Mar 27, 2026
CUT&RUN-direct Meers lab Version
- Arnold Federico1,
- Michael Meers1
- 1Washington University in St. Louis School of Medicine, Dept of Genetics
Protocol Citation: Arnold Federico, Michael Meers 2026. CUT&RUN-direct Meers lab Version. protocols.io https://dx.doi.org/10.17504/protocols.io.eq2lyj4jrlx9/v1
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: December 14, 2023
Last Modified: March 27, 2026
Protocol Integer ID: 92323
Keywords: direct meers lab version, 4x stop buffer, need for pce, pce, following protocol, protocol
Abstract
The following protocol is adapted from the CUT&RUN V.3 Protocol as written here: dx.doi.org/10.17504/protocols.io.zcpf2vn
This protocol is based on option 3 in the original, and makes use of a 4X STOP buffer without the need for PCE after the reaction
Guidelines
A typical protocol will take us three days, from cell prep to library prep.
3-Day split
1. Prepare beads and cells, incubate with primary antibody for target of interest O/N
2. Perform CUT&RUN, continue up until A-tail ligation and O/N Protein digestion
3. Perform cleanups, PCR, and quantifications/Tapestation
This protocol is written using 100k cells, if poor signals are obtained one may optimize the cell number or the primary antibody concentration, the original CnR protocol used up to 500K cells.
Troubleshooting
Before start
PLEASE READ BEFORE STARTING
*Reactions are originally written for 1.7 mL tubes to accommodate large cell numbers, however 0.6 mL tubes can be used when transferring tubes for primary antibody incubation and then on, requiring only 200uL per wash but other volumes should be maintained.
Reagents
Notes: Wear a mask and be careful when measuring out Digitonin as it is toxic! Using a slim spatula (one that will fit in a 1.7mL tube) measure 50 mg solid digitonin into a (tared) 1.7 mL tube (TAKE YOUR TIME). Then pipette 1mL of hot water into tube, no need to pipette mix! Take tube and vigorously vortex until well mixed
I recommend doing all of this on a paper towel so clean up is easier in case of spills. Digitonin crashes quickly out of solution. This should be made fresh and on day 1
Storage: Store binding buffer in 4 degree fridge up to 6 months
Note: Makes 0.05% buffer! Most washes use Digitonin wash buffer, not wash buffer. However, given that digitonin crashes out of solution easy, should only make as much Dig wash as needed per day. Comments in protocol for more details
Note: For day 1, this is the only time you will need Digitonin buffer, can just make this directly by adding 1mL Wash, 10uL 5% digitonin, and 4uL 0.5 M EDTA.
Note: Should be made before reaction, ie: During pA/G-MNase loading. each reaction takes 24 uL, this makes a little over 10 reactions worth. Can adjust accordingly as its a 50x dilution of 100mM CaCl2 in Dig wash buffer
Note: This will make enough for > 30 samples. Prepare stop buffer at the start of 30 min incubation for CUT&RUN on ice, have ready to go as soon as it's done! See notes in protocol for more details on this process
Cell Washes and Resuspension
For a streamlined experiment, prepare the following before starting
- 5% Digitonin
- Wash Buffer
- Binding Buffer
Gently homogenize cells in flask for even suspension, transfer small amount (0.5 mL) into a sterile eppendorf tube. Mix 10 uL of cells in tube with 10 uL of trypan blue. Do this again for 2 "replicates". Add 10 uL from each mix to both sides of a countess slide. Measure in countess/hemocytometer and take average for cell concentration.
Note
Make sure countess says *Trypan blue corrected! if not using trypan, make sure it says *Not/No trypan blue correction! Trypan assumes 1:1 dilution and calculates cell number. Ideally, cell viability is > 80%.
Transfer number of cells needed (100k per sample) into 15mL centrifuge tube and spin at 600 g for 3 minutes.
600 x g, Room temperature, 00:03:00
3m
Aspirate supernatant, resuspend in 1.0 mL room temperature Wash Buffer and transfer into 2 mL eppendorf tube.
Centrifuge again at 600 g for 3 minutes and remove supernatant. Add 1.0mL Wash buffer and resuspend for an additional wash. Perform one final centrifuge step, resuspending cells in 1.0 mL of wash buffer
600 x g, Room temperature, 00:03:00 X 2
Note
For low cell numbers (<100K) we often avoid this second wash step as we note cell loss/pellet smearing is common. Alternatively, one may prepare cells a head of time in a neutral buffer. The goal of these washes are to remove cell medium which lowers the efficiency of bead binding to cells.
3m
Bead prep and binding to cells
10m
Transfer 1.0 mL binding buffer into 1.7 mL tube (lo bind ). Add 5uL of Con-A beads per sample (5uL to 100K cells) into the binding buffer. Can perform over gentle vortexing, or flick for resuspension
Note
Make sure to mix Con-A beads very well before taking volume.
Place bead mixture on magnet stand, allow to clear ~ 2 min. Remove supernatant while on mag stand. Remove tube and add 1.0 mL of binding buffer, resuspend thoroughly
Place back on magnet stand, allow to clear ~ 2 min. Remove supernatant while on mag, remove tube and add equal volume of starting beads (5 uL per sample) resuspend thoroughly
Add all beads to resuspended cells. Can perform with gentle vortexing, or gently flick for resuspension. Incubate at room temp on rotator for 10 min 00:10:00
10m
Permeabilize cells and bind primary antibodies.
While cells are incubating, prepare antibody buffer and appropriate dilutions for 100 uL per sample. Mix thoroughly and keep on ice until use
Note
These should be optimized. A good starting point for histone mods is 1:100, and 1:50-1:25 for TFs
Example: If doing 6 samples, with two reps of two chromatin mods and a TF
Tube 1: 200uL antibody buffer, 2 uL mark 1 primary ab
Tube 2: 200uL antibody buffer, 2 uL mark 2 primary ab
Tube 3: 200uL antibody buffer, 4 uL TF primary ab
*MAKE SURE DIGITONIN WAS ADDED, native cells need to be permeabilized!
When cells are ready, gently mix and divide equally into labeled 1.7 mL eppendorf tubes for each sample using a wide bore pipette.
Note
Example: If doing 6 samples, should have 600k cells resuspended in ~1.0mL volume + 30 uL beads
Add 176 uL into 6 tubes labeled according to sample name
*This is where 0.6 mL tubes can be used
Add sample tubes to magnet stand, allow to clear. Resuspend in 100uL of appropriate antibody solution, ensure beads are homogenized by squirting the solution down beads and gently flicking/tapping.
Place resuspended sample tubes in rotator on cold room O/N
END DAY 1
Bind pA/G-MNase
1h
Note
This protocol does not utilize secondary antibody as it is typically not necessary for C&R as pA/G is fairly efficient at recognizing most primary antibodies we use. If secondary antibody step is required, refer to original protocol for 1 hr incubation: https://www.protocols.io/view/cut-amp-run-targeted-in-situ-genome-wide-profiling-14egnr4ql5dy/v3?preview_img=ED883866342911EEAFD10A58A9FEAC02&step=52
Before collecting tubes, dilute epicypher pA/G-MNase for 2 uL per reaction, in 100 uL volume of Dig-Wash Buffer. (ie: 6 samples, 600 uL Dig-Wash + 12 uL MNase). Keep on ice
Quick spin tubes to remove liquid from top, place on magnet stand and allow to clear. Remove supernatant.
Add 0.5 mL of Dig-wash buffer to sample, mix by flicking. Return back to magnet stand, allow to clear, remove supernatant and repeat again for a total of 2 washes.
Note
Given that we recommend preparation of Dig-wash as needed, for the Day 2 process, each sample will require 0.5 mL per wash for a total of 4 washes (2mL) ensure you have prepared enough wash buffer. On day 2 preparing a 10mL batch of Dig-wash will be enough for 4 samples plus other buffers that call for it.
Add 100 uL of Dig-Wash + pA/G-MNase to each tube being sure to homogenize beads. Flick/Tap gently to mix. Briefly spin if needed and incubate in cold room rotator for 1 hr.
01:00:00
1h
While incubating, use time to make 1X pA/G-MNase reaction mix
Chromatin Digestion and Release (Direct)
1h
Before removing tubes from cold room, place tube holder cold block on ice and allow to get down to temp. Keep in mind that there are holes for both 1.7 and 2 mL tubes, use the one for 1.7 (more snug)
Collect tubes, quick spin tubes to remove liquid from top, place on magnet stand and allow to clear. Remove supernatant.
Add 0.5 mL of Dig-wash buffer to sample, mix by flicking. Return back to magnet stand, allow to clear, remove supernatant and repeat again for a total of 2 washes
Remove all liquid, quick spin samples and place on cold block. Individually and quickly, add 24 uL of 1X pA/G-MNase reaction mix. Tap on tubes to mix well and leave in cold block. Immediately start 30 minute timer. 00:30:00 Use this time to make 4X stop buffer
Note
Time is important here, Set 30 min timer before adding anything, add 24uL to each tube, flick tubes, start timer.
30m
Add 8 uL of 4X STOP buffer to each reaction by submerging tip in bead solution. Flick/Tap to mix and spin down if necessary
Place samples on magnet stand and allow to completely clear. Transfer 30 uL of supernatant into new, labeled, 0.6 mL tubes.
Incubate samples in 37 C water bath for 30 minutes to release CUT&RUN fragments
00:30:00
30m
End Repair and Adapter Ligation (Library prep)
Note
Below is how CnR libraries are manually prepared in the Meers lab, one may use an alternative Illumina based kit such as NEBNext‱ Ultra if it is available/preferred. Here is a protodeveloped by Nan Liu in Stuart Orkin's lab (dx.doi.org/10.17504/protocols.io.wvgfe3w)
Prepare 4X End repair and A-tailing (ERA) Buffer:
Above is written per sample, multiply the amounts in the center column by number of samples (+1) to get the amount needed of each reagent. Keep on ice until ready for use
Add 10uL 4X ERA buffer to each 30 uL sample and mix by pipetting up and down
Place in thermocycler that has been pre-cooled to 12 C, run the "CnR_ERA" program
While samples are running, remove selected 1.5 uM TruSeq barcoded Y adapter from 4 C fridge and dilute 1:10 for 0.15 uM.
Note
Unlike C&T, C&R uses a y adapter which is annealed after end repair and A tailing. In the 4C, these are labeled as "IDT10 UDI #" in "CUT&RUN Adapters" Box. Where # represents the barcode that will be recognized with illumina seq.
One tube for one sample, try to not have overlapping barcode when pooling library. See sequencing sheet for example if necessary
Remove samples from thermocycler, add 5 uL of pre-specified adapter to respective sample.
Prepare "2X" rapid ligase solution using the 5X qiagen rapid ligase buffer and Qiagen ligase.
Per sample:
- 16 uL 5X Rapid Ligase buffer
- 4uL Ligase
- 20 uL of MilliQ water
Total = 40uL of 2X ligase solution
Multiply by the number of samples (+1)
Add 40 uL of 2X rapid ligase solution to each sample, mix well by pipetting
Place in thermocycler that has been pre-cooled to 20 C, run the program labeled "CnR_Ligate" in the Meers Lab folder.
Collect tubes, add 2 uL 10% SDS and 2 uL of Proteinase K. Incubate in 37 C waterbath O/N
END DAY 2
PCR Enrichment of CUT&RUN Libraries
20m
Remove Ampure beads from 4 C fridge, mix very well. Multiplying the number of samples by 300 uL will ensure you get enough beads. Add into smaller tube (Eppendorf or 15mL), place on RT rotator and allow to get to room temperature.
Note
This is still pre-pcr, perform these steps at your bench not the clean up bench
FIRST CLEAN UP: Add 1.0 X the volume of Ampure beads (e.g to 89 uL sample add 89 uL Ampure Beads). Mix well by pipetting up and down, incubate at RT for 5 minutes
00:05:00
5m
Place tubes on mag, remove all supernatant. While beads are on stand, add 200 uL of 80% EtOH to each sample, incubate for 30 seconds. Remove ethanol, repeat again for 2 total washes. Briefly spin down, replace on mag, remove any residual EtOH. Allow to air dry for 2-3 min
Remove tube from magnet, add 50uL 10mM Tris-HCl pH 8 (NO EDTA). Mix well, incubate at RT for 5 minutes
00:05:00
5m
Return samples to magnet, allow to clear. Transfer samples into NEW and labeled 0.6 mL tubes.
SECOND CLEAN UP: Add 1.2 X the volume of Beads to the transferred samples (60 uL). Mix well, and allow to incubate at RT for 5 minutes
00:05:00
5m
Place tubes on mag, remove all supernatant. While beads are on stand, add 200 uL of 80% EtOH to each sample, incubate for 30 seconds. Remove ethanol, repeat again for 2 total washes. Briefly spin down, replace on mag, remove any residual EtOH. Allow to air dry for 2-3 min
Add 25 uL of Tris-HCL pH 8 to all tubes, mix very well and allow to incubate at RT for 5 minutes
00:05:00
5m
From this volume, transfer 21 uL into a final set of tubes to be used for PCR. Excess volume ensures beads do not get pulled with supernatant.
Add 2 uL of Illumina F and R universal primers to each sample (0.4 uM final, each). Add 25uL of 2X NEB Next Hot Start PCR Mastermix. Place tubes in thermo cycler and run the protocol labeled "CnR_14cycl", letting come to temperature first.
Post-PCR Cleanup
15m
FIRST CLEAN UP: Add 1.1 X the volume of Ampure beads (e.g to 50 uL sample add 55 uL Ampure Beads). Mix well by pipetting up and down, incubate at RT for 5 minutes
Place tubes on mag, remove all supernatant. While beads are on stand, add 200 uL of 80% EtOH to each sample, incubate for 30 seconds. Remove ethanol, repeat again for 2 total washes. Briefly spin down, replace on mag, remove any residual EtOH. Allow to air dry for 2-3 min
Remove tube from magnet, add 50uL 10mM Tris-HCl pH 8 (NO EDTA). Mix well, incubate at RT for 5 minutes
00:05:00
5m
Transfer samples into NEW and labeled 0.6 mL tubes.
SECOND CLEAN UP: Add 1.2 X the volume of Beads to the transferred samples (60 uL). Mix well, and allow to incubate at RT for 5 minutes
00:05:00
5m
Place tubes on mag, remove all supernatant. While beads are on stand, add 200 uL of 80% EtOH to each sample, incubate for 30 seconds. Remove ethanol, repeat again for 2 total washes. Briefly spin down, replace on mag, remove any residual EtOH. Allow to air dry for 2-3 min
Add 28 uL of Tris-HCL pH 8 to all tubes, mix very well and allow to incubate at RT for 5 minutes
00:05:00
5m
Replace back on magnet stand and allow to clear. Transfer 25 uL of samples into another set of clean 1.7 mL tubes, and you're done!