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

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'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)

1. Dilute glacial acetic acid with ddH₂O to 0.02 M and autoclave it.
2. Dilute collagen (from rat tail) to 0.1 mg/mL with 0.02 M acetic acid.
3. Add this collagen solution into culture dishes to coat the dish bottom in a 37 °C cell incubator for overnight.
4. Before use, remove the collagen solution and wash the dish twice with 1x PBS. Collagen solution can be reused for 8-10 times.
5. Sacrifice an 8-12-week-old mouse. (Old mice have poor yields of MuSCs and myoblasts.)
6. Isolate and place all hindlimb muscles from the mouse into a 10-cm dish.
7. Use 1x PBS (with penicillin/streptomycin) to rinse the muscles 4 times. After each rinse, always transfer the muscles into a new plate.
8. After the final rinse, remove PBS and mince up the muscles with scissors.
9. 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.
10. To perform digestion, transfer the minced muscles into a tube in a 37 °C water bath or directly leave the minced muscles within the dish and place it in a 37 °C CO₂ cell incubator.
11. 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.
12. 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.
13. 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.
14. Run the mixture through a 70-µm 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.
15. Centrifuge at 350 x g for 10 min and discard the supernatant with vacuum.
16. Resuspend the cell pellet with 20 mL of 1x PBS and run the cells through a 40-µm strainer.
17. Centrifuge at 350 x g for 5 min and remove the supernatant with vacuum.
18. 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.
19. 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.
20. After another day, passage the cells: Remove the media, wash the cells once with 1x PBS, and trypsinize the cells.
21. 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.
22. 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.
23. 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.

2. Preparation of MuSC/Myoblast growth media (Example of preparing from 5 mice)

1. Sacrifice 5 young and healthy wild-type mice, isolate the spleens, and rinse the spleens once with 1x PBS (with penicillin/streptomycin).
2. Use scissors to remove the dark parts/spots on the spleens, if any. Discard the spleen if most of it has turned black.
3. Place all 5 spleens into a 40-µm strainer on the top of a 50 mL tube.
4. Open a sterile 1 mL syringe package, pull out the plunger, and discard the barrel.
5. In the 40-µm 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.
6. While grinding the spleens, occasionally use 1x PBS to wash off the dissociated spleen cells into the 50 mL tube underneath the strainer.
7. 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.
8. 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.
9. Fill up the 50 mL tube with 1x PBS to cease the lysing process of red blood cells.
10. Strain the cell suspension again with a 40-µm strainer, and then centrifuge at 500 x g for 10 min.
11. Resuspend the cells with 2 mL of RPMI-1640 full media (RPMI-1640 media with 10% FBS and penicillin/streptomycin) with very gentle pipetting.
12. 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.
13. Add 10 µL 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&Tag assays later on.
14. 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.
15. Another 24 h later, collect all the culture in each T75 flask and centrifuge at 600 x g for 3 min.
16. The supernatant is T-cell conditional media and can be stored at -20 °C to -80 °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 µm filter.
17. To make Ham's F10 full media, add 100 mL of FBS, 300 µL of bFGF stock, and 6 mL of penicillin/streptomycin into 500 mL of Ham's F10 media.
    NOTE: bFGF stock is 5 µg/mL and prepared by dissolving bFGF in Ham's F10 media.
18. Mix the Ham's F10 full media with filtered T-cell conditional media described above at 1:1 ratio to make MuSC/Myoblast growth media.

3. CUT&Tag with myoblasts (Example of 3 CUT&Tag reactions)

1. Preparation of key reagents
   1. 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.
   2. 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.
   3. 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&Tag.
   4. 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.
   5. Prepare the Binding buffer, which contains 20 mM HEPES pH 7.5, 10 mM KCl, 1 mM CaCl₂, and 1 mM MnCl₂.
   6. Prepare the Wash buffer, which contains 20 mM HEPES pH 7.5, 150 mM NaCl, 0.5 mM Spermidine, and 1x Protease inhibitor cocktail.
   7. 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.
   8. 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.
   9. 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.
   10. 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.
2. Concanavalin-A bead activation
   1. Mix 30 µL of Concanavalin-A beads (10 µL for each reaction) into 300 µL 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.
   2. Remove the tube from the magnetic rack, add 300 µL 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&Tag reaction.
   3. Put the 8-PCR tube stripe on a magnetic rack and wait till the liquid is clear.
   4. Aspirate the supernatant and remove the tubes from the magnetic rack. Add 10 µL of Binding buffer into each tube and mix well. Place the prepared beads on ice or at 4 °C till the cells are ready to proceed.
3. Binding cells onto Concanavalin-A beads
   1. 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.
   2. Adjust the cell density to around 7 x 10⁵ cells/mL in Wash buffer such that 100 µL of cell suspension contains roughly 70,000 cells, sufficient for one CUT&Tag reaction.
   3. Pipet 100 µL of the cell suspension into each tube of the 8-PCR tube stripe, which already contains 10 µL of prepared Concanavalin-A beads. Mix well and incubate on a "seesaw motion" shaker for 10 min at RT.
   4. Put the 8-PCR tube stripe on a magnetic rack. Wait till it clears, and remove the supernatant.
   5. 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.
4. Incubating the cells with a primary antibody
   1. For each sample, dilute 1 µL of H3K4me1 antibody into 50 µL 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 "seesaw motion" shaker to incubate at 4 °C for overnight.
      NOTE: As mentioned by Dr. Steven Henikoff¹, it was not necessary to use an IgG negative control in all CUT&Tag assays. Compared to ChIPs, CUT&Tag assays come with much lower or no background signal. Therefore, using IgG to perform a "negative control" CUT&Tag assay does not introduce any useful information. In contrast, using a CUT&Tag-validated antibody to perform a positive control is important to assess whether the CUT&Tag assay has worked.
5. Incubating the cells with a secondary antibody
   1. The next day morning, put the samples on the magnetic rack and wait till the samples clear.
   2. Remove the supernatant, which is the primary antibody. There is no need to wash the beads; Directly add 50 µL 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.
   3. Mix well and incubate on the "seesaw motion" shaker at RT for 1 h.
   4. Put the samples back on the magnetic rack, wait till the samples clear, and discard the supernatant.
   5. Add 200 µL of Dig-wash buffer into each tube and invert several times to wash off the excessive antibodies.
   6. 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.
6. Incubating the cells with pA/G-Tn5a
   1. 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 µL of 300-Dig buffer containing pA/G-Tn5a at 0.04 µM. Mix well and incubate on the seesaw motion shaker for 1 h at RT.
   2. Put on the magnetic rack. Wait till the samples clear and remove the supernatant.
   3. Add 200 µL of 300-Dig buffer into each sample, mix well, and place on the shaker to wash the sample for 3 min at RT.
   4. 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.
7. Cleavage and tagmentation of the genomes
   1. After the last wash, put the samples on the magnetic rack and thoroughly pipet the liquid off the tube. For each sample, add 50 µL of Tagmentation buffer. Mix well and incubate in a PCR machine at 37 °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.
8. Total genomic DNA purification
   1. After tagmentation, add another 150 µL of Tagmentation buffer for each sample. The total volume is now 200 µL.
   2. Then, for each sample, add 10 µL of 0.5 M EDTA, 3 µL of 10% SDS, and 2.5 µL of 20 mg/mL Proteinase K. Vortex to mix well and incubate at 55^°C for 2 h.
   3. Pipet each sample into 200 µL 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.
   4. Add 200 µL of chloroform into each tube and vortex for 10 s. Centrifuge at 18,000 x g for 5 min.
   5. Aspirate the top layer into a new 1.5 mL centrifuge tube. Then, for each sample, add 550 µL of 100% ethanol to precipitate the DNA at -80 °C for 1 h. To better visualize the DNA pellet, add 1 µL of 20 mg/mL glycogen.
   6. Centrifuge at max speed (>18,000 x g) for 15 min to pellet the DNA.
   7. Carefully pour off the supernatant, wash the glycogen/DNA pellet with 75% ethanol once, and centrifuge again at max speed for 5 min.
   8. Remove the 75% ethanol and resuspend the glycogen/DNA pellet with 21 µL of ddH₂O.
9. Library preparation and sequencing
   1. 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.
   2. Roughly determine the PCR cycle number in library-preparing PCR by the cell number used in the CUT&Tag assay. Normally, for a CUT&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&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/µL (determined by Qubit assay) in concentration and 20-30 µL 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.
   3. Use some commercially available DNA purification/size-selection beads to purify the library DNA and select the DNA of desirable sizes.
   4. 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&Tag has worked.
   5. Perform sequencing and data analysis as per the manufacturer's instructions.
      NOTE: CUT&Tag kits are commercially available from a handful of companies, and one example can be found in the Table of Materials file.