300-Dig buffer containing 20 mM HEPES pH 7.5, 300 mM NaCl, 0.5 mM Spermidine, 1x Protease inhibitor cocktail, and 0.1-0.5 mg/mL digitonin.</li>
		<li>Prepare the Tagmentation buffer by supplementing the 300-Dig buffer with 10 mM MgCl2. MgCl2 can activate the enzymatic activity of pA/G-Tn5a. 
		NOTE: Information for other materials is provided in the Table of Materials file.</li>
	</ol>
	</li>
	<li>Concanavalin-A bead activation
	<ol>
		<li>Mix 30 &#181;L of Concanavalin-A beads (10 &#181;L for each reaction) into 300 &#181;L of Binding buffer in a 1.5 mL centrifuge tube. Invert several times to mix well. Put the tube onto a magnetic rack. After the liquid is clear, remove the supernatant.</li>
		<li>Remove the tube from the magnetic rack, add 300 &#181;L of Binding buffer in the tube, mix well, and evenly split into 3 tubes of an 8-PCR tube stripe. Designate each tube with one CUT&#38;Tag reaction.</li>
		<li>Put the 8-PCR tube stripe on a magnetic rack and wait till the liquid is clear.</li>
		<li>Aspirate the supernatant and remove the tubes from the magnetic rack. Add 10 &#181;L of Binding buffer into each tube and mix well. Place the prepared beads on ice or at 4&#160;&#176;C till the cells are ready to proceed.</li>
	</ol>
	</li>
	<li>Binding cells onto Concanavalin-A beads
	<ol>
		<li>Trypsinize cultured mouse myoblasts off the plates, wash the cells once with RT 1x PBS and eventually resuspend the cells in Wash buffer at RT. Avoid over-trypsinization, as this can compromise the affinity of the cell membrane to Concanavalin-A beads.</li>
		<li>Adjust the cell density to around 7 x 105 cells/mL in Wash buffer such that 100 &#181;L of cell suspension contains roughly 70,000 cells, sufficient for one CUT&#38;Tag reaction.</li>
		<li>Pipet 100 &#181;L of the cell suspension into each tube of the 8-PCR tube stripe, which already contains 10 &#181;L of prepared Concanavalin-A beads. Mix well and incubate on a &#34;seesaw motion&#34; shaker for 10 min at RT.</li>
		<li>Put the 8-PCR tube stripe on a magnetic rack. Wait till it clears, and remove the supernatant.</li>
		<li>To assess whether the Concanavalin-A beads have efficiently sequestered the cells, check the supernatant under a microscope to see how many cells are left in the supernatant. After the Concanavalin-A bead binding step, very few or no cells should be observed in the supernatant.</li>
	</ol>
	</li>
	<li>Incubating the cells with a primary antibody
	<ol>
		<li>For each sample, dilute 1 &#181;L of H3K4me1 antibody into 50 &#181;L of Antibody buffer and add the diluted antibody into each sample tube. Mix the beads well with the diluted antibody by inverting the tubes several times. Put the samples on the &#34;seesaw motion&#34; shaker to incubate at 4&#160;&#176;C for overnight. 
		NOTE: As mentioned by Dr. Steven Henikoff1, it was not necessary to use an IgG negative control in all CUT&#38;Tag assays. Compared to ChIPs, CUT&#38;Tag assays come with much lower or no background signal. Therefore, using IgG to perform a &#34;negative control&#34; CUT&#38;Tag assay does not introduce any useful information. In contrast, using a CUT&#38;Tag-validated antibody to perform a positive control is important to assess whether the CUT&#38;Tag assay has worked.</li>
	</ol>
	</li>
	<li>Incubating the cells with a secondary antibody
	<ol>
		<li>The next day morning, put the samples on the magnetic rack and wait till the samples clear.</li>
		<li>Remove the supernatant, which is the primary antibody. There is no need to wash the beads; Directly add 50 &#181;L of diluted (1:100) secondary antibody into each sample tube. 
		NOTE: Secondary antibody is diluted with Dig-wash buffer at a 1:100 ratio. To allow more efficient pA/G-Tn5a recognition, secondary antibodies should not be conjugated with any molecules or groups such as HRP, biotin, or fluorophores.</li>
		<li>Mix well and incubate on the &#34;seesaw motion&#34; shaker at RT for 1 h.</li>
		<li>Put the samples back on the magnetic rack, wait till the samples clear, and discard the supernatant.</li>
		<li>Add 200 &#181;L of Dig-wash buffer into each tube and invert several times to wash off the excessive antibodies.</li>
		<li>Put it back on the magnetic rack, wait till it clears, and remove the supernatant. Repeat this washing twice more to completely remove excessive unbound antibodies.</li>
	</ol>
	</li>
	<li>Incubating the cells with pA/G-Tn5a
	<ol>
		<li>After the last wash, put the samples on the magnetic rack, wait till the samples clear, and remove all the liquid in the tube. For each sample tube, add 100 &#181;L of 300-Dig buffer containing pA/G-Tn5a at 0.04 &#181;M. Mix well and incubate on the seesaw motion shaker for 1 h at RT.</li>
		<li>Put on the magnetic rack. Wait till the samples clear and remove the supernatant.</li>
		<li>Add 200 &#181;L of 300-Dig buffer into each sample, mix well, and place on the shaker to wash the sample for 3 min at RT.</li>
		<li>Put on the magnetic rack. Wait till the samples clear and remove the supernatant. Repeat this washing twice more to remove excessive pA/G-Tn5a and lower the background.</li>
	</ol>
	</li>
	<li>Cleavage and tagmentation of the genomes
	<ol>
		<li>After the last wash, put the samples on the magnetic rack and thoroughly pipet the liquid off the tube. For each sample, add 50 &#181;L of Tagmentation buffer. Mix well and incubate in a PCR machine at 37&#160;&#176;C for 1 h. Do not use the lid-heating function of the PCR machine. 
		NOTE: Optionally, due to the material wearing during the following genomic DNA purification step, some spike-in DNAs can be added into the Tagmentation buffer so that the spike-in will show up in the final sequencing results and can be used to normalize the data. The method for preparing and using the spike-in DNA can be found elsewhere14.</li>
	</ol>
	</li>
	<li>Total genomic DNA purification
	<ol>
		<li>After tagmentation, add another 150 &#181;L of Tagmentation buffer for each sample. The total volume is now 200 &#181;L.</li>
		<li>Then, for each sample, add 10 &#181;L of 0.5 M EDTA, 3 &#181;L of 10% SDS, and 2.5 &#181;L of 20 mg/mL Proteinase K. Vortex to mix well and incubate at 55&#176;C for 2 h.</li>
		<li>Pipet each sample into 200 &#181;L of phenol/chloroform in a 1.5 mL centrifuge tube and vortex for 10 s. Centrifuge at 18,000 x g for 5 min. Transfer the top layer into a new 1.5 mL centrifuge tube.</li>
		<li>Add 200 &#181;L of chloroform into each tube and vortex for 10 s. Centrifuge at 18,000 x g for 5 min.</li>
		<li>Aspirate the top layer into a new 1.5 mL centrifuge tube. Then, for each sample, add 550 &#181;L of 100% ethanol to precipitate the DNA at -80&#160;&#176;C for 1 h. To better visualize the DNA pellet, add 1 &#181;L of 20 mg/mL glycogen.</li>
		<li>Centrifuge at max speed (&#62;18,000 x g) for 15 min to pellet the DNA.</li>
		<li>Carefully pour off the supernatant, wash the glycogen/DNA pellet with 75% ethanol once, and centrifuge again at max speed for 5 min.</li>
		<li>Remove the 75% ethanol and resuspend the glycogen/DNA pellet with 21 &#181;L of ddH2O.</li>
	</ol>
	</li>
	<li>Library preparation and sequencing
	<ol>
		<li>Set up the library-preparing PCR. To index the samples to be sequenced together, use a different N7XX Illumina primer for each sample and a universal N5XX Illumina primer. If indexing more than 12 samples, then different N5XX Illumina primers will be required as well. Illumina provides twelve N7XX primers and eight N5XX primers, so the total number of N7XX with N5XX combinations is 12 x 8 = 96. The PCR reaction setup is shown in Table 2.</li>
		<li>Roughly determine the PCR cycle number in library-preparing PCR by the cell number used in the CUT&#38;Tag assay. Normally, for a CUT&#38;Tag assay using 50,000-100,000 cells, 9-12 cycles are quite sufficient, and cell numbers between 10,000-50,000 might require up to 14 cycles. The PCR program is displayed in Table 3. 
		NOTE: For most CUT&#38;Tag assays, if the antibody has functioned properly, then a PCR cycle number between 9-14 should always generate a useful library for sequencing. A good library is normally 10-50 ng/&#181;L (determined by Qubit assay) in concentration and 20-30 &#181;L in volume. On the contrary, if the antibody did not work efficiently and specifically, then merely increasing the cycle number cannot save the experiments. Excessive cycle numbers do not benefit the results at all and instead introduce more PCR duplicates, which will later complicate the data analysis.</li>
		<li>Use some commercially available DNA purification/size-selection beads to purify the library DNA and select the DNA of desirable sizes.</li>
		<li>Dilute a small aliquot of the purified library with water and use it for qPCR with locus-specific primers to check enrichments as in ChIP-qPCR experiments. Before sequencing the library, this can help determine whether the CUT&#38;Tag has worked.</li>
		<li>Perform sequencing and data analysis as per the manufacturer&#39;s instructions. 
		NOTE: CUT&#38;Tag kits are commercially available from a handful of companies, and one example can be found in the&#160;Table of Materials file.</li>
	</ol>
	</li>
</ol>

Profiling Genomic Histone Marks in Mouse Myoblasts Using CUT&#38;Tag

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H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myogenic+transcriptional+activation+of+MyoD+mediated+by+replication-independent+histone+deposition.">Myogenic transcriptional activation of MyoD mediated by replication-independent histone deposition.</a> Proc Natl Acad Sci U S A. 108 (1), 85-90 (2011).</li><li>Kaya-Okur, H. S., Janssens, D. H., Henikoff, J. G., Ahmad, K., Henikoff, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+low-cost+chromatin+profiling+with+CUT&amp;Tag.">Efficient low-cost chromatin profiling with CUT&amp;Tag.</a> Nat Protoc. 15 (10), 3264-3283 (2020).</li></ol>

The authors have nothing to disclose.

The specific cell number required in a certain CUT&#38;Tag reaction completely relies on the enrichment of the histone marks/variants or chromatin-binding proteins that are to be tested. Normally for very enriched histone marks such as H3K4me1, H3K4me3, and H3K27ac etc., 25,000-50,000 myoblasts are quite sufficient for one CUT&#38;Tag reaction. However, some rare chromatin-binding proteins might require up to 250,000 cells. The cell number used in CUT&#38;Tag assays is critical, which, if not handled well normally causes...

CUT&#38;Tag assay was invented to compensate for some overt flaws of ChIPs1. The two major disadvantages of ChIPs are 1) the bias introduced when fragmenting chromatin and 2) the incompetence to work with low cell numbers. X-ChIP assays rely on either sonication or MNase digestion to get chromatin fragments, whereas N-ChIP mostly uses MNase digestion to get nucleosomes. Sonication shows a bias towards relaxed chromatin locations such as promoter regions2, and apparently, MNase digestion also works more efficiently on relaxed chromatin fibers. Moreover, some reported that MNase digestion also shows a DNA sequence-dependent bias3. Therefore, at the input preparation step of ChIP assays, it is impossible to get chromatin fragments from all kinds of genomic locations in a perfectly random manner. Moreover, ChIP assays normally generate higher background signals compared to CUT&#38;Tag and require over 10 folds more reads than CUT&#38;Tag to accentuate where the peaks are1,4,5. This explains why ChIP experiments have to start with tremendously more cells than CUT&#38;Tag. This is not a problem when studying cell lines as they can be repetitively passaged to achieve a very high cell number. However, the ChIP assay is definitely not a strong epigenetic tool to study rare or precious primary cell populations, although primary cells obviously hold more practical and medical implications.
While the long and complicated ChIP procedure discourages some researchers from learning or using this technique, people are more comfortable with easier assays such as immunocytochemistry (ICC) or immunofluorescence (IF). A CUT&#38;Tag assay essentially resembles the process of ICC and IF experiments but only takes place in a test tube. CUT&#38;Tag does not need fragmented chromatin to start with, and instead, the genome must be intact for antibody binding1. On the first day of a ChIP experiment, researchers normally spend up to 4 h preparing the fragmented chromatins from nuclei with sonication or MNase digestion before the short chromatin pieces can be mixed with antibody-beads4,5. In remarkable contrast, the first-day workload of a CUT&#38;Tag procedure is to just immobilize the cells to Concanavalin-A beads and then to add the primary antibody onto the cell-beads. This only requires ~40 min1.
It is worth mentioning that Cleavage Under Targets and Release Using Nuclease (CUT&#38;RUN) is an important alternative to CUT&#38;Tag. CUT&#38;RUN was established based on a similar working mechanism as CUT&#38;Tag. In CUT&#38;Tag, the antibodies guide the pA/G-Tn5 transposase to all the locations where the enzyme will each cut out a piece of chromatin and meanwhile tag it with library-making adaptors, while in CUT&#38;RUN, the role of pA/G-Tn5 is played by the pA/G-MNase which only performs the cutting part of the job6. Therefore, compared to CUT&#38;Tag, CUT&#38;RUN requires an additional step which is to glue the library-making adaptors onto the pA/G-MNase-fragmented DNA pieces7,8. Due to the high similarities between CUT&#38;Tag and CUT&#38;RUN, researchers familiar with CUT&#38;Tag will easily adapt themselves to performing CUT&#38;RUN proficiently. However, it should be noted that some minor differences exist between CUT&#38;Tag and CUT&#38;RUN. CUT&#38;RUN protocols normally use the physical concentration of salt (~150 mM) in the washing steps, while in CUT&#38;Tag, the 300-Dig wash buffer is high in salt. Therefore, CUT&#38;Tag is good at controlling the background when profiling histone marks/variants or transcription factors, as these proteins directly and strongly bind DNA1. CUT&#38;Tag may run into problems when profiling chromatin-associated factors that do not directly bind DNA and show weak affinity to the chromatin. High salt washing steps in CUT&#38;Tag may strip off chromatin-associated factors and cause no signals in the final output. Although there are successful cases where CUT&#38;Tag can be used to profile some non-histone/non-transcription factor proteins9, we still recommend CUT&#38;RUN over CUT&#38;Tag to profile weakly-bound chromatin-associated proteins.
After mammals reach adulthood, their skeletal muscle tissues still contain muscle stem cells. During muscle injury, these stem cells can be activated and undergo cell number expansion and differentiation to regenerate damaged muscle fibers10. These stem cells are known as Muscle stem/satellite cells (MuSCs). After MuSCs are isolated from animals or once activated by muscle injury, they start proliferating and become myoblasts.
To obtain MuSCs from mouse skeletal muscle digest, MuSC surface markers such as Vcam1 (Cd106), Cd34, and &#945;7-integrin (Itga7) are often used individually or in combinations to enrich MuSCs during fluorescence-activated cell sorting (FACS)11. It has been shown that Cd31-/Cd45-/Sca1-/Vcam1+ is probably the best marker combination to get &#62;95% pure MuSCs12. FACS can isolate pure MuSCs right after fresh muscles are digested. However, if the experimental design does not require pure MuSCs right at their isolation, pre-plating is more cost-effective than FACS to obtain &#62;90% pure myoblasts (MuSC progeny).
MuSCs freshly isolated from mice do not proliferate efficiently in Ham&#39;s F10 media supplemented with fetal bovine serum (FBS). To better expand the cell number of MuSCs and get sufficient myoblasts, bovine growth serum (BGS) should be used instead of FBS. However, if BGS is not available, Ham&#39;s F10 full media (containing about 20% FBS) can be mixed with an equal volume of T-cell conditional media to dramatically promote myoblast expansion13. Therefore, this protocol will also describe the preparation of T-cell media conditioned MuSC media.
Most importantly, this protocol provides a complete example of performing H3K4me1 CUT&#38;Tag assays on mouse myoblasts isolated from mouse hindlimb muscles. Please note that this protocol also applies to other cell types and histone marks and histone variants, and readers only need to optimize the cell numbers or antibody amounts for their cases based on the enrichment of the specific histone marks or variants they study.
In order to be used in CUT&#38;Tag or CUT&#38;RUN, the Tn5 or MNase needs to be fused with protein A or protein G to make pA-Tn5, pG-Tn5, pA-MNase, or pG-MNase. Apparently, both protein A and protein G can be fused onto these enzymes at the same time to generate pA/G-Tn5 or pA/G-MNase. Protein A and protein G show differential affinities to IgGs from different species. Therefore, fusing protein A and protein G altogether onto the enzyme can overcome this problem and make the enzyme compatible with antibodies from multiple species.

<table><tbody><tr><td>bFGF</td><td>R&amp;amp;D Systems</td><td>233-FB-025</td><td></td></tr><tr><td>Collagen</td><td>Corning</td><td>354236</td><td></td></tr><tr><td>Collagenase II</td><td>Worthington</td><td>LS004177</td><td></td></tr><tr><td>Concanavalin-A</td><td>Sigma-Aldrich</td><td>C5275</td><td></td></tr><tr><td>Concanavalin-A beads</td><td>Bangs Laboratories</td><td>BP531</td><td></td></tr><tr><td>Digitonin</td><td>Sigma-Aldrich</td><td>300410</td><td></td></tr><tr><td>Dispase II</td><td>Thermo Fisher Scientific</td><td>17105041</td><td></td></tr><tr><td>Fetal bovine serum</td><td>Hyclone</td><td>SH30396.03</td><td></td></tr><tr><td>H3K4me1 antibody&amp;nbsp;</td><td>abcam</td><td>ab8895</td><td></td></tr><tr><td>Ham&#039;s F10 media</td><td>Thermo Fisher Scientific</td><td>11550043</td><td></td></tr><tr><td>Hyperactive Universal CUT&amp;amp;Tag Assay Kit for Illumina&amp;nbsp;</td><td>Vazyme</td><td>TD903</td><td>This kit has been tested by us to function well</td></tr><tr><td>Magnetic rack for 1.5 mL EP tubes</td><td>Qualityard</td><td>QYM06</td><td></td></tr><tr><td>Magnetic rack for 8-PCR tube stripes</td><td>Anosun Magnetic</td><td>CLJ16/21-021</td><td></td></tr><tr><td>NEBNext High-Fidelity 2x PCR Master Mix</td><td>NEB</td><td>M0541L</td><td>For library-making PCR reaction</td></tr><tr><td>pA-Tn5</td><td>Vazyme</td><td>S603-01</td><td>Needs to be mounted with adaptors before use</td></tr><tr><td>Protease inhibitor cocktail</td><td>Sigma-Aldrich</td><td>5056489001</td><td></td></tr><tr><td>Proteinase K</td><td>Beyotime</td><td>ST535-100mg</td><td></td></tr><tr><td>RPMI-1640 media</td><td>Thermo Fisher Scientific</td><td>C11875500BT</td><td></td></tr><tr><td>Secondary antibody (Guinea Pig anti-rabbit IgG)</td><td>Antibodies-online</td><td>ABIN101961</td><td></td></tr><tr><td>Spermidine</td><td>Sigma-Aldrich</td><td>S2626</td><td></td></tr><tr><td>TruePrep Index Kit V2 for Illumina&amp;nbsp;</td><td>Vazyme</td><td>TD202</td><td>This kit provide Illumina N7XX and N5XX primers&amp;nbsp;</td></tr><tr><td>VAHTS DNA Clean Beads&amp;nbsp;</td><td>Vazyme</td><td>N411</td><td>Can substitute Ampure XP beads. Can purify CUT&amp;amp;Tag libraries and select DNA fragments by size</td></tr></tbody></table>

using cleavage under targets and tagmentation (cut&#38;tag) assay in mouse myoblast research

This protocol paper aims to provide the new researchers with the full details of using Cleavage Under Targets and Tagmentation (CUT&amp;Tag) to profile the genomic locations of chromatin binding factors, histone marks, and histone variants. CUT&amp;Tag protocols function very well with mouse myoblasts and freshly isolated muscle stem cells (MuSCs). They can easily be applied to many other cell types as long as the cells can be immobilized by Concanavalin-A beads. Compared to CUT&amp;Tag, chromatin immunoprecipitation (ChIP) assays are time-consuming experiments. ChIP assays require the pre-treatment of chromatin before the chromatic material can be used for immunoprecipitation. In cross-linking ChIP (X-ChIP), pre-treatment of chromatin involves cross-linking and sonication to fragment the chromatin. In the case of native ChIP (N-ChIP), the fragmented chromatins are normally achieved by Micrococcal nuclease (MNase) digestion. Both sonication and MNase digestion introduce some bias to the ChIP experiments. CUT&amp;Tag assays can be finished within fewer steps and require much fewer cells compared to ChIPs but provide more unbiased information on transcription factors or histone marks at various genomic locations. CUT&amp;Tag can function with as few as 5,000 cells. Due to its higher sensitivity and lower background signal than ChIPs, researchers can expect to obtain reliable peak data from merely several millions of reads after sequencing.

Before binding cells to Concanavalin-A beads, check the cell suspension under the microscope. Accordingly, after incubating the cells with Concanavalin-A beads, put the sample tubes on the magnetic rack, and the supernatant should be again observed using a microscope. This is to assess how efficiently the cells have been captured by Concanavalin-A beads. Wash buffer containing 7 x 105 cells/mL should look like Figure 1A under the microscope. In contrast, Figur...

Watch this Scientific Journal Video about Author Spotlight: Navigating Challenges and Innovations in Muscle Stem Cell Studies at JoVE.com

Author Spotlight: Navigating Challenges and Innovations in Muscle Stem Cell Studies

Researchers new to the epigenetic field will find CUT&amp;Tag a significantly easier alternative to ChIP assays. CUT&amp;Tag has tremendously benefited the epigenetic studies on rare and primary cell populations, generating high-quality data from very few cells. This protocol describes performing H3K4me1 CUT&amp;Tag assays on mouse myoblasts isolated from mouse hindlimb muscles.

Using Cleavage Under Targets and Tagmentation (CUT&#38;Tag) Assay in Mouse Myoblast Research

This protocol paper aims to provide the new researchers with the full details of using Cleavage Under Targets and Tagmentation (CUT&amp;Tag) to profile ...

using-cleavage-under-targets-tagmentation-cut-tag-assay-mouse

Research

JoVE Journal

Developmental Biology

558 Views.  Guangzhou Laboratory. Researchers new to the epigenetic field will find CUT&Tag a significantly easier alternative to ChIP assays. CUT&Tag has tremendously benefited the epigenetic studies on rare and primary cell populations, generating high-quality data from very few cells. This protocol describes performing H3K4me1 CUT&Tag assays on mouse myoblasts isolated from mouse hindlimb muscles.

Video: Author Spotlight: Navigating Challenges and Innovations in Muscle Stem Cell Studies

RiboTag Immunoprecipitation in the Germ Cells of the Male Mouse

Quantifying differences in mRNA abundance is a classic approach to understand the impact of a given gene mutation on cell physiology. However, characterizing differences in the translatome (the whole of translated mRNAs) provides an additional layer of information particularly useful when trying to understand the function of RNA regulating or binding proteins. A number of methods for accomplishing this have been developed, including ribosome profiling and polysome analysis. However, both methods carry significant technical challenges and cannot be applied to specific cell populations within a tissue unless combined with additional sorting methods. In contrast, the RiboTag method is a genetic-based, efficient, and technically straightforward alternative that allows the identification of ribosome associated RNAs from specific cell populations without added sorting steps, provided an applicable cell-specific Cre driver is available. This method consists of breeding to generate the genetic models, sample collection, immunoprecipitation, and downstream RNA analyses. Here, we outline this process in adult male mouse germ cells mutant for an RNA binding protein required for male fertility. Special attention is paid to considerations for breeding with a focus on efficient colony management and the generation of correct genetic backgrounds and immunoprecipitation in order to reduce background and optimize output. Discussion of troubleshooting options, additional confirmatory experiments, and potential downstream applications is also included. The presented genetic tools and molecular protocols represent a powerful way to describe the ribosome-associated RNAs of specific cell populations in complex tissues or in systems with aberrant mRNA storage and translation with the goal of informing on the molecular drivers of mutant phenotypes.

Here, we describe the immunoprecipitation of ribosomes and associated RNA from select populations of adult male mouse germ cells using the RiboTag system. Strategic breeding and careful immunoprecipitation result in clean, reproducible results that inform on the germ cell translatome and provide insight into the mechanisms of mutant phenotypes.

Quantifying differences in mRNA abundance is a classic approach to understand the impact of a given gene mutation on cell physiology. However, ...

Satellite Cell Isolation from Mouse Limb Muscles and Flow Cytometry Analysis

An Efficient Protocol for CUT&amp;RUN Analysis of FACS-Isolated Mouse Satellite Cells

Genome-wide analyses with small cell populations are a major constraint for studies, particularly in the stem cell field. This work describes an efficient protocol for the fluorescence-activated cell sorting (FACS) isolation of satellite cells from the limb muscle, a tissue with a high content of structural proteins. Dissected limb muscles from adult mice were mechanically disrupted by mincing in medium supplemented with dispase and type I collagenase. Upon digestion,&#160;the homogenate was filtered through cell strainers, and cells were suspended in FACS buffer. Viability was determined with fixable viability stain, and immunostained satellite cells were isolated by FACS. Cells were lysed with Triton X-100 and released nuclei were bound to concanavalin A magnetic beads. Nucleus/bead complexes were incubated with antibodies against the transcription factor or histone modifications of interest. After washes, nucleus/bead complexes were incubated with protein A-micrococcal nuclease, and chromatin cleavage was initiated with CaCl2. After DNA extraction, libraries were generated and sequenced, and the profiles for genome-wide transcription factor binding and covalent histone modifications were obtained by bioinformatic analysis. The peaks obtained for the various histone marks showed that the binding events were specific for satellite cells. Moreover, known motif analysis unveiled that the transcription factor was bound to chromatin via its cognate response element. This protocol is therefore adapted to study gene regulation in adult mice limb muscle satellite cells.

Author Spotlight: Cistrome Analysis in Mouse Muscle Stem Cells 

Presented here is an efficient protocol for the fluorescence-activated cell sorting (FACS) isolation of mouse limb muscle satellite cells adapted to the study of transcription regulation in muscle fibers by cleavage under targets and release using nuclease (CUT&amp;RUN).

Genome-wide analyses with small cell populations are a major constraint for studies, particularly in the stem cell field. This work describes an efficient ...

Modeling Myotonic Dystrophy 1 in C2C12 Myoblast Cells

Myotonic dystrophy 1 (DM1) is a common form of muscular dystrophy. Although several animal models have been established for DM1, myoblast cell models are still important because they offer an efficient cellular alternative for studying cellular and molecular events. Though C2C12 myoblast cells have been widely used to study myogenesis, resistance to gene transfection, or viral transduction, hinders research in C2C12 cells. Here, we describe an optimized protocol that includes daily maintenance, transfection and transduction procedures to introduce genes into C2C12 myoblasts and the induction of myocyte differentiation. Collectively, these procedures enable best transfection/transduction efficiencies, as well as consistent differentiation outcomes. The protocol described in establishing DM1 myoblast cell models would benefit the study of myotonic dystrophy, as well as other muscular diseases.

In this protocol, we present the procedures in establishing myotonic dystrophy 1 myoblast models, including optimized C2C12 cell maintenance, gene transfection/transduction, and myocyte differentiation.

Myotonic dystrophy 1 (DM1) is a common form of muscular dystrophy. Although several animal models have been established for DM1, myoblast cell models are ...

Medicine

Single Cell Transcriptional Profiling of Adult Mouse Cardiomyocytes

While numerous studies have examined gene expression changes from homogenates of heart tissue, this prevents studying the inherent stochastic variation between cells within a tissue. Isolation of pure cardiomyocyte populations through a collagenase perfusion of mouse hearts facilitates the generation of single cell microarrays for whole transcriptome gene expression, or qPCR of specific targets using nanofluidic arrays. We describe here a procedure to examine single cell gene expression profiles of cardiomyocytes isolated from the heart. This paradigm allows for the evaluation of metrics of interest which are not reliant on the mean (for example variance between cells within a tissue) which is not possible when using conventional whole tissue workflows for the evaluation of gene expression (Figure 1). We have achieved robust amplification of the single cell transcriptome yielding micrograms of double stranded cDNA that facilitates the use of microarrays on individual cells. In the procedure we describe the use of NimbleGen arrays which were selected for their ease of use and ability to customize their design. Alternatively, a reverse transcriptase - specific target amplification (RT-STA) reaction, allows for qPCR of hundreds of targets by nanofluidic PCR. Using either of these approaches, it is possible to examine the variability of expression between cells, as well as examining expression profiles of rare cell types from within a tissue. Overall, the single cell gene expression approach allows for the generation of data that can potentially identify idiosyncratic expression profiles that are typically averaged out when examining expression of millions of cells from typical homogenates generated from whole tissues.

Single cell expression profiling allows the detailed gene expression analysis of individual cells. We describe methods for the isolation of cardiomyocytes, and preparing the resulting lysates for either whole transcriptome microarray or qPCR of specific targets.

While numerous studies have examined gene expression changes from homogenates of heart tissue, this prevents studying the inherent stochastic variation ...

Biology

In Vitro and In Vivo Detection of Mitophagy in Human Cells, C. Elegans, and Mice

Mitochondria are the powerhouses of cells and produce cellular energy in the form of ATP. Mitochondrial dysfunction contributes to biological aging and a wide variety of disorders including metabolic diseases, premature aging syndromes, and neurodegenerative diseases such as Alzheimer's disease (AD) and Parkinson's disease (PD). Maintenance of mitochondrial health depends on mitochondrial biogenesis and the efficient clearance of dysfunctional mitochondria through mitophagy. Experimental methods to accurately detect autophagy/mitophagy, especially in animal models, have been challenging to develop. Recent progress towards the understanding of the molecular mechanisms of mitophagy has enabled the development of novel mitophagy detection techniques. Here, we introduce several versatile techniques to monitor mitophagy in human cells, Caenorhabditis elegans (e.g., Rosella and DCT-1/ LGG-1 strains), and mice (mt-Keima). A combination of these mitophagy detection techniques, including cross-species evaluation, will improve the accuracy of mitophagy measurements and lead to a better understanding of the role of mitophagy in health and disease.

In Vitro and In Vivo Detection of Mitophagy in Human Cells, C. Elegans, and Mice

Mitophagy, the process of clearing damaged mitochondria, is necessary for mitochondrial homeostasis and health maintenance. This article presents some of the latest mitophagy detection methods in human cells, Caenorhabditis elegans, and mice.

Mitochondria are the powerhouses of cells and produce cellular energy in the form of ATP. Mitochondrial dysfunction contributes to biological aging and a ...

Detection and Analysis of DNA Damage in Mouse Skeletal Muscle In Situ Using the TUNEL Method

Terminal deoxynucleotidyl transferase (TdT) deoxyuridine triphosphate (dUTP) nick end labeling (TUNEL) is the method of using the TdT enzyme to covalently attach a tagged form of dUTP to 3&#8217; ends of double- and single-stranded DNA breaks in cells. It is a reliable and useful method to detect DNA damage and cell death in situ. This video describes dissection, tissue processing, sectioning, and fluorescence-based TUNEL labeling of mouse skeletal muscle. It also describes a method of semi-automated TUNEL signal quantitation. Inherent normal tissue features and tissue processing conditions affect the ability of the TdT enzyme to efficiently label DNA. Tissue processing may also add undesirable autofluorescence that will interfere with TUNEL signal detection. Therefore, it is important to empirically determine tissue processing and TUNEL labeling methods that will yield the optimal signal-to-noise ratio for subsequent quantitation. The fluorescence-based assay described here provides a way to exclude autofluorescent signal by digital channel subtraction. The TUNEL assay, used with appropriate tissue processing techniques and controls, is a relatively fast, reproducible, quantitative method for detecting apoptosis in tissue. It can be used to confirm DNA damage and apoptosis as pathological mechanisms, to identify affected cell types, and to assess the efficacy of therapeutic treatments in vivo.

Detection and Analysis of DNA Damage in Mouse Skeletal Muscle In Situ Using the TUNEL Method

This video describes dissection, tissue processing, sectioning, and fluorescence-based TUNEL labeling of mouse skeletal muscle. It also describes a method for semi-automated analysis of TUNEL labeling.

Terminal deoxynucleotidyl transferase (TdT) deoxyuridine triphosphate (dUTP) nick end labeling (TUNEL) is the method of using the TdT enzyme to covalently ...

Cell-Type Specific Protein Purification and Identification from Complex Tissues Using a Mutant Methionine tRNA Synthetase Mouse Line

Understanding protein homeostasis in vivo is key to knowing how the cells work in both physiological and disease conditions. The present protocol describes in vivo labeling and subsequent purification of newly synthesized proteins using an engineered mouse line to direct protein labeling to specific cellular populations. It is an inducible line by Cre recombinase expression of L274G-Methionine tRNA synthetase (MetRS*), enabling azidonorleucine (ANL) incorporation to the proteins, which otherwise will not occur. Using the method described here, it is possible to purify cell-type-specific proteomes labeled in vivo and detect subtle changes in protein content due to sample complexity reduction.

This protocol describes how to perform cell-type-specific protein labeling with azidonorleucine (ANL) using a mouse line expressing a mutant L274G-Methionine tRNA synthetase (MetRS*) and the necessary steps for labeled cell-type-specific proteins isolation. We outline two possible ANL administration routes in live mice by (1) drinking water and (2) intraperitoneal injections.

Understanding protein homeostasis in vivo is key to knowing how the cells work in both physiological and disease conditions. The present protocol ...

Neuroscience

Identification of MyoD Interactome Using Tandem Affinity Purification Coupled to Mass Spectrometry

Skeletal muscle terminal differentiation starts with the commitment of pluripotent mesodermal precursor cells to myoblasts. These cells have still the ability to proliferate or they can differentiate and fuse into multinucleated myotubes, which maturate further to form myofibers. Skeletal muscle terminal differentiation is orchestrated by the coordinated action of various transcription factors, in particular the members of the Muscle Regulatory Factors or MRFs (MyoD, Myogenin, Myf5, and MRF4), also called the myogenic bHLH transcription factors family. These factors cooperate with chromatin-remodeling complexes within elaborate transcriptional regulatory network to achieve skeletal myogenesis. In this, MyoD is considered the master myogenic transcription factor in triggering muscle terminal differentiation. This notion is strengthened by the ability of MyoD to convert non-muscle cells into skeletal muscle cells. Here we describe an approach used to identify MyoD protein partners in an exhaustive manner in order to elucidate the different factors involved in skeletal muscle terminal differentiation. The long-term aim is to understand the epigenetic mechanisms involved in the regulation of skeletal muscle genes, i.e., MyoD targets. MyoD partners are identified by using Tandem Affinity Purification (TAP-Tag) from a heterologous system coupled to mass spectrometry (MS) characterization, followed by validation of the role of relevant partners during skeletal muscle terminal differentiation. Aberrant forms of myogenic factors, or their aberrant regulation, are associated with a number of muscle disorders: congenital myasthenia, myotonic dystrophy, rhabdomyosarcoma and defects in muscle regeneration. As such, myogenic factors provide a pool of potential therapeutic targets in muscle disorders, both with regard to mechanisms that cause disease itself and regenerative mechanisms that can improve disease treatment. Thus, the detailed understanding of the intermolecular interactions and the genetic programs controlled by the myogenic factors is essential for the rational design of efficient therapies.

MyoD is a myogenic transcription factor with a strong capacity to induce myogenic transdifferentiation of many fully differentiated non-muscle cell lines. The epigenetic mechanisms involved in this transdifferentiation are largely unknown. Here we describe a double-affinity purification method followed by mass spectrometry to exhaustively characterize MyoD partners.

Skeletal muscle terminal differentiation starts with the commitment of pluripotent mesodermal precursor cells to myoblasts. These cells have still the ...

Cryosectioning of Contiguous Regions of a Single Mouse Skeletal Muscle for Gene Expression and Histological Analyses

With this method, consecutive cryosections are collected to enable both microscopy applications for tissue histology and enrichment of RNA for gene expression using adjacent regions from a single mouse skeletal muscle. Typically, it is challenging to achieve adequate homogenization of small skeletal muscle samples because buffer volumes may be too low for efficient grinding applications, yet without sufficient mechanical disruption, the dense tissue architecture of muscle limits penetration of buffer reagents, ultimately causing low RNA yield. By following the protocol reported here, 30 &#956;m sections are collected and pooled allowing cryosectioning and subsequent needle homogenization to mechanically disrupt the muscle, increasing the surface area exposed for buffer penetration. The primary limitations of the technique are that it requires a cryostat, and it is relatively low throughput. However, high-quality RNA can be obtained from small samples of pooled muscle cryosections, making this method accessible for many different skeletal muscles and other tissues. Furthermore, this technique enables matched analyses (e.g., tissue histopathology and gene expression) from adjacent regions of a single skeletal muscle so that measurements can be directly compared across applications to reduce experimental uncertainty and to reduce replicative animal experiments necessary to source a small tissue for multiple applications.

Consecutive cryo-sections are collected to enable histological applications and enrichment of RNA for gene expression measurements using adjacent regions from a single mouse skeletal muscle. High-quality RNA is obtained from 20 - 30 mg of pooled cryosections and measurements are directly compared across applications.

With this method, consecutive cryosections are collected to enable both microscopy applications for tissue histology and enrichment of RNA for gene ...

Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion

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

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

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

Using Cleavage Under Targets and Tagmentation (CUT&Tag) Assay in Mouse Myoblast Research

Summary

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Chapters in this video

RiboTag Immunoprecipitation in the Germ Cells of the Male Mouse

An Efficient Protocol for CUT&RUN Analysis of FACS-Isolated Mouse Satellite Cells

Modeling Myotonic Dystrophy 1 in C2C12 Myoblast Cells

Single Cell Transcriptional Profiling of Adult Mouse Cardiomyocytes

In Vitro and In Vivo Detection of Mitophagy in Human Cells, C. Elegans, and Mice

Detection and Analysis of DNA Damage in Mouse Skeletal Muscle In Situ Using the TUNEL Method

Cell-Type Specific Protein Purification and Identification from Complex Tissues Using a Mutant Methionine tRNA Synthetase Mouse Line

Identification of MyoD Interactome Using Tandem Affinity Purification Coupled to Mass Spectrometry

Cryosectioning of Contiguous Regions of a Single Mouse Skeletal Muscle for Gene Expression and Histological Analyses

Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion