A fluorescence-based protocol to quantitatively titrate CUT&RUN buffer components

Summary

Cleavage under targets & release using nuclease (CUT&RUN) is a technique for identifying genomic sites where proteins or histone modifications are present in chromatin in permeabilized cells. Here, we present a fluorescence-based protocol to quantitatively titrate CUT&RUN buffer components, for efficient cell permeabilization and retention of target epitopes on chromatin. We describe steps for capturing cells on concanavalin A beads and using a fluorescently labeled secondary antibody to titrate concentrations of digitonin and NaCl in CUT&RUN buffers. We then detail procedures for fluorescence imaging to identify optimal conditions.

For complete details on the use and execution of this protocol, please refer to Lerner et al.¹

Graphical abstract

Highlights

• •A fluorescence-based approach to measure reagent delivery in CUT&RUN experiments
• •Titration of digitonin and NaCl for cell and target-specific epitope accessibility
• •Titrated CUT&RUN conditions can improve genomic profiling of transcription factors

Publisher’s note: Undertaking any experimental protocol requires adherence to local institutional guidelines for laboratory safety and ethics.

Cleavage Under Targets & Release Using Nuclease (CUT&RUN) is a technique for identifying genomic sites where proteins or histone modifications are present in chromatin in permeabilized cells. Here, we present a fluorescence-based protocol to quantitatively titrate CUT&RUN buffer components, for efficient cell permeabilization and retention of target epitopes on chromatin. We describe steps for capturing cells on concanavalin A beads and using a fluorescently labeled secondary antibody to titrate concentrations of digitonin and NaCl in CUT&RUN buffers. We then detail procedures for fluorescence imaging to identify optimal conditions.

Before you begin

Cleavage under targets & release using nuclease (CUT&RUN) is a method to profile the genomic targets of chromatin-associated proteins.² CUT&RUN provides an alternative to ChIP-seq in assessing protein-DNA interactions, with notable advantages of low cell number requirements, a high signal to noise, and profiling in native, crosslinking-free conditions.^3,4 While CUT&RUN works well for mapping histones and their modifications, due to their stable integration in chromatin, mapping transcription factors can provide challenges due to an overall lower nuclear abundance and transient and dynamic interactions with chromatin.

In a CUT&RUN experiment, cell and nuclear membranes are permeabilized with digitonin, permitting a primary antibody, targeting an epitope of interest, and protein A/G-MNase fusion protein to enter the nucleus. MNase is then activated by addition of CaCl₂, enabling the release of fragmented chromatin specifically around the target epitope, which can be recovered, assembled into a library, and sequenced. Two critical experimental parameters are 1) ensuring sufficient nuclear permeability for antibody entry, and 2) retaining target epitopes on the chromatin through successive salt washes. We provide a method using a fluorescent secondary antibody to titrate buffer conditions within the context of a CUT&RUN experiment, testing how concentrations of digitonin impact accessibility for antibody delivery and how varying concentrations of NaCl impact the release of proteins from chromatin.

Different cell types display varying sensitivities to digitonin, and thus performing the assay with a sufficient concentration of digitonin is critical. Current CUT&RUN protocols assess nuclear permeability using trypan blue staining.^2,5,6,7 However, we found that access of a small dye is not representative of allowing access to bulkier molecules such as antibodies within the assay. Notably, our protocol describes the specific steps to visualize how increasing amounts digitonin enable nuclear access to specific primary antibodies, as shown here with primary human fibroblasts.

Successive washes containing high salts can elute factors off the chromatin and into the nucleoplasm. Existing CUT&RUN protocols have not tested how salt concentrations impact the retainment of proteins on the chromatin, and within the nucleus. Yet titration of different salt conditions can improve the stability of proteins on the chromatin, making the assay compatible with a wider range of targets. We show how to test the effect of NaCl concentration in wash buffers on the retainment of transcription factors in the nucleus, with the transcription factor SOX2 as an example. We anticipate that our modifications to the original CUT&RUN protocol will expand the number of investigators that are successfully able to use the method to map transcription factors in chromatin.

Prepare wash buffers for titration

Timing: 1 h

The steps below describe preparation of wash buffers containing titrated NaCl or digitonin concentrations. Wash buffers prepared with titrated NaCl are used to test nuclear epitope retention while wash buffers prepared with titrated digitonin are used to test nuclear permeability.

• 1.
  Prepare digitonin-containing wash buffer (dig-wash) with varying concentrations of digitonin.

  • a.
    In a 1.5 mL microcentrifuge tube, prepare 1 mL of a 5% (w/v) digitonin stock solution in DMSO. The stock solution should be prepared at room temperature, with long-term storage at 4°C or ‒20°C.

    Note: The 5% digitonin stock can be stored at 4°C for one week, or ‒20°C for up to 6 months.

  • b.
    Add digitonin to the CUT&RUN wash buffer (20 mM HEPES pH 7.5, 150 mM NaCl, 0.5 mM Spermidine, cOmplete EDTA-free protease inhibitor) to achieve concentrations ranging from 0%–0.1%.

    Note: We suggest first testing digitonin concentrations varying by factors of 10, e.g., dig-wash buffers containing 0%, 0.001%, 0.01%, 0.1% digitonin. Subsequently, narrower concentration ranges can then be tested for a particular cell type.

• 2.
  Prepare wash and dig-wash buffer with varying concentrations of NaCl.

  • a.
    Modify CUT&RUN wash buffer and dig-wash buffer to contain 75 mM, 150 mM or 350 mM NaCl.

Note: Each sample will require 4 mL wash and dig-wash buffer for each NaCl or digitonin condition.

Note: Wash buffer containing 350 mM NaCl is higher than required for CUT&RUN, but it’s inclusion here is recommended to show how higher salt concentrations may solubilize proteins off chromatin and wash out of the nucleus.

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
Rabbit anti-H3K27me3 (1:100)	Cell Signaling Technology	Cat#C36B11; RRID: AB_2616029
Donkey anti-rabbit 488 secondary (1:500)	Thermo Fisher Scientific	Cat#A-21206; RRID: AB_2535792
Mouse anti-V5 (1:100)	Invitrogen	Cat#R960-25; RRID: AB_2556564
Donkey anti-mouse 488 secondary (1:500)	Thermo Fisher Scientific	Cat#A-21202; RRID: AB_141607
Chemicals, peptides, and recombinant proteins
Phosphate-buffered saline (PBS)	Sigma-Aldrich	Cat#806544
Dimethyl sulfoxide (DMSO)	Sigma-Aldrich	Cat#D8418-50ML
Digitonin	Sigma-Aldrich	Cat#D141-100MG
cOmplete, Mini, EDTA-free protease inhibitor cocktail	Millipore Sigma	Cat#11836170001
DAPI (4′,6-diamidino-2-phenylindole, dihydrochloride)	Thermo Fisher Scientific	Cat#D1306
Deposited data
SOX2 CUT&RUN	Lerner et al., 2023	GSE220570
Experimental models: Cell lines
Human BJ fibroblasts	ATCC	Cat#CRL-2522
Software and algorithms
Fiji is Just ImageJ (Fiji) version 2.0.0-rc/1.51f	ImageJ	https://imagej.net/
Other
1.5 mL microcentrifuge tubes	USA Scientific	Cat#4036-3204
Accutase	STEMCELL Technologies	Cat#07920
Magnetic concanavalin A beads	Antibodies-online	Cat#ABIN6952467
Magnetic rack for 1.5 mL Tubes	Invitrogen	Cat#12321D
Wide-field fluorescent microscope	Nikon	Nikon Eclipse TE2000

Materials and equipment

Bead activation buffer

Reagent	Final concentration	Amount
1 M HEPES pH 7.5	20 mM	200 μL
1 M KCl	10 mM	200 μL
1 M CaCl2	1 mM	20 μL
1 M MnCl2	1 mM	20 μL
ddH2O	N/A	9.58 mL
Total	N/A	10 mL

Note: Store at 4°C for up to 6 months.

Wash buffer (testing protein retention on chromatin)

Reagent	Final concentration	Amount
1 M HEPES pH 7.5	20 mM	200 μL
6.4 M spermidine	0.5 mM	0.78 μL
5 M NaCl	75 mM–350 mM	150 μL–700 μL
cOmplete mini protease inhibitor	1x	1 tablet
ddH2O	N/A	9.65 mL–9.05 mL
Total	N/A	10 mL

Note: Make fresh for each use.

Wash buffer (testing nuclear permeability)

Reagent	Final concentration	Amount
1 M HEPES pH 7.5	20 mM	200 μL
6.4 M spermidine	0.5 mM	0.78 μL
5 M NaCl	150 mM	300 μL
cOmplete mini protease inhibitor	1x	1 tablet
ddH2O	N/A	9.5 mL
Total	N/A	10 mL

Note: Make fresh for each use.

Dig-wash buffer (testing nuclear permeability)

Reagent	Final concentration	Amount
Wash buffer	N/A	10 mL
5% digitonin	0.001%–0.1%	2 μL–200 μL
Total	N/A	10 mL

Note: Make fresh for each use. Digitonin should be added to wash buffer at room temperature, with the dig-wash buffer subsequently stored on ice.

Dig-wash buffer (testing protein retention to chromatin)

Reagent	Final concentration	Amount
Wash buffer (with variable NaCl)	N/A	10 mL
5% digitonin	0.05%	100 μL
Total	N/A	10 mL

Note: Make fresh for each use. Digitonin should be added to wash buffer at room temperature, with the dig-wash buffer subsequently stored on ice.

Step-by-step method details

Our protocol follows the steps outlined in Skene et al., 2018⁴ through the primary antibody incubation, with the variation of using wash and dig-wash buffers with titrated concentrations of NaCl or digitonin. Our protocol next deviates by incubation with a fluorescent secondary antibody, measuring the impact of buffer conditions on reagent delivery to the nucleus and epitope retention on the chromatin.

Activate magnetic concanavalin A beads

Timing: 10 min

The following steps describe activation of Concanavalin A (conA) beads for cell capture in the assay.

Note: To ensure cell morphology is clearly visible during imaging, we suggest activating 10 μL of conA beads per 250,000 cells. If fewer cells are available, the volume of conA beads should be scaled down proportionally (e.g. 5 μL of conA beads per 125,000 cells).

• 1.Resuspend Concanavalin A (conA) bead stock by light vortexing.
• 2.Pipette 10 μL of conA beads per NaCl or digitonin condition to be tested into a chilled 1.5 mL microcentrifuge tube.
• 3.Place beads onto a magnetic rack and once the supernatant is clear, remove supernatant and resuspend in 1 mL chilled bead activation buffer by light pipetting.
• 4.
  Repeat step 3 and resuspend beads in 10 μL bead activation buffer per sample.

  • a.
    Leave activated conA beads on ice until use.

Capture cells on concanavalin A beads

Timing: 1 h

The following steps describe the dissociation of adherent fibroblasts from a 10 cm tissue culture dish. We suggest using 250,000 cells for each different wash buffer condition with titrated NaCl or titrated digitonin.

Note: Unless otherwise noted, all steps can be performed at room temperature.

• 5.Wash adherent cells on tissue culture plate twice with room temperature PBS.
• 6.Detach cells by pipetting 3 mL Accutase directly onto the tissue culture plate. Once the cells are successfully detached from the plate, count, and distribute 250,000 cells for each wash or dig-wash buffer condition into separate 1.5 mL microcentrifuge tubes.
• 7.Centrifuge cells in a rotor at 600 xg for 3 min.
• 8.Discard the supernatant and resuspend cells in 1 mL of the appropriate wash buffer based on NaCl concentration.
• 9.Repeat 1 mL wash and spin down at 600 xg for 3 min.
• 10.
  Capture cells onto magnetic conA beads, adding 10 μL beads/sample.

  • a.
    Allow beads to bind cells by rotating for 5–10 min.

Permeabilize cells and incubate with primary antibody

Timing: 1.5 h to overnight

The following steps describe the addition and incubation of bead-bound cells with a primary antibody of interest. The epitope targeted by the antibody chosen here will be visualized by fluorescent microscopy.

• 11.Place bead-bound cells on magnetic rack discard the supernatant and resuspend in 100 μL of the appropriate dig-wash buffer based on digitonin or NaCl concentration.
• 12.
  Add 1 μL of primary antibody to bead-bound cells, gently inverting tube until beads appear homogenous in solution.

  • a.
    If testing nuclear permeability with variable digitonin, add 1 μL H3K27me3 antibody.
  • b.
    If testing transcription factor solubility with variable NaCl, add 1 μL antibody specific for the transcription factor.

Note: we suggest staining for H3K27me3 when testing nuclear permeability due to this mark’s overall abundance and the availability of a highly-specific monoclonal antibody (see key resources table), providing clearly fluorescent cells. Other abundant histone modifications, such as H3K36me3 and H3K9me3, may be substituted.

• 13.Leave bead-bound cells containing the appropriate antibody for 1 h at room temperature or at 4°C overnight.

Conjugate primary antibody with fluorescent secondary antibody

Timing: 1.5 h

The following steps describe incubation with a fluorescently labeled secondary antibody (see key resources table). The secondary antibody must be species matched to the primary antibody, and compatible with fluorescent microscopy.

• 14.Place bead-bound cells on the magnetic rack. Once the supernatant is cleared, discard the antibody-containing buffer, and wash beads with 1 mL dig-wash buffer.

Note: For all subsequent washing steps, use the dig-wash buffer containing the appropriate concentration of digitonin or NaCl, consistent with prior washes

• 15.Repeat step 14.
• 16.To each sample, add a fluorescent secondary antibody 1:100, species matched to the primary antibody.

Note: Any fluorescent secondary antibody is compatible with this step and should be chosen based on your imaging system.

• 17.Leave sample in dark for 1 h at 4°C.
• 18.Place the bead-bound cells on the magnetic rack. Once the supernatant is cleared, discard the buffer containing the secondary antibody, and resuspend the bead-bound cells in 1 mL dig-wash buffer.
• 19.Repeat step 18.
• 20.After the second wash, resuspend bead-bound cells in 100 μL dig-wash buffer.

Note: To minimize photobleaching, we recommend performing washes as fast as possible or washing in a low-light area.

Image fluorescent cells

Timing: 30 min to 2 h

The following steps describe the assessment of buffer conditions enabling nuclear permeabilization or epitope retainment, through fluorescent microscopy.

• 21.Pipette 25 μL of cells onto a slide, and gently cover with coverslip.

Note: Cells do not necessarily need to be mounted or otherwise immobilized for imaging.

• 22.Image cells on both fluorescent channel and bright-field channel, maintaining consistent exposure between samples.

Note: We recommend a 4x or 10x magnification to capture a field of cells and detect possible heterogeneity of fluorescent signal. A 20x or 40x magnification is important to note cell morphology.

Note: All images should be saved as TIFF files for quantitative comparisons between different conditions.

Note: After imaging, the bead-bound cells are no longer required. The cells in dig-wash buffer and fluorescent dye can be kept for up to a day in the dark at 4°C, and re-imaged if further fields of view are required.

Image analysis

Timing: 30 min to 1 h

The following steps describe basic analysis of bright-field and fluorescent images.

• 23.Load bright-field and fluorescent images into an image processing software such as FIJI (Figure 1A).⁸
• 24.Overlay bright-field and fluorescent images to ensure signal is localized to the nucleus, and not autofluorescence from free-floating conA beads or other cellular debris (Figure 1B).
• 25.Create a panel montage of images, ordered from low concentration to high concentration (ex. 0% digitonin to 0.1% digitonin, or 75 mM NaCl to 350 mM NaCl) (Figure 1C).
• 26.Count total number of cells and fluorescent nuclei. Identify the minimal concentration of digitonin and NaCl that optimize number of fluorescing nuclei. These are the composition for wash buffers that should be employed when performing the complete CUT&RUN protocol, with the tested cell type and chromatin-associated protein.

Figure 1.

Analysis of images in FIJI

(A) Example phase contrast and fluorescent images for further analysis.

(B) FIJI commands to overlay phase contrast and fluorescent channels to confirm fluorescent signal is specific to nuclei.

(C) FIJI commands to create an image montage for side-by-side comparison of different washing conditions.

Expected outcomes

A titration of digitonin will show that increasing concentrations enable more efficient nuclear accessibility to antibodies, as measured by fluorescent nuclei (Figure 2). We suggest using the minimal concentration that show > 95% cell permeability, which can be calculated by (the number of fluorescent nuclei / total number of cells) ∗ 100. Using buffers containing the minimal concentration of digitonin for sufficient permeabilization will minimize cell lysis throughout the full protocol, including overnight antibody incubation and MNase digestion. In a variety of tested mammalian cell types, we find ranges between 0.01% and 0.1% digitonin are often suitable. Yet some cell types, such as myocytes that have high levels of nuclear lamins, may require significantly higher digitonin concentrations for proper permeabilization.

Figure 2.

Digitonin titration for cell permeabilization

(A) Phase contrast and GFP channel (anti-H3K27me3) imaging across digitonin titrated over 3 orders of magnitude, to identify ideal concentration for CUT&RUN experiments. Scale bars, 75 μm.

(B) Phase contrast and GFP overlay to compare H3K27me3 immunostaining in 0.1% vs. 0.01% digitonin conditions. White arrows indicate under-permeabilized cells. Scale bars, 20 μm.

Optimal concentration of NaCl will be dependent on the chromatin affinity for the specific target. For our titration of SOX2, we found that lowering NaCl from the original protocol of 150 mM–75 mM helped retain much stronger SOX2 nuclear signals (Figure 3). We have not attempted CUT&RUN with NaCl concentrations below 75 mM, as increasingly lower salt levels may lead to non-specific antibody binding and disrupting the osmotic balance within the cells.

Figure 3.

NaCl titration for SOX2 retainment on chromatin

(A) Phase contrast and GFP channel (anti-SOX2) testing impact of NaCl concentration of SOX2 chromatin retainment. Scale bars, 75 μm.

(B) Phase contrast and GFP overlay to compare SOX2 immunostaining in 75 mM vs. 150 mM NaCl conditions. White arrows indicate clusters of cells without SOX2 signal. Scale bars, 15 μm.

Optimizing the CUT&RUN assay by increasing the concentration digitonin in dig-wash buffer from 0.01% to 0.1% and decreasing the concentration of NaCl from 150 mM to 75 mM greatly improved the genomic signal-to-noise for SOX2,¹ identifying a greater number of peaks enriched for the SOX2 motif (Figure 4). The optimizations do not impact the ability to profile histone marks, likely due to their stability in the chromatin and overall abundance across the genome (Figure 4C).

Figure 4.

Impact of CUT&RUN optimizations on SOX2 profiling

SOX2 CUT&RUN data in optimized conditions presented as analyzed in Lerner et al., 2023.

(A) Summary of SOX2 peak calling and motif enrichment in starting vs. optimized buffer conditions.

(B) Heat map showing CUT&RUN signal of SOX2 in starting vs. optimized buffer conditions.

(C) Genome browser showing SOX2 and H3K27me3 signal in starting vs. optimized buffer conditions. SOX2 but not H3K27me3 is sensitive to these optimizations.

After optimization of buffer conditions, we have had success using the complete CUT&RUN protocol described in.⁴ We find that the only way to assess if the optimizations improve data quality is by sequencing. Assessing library quality with capillary electrophoresis methods such as a Bioanalyzer or Tape Station can be inconclusive, as CUT&RUN of both IgG (resulting in non-specific digestion) and chromatin-associated proteins often show an enrichment of fragments around 300 bp.

Limitations

CUT&RUN experiments that yield low-quality data may be due to reasons beyond improper buffer conditions. Antibody specificity is a key indicator of the success in a CUT&RUN experiment, and as such has been suggested to test specificity by immunostaining.² The inclusion of a control condition in which the target epitope is knocked down or out can discriminate non-specific from specific antibody binding. Further, while here we optimize for delivery of reagents to the chromatin, it has been reported that target sites bound by large protein complexes are insufficiently solubilized and released after digestion, which may necessitate further optimization.⁹

Troubleshooting

Problem 1

Clumped cells make it difficult to count the number of fluorescent nuclei (related to step 22).

Potential solution

Bead-bound cells show a propensity to clump together. Prior to imaging (related to step 21), lightly resuspending the cells by pipetting can homogenize the sample.

If resuspension does not separate cells, DAPI staining can distinguish individual nuclei. After step 17, cells can be incubated in dig-wash buffer containing 1 μg/mL DAPI for 10 min, prior to final washes.

Problem 2

Weak fluorescent signal impacts the ability to identify properly permeabilized nuclei with intact target epitopes (related to step 26).

Potential solution

Weak signals can occur due to low abundance of the target, as well as poor antibody specificity. Inclusion of an IgG control to test for non-specific fluorescent background, in a side-by-side comparison with the target condition, allows a quantitative assessment of signal intensity. The target condition should have higher signal than the IgG control. If no difference between IgG and the target condition are observed, further testing if the antibody provides clear signal with traditional immunofluorescence is recommended.

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Kenneth Zaret (zaret@pennmedicine.upenn.edu).

Technical contact

Questions regarding the use and analysis of the protocol can be directed to the technical contact, Andrew Katznelson (akatznel@pennmedicine.upenn.edu).

Materials availability

This study did not generate new, unique reagents.

Data and code availability

Original data have been deposited to Mendeley Data: https://doi.org/10.17632/zjbw23wz76.1.