CUT&RUN is a powerful technique for determining protein localization on chromatin, and here we describe a single-cell version of this technique in a manual 96-well format. Most protein localization techniques, like ChIP-seq, require high cell input and take approximately three days to complete. CUT&RUN, because of the high signal and low background, has been optimized for low-cell and single-cell application, and can be completed in a single day.
We envision that this technique can be applied to almost any biological sample and can be especially powerful to determine factor localization in rare and low-abundance samples, such as precious patient samples. Before performing any single-cell experiments, test the antibody and technique on a high number of non-precious cells. The main limitation of this experiment is the dependence on a robust antibody.
Demonstrating the procedure is Santana Lardo, an experienced research specialist from my laboratory. After sorting single cells and binding nuclei to beads, place the 96-well plate onto a 96-well magnetic rack and allow the beads to bind for a minimum of five minutes. Then, remove and discard the supernatant.
Add 100 microliters of blocking buffer to the nucleus-bound beads, mix with gentle pipetting, and incubate for five minutes at room temperature. Place the plate on a 96-well magnetic rack and allow the supernatant to clear for a minimum of five minutes. Then, remove and discard the supernatant without disturbing the beads.
Remove the plate from the magnetic rack and resuspend the beads in 100 microliters of wash buffer per reaction with gentle pipetting. Place the plate back on the 96-well magnetic rack, allow the supernatant to clear, and then remove and discard the supernatant. Resuspend the beads in 25 microliters of wash buffer per reaction with gentle pipetting.
Make a primary antibody master mixture by aliquoting 25 microliters of wash buffer per reaction and then add 0.5 microliters of antibody per reaction. Add 25.5 microliters of the antibody wash buffer mix, followed by gentle pipetting to each sample being treated with an antibody targeting the protein of interest. Incubate for one hour at room temperature.
Then, again place the samples back on the 96-well magnetic rack. Once the supernatant is clear, remove and discard it without disturbing beads. After removing the plate from the magnetic rack, wash the beads with 100 microliters of wash buffer and resuspend by pipetting.
After placing the plate back on a 96-well magnetic rack and discarding supernatant as demonstrated previously, resuspend each sample in 25 microliters of wash buffer. Make a protein A MNase master mixture by adding 25 microliters of wash buffer and an optimized amount of protein A MNase per reaction. Add 25 microliters of protein A MNase master mixture to each sample, including the control samples.
Incubate the samples for 30 minutes at room temperature. Place the plate on a 96-well magnetic rack. Allow the supernatant to clear for a minimum of five minutes and then remove and discard the supernatant without disturbing the beads.
After removing the plate from the magnetic rack, wash the beads with 100 microliters of wash buffer and resuspend by gentle pipetting. Place the plate on a 96-well magnetic rack and discard the supernatant as demonstrated previously. Then, remove the samples from the magnetic rack and resuspend the beads in 50 microliters of wash buffer by gentle pipetting.
Equilibrate the samples to zero degrees Celsius in an ice water mixture for five minutes and then remove the samples from the ice water bath. Add one microliter of 100-millimolar calcium chloride using a multichannel pipette. Mix well three to five times with gentle pipetting using a large-volume multichannel pipette and then return the samples to zero degrees Celsius.
Start a 10 minute timer as soon as the plate is back in the ice water bath. Stop the reaction by pipetting 50 microliters of a solution containing twice the concentration of RSTOP+and buffer into each well in the same order as the calcium chloride was added. Incubate the samples for 20 minutes at 37 degrees Celsius.
Spin the plate at 16, 000 times G for five minutes at four degrees Celsius. Place the plate on a 96-well magnetic rack. Allow the supernatant to clear for a minimum of five minutes.
Then, transfer supernatants to a fresh 96-well plate and discard the beads. For DNA extraction, add one microliter of 10%SDS and 0.83 microliters of 20 milligrams per milliliter proteinase K to each sample and mix the samples by gentle pipetting. Incubate the samples for 10 minutes at 70 degrees Celsius.
Return the plate to room temperature. Add 46.6 microliters of five-molar sodium chloride and 90 microliters of 50%PEG4000 and mix by gentle pipetting. Add 33 microliters of polystyrene magnetite beads to each sample and incubate for 10 minutes at room temperature.
Place the plate on a magnetic rack and allow the supernatant to clear for around five minutes. Then, carefully discard the supernatant without disturbing the beads. Rinse twice with 150 microliters of 80%ethanol without disturbing the beads.
Spin the plate briefly at 1000 times G for 30 seconds. Place the plate back on a 96-well magnetic rack and remove all residual ethanol without disturbing the beads. Air dry the samples for around two to five minutes.
Resuspend the beads in 37.5 microliters of 10-millimolar TRIS hydrochloride of pH 8 and incubate for five minutes at room temperature. Place the plate back on a magnetic rack and allow the beads to bind for five minutes. Transfer 36.5 microliters of the supernatant to a fresh thermocycler-compatible 96-well plate and discard the beads.
After performing cell quality assessment, cell appearance and percentage were examined, demonstrating that low-quality embryonic stem cells should not be used. Single-cell sorting was performed using Hoechst 33342 stain, and the test cells were counted to assure either zero or one cell is found in each well. Agarose gel analysis revealed a low molecular weight ladder and individual single-cell uliCUT&RUN libraries.
Suboptimal and optimal library analysis show a low-molecular weight ladder, suboptimal library due to inefficient Mnase digestion, and successful library with appropriate digestion. Fragment analyzer distribution shows that the expected size of single-cell ranges from 150 to 500 base pairs. However, in large proteins, DNA will have around 270 base pairs.
Protein occupancy using single-locus genome browser depicting high cell number or negative control and single-cell CTCF uliCUT&RUN revealed that single-cell largely represented strong peaks, similar to high-cell uliCUT&RUN. Heat maps of single-cell CTCF or negative control revealed a clear pattern of read enrichment directly over established CTCF binding sites in the cells. One-dimensional heat maps show a higher read density over established CTCF binding sites in single-cell libraries relative to surrounding regions and no-antibody controls, demonstrating antibody-specific enrichment at established binding sites.
Equilibrating the bead-bound nuclei on the ice water bath and not overheating the sample upon calcium chloride addition is important to prevent over-digestion of the chromatin.
