For the best experience on the Abcam website please upgrade to a modern browser such as Google Chrome
See how two new chromatin profiling methods, CUT&RUN and CUT&Tag, overcome many of the drawbacks of conventional ChIP methods with our comprehensive guide written by Claudia Semprich (Babraham Institute, UK).
Published September 28, 2020
The Henikoff lab has recently developed two new chromatin profiling methods: Cleavage Under Targets and Release Using Nuclease (CUT&RUN) and Cleavage Under Targets and Tagmentation (CUT&Tag) (Kaya-Okur et al ., 2019; Skene et al ., 2018; Skene and Henikoff, 2017). These techniques provide an exciting advance because they overcome many of the drawbacks of conventional and widely used chromatin immunoprecipitation (ChIP) methods.
CUT&RUN is a genome-wide extension to chromatin immunocleavage (ChIC), which is a method developed by the Laemmli lab (Schmid et al ., 2004). ChIC uses a Protein A-MNase fusion protein to cleave DNA regions associated with target proteins and recognized by specific antibodies. However, ChIC is limited to loci-specific analysis using Southern blotting. A related method, chromatin endogenous cleavage (ChEC), uses a fusion between the protein of interest and MNase to analyze the protein’s binding sites genome-wide (Schmid et al ., 2004; Zentner et al ., 2015). An obvious drawback of this method is the need to generate specific fusion proteins for each protein of interest. CUT&RUN is a major advancement over these two methods as it uses a recombinant Protein A/G-MNase tethered to the location of a protein of interest by an antibody (Meers et al ., 2019; Skene and Henikoff, 2017).
Importantly, this method recovers MNase-digested fragments and therefore is compatible with a sequencing-based genome-wide analysis of protein occupancy and histone modification positioning. Advantages over ChIP-based methods include improved method simplicity achieved by the use of magnetic beads to immobilize nuclei, the compatibility with fresh and frozen tissue samples, and a shortened protocol (1–2 days) to generate material suitable for preparing DNA sequencing libraries. Furthermore, due to the in situ targeted cleavage of DNA on both sides of the protein of interest, only on-target DNA fragments are released from the nucleus and collected, leaving the off-target sequences behind. Thus, CUT&RUN produces very little background signal compared to ChIP. Since its development, CUT&RUN has been adapted for a variety of experimental setups, including automation for high-throughput epigenetic profiling (AutoCUT&RUN) (Janssens et al ., 2018), profiling of insoluble chromatin such as centromeric regions with CUT&RUN. Salt (Thakur and Henikoff, 2018) and CUT&RUN.ChIP for examining specific protein components within complexes released by CUT&RUN digestion (Brahma and Henikoff, 2019). Most strikingly, the low input requirements and the high signal-to-noise mean that CUT&RUN is compatible with single-cell analysis, for example, to investigate transcription factor occupancy in individual mouse embryo cells (Hainer and Fazzio, 2019).
CUT&Tag was published in 2019 by the Henikoff lab (Kaya-Okur et al ., 2019) as a variation of the CUT&RUN protocol that allows for quicker library preparation and easier automation. CUT&Tag uses antibody-guided tagmentation of native or lightly fixed chromatin to identify the location of target protein occupancy genome-wide. CUT&Tag uses a hyperactive Tn5 transposase preloaded with DNA adaptors and fused with Protein A. This fusion protein binds to primary antibodies, fragments the DNA in the vicinity of the antibody and inserts short tags with sequence adaptors (tagmentation). After recovering the tagmented DNA fragments, PCR amplification uses primers, which recognize sequences within the added tags, to generate next-generation sequencing (NGS) libraries. The frequency of sequence reads at a particular region corresponds to the location of target protein occupancy or histone modifications.
CUT&RUN and CUT&Tag are simple, versatile, and powerful methods to profile DNA-protein interactions and should be in every molecular biologist’s toolkit. Both methods can aid the genome-wide identification of specific gene and cis-regulatory elements marked by histone modifications or bound by a protein of interest, thereby providing insight into chromatin-based mechanisms of gene control. Due to the very low amount of starting material needed, CUT&RUN and CUT&Tag are uniquely positioned to investigate rare cell types. On top of that, the easy-to-use and time-saving protocol allows for a quick turnaround, enabling the researcher to do multiple parallel analyses and, therefore, get a more comprehensive insight into the complexity of genome regulation.
Although in its infancy, CUT&RUN has been used to profile DNA-protein interactions in a variety of cell lines and tissue samples and a multitude of model organisms, including yeast, plants, and animal cells. So far, all these studies focused on mapping histone modifications and transcription factor binding sites. CUT&Tag has been reported for the analysis of histone modifications, RNA Polymerase II, and transcription factors both in low cell number samples and single cells (Kaya-Okur et al ., 2019).
The first report of CUT&RUN profiled the histone modification H3K27me3 in a human cell line and H2A in yeast cells (Skene and Henikoff, 2017), and similarly, CUT&Tag was first used to map histone modifications (H3K27me3, H3K27ac, H3K4me1, H3K4me3, H3K4me3) in human cell lines (Kaya-Okur et al ., 2019). Comparison of the equivalent data sets generated by ChIP-Seq, CUT&RUN, and CUT&Tag demonstrated close similarities between the three different chromatin-profiling techniques in discovering ‘peak’ regions. However, due to the much-improved signal-to-noise ratio in CUT&RUN and CUT&Tag compared to ChIP, substantially fewer sequencing reads were required, and the new methods are more sensitive and provide base-pair resolution (Kaya-Okur et al ., 2019).
Over the last decade, ChIP-seq has been the predominant method to identify the genome-wide location of transcription factor binding sites. CUT&RUN and CUT&Tag have similar capabilities, but compared to ChIP, these new methods are more cost-effective, quicker to use, require only a fraction of starting material including single cells, have a more favorable signal-to-noise ratio, detect more defined ‘peaks’, and are compatible with automation. Both methods are compatible with unfixed, native chromatin, even for transcription factor profiling, and therefore overcome some of the difficulties associated with cross-linked ChIP protocols. CUT&RUN and CUT&Tag have been used to profile multiple transcription factors, such as CTCF and pluripotency factors, and large chromatin-associated complexes, such as Polycomb Repressive Complexes and chromatin remodellers.
|Starting material||Millions of cells in a standard protocol, with some low-input protocols reported|
No more than 500,000 cells, with a standard of 50,000 cells. Availability of low input and single-cell protocols, compatible with automation.
No more than 500,000 cells, with a standard of 50,000 cells. Availability of low input and single-cell protocols, compatible with automation.
Standard for cross-linked protocols with formaldehyde, sometimes double fixation with DSG and formaldehyde
|Not necessary or recommended||Not necessary or recommended|
|Nuclei isolation||Recommended in most protocols||Not necessary but compatible with methods||Not necessary but compatible with methods|
Used in most cross-linked protocols to fragment and solubilize the chromatin
|Not necessary||Not necessary|
|Lysis||Takes place before antibody-based immunoprecipitation||Takes place after antibody guided DNA fragmentation by MNase||Takes place after antibody guided DNA tagmentation by Tn5|
No enzymatic cleavage in conventional cross-linked protocols, only in the native protocol using MNase to fragment DNA before immunoprecipitation
|In situ DNA fragmentation using MNase or target regions identified with a specific antibody||In situ DNA tagmentation using Tn5 or target regions identified with a specific antibody|
|Protocol length||About 1 week||Generally, 1 to 2 days||Generally, 1 to 2 days|
|Secondary antibody||Not generally used||Not generally used, unless necessary due to low abundant target epitope||Generally used as bridging antibody between specific antibody and Protein A-Tn5 fusion protein|
|Sequencing library prep||End repair and adaptor ligation protocols necessary|
End repair and adaptor ligation protocols for low-input DNA samples necessary
|Single PCR amplification due to adapter integration by the used Tn5|
|Sequencing depth||~ 20 million reads||~ 8 million reads||~ 2 million reads|
|Cost||Relatively high cost per sample||Low cost per sample||Low cost per sample|
During CUT&RUN, an antibody against the protein of interest (such as a histone with a specific modification or a transcription factor) is used to guide Protein A/G-tagged
MNase to the region in the genome where the protein is located. Activating the MNase then excises only those DNA sequences that are close to the protein of interest, and the small fragments are released from the nucleus. These DNA fragments are collected and used to generate NGS libraries with a low-input DNA library prep kit.
Figure 1. Schematic of the CUT&RUN protocol. Nuclei are attached to magnetic Concanavalin A beads to allow ease of handling and safe liquid removal after each wash step. Nuclei are permeabilized and simultaneously incubated with an antibody against the protein of interest. Protein A/G-MNase fusion protein binds to the antibody against the protein of interest. When Ca2+ is added, the MNase cleaves the DNA on both sides of the formed complex and release DNA fragments that diffuse out of the nucleus. The DNA can be extracted and used in an end-repair and adapter ligation-based DNA library preparation. NGS informs the binding profile of the protein of interest via the frequency of sequences in a particular region.
CUT&RUN typically uses fresh, unfixed samples as starting material, but protocol adjustments can allow the use of samples cryopreserved in 10% DMSO (Janssens et al ., 2018; Skene et al ., 2018).
Very little material is necessary to perform the protocol, and the general recommendation is to start with less than 500,000 cells (mammalian).
Cells or nuclei released with the Henikoff recommended nuclear extraction buffer are bound to Concanavalin A magnetic beads, which have unique saccharide-binding properties. Other nuclear extraction protocols can be used providing their compatibility (see suggestions below about the use of Triton X-100 containing buffers). Using magnetic stands allows for easy washing of the samples during the rest of the protocol.
The nuclei bound to Concanavalin A beads are permeabilized and simultaneously incubated with an antibody against the protein of interest (primary antibody) in a buffer containing digitonin and EDTA. EDTA allows the rapid cessation of cell metabolism and thereby inhibits endogenous DNAse activity, preserving the chromatin and reducing overall background signal. The duration of this step can be adapted by the user; generally, ranging from 2 h to overnight. Antibody dilution of 1:100 or 0 .5 1 .0 µg is recommended as a starting point, but the amount of antibody used should be optimized. It is advised to always include positive (α-H3K27me3) and negative control (isotype control IgG) samples.
Of note, Protein A and Protein G have different binding efficiencies to different antibody species (see here). Even though antibody compatibility is increased by using Protein A/G over Protein A alone, it might be necessary to use a secondary antibody in some cases. A secondary antibody guides MNAse to the target area by increasing the number of Protein A/G binding areas and thereby helps to recover low abundant target sequences.
To allow Protein A/G-MNase to be directed to the antibody-bound genomic target regions, the fusion protein is diluted in a wash buffer containing digitonin and incubated with the nuclei. Unbound enzyme fusion is then washed away, and MNase activity is activated at 0°C by adding Ca2+. Even though the cleavage itself is not particularly temperature-sensitive, the subsequent diffusion of the cleaved DNA fragments is sensitive, and a temperature rise would result in higher background. The digestion time can be adjusted if the final material contains a disproportional amount of high molecular weight fragments.
The MNase activity is stopped by the addition of a STOP buffer containing EGTA. This buffer can optionally contain heterologous spike-in DNA to help calibrate the CUT&RUN profiles during data processing. The MNase generated fragments are released from the nuclei by increasing the incubation temperature and cleaned up using phenol/ chloroform/isoamyl alcohol extraction followed by ethanol precipitation.
A variation of the protocol can further restrict the premature release of the MNase-fusion protein from its binding site, its diffusion, and potential for non-specific cleavage.
This version of the protocol uses a combination of low salt buffers and a high Ca2+ concentration to activate MNase activity and is ideal for targets mainly found in active open chromatin, but it can also be used when an antibody shows high levels of background signal.
Following clean-up of the cleaved DNA fragments, standard end repair and adapter ligation methods are used to generate low input DNA libraries according to the manufacturer’s guidelines. The Henikoff lab originally recommended a TRUseq library preparation approach, and besides, there have now been multiple reports using the NEBNext® Ultra™ II DNA Library Kit. Alternatively, similar to the automated CUT&RUN protocol (Jenssens et al . 2018), the phenol/chloroform/isoamyl alcohol extraction steps can be omitted, and the released DNA fragments can be directly used in the end repair and adapter ligation-based protocol.
The size distribution and concentration of libraries can be determined by running capillary electrophoresis (eg Bioanalyzer or TapeStation). Multiple libraries can be pooled together to obtain about 8 million paired-end sequencing reads per library. Due to the low background of CUT&RUN libraries, 8 million paired-end reads are enough to profile histone modifications and even transcription factors.
Most conventional ChIP-seq data analysis tools can analyze CUT&RUN data. There are a few analysis tools, such as the SEACR peak caller (Meers et al ., 2019b), which are specifically designed by the Henikoff lab for CUT&RUN data. For calibrated CUT&RUN, the heterologous spike-in DNA added with the STOP buffer can be used to normalize signals across samples. Alternatively, E. coli DNA carried over from the recombinantly-produced Protein A/G-MNase can be used as a spike-in.
During CUT&Tag, an antibody is used to guide Protein A-tagged Tn5 transposase to the area of the genome where the protein of interest (such as a histone with a specific modification or a transcription factor) is located. Tn5 fragments and tags target DNA with specific, pre-defined nucleotide sequences (adapters). The tagmented DNA can be easily recovered and PCR amplified to generate NGS libraries. The entire protocol, depending on the duration of antibody incubations, can be performed in just one day.
Figure 2. Schematic of the CUT&Tag protocol. Nuclei are attached to magnetic Concanavalin A beads to allow ease of handling and safe liquid removal after each wash step. Nuclei are permeabilized and simultaneously incubated with the primary antibody against the protein of interest before being incubated with a secondary antibody that recognizes the primary antibody. Protein A-Tn5 fusion protein binds to the antibody complex formed on the protein of interest. When Mg2+ is added, the Tn5 tagments the DNA on both sides of the formed complex, releasing tagmented DNA fragments that diffuse out of the nucleus. The DNA is extracted and used in a PCR amplification-based DNA library preparation. NGS informs the binding profile of the protein of interest via the frequency of sequences in a particular region.
The protocol can be separated into five parts:
CUT&Tag typically uses fresh, unfixed samples as starting material but the protocol has been adapted to allow the use of frozen samples and more recently for samples lightly fixed with formaldehyde after nuclei extraction. A fixation step also reduces the tendency of nuclei to clump together during the protocol. When using fixed and cryopreserved samples generated with buffers containing Triton X-100, bead clumping can be further reduced by removing digitonin from all buffers. It is of note, however, that epitope fixation can interfere with antibody binding in some cases.
The starting material is generally recommended to be below 500,000 cells (mammalian). For fresh or frozen tissue, nuclei preparation is similar to that in the CUT&RUN protocol (Janssens et al ., 2018; Skene et al ., 2018).
As for CUT&RUN, nuclei are bound to Concanavalin A magnetic beads, allowing for easy washing with the help of magnetic stands. However, this step can be omitted, and the whole protocol can be performed with gentle centrifugations after each wash step followed by careful removal of the supernatant without disturbing the pelleted nuclei.
The nuclei bound to Concanavalin A beads are simultaneously permeabilized and incubated with an antibody against the protein of interest (primary antibody) in a buffer containing digitonin. The duration of this step can be adjusted, starting from as little as 2 h up to 5 days. The recommended antibody dilution is typically between 1:50 and 1:100 (or 0 .5 - 1 .0 µg) but the amount of antibody used should be optimized. The inclusion of positive (α-H3K27me3) and negative control (isotype control IgG) samples is advised.
After primary antibody incubation and a quick wash with digitonin containing wash buffer, the nuclei are incubated with a secondary antibody that is directed against the primary and acts as a bridging antibody as well as increases the amount of Protein A binding sites. This step can be omitted in the CUT&RUN protocol but is required in the CUT&Tag protocol to increase signal. Of note, Protein A does not bind to all antibody classes with the same efficiency, therefore compatibility of the secondary antibody with Protein A needs to be checked beforehand.
For binding of the Protein A-Tn5 transposase to the antibody of interest, the adapter-loaded Tn5 fusion protein is diluted in a high salt buffer containing digitonin and is incubated with the nuclei. Increasing the salt concentration during this step helps to reduce off-target tagmentation, primarily of accessible genomic regions resulting in ATAC-like peaks.
After incubation with the Tn5 fusion, the nuclei are washed to remove unbound Tn5, and tagmentation is activated by incubation at 37 °C in the high salt digitonin buffer supplemented with Mg2+.
Tagmentation is stopped by adding EDTA, and the nuclei are lysed with SDS and Proteinase K. The DNA fragments can then be cleaned up in a variety of ways, with the original protocol recommending phenol/chloroform/isoamyl alcohol extraction followed by ethanol precipitation. Alternatively, AMPure Beads can be used to clean up the DNA fragments. It is recommended to avoid using any carrier, such as glycogen, in the precipitation step as it can reduce the efficiency of the following PCR reaction.
As the Tn5 introduced compatible sequences into the DNA fragments during the tagmentation process, a simple PCR reaction using a universal i5 primer and barcoded i7 primer can generate sequencing libraries, thus making DNA library preparation extremely quick and easy. Due to the similarities with ATAC-seq, the primer sequences described in the ATAC protocol can be used (Buenrostro et al ., 2013). The cycling program needs to be adjusted for the user’s machine. A very short annealing time is necessary for the PCR to work, and in slow ramping cyclers, the annealing step can be omitted because the cool-down period between denaturation and elongation is long enough for annealing to occur. When using a fast ramping machine, an annealing step must be included. In general, it is recommended to not exceed 12 to 14 amplification cycles, otherwise, the complexity of the library will be reduced together with high levels of PCR duplication.
Capillary gel electrophoresis (eg Bioanalyzer or TapeStation) allows the evaluation of the CUT&Tag library after PCR amplification. Failed CUT&Tag experiments are often characterized by a lack of nucleosomal laddering in the positive control sample.
However, observing only a very weak signal in a transcription factor CUT&Tag library is common, and if that happens, it is still worth proceeding. After evaluation by gel electrophoresis, the library can be cleaned up and concentrated using SPRI beads. Multiple libraries can be pooled together to obtain about 2 million paired-end sequencing reads per library.
Most conventional ChIP-seq data analysis tools can analyze CUT&Tag data as well. There are a few analysis tools, such as the SEACR peak caller (Meers et al ., 2019b), which are specifically designed by the Henikoff lab for CUT&RUN and CUT&Tag data. To calibrate CUT&Tag data, E. coli DNA carried over from the recombinantly-produced Protein A-Tn5 can be used just like an ordinary spike-in.
In general, sample preparation is straightforward for CUT&RUN and CUT&Tag as both methods can use fresh, unfixed samples. A single-cell suspension needs to be achieved in a way that is suitable for the cell type used. This can include using dissociation reagents like Accutase™, scraping cells off cell culture dishes, or mechanically dissociating tissues.
For ease of collection, samples can be cryopreserved in 10% DMSO and frozen down in a Mr. Frosty isopropyl alcohol chamber. Sample fixation is not required for either of the methods, however, if the beads clump during washes, then a light fixation of the sample using 0 .1% formaldehyde for only 2 min at room temperature before antibody incubation is beneficial.
As for ChIP, not all antibodies will work for CUT&RUN and CUT&Tag. Most antibodies are not yet tested for compatibility with these newer methods, therefore testing and optimization will be required by the end user. ChIP-grade antibodies seem to largely work for CUT&RUN and CUT&Tag, especially when shown to work for native ChIP .
When trialing antibodies that are not labeled as ChIP grade, assuming specificity is fully characterized, then it’s good to start with choosing antibodies that recognize the native form of the protein of interest; for example, antibodies that have been shown to work in immunoprecipitation or immunocytochemistry. Equally, when considering the concentration of antibody to use in a CUT&RUN or CUT&Tag experiment, the concentration recommended for immunofluorescence-based methods seems to be a good starting point. Compatibility to Protein A and Protein G needs to be checked, and then appropriate secondary antibodies need to be used if necessary.
As with any other experiment type, including the correct controls is critical to ensure the experiment has worked as expected as well as to easily pinpoint areas to troubleshoot should the experiment fail.
Unlike ChIP, no input sample needs to be included in the protocols as a non-antibody guided MNase treatment or tagmentation would simply result in the identification of accessible chromatin. An IgG control should be included to get the background from the sample and set the baseline for the experiment. In terms of antibody controls, the Henikoff lab recommends using H3K27me3 as a positive control for CUT&RUN and CUT&Tag experiments. The use of total unmodified histone controls, such as total H3, can be considered to allow the proportional representation of histone modifications.
In the initial versions of the protocols, the digitonin concentration used in the wash buffers needed to be tested to ensure efficient permeabilization of nuclei. This test might not be necessary if you follow improvements to the protocols, specifically by adding NP40 to the wash buffers. However, the efficiency of nuclei permeabilization should be kept in mind when using either technique on new material.
It is important to titrate the amount of antibody used per reaction, starting with recommended dilutions for ChIP or immunofluorescence assays.
Primary antibody incubation for 1 h at room temperature is sufficient but this can be extended to 1 to 5 days in the cold room. This step can be optimized for the antibodies used in each experiment, keeping in mind that prolonged incubation might lead to an increase in background signal and, therefore, a less favorable signal-to-noise ratio overall.
The use of a secondary antibody is highly recommended for CUT&Tag and can be considered for CUT&RUN if experiencing low recovery with primary antibodies alone. Another reason to include a secondary antibody is to avoid an unfavorable pairing of Protein A/G with the primary antibody, which can be achieved by using a more favorable secondary antibody class.
MNase digestion and Tn5 tagmentation time can be adjusted depending on the protein of interest. Less abundant proteins may need more time to allow for the full recovery of all sites. It is important to keep in mind that prolonged digestion or tagmentation might result in untargeted cleavage and, therefore, higher background signals.
The number of cells used per experiment should not exceed 500,000 cells. Since the number of beads in the protocol is optimized for the use of 50,000 to 500,000 cells, there should be no need to increase the number of beads recommended in the protocol.
It is recommended to use 10% DMSO in an appropriate buffer or media and slowly freeze using a Mr. Frosty isopropyl alcohol chamber. Flash freezing is not recommended.
Depending on the fixation and cryopreservation conditions used for the ChIP sample, it might be possible. However, it is unlikely to work with the standard protocols because cell numbers for ChIP samples are likely to exceed recommendations for CUT&RUN and CUT&Tag samples and the strong fixation conditions (eg double fixation and quenching) used will impair MNase or Tn5 activity. The problem of cell numbers in ChIP samples exceeding the recommended cell starting material for CUT&RUN or CUT&Tag can be easily circumvented by splitting the sample into multiples. However, the other problems of fixation and freezing of samples remain. Fixation used in ChIP protocols is likely too harsh to be compatible with either CUT&RUN or CUT&Tag, as it will most likely impair the ability of the MNase or Tn5 to fragment/tagment the target sequences, and multiple reports have hinted at this incompatibility. Additionally, there is a possibility of epitope masking due to fixation. Sample fixation is, therefore, not recommended. When cryopreserving cells for CUT&RUN or CUT&Tag, the use of 10% DMSO and a Mr. Frosty isopropyl alcohol chamber to gently freeze the samples is required. Flash freezing included in most ChIP protocol is not recommended.
The problem of clumping can be caused by using too high a ratio of cells to beads or the lysis of nuclei/cells in the digitonin containing buffer, which would release DNA and lead to clumping. The first thing would be to check that the recommended number of cells is not exceeded. Furthermore, a light fixation with formaldehyde (0 .1% formaldehyde for 2 min at room temperature) before incubation with the antibody is recommended to reduce the clumping of beads in digitonin containing wash buffers. However, keep in mind that fixation could affect epitope availability and, therefore, will need to be tested for each antibody.
The addition of EDTA in the permeabilization and antibody incubation buffer is recommended as it chelates Mg2+ and thereby stops any ATP-dependent cellular processes, including replication and chromatin remodeling, and it will also stop endogenous DNases.
Both fusion proteins were initially provided by the Henikoff lab. There are protocols on how to make your own fusion proteins (Kaya-Okur et al ., 2019; Meers et al ., 2019a), whereas the plasmids, Protein A-MNase, Protein A/G-MNase, and Protein A-Tn5 are all commercially available from reagent suppliers.
Increasing the salt concentration in the tagmentation buffer as recommended in the standard protocol should solve this problem. In some cases, depending on the protein of interest, there could be artefactual detection of what looks like ATAC peaks as shown in Kaya-Okur et al 2019.
Buenrostro, J .D ., Giresi, P .G ., Zaba, L .C ., Chang, H .Y ., Greenleaf, W .J . Transposition of native chromatin for fast and sensitive epigenomic profiling of open chromatin, DNA-binding proteins and nucleosome position. Nat Methods 10, 1213–1218 (2013).