CUT&RUN for Chromatin Profiling in Caenorhabditis elegans

Abstract

Cleavage Under Targets and Release Using Nuclease (CUT&RUN) is a recently developed chromatin profiling technique that uses a targeted MNase cleavage strategy to obtain high-resolution binding profiles of protein factors or for the mapping of histones with specific post-translational modifications. Due to its high sensitivity, CUT&RUN allows for quality binding profiles to be obtained with only a fraction of the starting material and sequencing depth typically required for other chromatin profiling techniques such as Chromatin Immunoprecipitation (ChIP). Although CUT&RUN has been widely adopted in multiple model systems, it has rarely been utilized in C. elegans, a model system of great importance to genomic research. Cell dissociation techniques, which are required for this approach, can be challenging in C. elegans due to the toughness of the worm’s cuticle and the sensitivity of the cells themselves. Here, we describe a robust CUT&RUN protocol for use in C. elegans to determine the genome-wide localization of protein factors and specific histone marks. With a simple protocol utilizing live, un-crosslinked tissue as starting material, performing CUT&RUN using worms has the potential to produce physiologically relevant data at a higher resolution than ChIP. Our protocol involves a simple dissociation step to uniformly permeabilize worms while avoiding sample loss or cell damage, resulting in high quality CUT&RUN profiles with as few as 100 worms and detectable signal with as few as 10 worms. This represents a significant advancement for the C. elegans field, which typically uses thousands or hundreds of thousands of worms for a single ChIP experiment. With a detailed description of worm growth, sample preparation, CUT&RUN workflow, library preparation for high-throughput sequencing, and a basic overview of data analysis, the protocols described here make CUT&RUN simple and accessible for any worm lab.

Basic Protocol 1: Growth and synchronization of C. elegans

Basic Protocol 2: Worm dissociation and sample preparation

Basic Protocol 3: CUT&RUN chromatin profiling

Alternate Protocol 1: CUT&RUN with secondary antibody incubation

Basic Protocol 4: CUT&RUN library preparation for Illumina high-throughput sequencing

Basic Protocol 5: Sample basic data analysis using Linux

INTRODUCTION:

Methods to study the interaction between DNA and chromatin-associated proteins are critical for an in-depth understanding of chromatin biology. The genome-wide DNA binding profile of histone marks, transcription factors, and chromatin factors can reveal not only mechanisms of transcriptional regulation, but also complex interactions between chromatin-associated proteins, for instance, whether a certain transcription factor is necessary for the recruitment of a chromatin modifier. The most commonly used approach to obtain such profiles in a genome-wide manner is Chromatin Immunoprecipitation followed by sequencing (ChIP-seq), which remains a staple in the chromatin biology field to this day, and has been widely applied in organisms from yeast to humans (Ostrow et al., 2015; Chen et al., 2018; Pu & Lee, 2020; Sen et al., 2021; Tran et al., 2012; Soares & Castro, 2018; Sullivan & Santos, 2020). ChIP approaches, however, are often limited in their resolution, requiring both large amounts of chromatin as starting material and high sequencing depth to obtain a good signal-to-noise ratio. Additionally, ChIP protocols typically include a crosslinking step, which can lead to artifacts in the sequencing analysis (Baranello et al., 2016). ChIP protocols themselves are often technically difficult to perform and require specialized equipment and expertise.

Cleavage Under Targets and Release Using Nuclease (CUT&RUN) is a chromatin profiling technique recently developed by the Henikoff Lab (Skene & Henikoff, 2017). The approach allows for high-resolution genome-wide localization information to be obtained for protein factors and histone modifications from minimal starting material without crosslinking. In CUT&RUN, live cells or tissue are used as starting material. Cells are then mixed with beads coated with the lectin Concanavalin A (ConA), which binds to glycoproteins on the cell surface and captures the cells, allowing for easy wash steps and minimal sample loss throughout the procedure. Cells are then incubated with an antibody to target the chromatin-bound protein of interest (e.g. transcription factor or specific histone modification). After primary antibody incubation, cells are incubated with a Protein-A-Protein-G-MNase (pAG-MNase) fusion protein which, through the binding properties of proteins A and G, binds the target antibody. Upon the addition of calcium, the MNase cleaves DNA at the specific sites where it is bound, resulting in the production of cleaved DNA fragments corresponding to the binding sites of the protein of interest. Cleaved DNA fragments are released from the nucleus and float out of the cells during a 37 °C incubation period, leaving uncut genomic DNA behind to be discarded with cellular debris. The cleaved DNA fragments of interest are then isolated and subjected to library preparation for high-throughput sequencing to reveal the specific genome-wide localization information for the chromatin-associated protein being surveyed (Skene & Henikoff, 2017) (See Figure 1 for overview). Because only DNA regions of interest are cleaved and purified in this procedure, CUT&RUN experiments can result in data with a high signal-to-noise ratio, with only 5 million sequencing reads required for mammalian cells (Skene & Henikoff, 2017). In contrast, a typical ChIP experiment, which involves sonicating the entire genome before immunoprecipitation, requires a minimum of 20 million reads for high quality results (Landt et al., 2012).

Figure 1. CUT&RUN workflow described in this article.

To perform CUT&RUN using C. elegans, (Top Left) synchronized worms are grown and (Top Middle) dissociated into a worm chunk/cell mixture using a Cuticle Disruption Buffer and mechanical douncing in Wash Buffer. The worm/cell mixture is (Top Right) bound to Concanavalin A-coated (ConA) beads to facilitate the ease of the CUT&RUN protocol. The worm/cell mixture is permeabilized with digitonin and (Middle Left) incubated with a primary antibody against the chromatin-bound protein of interest. After primary antibody binding, (Middle) worms are incubated with a pAG-MNase fusion protein, which binds to the primary antibody. With the addition of calcium (Middle Right), MNase cleaves DNA at the specific sites surrounding the protein of interest, and (Bottom Left) after releasing cleaved fragments to flow out of the cells at 37 °C, DNA is isolated and subjected to library preparation for (Bottom Middle) high-throughput sequencing. The resulting DNA sequencing files are (Bottom Right) analyzed to identify regions of enrichment, corresponding to the binding profile of the chromatin-bound protein of interest. Circled numbers correspond to the Basic Protocol number in which the pictured step is performed.

The CUT&RUN workflow is simple and reproducible, and has been used successfully in many model organisms, including Saccharomyces cerevisiae and Drosophila melanogaster, and in human cells (Skene & Henikoff, 2017; Skene et al., 2018), all with well-documented protocols available, especially for use in mammalian cells (Janssens & Henikoff, 2019; Hainer & Fazzio, 2019; Liu, 2021). However, the technique has not been broadly adopted for use in the important model organism Caenorhabditis elegans —having only recently been utilized for the first time (Methot et al., 2021) —, likely because of the lack of a detailed protocol and the difficulty of dissociating worms into healthy cells. C. elegans is a widely used model organism for studying development, aging, behavior, gene regulation, chromatin biology, and more (Corsi et al., 2015; Sulston & Horvitz, 1977; Kenyon, 2005; González-Aguilera et al., 2014; Fischer, 2010; Rechavi & Lev, 2017; Sengupta & Samuel, 2009; Caldwell et al., 2020; Wenzel et al., 2011), and detailed studies of chromatin biology in these contexts would be instrumental. ChIP-seq has been previously applied to C. elegans in numerous studies, however, ChIP approaches are notoriously challenging in C. elegans because of the animal’s touch cuticle, which makes permeabilization difficult, and relatively low DNA content per body weight of C. elegans, limiting the amount of DNA available for library preparation (Sen et al., 2021). In practice, this means that for one ChIP experiment, researchers typically must start with thousands, or even hundreds of thousands, of worms to obtain enough DNA for sequencing. This may limit the types of experiments amenable to ChIP approaches, as certain samples are difficult to collect in large numbers, for example, aged worms, balanced strains, sorted worms, or dissected tissue. Because CUT&RUN allows for high-resolution binding profiles from limited starting material, the C. elegans community could greatly benefit from adopting this approach.

Here, we describe a protocol for CUT&RUN in C. elegans, which can produce high quality genome-wide profiles of histone marks with as few as 100 worms, and can generate detectable signal with as few as 10 worms. We describe five in-depth protocols to adapt the CUT&RUN approach to C. elegans. In Basic Protocol 1, we describe our method for growing synchronized worm populations for use in CUT&RUN. In Basic Protocol 2, we describe the steps for worm dissociation and sample preparation that are necessary before starting a standard CUT&RUN protocol. This step is critical for C. elegans, as the worm’s notoriously tough cuticle must be softened, and worms must be somewhat dissociated, to ensure equal permeabilization by antibodies. In contrast to traditional nuclei or cell isolation approaches for CUT&RUN sample preparation, we describe a simple approach for gently permeabilizing worms that results in limited cell loss, allowing for maximum sample recovery from very few worms. In Basic Protocol 3, we describe the CUT&RUN protocol itself. Our protocol closely follows the traditional Henikoff Lab CUT&RUN protocol developed for cells (Skene et al., 2018; Janssens & Henikoff, 2019), with the exception that we typically perform the protocol with a combination of isolated cells and worm chunks, for convenience and sample preservation. In Basic Protocol 4, we describe library preparation of CUT&RUN fragments and quality control checks for Illumina sequencing. For library preparation, we and others (Liu et al., 2018; Liu, 2021) utilize the NEBNext^® Ultra^™ II DNA Library Prep Kit for Illumina^® with a few minor adaptions for the low input material and short fragments found in CUT&RUN. In Basic Protocol 5, we describe a general guideline for CUT&RUN data analysis using Linux to visualize the data and determine whether the process was successful.

With these protocols, we hope to provide users with an accessible way to introduce CUT&RUN into their own C. elegans labs, and help facilitate future studies of chromatin-associated proteins in worms.

BASIC PROTOCOL 1: Growth and Synchronization of C. elegans

Here, we describe our standard growth and synchronization protocol to prepare worms for use in CUT&RUN. Because of the high signal-to-noise ratio obtained from CUT&RUN, most CUT&RUN reactions probably only require 100 worms or fewer. However, we typically recommend growing 1,000–3,000 worms for each antibody being used for the first time, as we have found that this number of worms is sufficient for high quality DNA binding profiles for both chromatin factors and histones harboring specific post-translational modifications. Once you are confident the reaction is working, the worm number could be decreased down to 100 worms, or even fewer. The method of worm synchronization described here is the one that is the simplest and most convenient for us, but individual labs may prefer to prepare worms differently depending on their standard practices or the specific regimens required for their experiments. For instance, RNAi bacteria can be substituted for OP50, or precise egg lays can be done when limited plate density and exact synchronization are of concern.

Materials

Worm strain(s) for use in CUT&RUN (Standard wild-type N2 or any other strain of interest. Strains can be ordered from the Caenorhabditis Genetics Center (CGC) at https://cgc.umn.edu/)

NGM plates (6-cm and 15-cm) (see Reagents and Solutions) seeded with OP50-strep resistant Escherichia coli (see Reagents and Solutions)

Sterile M9 buffer (see Reagents and Solutions), room temperature

Sterile M9 buffer + 0.05% (v/v) Tween-20, room temperature. To prepare, add 250 μL of Tween-20 (e.g. Fisher Scientific, cat #BP337-500) to 500 mL of M9 Buffer

Worm pick

Standard stereomicroscope (e.g. Olympus SZ51)

Worm incubator set at desired growth temperature (typically 20 °C, but 15–25 °C can also be used) (e.g. Percival Scientific, Cat #I-36NL)

Standard benchtop pipette set (P20-P1000)

1.5-mL tubes, DNase- and RNase-free (e.g. LPS, cat #L211511-CS)

Mini centrifuge (e.g. Fisher Scientific, cat #12-006-901)

Laminar flow hood (e.g. Nuaire, AireGard NU-340)

Protocol Steps

Growth of Synchronized Worms

• 1From a well-fed normal maintenance plate of your desired worm strain, pick 3–7 gravid adults to a new 6-cm NGM plate seeded with 0.2 mL of OP50. Incubate overnight for 12–20 hours at the desired growth temperature, typically 20 °C.

  For many experiments, you may desire to use multiple strains, for instance, if comparing binding profiles between wild-type worms and mutants. If you are obtaining a binding profile for a chromatin factor or transcription factor using a tagged-strain (e.g. DAF-16::GFP), we also recommend processing N2 wild-type worms in parallel, so that the antibody background can be obtained in a strain lacking the epitope tag (i.e. anti-GFP antibody in N2 worms). When beginning Step 1, you should grow all strains that you plan to use for the experiment in parallel.

  The number of gravid adults chosen depends on the worm strain used. For wild-type N2 worms, 3 gravid adults is more than enough, but for worms with a reduced brood size, you may wish to increase the number of worms initially picked. For a typical experiment with N2 worms, one plate will be enough, but if large numbers of worms are desired for CUT&RUN (>6,000, for example, when using one genotype for more than 6 antibodies against histone marks or more than 2–3 antibodies against chromatin/transcription factors), consider preparing multiple plates at this stage, each with 3–7 worms.

• 2The next day, remove the gravid adults from these plates and return plates to the incubator. The plates should now contain hundreds of embryos and young larvae.

  This ensures that embryos are relatively synchronized, and plates do not get overcrowded and starve before step 3.

• 3Grow embryos until they reach Day 1–2 of adulthood and the plates are again full of embryos. Ensure the worms are well fed during this time.

  The time required for this step depends on the temperature at which the worms are kept, but typically, at 20 °C and with N2 worms (see below), it will take 4 days from the initial picking of 3–7 gravid adults (step 1) to the stage when there are thousands of relatively synchronized embryos (the end of step 3). At 16 °C, in our hands, this typically takes ~6 days for most strains.

  The time required for this step will also depend on the worm strains used, since some strains will grow more slowly than others. The timing mentioned above is for wild-type N2 worms.

Embryo Collection

• 4Wash adult worms off of the full plates using room temperature, sterile M9 buffer. Do this by pouring or pipetting 1–2 mL of M9 onto the edge of the worm plates, but be careful not to add it directly to the bacterial lawn, to avoid removing the embryos, which will be primarily present on the bacterial lawn. Tilt the plate to cover the entire plate with M9, then tilt the plate to one side and either pour off the adults into a waste beaker or remove them by pipetting. Discard the adult worms.

  Only adult worms and larvae will begin to thrash in the M9, allowing for adults to be easily removed from the plate and leaving behind embryos which stick to the bacterial lawn.

  If tighter synchronization is needed, for example, in a study across precise developmental stages, consider doing a large-scale synchronized egg-lay for 3–4 hours using the adults obtained in step 4 rather than discarding them.

• 5Repeat Step 4 one or two additional times until you can see under the microscope very few, if any, adult worms remaining on the plates.
• 6Pipette 750 μL of M9 buffer directly onto the bacterial lawn and use this liquid to forcefully resuspend the bacterial lawn via vigorous pipetting. Pipette the liquid containing the bacteria and embryos into a sterile 1.5-mL tube. If you are using multiple genotypes of worms, be careful to keep the embryos from distinct genotypes in separate tubes.

  This will release the embryos that stick to the bacterial lawn, allowing you to collect large populations of relatively synchronized embryos.

• 7Repeat step 6 and transfer the embryos to the same 1.5-mL tube.
• 8Quick-spin collected embryos in a mini centrifuge for ~5 seconds. Remove the supernatant and resuspend the embryo pellet in 1 mL of M9 buffer supplemented with 0.05% (v/v) Tween.

  Tween supplementation prevents worms or embryos from sticking to the plastic pipette tip, which can occur once the bacteria is removed and prevents accurate counting in step 11 and leads to worm loss.

• 9(Optional) Wash embryos in M9 buffer supplemented with Tween another 1–3 additional times to remove bacteria, spinning for ~5 seconds in between each wash. This is especially recommended if embryos will be seeded on a plate containing different bacteria in the next step (for example, RNAi plates).

Seeding Embryos for CUT&RUN

• 10Resuspend embryos thoroughly by pipetting the entire volume of liquid with a 1-mL pipette several times.

  This is essential to ensure accurate counting and avoid embryo clumping, which sometimes occurs after spinning.

• 11Pipette 5–10 μL of the embryo suspension onto a plate and count the embryos present. If you are using multiple genotypes of worms, repeat this on separate plates for each genotype.

  Sometimes, a few L1 stage larvae will also be present after wash steps. Worms will still be relatively well synchronized, as they will reach adulthood on the same day. If older larvae or adults are still present in the embryo mixture, settle the worms for 1–3 minutes by placing the tube in a normal microcentrifuge rack. Adults and older larvae will visibly settle to the bottom of the tube after a few minutes, and most embryos will not. Keep the top of the embryo suspension and re-count the embryos present in 5–10 μL before proceeding.

• 12Seed 15-cm bacteria-containing plates with the desired number of worms for each genotype, estimating the embryos needed based on the counts above. Typically, aim for up to 3,000 worms per 15-cm plate. Depending on the number of CUT&RUN reactions desired (i.e. the number of antibodies planned for each genotype) and the target of interest, consider seeding multiple plates with 3,000 worms.

  For example, if you count 50 embryos in 5 μL, you will need to seed 300 μL of embryo suspension to obtain roughly ~3000 Day 1 adult worms for CUT&RUN per plate.

  Although CUT&RUN can work well with as few as 100 worms, for an initial trial, we recommend using ~1,000–3,000 worms for histone marks and up to ~3,000 worms for transcription or chromatin factors, to ensure high enough DNA yield.

  The number of worms seeded per plate will also vary depending on the age of the worms to be collected for CUT&RUN. One 15-cm plate easily feeds up to 3,000 worms until Day 1 of adulthood, however, it would not feed them well after Day 1. Alternatively, if larval stage worms are desired for CUT&RUN, more than 3,000 worms can be seeded per plate since larvae are less likely to starve the plate. Avoid seeding too many worms (>4,000 for Day 1 adult growth) per plate, as this can lead to overcrowding and starvation.

  Depending on the experimental design, you can seed worms on plates containing any desired bacteria, such as RNAi plates.

• 13Allow plates to dry for a few minutes in the hood. Incubate at the desired temperature until Day 1 of adulthood (typically ~3 days at 20 °C, ~4 days at 16 °C, or ~2 days at 25 °C, with N2) or until desired developmental stage, then proceed to Basic Protocol 2.

BASIC PROTOCOL 2: Worm Dissociation and Sample Preparation

Here, we describe how to dissociate the worms prepared in Basic Protocol 1 for use as starting material for CUT&RUN. Worms are washed off of plates and exposed to a buffer containing SDS and DTT to weaken their cuticle, as previously described (Zhang et al., 2011; Kaletsky et al., 2016; Kaletsky & Murphy, 2020; Methot et al., 2021). Then, after washing out the SDS and DTT buffer, weakened worms are gently dounced directly in CUT&RUN Wash Buffer. The result is a mixture of isolated cells and worm chunks that are ready for use in CUT&RUN (Basic Protocol 3).

We also include four optional steps to test for uniform permeability of the worm/cell mixture, by using the fluorescent dye Propidium Iodide. Permeability of cells is an important factor for CUT&RUN, because cells must be permeabilized enough to allow for the antibody to flow into them (Basic Protocol 3). These optional steps can be used to optimize the dissociation protocol, which may be necessary if you are using worms of a different age or genetic background from those described here (i.e. wild-type worms at Day 1 of adulthood). The optimization steps are intended to be performed before a “real” CUT&RUN reaction, thus, if optimization is desired, it is recommended to follow Basic Protocol 2 one time as a trial and test for permeability, without proceeding to CUT&RUN in Basic Protocol 3. After determining the optimal conditions, users can return to Basic Protocol 1 and grow new worms that can be used for a CUT&RUN experiment (by leaving out the optional steps in Basic Protocol 2 and proceeding directly to Basic Protocol 3).

Materials

Worms prepared in Basic Protocol 1, Step 13

Sterile M9 Buffer (see Reagents and Solutions), room temperature

Sterile M9 Buffer + 0.05% (v/v) Tween-20, room temperature. To prepare, add 250 μL of Tween-20 (e.g. Fisher Scientific, cat #BP337-500) to 500 mL of M9 Buffer

Cuticle Disruption Buffer (see Reagents and Solutions), room temperature

Wash Buffer (see Reagents and Solutions), room temperature

(Optional for Optimization) Optimization Digitonin-Wash (Dig-Wash) Buffer, containing 0.02% (w/v) digitonin (see Reagents and Solutions), 4 °C

(Optional for Optimization) Propidium Iodide (PI) Solution, 1 mg/mL (see Reagents and Solutions)

Sterile 50-mL tubes (e.g. VWR, Cat #89039-660)

Water aspirator or other aspirator. Pipettes can be used if not available.

Low-bind 1.5-mL tubes (e.g. Fisher Scientific Low Retention tubes, cat #02-681-320)

Standard pipettes and tips (P1000)

(Optional) Tube nutator (e.g. VWR, cat #10127-872)

Mini centrifuge (e.g. Fisher Scientific, cat #12-006-901)

Tissue grinder (pestle) for 1.5-mL tubes, one for each genotype (e.g. Fisher, cat #12-141-363)

(Optional for Optimization) Fluorescent microscope (e.g. Leica DM5000 B)

Protocol Steps

Collecting Worms for CUT&RUN

• 1Wash synchronized worms from Basic Protocol 1 into sterile 50-mL tubes using M9 + 0.05% (v/v) Tween, using one 50-mL tube per genotype. To do this, pour ~5 mL of M9+0.05% (v/v) Tween onto each 15-cm plate, avoiding the bacterial lawn. Tilt the plate to cover the entire plate with liquid and collect the worms at one corner. Carefully tilt the 15-cm plate to pour the worms into a 50-mL tube.

  M9+0.05% (v/v) Tween should be used, to prevent worms from sticking to the tube, or, in later steps, to the pipette tip.

• 2Repeat step 1 to ensure that the vast majority of worms are successfully washed off the plate.

  At this point, if you check the 15-cm plate after washing, you should see very few worms left (less than 10%). If too many worms remain, wash an additional time.

• 3Set the 50-mL tubes in a rack and allow worms to settle for ~5 min at room temperature. When all the worms settle at the bottom of the tube, remove the supernatant with a water aspiration system or similar, leaving ~2 mL or less behind, taking care not to aspirate the worms.
• 4Transfer worms into 1.5-mL low-bind tubes with a 1-mL pipette, using one 1.5-mL tube per genotype. Worm pellet should be between 50-and 250 μL.

  If more than 250 μL of worms are needed (i.e. if one genotype is being used for CUT&RUN reactions with many different antibodies), split the worms into multiple tubes containing less than 250 μL each and process them in parallel.

  Low-bind tubes should be used throughout the worm dissociation protocol to reduce worms and cells from sticking to the side of the plastic tube.

• 5Wash worms several times with 1 mL of M9+0.05% (v/v) Tween buffer each time, collecting worms by a 5 second quick-spin in a mini centrifuge in between washes.

  Wash until the supernatant appears clear of bacteria. This usually requires < 5 washes. Avoid over-washing, as this will lead to worm loss.

  If undesired embryos or larvae are present, take advantage of the tendency of adult worms to settle to the bottom of a tube before embryos and larvae. Allow the worm mixture containing adults, larvae, and embryos to settle in a 1.5-mL tube rack on the bench for 1–5 minutes, until ~95% of adult worms have settled to the bottom of the tube. Remove and discard the supernatant, which contains only larvae and embryos, while being careful not to remove the adults at the bottom of the tube. Add another 1 mL of M9 + 0.05% (v/v) Tween buffer and re-settle until the supernatant appears mostly clear of larvae (typically 3–5 times).

Cuticle Dissociation of Synchronized Worms

• 6Wash worms once in 1 mL of room temperature Cuticle Disruption Buffer. Invert 5 times to mix. Quick-spin worms for 5 seconds in a mini centrifuge to collect them at the bottom. Remove and discard supernatant.
• 7Add 1 mL of room temperature Cuticle Disruption Buffer and incubate at room temperature for 6–8 minutes. Gently invert the tube frequently during incubation, or incubate the tube on a nutating mixer at a low speed.

  This process will weaken the cuticle, and allows worms to be broken down more easily by douncing later on.

  During incubation, you should see the head of the worm become slightly blunt as a result of the weakened cuticle, and worms should mostly stop moving by the end of the incubation. Some worms will explode from their vulva, however, the majority of worms should not rupture at this step. Hold the tube under a microscope during the incubation or check a few microliters of the reaction on a slide, to check progress.

  Note that this step will need to be optimized for different genotypes and different stages of worms, as the cuticle changes with development and aging, and cuticle mutants may alter dissociation. The time given here (6–8 minutes) is for day-1 adult wild-type (N2) worms.

• 8Quick-spin 5 seconds in a mini centrifuge to collect worms and discard the Cuticle Disruption Buffer.
• 9Wash worms three times in 1 mL of M9 Buffer. To wash, add 1 mL of M9 Buffer, invert the tubes to mix, and quick-spin to collect worms before removing the supernatant. Work as quickly as possible to remove worms from any trace of the Cuticle Disruption Buffer.

  Typically, the supernatant appears mostly clear but with some bacteria, due to some number of worms exploding at this stage. If the supernatant is very cloudy, consider reducing the cuticle disruption time to reduce worm loss and possible cell damage.

Dounce Worms to Increase Permeability

• 10Wash worms twice in 1 mL of room temperature Wash Buffer. To wash, add 1 mL of Wash Buffer, invert the tubes to mix, and quick-spin to collect worms before removing the supernatant.
• 11Resuspend the worms in Wash Buffer, adding 150 μL of Wash Buffer per desired CUT&RUN reaction, or up to 500 μL total.

  For example, if you plan to use one genotype for three reactions (e.g. one reaction with an anti-H3K4me3 antibody, one reaction with an anti-H3K9me3 antibody, and another reaction with an anti-H3 antibody for normalization), resuspend the worms in 450 μL. If you plan to use one genotype for more than three reactions (i.e. with more than three different antibodies), resuspend the worms in just 500 μL of Wash Buffer.

• 12Gently dounce worms with a tissue grinder ~200 times, taking a break after each set of 50 dounces. Rotate between genotypes (if multiple genotypes are used) to let worms settle to the bottom.

  By the end of the 200 dounces, there will be a mix between whole worms, worm chunks, and worm cells (See Figure 2 for example). The solution should be cloudy with cells and bacteria. This cell/worm chunk mixture will be used as input material for CUT&RUN.

• 13Adjust the volume of Wash Buffer, if necessary, so that there is now approximately 150 μL of Wash Buffer per desired CUT&RUN reaction.

  For example, for one genotype that will be used in 8 CUT&RUN reactions (i.e. 8 antibodies), increase the volume from ~500 μL to ~1.2 mL.

  Unless absolutely necessary for experimental design (i.e. if adapting this protocol for use with an additional step such as FACS), do NOT spin after this step, as any spinning could lead to both a loss or damage of individual cells that have been released from the dissociated worms and to decreased signal in CUT&RUN or loss of signal from a specific cell population.

Figure 2. C. elegans are permeable to Propidium Iodide after cuticle disruption and douncing in Wash Buffer.

Differential Interference Contrast (DIC) (left) or fluorescent (right) images of Day 1 adult wild-type N2 worms stained with Propidium Iodide, a dye that only enters permeabilized cells. Worms were dissociated as in Basic Protocol 2, then incubated in Digitonin-Wash Buffer containing a final concentration of 0.01% (w/v) digitonin overnight at 4 °C. The next day, Propidium Iodide was added to a final concentration of 25 μg/mL and worms were imaged for red fluorescence. A and B represent examples of worms imaged at 100X total magnification, where both intact worms and worm chunks can be seen to be permeable to Propidium Iodide. C and D show worms imaged at 200X total magnification, where arrows represent examples of individual cells permeable to Propidium Iodide either inside larger worm chunks (white arrows) or free-floating in the dounced solution (orange arrows). # symbols indicate examples of embryos, which often do not stain with Propidium Iodide.

Optimization (Optional)

Steps 14–17 are optional for optimization purposes, and should be performed once prior to a real CUT&RUN protocol, if desired, to check for uniform permeability of the worm mixture. If this optimization is not desired/needed, proceed directly from step 13 to step 18 of Basic Protocol 2. If users, instead, will perform the optimization steps, stop the protocol after step 17 of Basic Protocol 2 and do not proceed to Basic Protocol 3 until optimal permeabilization has been achieved, and new samples of worms have been prepared (by repeating Basic Protocol 1) and then processed as in Basic Protocol 2 with the optimal conditions.

• 14To check if the worm mixture is uniformly permeabilized, add an equivalent volume of the Optimization Dig-Wash Buffer to the sample volume.

  For example, if the worms are in 150 μL of Wash Buffer, add 150 μL of Optimization Dig-Wash Buffer.

  The Optimization Dig-Wash Buffer is made with double the concentration of digitonin (0.02% (w/v)) used in the normal Dig-Wash Buffer (0.01% (w/v)) so that, when added in equal volume to Wash Buffer containing no digitonin, the final concentration of digitonin (0.01% (w/v)) will be the same as if worms were added directly into Dig-Wash Buffer in Basic Protocol 3. This is necessary, because, unlike during the real CUT&RUN protocol, we do not recommend using the ConA beads during the optimization process due to their high fluorescent properties, nor do we recommend spinning worms down and resuspending them in Dig-Wash Buffer, as that could lead to cell loss or damage.

• 15Mix well by pipetting and place the tube on a tube nutator at around 1.5–2 rpm overnight (12–18 h) at 4 °C.
• 16Add the Propidium Iodide Solution to the worm mixture for a final concentration of 25 μg/mL. Place the tube on a tube nutator at around 1.5–2 rpm and incubate for 10 minutes. Cover the tube during incubation so the sample is in the dark.

  For example, add 7.5 μL of the 1 mg/mL Propidium Iodide solution to a 300 μL sample for a final concentration of 25 μg/mL.

• 17Visualize red fluorescence with a fluorescent microscope, which will indicate the permeability of cells to Propidium Iodide. See Figure 2 for an example of worm chunks and cells nearly uniformly permeable to Propidium Iodide.

  Since Propidium Iodide only enters dead or permeabilized cells, red fluorescence will indicate the cells are permeable to the dye and permeable enough for use in CUT&RUN.

  If worm chunks are not uniformly permeabilized after the douncing protocol described above, try checking for permeability after a variable number of dounces (e.g. 100 dounces, 150 dounces, 200 dounces, 250 dounces, 300 dounces). The optimum number of dounces for CUT&RUN will be the minimum number needed to achieve nearly uniform permeabilization (See Figure 2 for example). Avoid over-douncing as this could lead to cell damage and high background in CUT&RUN.

• 18Proceed to Basic Protocol 3.

BASIC PROTOCOL 3: CUT&RUN Chromatin Profiling

Here, we describe the actual CUT&RUN protocol, which starts with C. elegans tissue from Basic Protocol 2 and ends with purified fragments of factor-bound DNA ready to be used in library preparation for high-throughput sequencing. In this protocol, cells and worm chunks are first bound to ConA beads, to aid in immobilization and wash steps, and then sequentially incubated with a) an antibody against the factor of interest, b) pAG-MNase, to bind the primary antibody, and c) calcium, to allow for pAG-MNase cleavage of DNA. Cells are then incubated at 37 °C to allow for release of cleaved DNA. Cleaved DNA is then purified and ready to be used in library preparation. Our CUT&RUN protocol closely follows the standard protocol for cells described by the Henikoff Lab (Skene et al., 2018; Janssens & Henikoff, 2019), with a few minor adjustments. For convenience, we perform the reactions in 0.2-mL PCR strip tubes to allow for multichannel pipetting during all wash and mixing steps. This allows for up to 16 samples to be easily and uniformly processed simultaneously.

Materials

Worm suspension from Basic Protocol 2

Activated Concanavalin A-coated (ConA) beads (Bangs Laboratories, cat #BP531) (see Reagents and Solutions), on ice

Antibody Buffer, containing 0.01% (w/v) digitonin (see Reagents and Solutions), 4 °C

Primary antibody against target of interest, for example transcription factor or histone mark, (e.g. anti-H3K4me3 antibody, Epicypher, cat #13-0041)

Control antibody against species-matched IgG or experiment-appropriate control like H3, (e.g. anti-H3 antibody, abcam, cat #ab1791)

Digitonin-Wash (Dig-Wash) Buffer, containing 0.01% (w/v) digitonin (see Reagents and Solutions), 4 °C

pAG-MNase, 20X stock (Epicypher, cat #15-1116 or similar)

NaCl (e.g. VWR, J.T. Baker, cat #JT4058-1)

100 mM CaCl₂

• Mix 5 μL of 1 M CaCl₂ (Moore, 1996) with 45 μL of ultrapure H₂O (e.g. Invitrogen, cat #10977-015)

  2X Calcium Reaction Buffer, containing 4 mM CaCl₂, on ice

• Mix 40 μL of 100 mM CaCl₂ with 960 μL of Dig-Wash Buffer

STOP Buffer, 2X (see Reagents and Solutions), 4 °C

10% (w/v) SDS (see Reagents and Solutions)

10 mg/mL Proteinase K (see Reagents and Solutions)

Phenol:Chloroform:Isoamyl Alcohol 25:24:1, Saturated with 10 mM Tris, pH 8.0, 1 mM EDTA (e.g. Sigma-Aldrich, cat #P3803)

Chloroform (e.g. Sigma-Aldrich, cat #C2432)

2 mg/mL GlycoBlue

• Prepare 45 μL by diluting 6 μL of GlycoBlue (e.g. ThermoFisher, cat #AM9515) in 39 μL of ultrapure H₂O (e.g. Invitrogen, cat #10977-015) right before use

100% Ethanol (e.g. VWR, cat #71001-866)

1 mM Tris-HCl, pH 8.0, 0.1 mM EDTA (see Reagents and Solutions), room temperature

dsDNA quantification reagents (Qubit^™ DNA High Sensitivity reagents or similar) (e.g. ThermoFisher, cat #Q32851 and Q32856)

Standard pipettes (P10-P1000)

1.5-mL Tube rotator (room temperature) (e.g. Fisher Scientific, cat #22-505-001)

PCR strip tubes, DNase- and RNase-free (e.g. VWR, cat #12000-206)

Multichannel pipette capable of pipetting 50–200 μL

Magnet stand for PCR tubes (e.g. NEBNext^® Magnetic Separation Rack for 24 PCR tubes, NEB, cat #S1515S)

Tube nutator at 4 °C (e.g. VWR, cat #10127-872)

Multichannel pipette reservoir, on ice (e.g. Corning, cat #4870)

PCR-cooler block, kept at −20 °C (e.g. Eppendorf, cat #022510525)

Ice/water bath (see Reagents and Solutions)

PCR machine with adjustable heated lid (e.g. BioRad T100 cat #1861096)

1.5-mL tubes, DNase- and RNase-free (e.g. LPS, cat #L211511-CS)

Heater block at 50 °C (e.g. Fisher Scientific, cat #88-870-003)

Vortex (e.g. Sigma Aldrich, cat #Z258415)

Phase-lock tubes, MAXtract, High Density, 1.5-mL (Qiagen, cat #129046)

Benchtop microcentrifuge for 1.5-mL tubes at room temperature and at 4 °C (e.g. Eppendorf, cat# 5418R)

−80 °C freezer (or −20 °C if not available)

Mini centrifuge (e.g. Fisher Scientific, cat #12-006-901)

dsDNA quantification instrument (Qubit^™ Fluorometer [e.g ThermoFisher, cat #Q33238] or similar)

Protocol Steps

Binding Worms/Cell Mixture to Activated ConA Beads

• 1To the worm/cell mixture obtained in Basic Protocol 2, containing roughly 150 μL per desired CUT&RUN reaction, add 15 μL of activated ConA beads per reaction and mix well by pipetting.

  For instance, for one genotype which will be used for 3 reactions (i.e. 3 antibodies) and, thus, contains ~450 μL of Wash Buffer, add 45 μL of activated beads.

• 2Rotate on tube rotator for 10 minutes at room temperature.
• 3Mix well and divide equally into 0.2-mL PCR tubes, one for each reaction/antibody to be used.

  For example, for one genotype which will be used for 3 reactions (i.e. antibodies) and contains roughly 450 μL of Wash Buffer, aliquot 150 μL of the worm/cell mixture to each of three 0.2-mL tubes (e.g. one tube to be used with an anti-H3K4me3 antibody, one tube to be used with an anti-H3K9me3 antibody, and one tube to be used with an anti-H3 antibody).

  Note that one tube should always be used as a negative control (for example, species-matched IgG, H3, or antibody background in wild-type worms lacking an epitope-tag).

  Note that while each CUT&RUN reaction consists of a pool of hundreds of worms, the reaction is still considered only one biological replicate. One biological replicate will be sufficient to determine, in test runs,if the CUT&RUN protocol is functional for your target of interest. However, for real experiments, at least two biological replicates should be collected. Biological replicates should be collected from worms grown on different days to account for biological variability. Thus, Basic Protocols 1–3 will need to be completed two separate times to produce two biological replicates.

Binding Primary Antibody

• 4Pipette to mix again and place tubes on the magnet stand to clear for 5 minutes. After that, discard the majority of the liquid.

  The majority of the cells and worm chunks should now be bound to the magnetic beads (See Figure 3A for an example, but note that, at this stage, the worms are sometimes difficult to see because the solution is still cloudy with bacteria). However, because worm chunks are heavy, some will settle to the bottom of the tube instead of sticking to the side with the beads. Because of this, leave 10–20 μL of liquid at the bottom for all magnetic wash steps.

  We recommend mixing by pipetting with multichannel pipettes for each mixing step, for faster and more uniform mixing.

• 5Add Antibody Buffer along the side of the tube so that the final solution volume in the tube is 100 μL. Pipette to mix.

  For example, if you left 15 μL of Wash Buffer in Step 4 to avoid worm loss, add 85 μL of Antibody Buffer to obtain a final volume of 100 μL in each tube.

• 6Add 1 μL of the primary antibody to get a final concentration of 1:100.

  Antibody concentration can be adjusted based on optimized concentrations, but 1:100 is the standard recommendation. Use 1 μL of antibody directly from the antibody tube (not previously diluted).

• 7Mix well by pipetting and place the tubes on a tube nutator at around 1.5–2 rpm overnight (12–18 h) at 4 °C.

  Ideally, the mixing by pipetting should be done immediately before placing the tubes on the nutator, to avoid worm chunks settling to the bottom of the tube. If possible, do this step in a 4 °C room that has the nutator, so the tubes can be immediately placed on the nutator after mixing.

  Depending on the nature of the primary antibody, a secondary antibody may be required. See Alternate Protocol 1.

Figure 3. Setup of key CUT&RUN steps.

A) Worm chunks and cells (visible in each tube as white specks, indicated by black arrow) are bound to (brown) magnetic beads and immobilized using a magnet stand for all wash steps. B) Setup of ice/water bath (see Reagents and Solutions) for critical calcium addition step in Basic Protocol 3. Note that the ice/water bath should contain a large amount of water, as visible in B. C) The final ice/water bath setup, including PCR cooler block and thermometer.

Binding pAG-MNase

• 8The next morning, place a reservoir for multichannel pipetting on ice and fill it with Dig-Wash Buffer.
• 9Remove samples from the tube nutator, pipette to mix again, and then place the tubes on the magnet stand to clear for 5 minutes.
• 10Remove the majority of the liquid, leaving 10–20 μL behind to avoid worm loss. Add 200 μL of cold Dig-Wash Buffer. Then, remove tubes from the magnet stand and fully resuspend by pipetting. Place the tubes back on the magnet stand for 5 minutes (See Video 1 for an example).
• 11Repeat step 10 for a total of two washes. Remove liquid, again leaving 10–20 μL behind.
• 12Add Dig-Wash Buffer to each tube for a final volume of 50 μL. Pipette to mix.

  Again, add up to 50 μL. So, if 15 μL were left to preserve worms, add 35 μL of Dig-Wash Buffer, for a final volume of 50 μL.

• 13Add 2.5 μL of pAG-MNase.

  This volume would need to be adjusted if pAG-MNase is purchased from a supplier with a different stock concentration, or if purified by the lab.

• 14Pipette to mix and immediately place the tubes on a nutator at 4 °C for 1 hour.

  During this time, prepare the ice/water bath needed for step 19 (see Reagents and Solutions).

• 15Mix again by pipetting and place tubes on a magnet stand for 5 minutes to separate.
• 16Remove the majority of liquid from the tubes, again leaving 10–20 μL behind. Add 200 μL of cold Dig-Wash Buffer. Then, remove tubes from the magnet stand and fully resuspend by pipetting. Place the tubes back on the magnet stand for 5 minutes.
• 17Repeat step 16 for a total of two washes. Remove liquid, again leaving 10–20 μL behind.

MNase Digestion by Addition of Calcium

• 18Add Dig-Wash Buffer to each tube for a final volume of 50 μL. Pipette to mix.
• 19Place the tubes into the PCR cooler block in the prepared ice/water bath (See Figure 3 for an example of experimental setup). Place the 2X Calcium Reaction Buffer in the ice/water bath. Let the tubes chill for 2–5 minutes.

  If a 4 °C room is available, move the entire ice bucket to the room and perform steps 20–23 there.

• 20Add 50 μL of the 2X Calcium Reaction Buffer to each tube while the tubes are sitting in the 0 °C block.
• 21Quickly remove the tubes from the block, pipette to mix, and place the tubes back in the 0 °C block.
• 22Incubate at 0 °C for 30 minutes.

  Do not touch the tubes during this time, as keeping the temperature close to 0 °C is very important to avoid excessive MNase activity and high background in CUT&RUN. Monitor the temperature, to make sure it stays consistent throughout the incubation.

• 23Add 100 μL of the 2X STOP Buffer to each tube and mix by pipetting.

  The STOP Buffer will stop the MNase cleavage reaction.

Chromatin Release

• 24Incubate samples for 30 minutes at 37 °C in a thermocycler with the heated lid set to 40 °C.

  This allows for cleaved DNA fragments of interest to be released from chromatin within the nucleus. The cleaved fragments are now able to float out of the permeabilized cells and into the solution, while chromatin that was not cleaved by MNase remains within the cells.

• 25Mix tubes by pipetting, then place tubes on a magnet stand to clear for 5 minutes.
• 26Carefully transfer the unbound liquid supernatant (~200 μL) containing the CUT&RUN fragments of interest to a new 1.5-mL tube. Avoid transferring beads or worms.

  CUT&RUN DNA fragments of interest are now in the supernatant, and worms and cells containing uncut DNA fragments are stuck to the magnet or at the bottom of the tube, and can be discarded.

  Since some worms settle to the bottom, leave 10–20 μL of the liquid at the bottom of the tube to avoid transferring large worm chunks.

• 27Place the tubes in a tube rack for 1–5 minutes to allow any remaining large worm chunks to settle to the bottom, then take the top ~90% of the sample and transfer into a fresh 1.5-mL tube.

  It is important to avoid carrying over worm chunks, which can still be present after the initial supernatant transfer in step 26, because this could introduce additional genomic DNA and/or high background to your experiment. Check under a microscope at this stage to make sure you do not have large worm chunks in your tube after the second transfer. If you see large chunks, settle and transfer again.

DNA Extraction

• 28Add 2 μL of 10% (w/v) SDS and 5 μL of 10 mg/mL Proteinase K solution to each sample. Mix by inversion. Incubate for 1 hour at 50 °C in a heat block or similar.

  This will denature the proteins also present with the extracted DNA.

  During this time, prepare phase-lock tubes by spinning them down at 16,000 rcf for 30 seconds, per the manufacturer’s instructions.

• 29Add an equal volume of Phenol:Chloroform:Isoamyl Alcohol to each sample. Mix by vortexing on high for ~2 seconds and transfer each sample to a separate phase-lock tube.

  For example, for a sample volume of ~200 μL, add 200 μL of Phenol:Chloroform:Isoamyl Alcohol.

• 30Centrifuge at 16,000 rcf for 5 minutes at room temperature.
• 31To each phase-lock tube containing your sample, add the same volume of chloroform as the starting sample volume. Mix by inversion or shaking the tube 10 times.

  For example, for an initial sample volume of ~200 μL, add 200 μL of chloroform.

  Do NOT vortex the phase-lock tubes.

• 32Centrifuge at 16,000 rcf for 5 minutes at room temperature.
• 33Transfer the aqueous top layer above the phase-locked gel to a new 1.5-mL tube containing 2 μL of 2 mg/mL GlycoBlue.

  Since the gel from the phase-lock tubes physically separates the aqueous layer from the interphase and organic layers, there is no risk of contamination. Therefore, transfer the entire aqueous phase to maximize DNA recovery. Be careful not to pierce the phase-lock gel with a pipette tip.

• 34Add ~2.5 volumes of 100% EtOH and mix by vortexing or tube inversion.

  For example, for a sample volume of ~200 μL, add 500 μL of EtOH.

• 35Chill at −80 °C overnight (~12–20 h).

  A shorter incubation time could also be used, although an overnight incubation may increase the recovery of DNA, especially for samples with low DNA content. This may be especially important if starting with low worm numbers or performing CUT&RUN for a protein factor like a transcription factor that has few expected binding sites or acts in a tissue-specific manner.

• 36Centrifuge samples at 16,000 rcf for 20 minutes at 4 °C.
• 37Remove liquid by pipetting, taking care not to aspirate the pellet.

  At this stage, the pellet is often gel-like and could be difficult to see. If removing all the liquid confidently will be too difficult because of the transparency of the pellet, remove most of the liquid and leave some behind.

• 38Add 1 mL of 100% EtOH to each tube.
• 39Centrifuge samples at 16,000 rcf for 5 minutes at 4 °C.
• 40Carefully remove all liquid. Do this first with a P1000, then quick-spin in a mini centrifuge, and then remove any residual EtOH with a P20 or P10.

  At this stage, the pellet is usually “fluffier” and easier to see than before the EtOH rinse.

• 41Air dry the pellet up to 5 minutes at room temperature.

  Ensure that the pellets are completely dry and all EtOH is removed before proceeding to the next step.

• 42When the pellet is dry, dissolve in 15 μL of 1 mM Tris-HCl, pH 8.0, 0.1 mM EDTA and transfer to a 0.2-mL PCR strip tube.
• 43Quantify DNA using 1 μL of the sample and dsDNA specific reagents, typically Qubit^™ DNA High Sensitivity reagents, following the manufacturer’s instructions.

  Typical values obtained from Qubit^™ range from 0.5 ng/μL to 3 ng/μL per sample, but even a sample containing too little DNA to be detected on high sensitivity Qubit^™ can still be successfully used to create a CUT&RUN library. DNA concentration here will depend on the number of worms used as well as on the abundance of the protein factor of interest.

  Because DNA concentration is typically quite low, and accurate quantification is desired, Nanodrop is not recommended as a suitable alternative to Qubit.

  At this point, fragment size analysis (e.g. Bioanalyzer or Tapestation) is not typically recommended because the percent of the sample that contains the DNA fragments of interest may be low compared to genomic DNA at this stage, since some genomic DNA contamination is expected before library preparation. Due to this, it is recommended to proceed directly to library preparation, which will preferentially amplify CUT&RUN fragments of interest, before fragment size analysis.

• 44Proceed to Basic Protocol 4 or store samples at −20 °C until use.

  If multiple biological replicates will be collected for one target, it is a good idea to keep the samples from the first replicate frozen at −20 °C until Basic Protocols 1–3 have been completed for the second replicate. Then, proceed to library preparation (Basic Protocol 4) with all replicates simultaneously, to minimize technical variability produced by preparing sequencing libraries at different times.

ALTERNATE PROTOCOL 1: CUT&RUN with Secondary Antibody Incubation

Depending on the choice of primary antibody in Basic Protocol 3, a secondary antibody may be needed to increase the efficiency of pAG-MNase recognition. This is because Protein A and G recognize antibodies of certain species better than others (See Table 7.6 in The Molecular Probes® Handbook, also available at https://www.thermofisher.com/us/en/home/references/molecular-probes-the-handbook/tables/binding-profiles-of-protein-a-and-protein-g.htm), so if the primary antibody is from a species less well recognized, addition of a secondary antibody before the binding of pAG-MNase may be needed. In most cases, a secondary antibody will not be necessary, as long as the recombinant pAG-MNase fusion protein is used (instead of the original pA-MNase fusion protein, which only included protein A and had a more limited binding capability). However, there may be instances in which you still find the results for antibodies produced in certain species can be improved by the use of a secondary antibody (e.g. mouse). Here, we describe the steps needed to bind a secondary antibody, which include two wash steps in Dig-Wash Buffer (identical to those in Basic Protocol 3), and a one-hour secondary antibody incubation step at 4 °C. Alternate Protocol 1 is to be completed in addition to the entirety of Basic Protocol 3, and would be completed after Basic Protocol 3 Step 8 and before Step 9 (i.e. after overnight incubation with the primary antibody and before pAG-MNase binding). After completing Alternate Protocol 1, the chromatin-bound protein of interest will have both a primary and secondary antibody bound to it, and cells will be ready for incubation with pAG-MNase and the completion of Basic Protocol 3.

Additional Materials to Basic Protocol 3

Secondary antibody, preferably produced in rabbit (e.g. Rabbit anti-Mouse IgG, abcam, cat #ab46540)

Protocol Steps

Binding Secondary Antibody

1. Follow Basic Protocol 3 steps 1–8.

2. Remove samples from the tube nutator, pipette to mix again, and then place tubes on the magnet stand to clear for 5 minutes.

   If only some of the samples need a secondary antibody incubation (e.g. half of the samples use a rat antibody and require a secondary antibody for strong pAG-MNase binding, while the other half of the samples use a rabbit antibody and do not require a secondary antibody to bind pAG-MNase), remove only the half of the samples that require the secondary antibody. Leave the remaining samples rocking at 4 °C until Alternate Protocol 1 is completed, then, when returning to Basic Protocol 3, Step 9, remove all samples and begin Step 9 with all samples together.

3. Remove the majority of the liquid, leaving 10–20 μL behind, to avoid worm loss. Add 200 μL of cold Dig-Wash Buffer. Then, remove tubes from the magnet stand and fully resuspend by pipetting. Place the tubes back on the magnet stand for 5 minutes.

4. Repeat step 3 for a total of two washes. Remove liquid, again leaving 10–20 μL behind.

5. Add Dig-Wash Buffer to each tube for a final volume of 150 μL. Pipette to mix.

6. Add 1.5 μL of secondary antibody for a final dilution of 1:100, or use optimized antibody concentration.

   Antibody dilution can be optimized based on experimental results. For example, if the CUT&RUN profile obtained with 1:100 diluted antibody has high background, an antibody dilution series could be tested and sequenced, or the antibody concentration that works best in immunofluorescent staining can be used.

7. Pipette to mix and immediately place tubes back on the tube nutator to incubate for 1 hour at 4 °C.

8. Return to step 9 in Basic Protocol 3 and follow all remaining steps.

BASIC PROTOCOL 4: CUT&RUN Library Preparation for Illumina High-Throughput Sequencing

Here, we describe the process of preparing libraries from the DNA fragments obtained from CUT&RUN in Basic Protocol 3, for use in Illumina sequencing. We and others (Liu et al., 2018; Liu, 2021) recommend using the NEBNext^® Ultra^™ II DNA Library Prep Kit for Illumina^® from New England BioLabs due to its high performance with even very low starting amounts of DNA. The protocol described below largely follows the kit’s instructions for library preparation, with a few minor adjustments because of the small fragment sizes critical for CUT&RUN. As described elsewhere (Liu et al., 2018; Liu, 2021), we perform two rounds of size selection and purification after library preparation using Ampure XP beads, in order to preferentially enrich CUT&RUN fragments of interest while depleting large genomic DNA fragments and unbound adaptor and primer dimer.

Materials

CUT&RUN DNA sample from Basic Protocol 3 (typically 7 μL, approximately half of the remaining sample after Qubit)

Ultrapure DNase- and RNase-free Distilled H₂O (e.g. Invitrogen, cat #10977-015)

NEBNext^® Ultra^™ II End Prep Enzyme Mix (NEB, cat # E7645S/L)

NEBNext^® Ultra^™ II End Prep Reaction Buffer (cat # E7645S/L)

NEBNext^® Adaptors for Illumina (NEB, cat #E7335S/L)

NEBNext^® Ultra^™ II Ligation Master Mix (NEB, cat #E7645S/L)

NEBNext^® Ligation Enhancer (NEB, cat #E7645S/L)

USER^® Enzyme (NEB, cat #E7335S/L)

Ampure XP beads (e.g. Beckman Coulter, cat #A63881)

80% Ethanol, made fresh

• Mix 16 mL of 100% Ethanol (e.g. VWR, cat #71001-866) with 4 mL of ultrapure H₂O (e.g. Invitrogen cat #10977-015)

Tris-HCl, 10 mM

• Mix 4.95 mL of ultra-pure H₂O (e.g. Invitrogen 10977–015) with 50 μL of 1 M Tris-HCl, pH 8.0 (Moore, 1996)

NEBNext^® Ultra^™ II Q5^® Master Mix (NEB, cat # E7645S/L)

Index Primer for Illumina (unique for each sample) (NEB, cat #E7335S/L, E7500S/L, E7710S/L, E7730S/L)

Universal PCR Primer for Illumina (NEB, cat #E7335S/L, E7500S/L, E7710S/L, E7730S/L)

dsDNA quantification reagents (Qubit^™ or similar) (e.g. ThermoFisher, cat #Q32851 and Q32856)

Standard pipettes (P10-P200)

PCR strip tubes, DNase- and RNase-free (e.g. VWR, cat# 12000–206)

Multichannel pipette capable of pipetting 50–200 μL

Mini centrifuge for PCR tubes (e.g. Fisher Scientific, cat #12-006-901)

PCR machine with adjustable heated lid (e.g. BioRad T100, cat #1861096)

Magnet stand (e.g. NEBNext^® Magnetic Separation Rack for 24 PCR tubes, NEB, cat #S1515S)

Multichannel pipette reservoir (e.g. Corning, cat #4870)

dsDNA quantification instrument (Qubit^™ Fluorometer [e.g ThermoFisher, cat #Q33238] or similar)

Fragment size analysis equipment (Agilent Fragment Analyzer or similar)

Protocol Steps

End Preparation of CUT&RUN Fragments

• 1Using the CUT&RUN DNA fragments obtained in Basic Protocol 3, place 1–20 ng of CUT&RUN DNA in a new PCR strip tube. Dilute up to 50 μL in ultrapure H₂O.

  Generally, use 7 μL, i.e. about half of the volume obtained from CUT&RUN after Qubit quantification. If DNA concentration is too low to be accurately quantified, use between half and all of the CUT&RUN fragments to generate the library.

• 2Add 3 μL of NEBNext^® Ultra^™ II End Prep Enzyme Mix and 7 μL of NEBNext^® Ultra^™ II End Prep Reaction Buffer for a total of 60 μL per tube.
• 3Mix by pipetting up and down 10 times and quick-spin in a mini centrifuge.

  Multichannel pipetting is again recommended for mixing, as it decreases time and increases uniformity.

• 4Place the tubes in a thermocycler with the heated lid set to ≥ 75 °C and incubate as follows:

  • 30 minutes at 20 °C

  • 30 minutes at 65 °C

  • Hold at 4 °C

Adaptor Ligation

• 5Dilute NEBNext^® Adaptors for Illumina to 1 in 25 or to desired dilution in ultrapure H₂O.

  Adaptor dilution should be adjusted according to experimental results and amount of adaptor present in Bioanalyzer size analysis after library preparation. For example, if using 1 in 25-diluted adaptors frequently results in libraries for a certain protein factor that contain more than 5–10% of adaptor dimer, then dilute adaptors further the next time library preparation is done for the same protein factor. In general, we typically use 25-fold diluted adaptors for 3–20 ng DNA, and 50-fold diluted adaptors when DNA concentration is undetectable after CUT&RUN. However, avoid diluting adaptors too much, because if adaptors are not in excess, this can result in low complexity libraries.

• 6To the 60 μL End Prep Reaction Mixture from step 4, add 30 μL of NEBNext^® Ultra^™ II Ligation Master Mix, 1 μL of NEBNext^® Ligation Enhancer, and 2.5 μL of diluted adaptors for a total volume of 93.5 μL per sample.

  Make sure to mix the Ultra^™ II Ligation Master Mix by pipetting before adding it to the reaction.

• 7Pipette to mix well. Do a quick-spin.
• 8Place tubes in a thermocycler and incubate at 20 °C for 15 minutes with the heated lid off or open.

  Now would be a good time to take out the Ampure XP beads to warm to room temperature.

• 9Add 3 μL of the USER^® Enzyme per reaction and mix well by pipetting.
• 10Place tubes in a thermocycler and incubate at 37 °C for 15 minutes with the heated lid set to ≥ 47 °C.

  Samples can safely be stored overnight at −20 °C before proceeding.

Cleanup of Adaptor-Ligated DNA

• 11Add 169 μL (1.75X) of resuspended, room temperature Ampure XP beads to the adaptor-ligated DNA fragments from Step 10. Mix well by pipetting up and down at least 10 times.

  This high ratio of beads relative to volume of the DNA solution will bind all but extremely small DNA fragments.

  Ensure the beads are at room temperature and thoroughly resuspended before mixing them with the DNA.

• 12Incubate for 5 minutes at room temperature.

  Prepare fresh 80% Ethanol during this incubation (or when convenient) and place into a multichannel reservoir at room temperature.

• 13Place tubes on a magnet stand for 5 minutes to allow separation of beads from the supernatant. Then, using a multichannel pipette, remove and discard supernatant.

  Do NOT discard the beads, which contain the CUT&RUN fragments of interest.

  Multichannel pipetting is recommended for all steps on the magnet stand, for faster and more uniform library preparation.

• 14With the tubes still on the magnet stand, add 200 μL of 80% EtOH to each reaction. Incubate at room temperature for 30 seconds, then remove and discard the supernatant.
• 15Repeat step 14 for a total of two washes. Remove all residual EtOH, using a P10 pipette if necessary.
• 16Air dry the beads for up to 5 minutes on the magnet stand.

  Beads should appear dry and matte, not shiny. Be careful not to over dry the beads to the point of cracking.

• 17Carefully remove the tubes from the magnet stand and add 17 μL of 10 mM Tris-HCl to elute the DNA from the beads.
• 18Mix well by pipetting up and down at least 10 times and incubate at room temperature for 5 minutes.
• 19Place the tubes on the magnet stand for 5 minutes and transfer the liquid (now containing eluted CUT&RUN DNA fragments of interest) to a new PCR tube. Discard the beads.

  Samples can safely be stored overnight at −20 °C.

PCR Enrichment of Adaptor-ligated DNA

• 20
  Combine the following reagents in a PCR tube and pipette to mix.

  15–17	μL Adaptor-ligated CUT&RUN DNA fragments (from Step 19)
  25 μL	NEBNext® Ultra™ II Q5® Master Mix
  2.5 μL	Index Primer for Illumina*
  2.5 μL	Universal PCR Primer for Illumina
  Up to 50 μL	Nuclease-free H2O

  ^*

  Index primer should be different for each sample that will be pooled together in one sequencing lane. Keep careful notes of which index primer is added to each reaction.
• 21
  Do a quick-spin. Then, run the following protocol to amplify libraries in a thermocycler:

  Temperature	Time	Cycles
  98 °C	30 sec	1
  98 °C	10 sec	10–16*
  65 °C	45 sec
  65 °C	5 min	1
  4 °C	Hold

  ^*

  Cycle number should be adjusted based on experimental results, including quantity of library obtained and CUT&RUN profile generated. We typically use 14 cycles for 3–15 ng input DNA. Increasing cycles may result in excessive PCR duplicates, which show up as duplicated sequences in the data analysis, and may decrease data quality. Too few cycles would result in insufficient DNA quantities to sequence at the end of the protocol. In general, the fewer cycles the better, but enough cycles must be run to produce enough DNA for sequencing.

Cleanup of PCR Reaction

• 22Add 26.5 μL (~0.5X) of resuspended, room temperature Ampure XP beads to each PCR-amplified reaction mix. Mix well by pipetting up and down at least 10 times and incubate at room temperature for 5 minutes.

  This low ratio of beads relative to the volume of the DNA solution will bind only large DNA fragments that you do not wish to sequence. Thus, this step acts as a size selection to eliminate any remaining genomic DNA or other large fragments.

• 23Place on a magnet stand for 5 minutes.
• 24Transfer the supernatant, containing CUT&RUN DNA of interest (~70 μL), to a new PCR tube. Discard the beads, which now contain large DNA fragments.

  As this is different from all the other Ampure XP cleanup steps, be careful NOT to discard the supernatant at this step.

• 25Add 33.5 μL (final concentration 1.2X) of Ampure XP beads to the supernatant from step 24, which contains the CUT&RUN fragments of interest. Mix well by pipetting up and down at least 10 times and incubate samples at room temperature for 5 minutes.

  To calculate the final Ampure XP concentration, keep in mind the PEG from the beads added in step 22 is still present with the DNA sample, even though the beads themselves were discarded in step 24. Thus, calculate the volume needed for a final 1.2X cleanup with the following formula:

  1.2[i.e. desired ratio] x 50 μL [i.e. initial sample volume] – 26.5 μL [i.e. volume of PEG already in the sample] = 33.5 μL beads. This should bind DNA fragments over 200 bp and deplete residual primer and adaptor dimer from the sample.

• 26Place samples on the magnet stand for 5 minutes, then remove and discard the supernatant.
• 27With tubes still on the magnet stand, add 200 μL of 80% EtOH to each reaction. Incubate at room temperature for 30 seconds, then remove and discard the supernatant.
• 28Repeat step 27 for a total of 2 washes. Remove all residual EtOH, using a P10 pipette if necessary.
• 29Air dry the beads for less than 5 minutes on the magnet stand.

  This air-dry step is typically faster than that of step 16 because the volume of beads is smaller. Be careful not to over dry the samples.

• 30Carefully remove tubes from the magnet stand and add 17 μL of 10 mM Tris-HCl to elute DNA from the beads. Mix well by pipetting up and down at least 10 times. Incubate for 5 minutes at room temperature.
• 31Place the tubes on the magnet stand for 5 minutes and transfer the supernatant, containing CUT&RUN libraries, to a new PCR tube. Discard the beads.

  Libraries can safely be stored at −20 °C indefinitely until use.

• 32Quantify dsDNA in the library using Qubit^™ DNA High Sensitivity reagents or similar following the manufacturer’s instructions.

  Typical values obtained by Qubit^™ after library preparation range from 1–70 ng/μL, depending on the target of interest, number of worms used, and number of PCR cycles used. We typically use 14 PCR cycles and obtain values on the lower end (1–20 ng/μL) for chromatin factors (starting with ~3000 worms/antibody), and values on the higher end (20–70 ng/μL) for histones harboring specific post-translational modifications (starting with ~1000 worms/antibody). However, if small numbers of worms are used (e.g. 100 worms), expect low DNA quantity after library preparation (less than 2 ng/μL is not unusual). Note that low DNA quantity does not, in our hands, necessarily mean poor CUT&RUN sequencing results. As long as enough DNA is obtained for sequencing, the experiment can go ahead as planned.

• 33Examine size distribution of the CUT&RUN libraries using Bioanalyzer or similar following the manufacturer’s instructions.

  The majority of fragments should be between 180–600 bp, and could show variable distribution based on the target. See Figure 4 and the Understanding Results section for more information.

• 34If adaptor or primer dimer represents more than 5–10% of the library, repeat a size selection with 1.2X Ampure XP beads.

  To tell if adaptor or primer dimer represents more than 5–10% of the library, first identify if there is a peak on the Bioanalyzer trace corresponding to the expected primer or adaptor dimer size (roughly 54 bp or 137 bp, respectively, see Figure 4). If a peak appears at this size, determine if the concentration of this peak relative to the total concentration of the sample, as estimated by the Bioanalyzer, exceeds 5–10%. If it does, proceed with the cleanup.

  To perform a 1.2X Ampure XP cleanup with prepared libraries, add water to libraries to standardize the volume to 20 μL total. Then, add 24 μL of resuspended beads and repeat steps 12–19 of the library preparation protocol.

• 35Pool samples at equal molarity or as desired and submit for 2×32 paired-end sequencing with Illumina NextSeq 500 or similar.

  Pool to the desired concentration of your sequencing facility; in our case, we pool to at least 4 nM total.

  If some samples are less important than others (e.g. a test of a new antibody or an extra negative control that will not be used in downstream analysis), they could be pooled to purposely take up a smaller proportion of reads than the other samples, if desired.

  The number of libraries that can be pooled together depends on the desired sequencing depth – aim for at least 10 million reads per sample.

• 36Proceed to data analysis (Basic Protocol 5).

Figure 4. Sample Bioanalyzer results for CUT&RUN libraries.

Examples of CUT&RUN libraries subjected to Bioanalyzer fragment size analysis after library preparation. A and B show libraries prepared in parallel using antibodies against either (A) H3K4me3 (Epicypher, cat #13-0041), which shows a nucleosomal laddering pattern as expected, or (B) H3 control (abcam, cat #ab1791). Some primer dimer (~54 bp, blue arrow), adaptor dimer (~137 bp, red arrow), and large genomic DNA fragments (~1,500 bp, purple arrow) are detectable, but represent only a small fraction of the final library and will not interfere with the sequencing. C and D show the same library, which originally (shown in C) had high adaptor content. The library was re-cleaned with a 1.2X Ampure XP size selection, and adaptor content decreased substantially (shown in D) without affecting the library itself.

BASIC PROTOCOL 5: Sample Basic Data Analysis using Linux

After Illumina sequencing (Basic Protocol 4), genomic data will be returned as .fastq files, which need to be processed in order to obtain binding profiles from CUT&RUN. The most basic analysis steps include quality control, adaptor trimming, alignment of CUT&RUN reads to the C. elegans genome, peak calling, and data visualization. Here, we provide a simple example of steps needed to visualize CUT&RUN data to determine if your experiment worked. This protocol is only intended as a general guideline for the steps that need to be taken in order to process CUT&RUN data, rather than a comprehensive step-by-step protocol to be exclusively followed. The individual programs used or preferences selected for each program may vary based on your data and preferences and, thus, we do not include the actual code needed to run each program in this protocol. Instead, the manual for each software program mentioned should be consulted to determine the correct command line to use for each step outlined below.

Materials

Computer equipped with Linux and processing power for NGS data

Fastq files representing CUT&RUN profile, produced with experimental antibody in Basic Protocol 4

Fastq files representing CUT&RUN background, produced with control antibody in Basic Protocol 4

Reference C. elegans genome (WBcel235/cel11) (available, for example, at http://useast.ensembl.org/Caenorhabditis_elegans/Info/Index)

NGS Software for use within the Linux environment, for example:

• Quality check software (e.g. fastqc (version 0.11.8), (https://www.bioinformatics.babraham.ac.uk/projects/fastqc/))

• Adaptor-trimming software (e.g. cutadapt (version 3.4) (Martin, 2011))

• Genomic alignment software (e.g. bowtie2 (version 2.4.3) (Langmead & Salzberg, 2012))

• Sorting and Indexing Software (e.g. samtools (version 1.14) (Li et al., 2009))

• Bigwig-Generating Software (e.g deeptools (version 3.3) (Ramírez et al., 2014))

• Peak-Calling Software (e.g. MACS2 (version 2.1.4) (Zhang et al., 2008))

Data Visualization Software (e.g. IGV (Robinson et al., 2011))

Protocol Steps

Processing CUT&RUN Reads

1. In the Linux environment, quality check samples with fastqc following the manual’s instructions.

   Fastqc will generate various quality scores for different aspects of the data (e.g. number of reads, quality score of reads, duplication rate, etc.). While the sequencing samples may not score perfectly in every section, the information generated by fastqc can provide an idea of the success of the sequencing itself. In particular, take note of the sequence counts (aiming for at least 5–10 million reads per sample), mean sequence quality of the samples (Phred Score should be above 20), and duplication rate (ideally below 50% duplicated sequences). If samples score poorly, that does not necessarily mean the experiment failed, but the information can be considered later on if it is determined at the end of Basic Protocol 5 that the sequencing did not work as desired.

2. Trim adaptors for paired-end sequencing with cutadapt or a similar tool, following instructions outlined in the manual.

3. Quality check samples again as in Step 1.

   Take particular note of the sequence counts, which should now contain identical numbers of reads for both paired-end sequencing files corresponding to one sample, and to the adaptor content, which should no longer be detectable.

4. Create an indexed genome file for the desired aligner using the reference C. elegans genome WBcel235/ce11. For example, use the build function in bowtie2 following the instructions outlined in the bowtie2 manual.

5. Align trimmed fastq files to the C. elegans genome using bowtie2 or a similar aligner, following the instructions outlined in the manual.

   This will generate a sam file of aligned reads.

6. Convert sam to bam files using samtools view, following the instructions outlined in the manual.

7. Sort and index bam files using samtools sort and samtools index, respectively. Follow the instructions outlined in the Samtools manual.

8. Convert sorted bam files into bigwig files to visualize tracks in IGV. Use either bamCompare to compare two files while normalizing them to each other (e.g. H3K4me3 vs H3), or bamCoverage to visualize a single file. Follow the directions in the deepTools manual.

   When using bamCompare to compare two different CUT&RUN sequencing files, the log2 ratio of the samples will be calculated. When the resulting bigwig file is opened in IGV in step 10, any ratio more than zero is displayed as a positive value, and any ratio less than zero is displayed as a negative value. Thus, if H3K4me3 vs H3 is plotted, regions above a flat line represent regions where H3K4me3 is enriched relative to H3, whereas regions below a flat line indicate regions where H3K4me3 is depleted relative to H3. See Figures 5–7 for examples.

   When visualizing a bigwig file generated using bamCoverage to examine a single CUT&RUN sequencing file, all values will be positive, since the file is not being compared to any other. See Figure 8 for an example.

9. Call peaks using MACS2 or similar peak-calling software (e.g. HOMER) following standard instructions. Ideally, call peaks against a background file, like H3, IgG, or, if possible, antibody background. Antibody background would be represented by the CUT&RUN profile produced by using an antibody which does not have a target in the worm strain profiled (e.g. anti-GFP antibody in N2 wild-type worms). Narrow or broad peaks should be called depending on the expected distribution of the mark profiled.

   For example, H3K4me3 usually marks sharp, distinct regions, so narrow peaks are used. However, H3K9me3 typically occupies wider regions, so broad peaks are used.

10. Visualize Bigwig files and peaks in IGV. Be sure to examine both the genome-wide view, which will give you an overall idea of the distribution of your data, as well as close-up views of individual chromosomes, parts of chromosomes, and individual peaks, which will give you an idea of the resolution of your data.

    Since peaks are called by identifying regions of high enrichment relative to background, you should see that peaks overlap well with regions of enrichment relative to control identified in BigWig files (See Figure 7).

    When you are first trying the technique, examine Bigwig files of ChIP-seq data in parallel, if available, to ensure that the majority of peak regions found by CUT&RUN are the regions expected based on previous experiments (i.e. H3K4me3 peaks should be found at the same regions of the genome using either ChIP-seq or CUT&RUN).

    If the technique worked somewhat but failed to produce high-resolution data, you may see the expected profile at the genome-wide view, but you may find high regions of background where you expect to find depleted regions, and/or unclear distinctions between peak regions (See Figure 5, C&R 10w, for an example of low-resolution data, compared to C&R 100w and C&R 1000w, which are examples of high-resolution data).

Figure 5. H3K4me3 CUT&RUN shares expected ChIP-seq binding pattern and can be detected with as few as 10 worms.

Shown are IGV screenshots of ChIP-seq data for H3K4me3 generated by our lab (Wang et al., 2018) compared to CUT&RUN data for H3K4me3 generated with 1000 worms (C&R 1000w), 100 worms (C&R 100w), or 10 worms (C&R 10w). Both ChIP-seq and CUT&RUN data were generated using wild-type worms of the N2 strain using either L4 worms (ChIP) or Day 1 adults (CUT&RUN). Data are plotted as the log2 fold change of H3K4me3 vs H3 control, thus blue regions represent enrichment of H3K4me3, and red regions represent depletion of H3K4me3 relative to H3 levels in the genome. As expected, sites of enrichment or depletion for H3K4me3 identified by CUT&RUN match the expected ChIP-seq profile well, and the CUT&RUN data are high resolution, even with only 100 worms when looking at a whole genome-view (A) or zoomed-in peak regions (B, C). Signal can still be detected from only 10 worms, although the resolution decreases. CUT&RUN tracks represent data from one biological replicate, while ChIP-seq data represent two merged biological replicates.

Figure 7. CUT&RUN captures the expected binding profile of H3K4me3 readers SET-9 and SET-26 in C. elegans.

Comparison of the ChIP-seq profile for the paralog H3K4me3 readers SET-9 and SET-26, previously generated by our lab (Wang et al., 2018), to CUT&RUN profile for SET-9 and SET-26, either A) genome-wide or B) at an individual peak level. Both ChIP and CUT&RUN data were generated using our lab’s set-9::GFP set-26::GFP endogenous CRISPR-tagged strain reported in Wang et al. (2018), and both profiles were immunoprecipitated with the same anti-GFP antibody (abcam, cat #ab290). Data are displayed as the log2 fold change of antibody binding in the SET-9/26-tagged strain against GFP antibody background binding in wild-type worms (lacking GFP). Overall, the binding pattern captured by CUT&RUN matches the expected ChIP-seq profile, and the individual peaks are typically higher resolution in the CUT&RUN data, with a sharper cutoff between enriched and depleted regions. Narrow peaks were called with MACS2 using N2 background antibody binding as control. CUT&RUN tracks represent data from one biological replicate, while ChIP-seq data represent two merged biological replicates. C&R, CUT&RUN.

Figure 8. Antibody background is an important consideration for CUT&RUN.

A and B show examples of regions that show enrichment in both binding profiles for a specific factor and antibody background, and thus, do not represent real regions of factor-enrichment. The top track shows the log2 fold change for the H3K4me3 readers SET-9/26 relative to antibody background (GFP antibody binding in set-9::GFP set-26::GFP tagged strain vs GFP antibody binding in wild-type worms lacking GFP). The middle track shows the un-normalized SET-9/26 binding profile, while the bottom track shows the un-normalized GFP antibody background in wild-type worms lacking the epitope, both displayed in log scale. Without taking into account the antibody background, the regions highlighted by the green arrows could have been called as regions of SET-9/26 enrichment, when they are in fact just non-specific antibody binding. CUT&RUN tracks represent data from one biological replicate.

REAGENTS AND SOLUTIONS:

Activated Concanavalin A-coated (ConA) beads

• Add 1.5 mL of Binding Buffer to a 2-mL tube

• Resuspend ConA beads (Bangs Laboratory, #BP531) by pipetting or shaking container

• Add 15 μL beads/reaction to the 2-mL tube containing Binding Buffer

  • e.g. 240 μL beads for 16 reactions

• Place the tube on a magnet stand (e.g. ThermoFisher Invitrogen #CS15000) for 2 minutes until cleanly separated

• Remove and discard liquid

• Remove tube from magnet stand, add 1.5 mL of Binding Buffer

• Pipette to mix and place tube back in the magnet stand for 2 minutes to separate

• Completely remove and discard liquid

• Resuspend the beads in Binding Buffer, using the same volume as the original volume of beads

  • e.g. 240 μL Binding Buffer for 16 reactions

• Keep on ice until use

• Use same day

Antibody Buffer, containing 0.01% (w/v) digitonin

• 2 mL of Dig-Wash Buffer

• 8 μL of 0.5 M EDTA (Moore, 1996)

• Keep on ice, use same day

Binding Buffer, 20 mM HEPES, 10 mM KCl, 1 mM CaCl₂, 1 mM MnCl₂

• 19.36 mL of ddH₂O

• 400 μL of 1 M HEPES, pH 7.5

• 200 μL of 1 M KCl (Moore, 1996)

• 20 μL of 1 M CaCl₂ (Moore, 1996)

• 20 μL of 1 M MnCl₂

• Store at 4 °C for up to 6 months

Bovine Serum Albumin (BSA), 10% (w/v)

• 1 g of BSA (e.g. Sigma-Aldrich, cat #A7906)

• 10 mL of ddH₂O

• Mix to dissolve

• Store in aliquots at −20 °C for up to 1 year

Cholesterol, 5 mg/mL

• 0.5 g of cholesterol (e.g. Sigma-Aldrich, cat #C8667)

• Add ~80 mL of 95% EtOH

• Stir until completely dissolved

• Adjust volume to 100 mL with 95% EtOH

• Store at room temperature for up to 6 months

Concentrated OP50 Bacterial Culture for Seeding NGM Plates

• Grow an overnight culture of streptomycin-resistant OP50 Escherichia coli (purchase at the CGC, OP50–1 strain https://cgc.umn.edu/strain/OP50-1) in LB supplemented with 30 μg/mL streptomycin for ~16 h, shaking at ~250 rpm at 37 °C

  • For example, supplement a 200-mL culture with 100 μL of 60 mg/mL streptomycin

• Spin down bacteria (~3,000rpm for ~30 minutes) in sterile tubes (e.g. VWR, Cat #VWRU89039-660) in a centrifuge (e.g. Eppendorf, cat #5702) and concentrate 5X for 6-cm plates or 25X for 15-cm NGM plates

Cuticle Disruption Buffer, 200 mM DTT, 0.25% (w/v) SDS, 20 mM HEPES pH 8.0, 3% (w/v) sucrose

• 25 mL of `ddH₂O

• 125 mg of SDS (e.g. Sigma-Aldrich, cat #L4509-500G)

  • Caution: Be careful when weighing out SDS powder, wear a mask as a precaution

• 1.5 g of sucrose (e.g. VWR, cat #JT4072-1)

• 1.54 g of DTT (e.g. Fischer Scientific, cat #BP172-5)

• 1 mL of 1 M HEPES, pH 8.0

• Mix to dissolve

• Add ddH₂O to 50 mL

• Freeze in aliquots at −20 °C for up to 1 year

• Thaw at room temperature

Digitonin, 5% (w/v)

• 50 mg of digitonin powder, high purity (EMD Millipore, cat #300410)

• 1 mL of boiling ddH₂O

• Add the boiling water to the digitonin powder and vortex on high until fully dissolved

• Keep on ice.

• Store at 4 °C for up to 1 week. If precipitation occurs, reheat and vortex to re-dissolve digitonin

• Caution: Be careful when weighing out the digitonin powder, wear a mask as a precaution

Digitonin-Wash (Dig-Wash) Buffer, containing 0.01% (w/v) digitonin

• Mix 20 mL of Wash Buffer with 40 μL of 5% (w/v) digitonin

EGTA, 200 mM

• 7.6 g of EGTA powder (e.g. Millipore cat #324626)

• Add ddH₂O to 90 mL

• Adjust pH to 7.5–8 with NaOH pellets (e.g. Fisher Scientific, cat #S318-500)

• Adjust final volume to 100 mL with H₂O

• Autoclave 30 minutes and store at room temperature for up to 1 year

HEPES, 1 M, pH 7.5 or pH 8.0

• 59.58 g of HEPES powder (Millipore Sigma, cat #5310-OP)

• 150 mL of ddH₂O

• Adjust pH to 7.5 or 8.0 (make separately) with NaOH pellets (e.g. Fisher Scientific, cat #S318-500)

• Adjust final volume to 250 mL with ddH₂O

• Mix, filter-sterilize, and store at 4 °C for up to 1 year

Ice/water Bath (See Figure 3 for example setup)

• Remove a PCR cooler block (e.g. Eppendorf, cat #022510525) from the −20 °C freezer and place it on the bench to warm up for a few minutes

• Sprinkle NaCl granules to the bottom of an ice bucket

• Add 1–2 inches of ddH₂O and swirl to dissolve the salt

• Add ice to fill approximately half of the ice bucket

• Mix in the ice with the salty water

• Add a second layer of salty water by sprinkling more NaCl granules and adding an additional 1–2 inches of ddH₂O

• Fill the ice bucket to the top with ice and mix

• Firmly position the PCR cooler block within the ice/water bath so that the top of the block is level with the top of the ice

• Put a thermometer inside the ice/water bath to monitor the temperature, which should be ~0 °C

• Cover the PCR cooler block with an ice bucket lid and place in a 4 °C room or fridge if possible

Lysogeny Broth (LB)

• 100 g of LB powder (e.g. US Biological, cat #L1505)

• 5 mL of 1 N NaOH

• ddH₂O up to 5 L

• Autoclave for 30 minutes and store at room temperature for up to 6 months

M9 Buffer

• 3 g of potassium phosphatic monobasic (KH₂PO₄) (e.g. Fisher Scientific, cat #P285-500)

• 11.32 g of sodium phosphate dibasic (Na₂HPO₄) (e.g. VWR, cat #JT4062-01)

• 5 g of NaCl (e.g. VWR, cat #MK758106)

• 1 L of ddH₂O

• Stir to dissolve

• Add 1 mL of 1 M MgSO₄ (Moore, 1996) and stir

• Autoclave for 30 minutes and store at room temperature for up to 6 months

Manganese Chloride Solution (MnCl₂), 1 M

• 19.7 g of MnCl₂*4H₂O (e.g. EMD Millipore, cat #105927)

• ddH₂O to 100 mL

• Autoclave for 30 minutes and store at room temperature for up to 1 year

NGM Plates

• 945 mL of ddH₂O

• 21 g of agar (e.g. VWR, BD 214040, cat #90000-764)

• 2.5 g of peptone (e.g. Fisher Scientific, Gibco 211820, cat #DF0118-07-2)

• 3 g of NaCl (e.g. VWR, cat #MK758106)

• Add above ingredients and mix until most of the powder dissolves

• Add 1 mL of 5 mg/mL cholesterol and stir to mix

• Autoclave for 30 minutes

• Add 1 mL of 1 M MgSO₄ (Moore, 1996) and 25 mL of 1 M KPO₄

• Stir until the flask has cooled to around 55 °C

• Add 1 mL 1 M CaCl₂ (Moore, 1996) and 0.5 mL 60 mg/mL streptomycin (or appropriate antibiotic)

• Mix well and pour into sterile 6-cm or 15-cm petri dishes (e.g. VWR, cat #25384-092 (6-cm) and #25384-326 (15-cm))

• Seed with 200 μL or 1 mL of concentrated OP50 culture, for 6-cm or 15-cm plates, respectively

• Allow to dry at room temperature for 1–2 days

• Store at 4 °C when dry for up to 3 months

Optimization Digitonin-Wash (Dig-Wash) Buffer, containing 0.02% (w/v) digitonin (Optional for Optimization)

• Mix 20 mL of Wash Buffer with 80 μL of 5% (w/v) digitonin

Potassium Phosphate Buffer (KPO₄), 1 M

• 53.93 g of potassium phosphate monobasic (KH₂PO₄) (e.g. Fisher Scientific, cat #P285-500)

• 21.57 g of potassium phosphate dibasic (K₂HPO₄) (e.g. Fisher Scientific, cat #P288-500)

• Add 300–400 mL of ddH₂O and stir until completely dissolved

• Add ddH₂O up to 500 mL

• Stir, autoclave for 30 minutes, and store at room temperature for up to 1 year

Propidium Iodide (PI) Solution, 1 mg/mL (Optional for Optimization)

• 10 mg of Propidium Iodide (e.g. Invitrogen, cat #P1304MP)

• 10 mL of ddH₂O

• Dissolve

• Store at 4 °C for up to 6 months protected from light

• Be careful when handling Propidium Iodide as it is a potential mutagen

Proteinase K, 10 mg/mL

• 100 mg of Proteinase K (e.g. Sigma-Aldrich, cat #P2308)

• ddH₂O up to 10 mL

• Freeze aliquots at −20 °C for up to 1 year

Sodium Dodecyl Sulfate (SDS), 10% (w/v)

• 10 g of SDS powder (e.g. Sigma-Aldrich, cat #L4509-500G)

• Dissolve in 80 mL ddH₂O

• Adjust final volume to 100 mL with ddH₂O

• Filter-sterilize and store at room temperature for up to 1 year

Sodium Hydroxide (NaOH), 1 N solution

• 4 g of NaOH (e.g. Fisher Scientific, cat #S318-500)

  • Weigh out carefully as NaOH is corrosive

• Add ~75 mL of ddH₂O

• Mix until completely dissolved

• Add ddH₂O to 100 mL

• Do NOT autoclave

• Keep at room temperature for up to 1 year

• Caution, solution is a very strong base

Spermidine, 2 M

• 34.37 μL of ultrapure H₂O

• 15.63 μL of 6.4 M (undiluted, neat liquid) spermidine (e.g. Sigma-Aldrich, cat #S0266)

• Store at −20 °C for up to 1 month

STOP Buffer, 2X (340 mM NaCl, 20 mM EDTA, 4 mM EGTA, 0.05% (w/v) digitonin, 100 μg/ml RNase A, 50 μg/mL Glycogen)

• 1.709 mL of ddH₂O

• 136 μL of 5 M NaCl (Moore, 1996)

• 80 μL of 0.5 M EDTA (Moore, 1996)

• 40 μL of 0.2 M EGTA

• 20 μL of 5% (w/v) digitonin

• 10 μL of 10 mg/mL RNase A, DNase- and protease-free (Thermo Scientific, cat #EN0531)

• 5 μL of 20 mg/mL Glycogen, RNA-grade (e.g. Thermo Scientific, cat #R0551)

• Store at 4 °C for up to 1 week

Streptomycin Solution, 60 mg/mL

• 3 g of Streptomycin Sulfate (e.g. Fisher BioReagents cat #BP910-50)

• 50 mL of ddH₂O

• Mix to dissolve

• Filter-sterilize and store in aliquots at −20 °C for up to 1 year

Tris-HCl, 1 mM pH 8.0 with 0.1 mM EDTA

• 4.994 mL of ultrapure H₂O (e.g. Invitrogen 10977–015)

• 5 μL of 1 M Tris-HCl, pH 8 (Moore, 1996)

• 1 μL of 0.5 M EDTA (Moore, 1996)

• Store at room temperature for up to 1 year

Wash Buffer (20 mM HEPES, 150 mM NaCl, 0.5 mM Spermidine, 0.1% (w/v) BSA)

• 46.5 mL of ddH₂O

• 1 mL of 1 M HEPES, pH 7.5

• 1.5 mL of 5 M NaCl (Moore, 1996)

• 12.5 μL of 2 M Spermidine

• 500 μL of 10% (w/v) BSA

• 1 tablet, cOmplete^™, EDTA-free Protease inhibitor cocktail (Millipore Sigma, cat #11873580001)

• Mix evenly and dissolve tablet

• Store at 4 °C for up to 1 week, but add the protease inhibitor tablet on Day 1 of CUT&RUN

COMMENTARY

BACKGROUND INFORMATION:

The interplay between DNA, the proteins that package it, and the modifications that adorn it, helps determine the accessibility of DNA and, ultimately, the transcription of genes. To understand the relationship between these factors, techniques to assess the genome-wide binding pattern of protein factors of interest are critical. Such methods can help us understand, for instance, which genes are directly regulated by a transcription factor, or which histone modifications are perturbed by disease. The traditional approach to explore protein-DNA interactions is chromatin immunoprecipitation (ChIP), which remains a powerful tool to understand protein-DNA interactions. ChIP approaches, however, are limited by the large amount of starting material required, making it more difficult to perform with rare samples or in model organisms with relatively low DNA content (Sen et al., 2021).

C. elegans has been a critical model organism for the chromatin biology field. Chromatin factors and histone modifications are well conserved from worms to humans (Wenzel et al., 2011), and researchers have identified numerous histone modifications, chromatin factors, and transcription factors that are associated with development, aging, and more (Cui & Han, 2007; González-Aguilera et al., 2014). ChIP-seq, which allows for a genome-wide localization analysis of proteins of interest and has been utilized in numerous organisms (Ostrow et al., 2015; Chen et al., 2018; Pu & Lee, 2020; Sen et al., 2021; Tran et al., 2012; Soares & Castro, 2018; Sullivan & Santos, 2020), has been less commonly applied in C. elegans due to the significant undertaking of this experiment. For instance, a recently published optimized ChIP-seq protocol recommends using ~600,000 worms for one experiment and, thus, necessarily requires specialized equipment and numerous complex steps to produce quality C. elegans ChIP-seq libraries (Sen et al., 2021).

CUT&RUN represents a significant advancement in the field of chromatin biology, with the ability to generate high-resolution DNA binding profiles from only ~100 human cells and with only 5 million reads (Skene et al., 2018). The CUT&RUN protocol is simple and quick, similar in principle to an immunostaining experiment followed by DNA purification. The Henikoff Lab first developed CUT&RUN in 2017 for use with purified nuclei (Skene & Henikoff, 2017), but has subsequently optimized it for use with intact cells or tissue samples (Skene et al., 2018), making it more accessible for various model systems. The technique quickly became popular in the chromatin biology field because of the ease of use, high quality data with low starting material, and the use of live nuclei without crosslinking, preserving chromatin in a more “natural” state throughout the procedure, compared to many ChIP protocols (Skene & Henikoff, 2017). The C. elegans field, however, has been slow to adopt CUT&RUN, with very few labs having successfully utilized it thus far (Methot et al., 2021). Adopting CUT&RUN for use in C. elegans can be challenging because of the animal’s tough outer cuticle, which forms a protective layer around the organism (Page & Johnstone, 2007) and makes it resistant to standard dissociation protocols. Additionally, C. elegans cells are notoriously sensitive and tricky to culture, and intact cells and nuclei are critical for CUT&RUN success (Skene & Henikoff, 2017).

CUT&RUN experiments in worms consist of the following crucial steps: worm dissociation, antibody binding, pAG-MNase binding, pAG-MNase cleavage, DNA purification, and library preparation. In our hands, performing worm dissociation while preserving cell survival and avoiding sample loss has been the most critical parameter for CUT&RUN success. One option for sample preparation in CUT&RUN is to prepare purified nuclei, as originally described (Skene & Henikoff, 2017). However, more recent CUT&RUN protocols typically utilize cells rather than purified nuclei, eliminating the need for precise nuclear isolation and, thus, simplifying the approach (Skene et al., 2018). CUT&RUN can even work well with intact tissue samples (Uyehara & McKay, 2019), which was particularly appealing to us as it had the potential to allow for maximum sample retention and minimum starting material. Because of this, we chose to optimize a C. elegans cell dissociation protocol for use with CUT&RUN. As previously described (Zhang et al., 2011; Kaletsky et al., 2016; Kaletsky & Murphy, 2020; Methot et al., 2021), we use a Cuticle Disruption Buffer composed of SDS and DTT as a first step to weaken the worm’s outer cuticle, which we find to be critical in allowing for sample permeabilization during the subsequent douncing step. After weakening the cuticle, we wash the worms and place them directly in CUT&RUN Wash Buffer, where we dounce them to increase permeabilization, which allows the antibody to enter the cells successfully during CUT&RUN. By douncing worms directly in CUT&RUN Wash Buffer and immediately binding the resulting worm/cell lysate to ConA beads, we eliminate the need for any spinning or filtration steps which could result in loss or damage to cells. Additionally, we have found the process of completely dissociating worms into cells to be either damaging to cells (if homogenized too much) or very inefficient. Thus, by eliminating the need for complete worm dissociation into a single-cell suspension, we simplify the sample preparation process and ensure healthy cells for use in CUT&RUN.

In contrast to the 100–3000 worms used in the CUT&RUN protocol outlined here, a recently published CUT&RUN protocol in C. elegans that involves several wash steps and cell sorting prior to bead binding calls for the utilization of 200,000 worms (Methot et al., 2021). While this particular technique may work well when cell sorting is required, our protocol allows for CUT&RUN to be performed with only 100 worms or less as starting input. Interestingly, other CUT&RUN protocols in C. elegans have, instead, dissociated worms into single-cell suspensions using the protease cocktail Pronase (Methot et al., 2021), as has been previously described for cell culture and FACS sorting in C. elegans (Kaletsky et al., 2016; Kaletsky & Murphy, 2020). However, we have found this step to be too harsh in our hands, leading to poor cell survival and unacceptably high background in CUT&RUN. One possibility is that the Pronase digestion worked well when coupled with exclusion of dead cells by FACS sorting analysis, keeping cells on ice, and fixation of cells with 100% Ethanol, as utilized by Methot et al. (2021). In contrast, in our protocol, we maintain cells at room temperature until antibody addition, to minimize stress on chromatin, and we do not fix the cells to maintain native chromatin, following the original recommendation by the Henikoff Lab (Skene & Henikoff, 2017; Skene et al., 2018). We find that under these traditional CUT&RUN conditions, Pronase digestion does not work well for most antibodies in our hands.

The remainder of our CUT&RUN steps resembles traditional CUT&RUN protocols published elsewhere (Skene et al., 2018). Notably, we and others (i.e. EpiCypher’s CUTANA^™ CUT&RUN Protocol) prefer to perform the CUT&RUN workflow in 0.2-mL PCR-strip tubes coupled with multi-channel pipetting for wash steps, for ease of use and increased uniformity. This simple modification allows for up to 16 samples to be easily processed in parallel without substantially increasing the time required for each step. Because of our inclusion of worm chunks in addition to released cells, we also recommend modifying wash steps in the CUT&RUN protocol to leave an extra 10–20 μL of buffer at the bottom of the tube at each wash step. This is critical to retain the sample, as some proportion of worms will settle to the bottom of the tube at each wash step rather than sticking to the side of the tube along with the magnetic beads. While it may be tempting to do away with the magnetic beads altogether and only use settled worm material, we do not recommend this strategy. Using a combination of worm chunks and released cells ensures that the majority of cells from the animal are captured, either by the beads or at the bottom of the tube. Using only worm chunks would lead to the loss of any cells that get released from the worms, which will happen with douncing and each subsequent wash step, whereas the inclusion of the beads allows for maximum sample retention.

CRITICAL PARAMETERS:

Antibody and Protein of Choice:

Antibody choice is critical for CUT&RUN success. Five factors must be considered when choosing antibodies for CUT&RUN. First, the antibody must, of course, be specific to the target of interest. Many antibodies are now tested for specificity, and assays used to check for specificity can be obtained by the company producing each antibody. It is also advisable, when first testing a new antibody —particularly for a new protein of interest—, to test several different antibodies, to ensure they all give a consistent biologically meaningful result. For instance, if one wants to obtain the binding profile for a GFP-tagged chromatin-associated protein and has not used an anti-GFP antibody in CUT&RUN before, we recommend trying two to three anti-GFP antibodies to ensure they all give a consistent binding profile for the protein of interest. Secondly, the species in which the antibody is produced is important for CUT&RUN. pAG-MNase contains both protein A and protein G, which bind antibodies from some species more efficiently than others (See Table 7.6 in The Molecular Probes® Handbook, also available at https://www.thermofisher.com/us/en/home/references/molecular-probes-the-handbook/tables/binding-profiles-of-protein-a-and-protein-g.htm). Standard CUT&RUN protocols typically prefer to use antibodies produced in rabbit, which is strongly bound by Protein A. If antibodies produced in rabbit are not available, other species can be used. However, if using a species which is not bound by Protein A/G as strongly, a secondary antibody should be used in addition to the primary (for example, Rabbit anti-Mouse), which will increase the recognition of the protein of interest by Protein A/G (See Alternate Protocol 1).

The antibody must also work well with the CUT&RUN workflow. While many antibodies have been validated as “ChIP-grade” for use in ChIP-seq, these antibodies may not necessarily work well in CUT&RUN due to the differences between the techniques, particularly because ChIP usually involves crosslinking, while CUT&RUN usually does not. Thus, using antibodies specifically validated in CUT&RUN is recommended. If not available, antibodies that work well in ChIP and/or immunofluorescence can also be used for CUT&RUN, although additional optimization may be needed. Antibody optimization can also be performed using a modified immunofluorescence protocol, by staining worms with the primary antibody in CUT&RUN Antibody Buffer overnight, a secondary fluorescent antibody for 1–2 hours the next morning, and evaluating signal-to-noise ratio with a fluorescent microscope. Antibody concentration must also be considered. Typically, we use the standard 1:100 dilution of primary antibody in CUT&RUN Antibody Buffer. However, depending on target abundance and antibody binding, you may need to perform a dilution series, testing antibody concentration from 1:1000–1:10 and evaluating which antibody concentration provides you with the highest resolution results while minimizing background. This can be done first with immunofluorescent staining as described above, and the optimal antibody concentration determined from the staining can be used in CUT&RUN. If the antibody does not perform as expected in the CUT&RUN assay, the dilution series can be repeated in CUT&RUN itself, and the sequencing results can be compared across antibody dilutions.

The proper control for the antibody must also be determined. Typical CUT&RUN protocols use IgG as a control for antibody binding, and normalize the binding of the target of interest to IgG. If this approach is utilized, ensure the IgG is from the same species as the antibody against the target of interest (e.g. rabbit IgG for a rabbit antibody). For histone marks, we also typically perform an anti-H3 CUT&RUN experiment and normalize the histone mark/total H3, which allows us to control for background histone levels throughout the genome. When using a tagged strain for CUT&RUN experiments (e.g. DAF-16::GFP), antibodies can be used in the target worm strain and in a negative control strain not containing the epitope tag (e.g. N2). The signal for GFP antibody in DAF-16::GFP can then be normalized to the GFP signal in N2 to nicely control for the background of the specific antibody, which can be important, as every antibody likely has some level of non-specific background.

Finally, the choice of protein target is important for CUT&RUN. When you are first learning the technique, make sure to include a positive control that you know should work, such as a histone mark. H3K4me3 provides a good positive control, since peak regions are very distinct and the Epicypher antibody works well for CUT&RUN (Epicypher, cat #13-0041). Make sure to test out antibodies against several known targets when first trying the technique, as we have found that when cells are damaged, we are still able to get good profiles for certain histone marks but not others. Testing a variety of active and repressive marks (3–4 total should suffice) will ensure the technique is working as desired. In particular, we have found H3K4me3 profiles to provide a good quality metric for CUT&RUN success (i.e. if cells are damaged, the H3K4me3 profile is likely to have very high background). While sequencing with multiple antibodies may sound expensive, an initial CUT&RUN test with numerous antibodies can be run at one time and pooled together for sequencing in one lane, so the only major cost difference between sequencing one sample and eight samples will be antibodies, pAG-MNase, and library preparation reagents, as most of the working solutions for CUT&RUN will cost the same whether one sample or sixteen samples are run in parallel. After you know the technique is working, you can begin to test your protein of interest, including transcription factors or chromatin factors. Keep in mind the success of the technique will also depend on the factor you choose. For example, if a factor binds more loosely to chromatin, it may be more difficult to obtain a good quality profile, especially since CUT&RUN does not typically use crosslinking.

Growth and Synchronization of C. elegans

The methods used for growing worms vary between labs, but any method to grow the necessary number of worms can be used to prepare worms for CUT&RUN. Here, we present our preferred method of growing large numbers of synchronized worms, which is simple, quick, and synchronizes worms within one day of growth. Many labs bleach gravid adult worms and starve the resulting embryos to obtain highly synchronized L1s that they then seed, when highly synchronized worms are needed (Stiernagle, 2006). However, bleaching and starvation can both induce stress, so if the phenotype being studied could be stress-sensitive, we do not recommend bleaching without first testing if this impacts the phenotype. The method of worm preparation described here avoids any bleaching or starvation, limiting the stress induced by the procedure.

The number of worms required for CUT&RUN will depend largely on the target of interest. Typically, we prepare 1,000 worms per antibody when targeting histone marks, and 3,000 worms per antibody when targeting chromatin factors. Certain targets (e.g. transcription factors expressed only in one tissue) may require more worms, and many targets will require fewer worms. We have obtained detectable signal from CUT&RUN for H3K4me3 with as few as 10 worms (See Figure 5). While we obtained only about half the number of H3K4me3 narrow peaks from 10 worms compared to 100 or 1000 worms, there are certain situations in which starting with a low amount of cellular material may be necessary (e.g. if attempting to use dissected tissue or FACS-sorted cells), and in those situations, it could still be possible to obtain some binding information with CUT&RUN.

As in every worm experiment, the strain, age, temperature, and bacteria should be carefully considered before the experiment. Different strains may grow more slowly or have cuticle phenotypes that impact dissociation. Additionally, because the cuticle is secreted during each larval stage (Page & Johnstone, 2007) and weakens with age (Wolkow et al., 2017), different ages can have different cuticle strengths, meaning the cuticle disruption step may need to be optimized separately for different ages. Because of this, we also recommend against using mixed-stage worms, since cuticle disruption will occur unequally across different age groups. As the protocol described here uses bulk worm samples rather than individual cell types or dissected tissues, the age of the worms used will also determine which tissues will contribute to the final CUT&RUN signal. For example, CUT&RUN signal from gravid adult worms will be a combined result from adult, oocyte, and embryonic signals. If embryos are not desired, different ages can be used, such as larval stage or aged worms, or germline-less mutants can also be employed. However, in our hands, embryos are typically not permeabilized after our regimen, suggesting that the majority of the CUT&RUN signal is not from embryos, even in Day 1 adults (See Figure 2).

Worm Dissociation and Sample Preparation

The efficiency of worm dissociation can be variable depending on the strain or age of worms used. The cuticle disruption time and number of dounces used here were optimized for use in wild-type worms at Day 1 of adulthood, but these conditions may need to be re-optimized for your worms of interest. During the cuticle disruption step, a small fraction of worms may explode by the end of the incubation, but the majority should remain intact. If your strain or age group begins to explode before the end of the incubation, re-optimize the cuticle disruption time by checking worms under the microscope every ~30 seconds, making sure to stop before the majority of worms have exploded but after they mostly stop moving and show a blunted head. Similarly, by the end of the douncing steps in Wash Buffer, you should see many worms in pieces or exploding, and the solution should be cloudy with cells and bacteria. If you see that the worms are completely broken down by the end of 200 dounces, consider re-optimizing by checking for dissociation every 25–50 dounces, as too much douncing may damage cells and lead to high background in CUT&RUN. If worms are not at all broken down by the end of 200 dounces, continue douncing and check every 25–50 dounces for dissociation. It can also be helpful to check for permeabilization of worms at the end of the douncing regimen, as described in Steps 14–17 of Basic Protocol 2, especially if the signal is either too low or the background is too high in CUT&RUN.

Another consideration for CUT&RUN is digitonin concentration. Here, we use a very low percentage of digitonin in the Antibody Buffer and Digitonin-Wash Buffers (0.01% (w/v)), because we have found that increasing digitonin concentration above 0.01% (w/v) did little to increase the permeabilization of the worm cells. In our hands, douncing made the biggest difference in terms of uniform Propidium Iodide staining while allowing for cells to stay intact. However, digitonin concentration can be re-optimized in your lab by performing the Propidium Iodide staining described above with varying digitonin concentrations.

During the worm dissociation and sample preparation steps, all buffers should be at room temperature, to minimize stress on chromatin (Skene et al., 2018). We strongly recommend using M9+Tween buffer for the initial wash steps, as Tween prevents worms from sticking to the plastic pipette tips and tubes. This is especially critical if working with low numbers of worms. Similarly, although inclusion of BSA is typically optional in the CUT&RUN Wash Buffer, we strongly recommend its inclusion, as it prevents worms from sticking to the tube during the douncing steps as well as to the pipette tips later on in every mixing step of CUT&RUN.

During worm dissociation, it is best to work as quickly as possible to minimize the time from when worms are on their plates until the worm/cell mixture is in Antibody Buffer, as the EDTA will help stabilize the cells from any further changes (Skene et al., 2018). It is especially essential to work quickly during the wash steps after the cuticle disruption step, as keeping the worms in the SDS/DTT mixture for too long can lead to increased worm exploding, cell loss, or cell damage.

CUT&RUN Chromatin Profiling

As outlined in the original CUT&RUN protocol (Skene & Henikoff, 2017), the MNase cleavage step is one of the most critical steps to achieve success and maintain low background. Calcium should only be added once the ice/water bath has reached a temperature of 0 °C and the tubes have cooled down sufficiently. See Figure 3 for a depiction of the experimental setup of this crucial step.

During the CUT&RUN process, it is essential to remember that not all worm chunks will successfully bind to the magnet, so during each wash step you should leave 10–20 μL at the bottom of the tube to ensure sample preservation. Adjust the volume of the subsequent incubation steps accordingly, as instructed in Basic Protocol 3. Also, because worm chunks tend to settle to the bottom of the tube, we recommend mixing by pipetting immediately before incubation steps on the tube nutator (i.e. overnight antibody incubation, pAG-MNase incubation), to ensure the sample is properly mixed during incubation. Similarly, it is important that at the final DNA release step (Basic Protocol 3, step 26), you ensure that you do not carry over any worm chunks that have fallen to the bottom of the tube into your DNA isolation and purification steps. This is easily achieved by a second transfer (Basic Protocol 3, step 27), since large worm chunks settle to the bottom of a tube within ~30 seconds. By transferring the DNA supernatant twice and checking under a microscope, you can ensure that no large worm chunks are carried to the DNA isolation stage, decreasing the risk of high background from extra uncut DNA.

If you want to use CUT&RUN to quantitatively compare amounts of binding between different samples, the inclusion of a spike-in control is necessary. Without a spike-in, profiles can be compared to each other relative to an internal control (such as H3, for example), but these comparisons will still be qualitative, rather than quantitative, in nature. Frequently, CUT&RUN users use the amount of E. coli DNA remaining from pAG-MNase preparation as a normalization for CUT&RUN (Meers et al., 2019), however, due to the presence of E. coli DNA in the C. elegans gut from its use as a food source, we do not believe this is an appropriate normalization control for worms. If you would like to include a spike-in control for your experiment, we recommend using Drosophila or Saccharomyces cerevisiae micrococcal nuclease-treated chromatin in the STOP Buffer, as originally described (Skene & Henikoff, 2017; Skene et al., 2018), or including cells from another organism in your original sample, as described by the Henikoff Lab (Sarthy et al., 2020).

Finally, if you do not obtain any detectable amounts of DNA after the CUT&RUN protocol, this does not necessarily mean the procedure has failed. When using a small number of worms (i.e. 100 or less), it will not be surprising to obtain non-quantifiable DNA even from an abundant histone mark. Proceed to library preparation with between half and the entire sample, using at least 14 cycles of PCR amplification. You should obtain detectable DNA after library prep if the procedure was successful. If using a small number of worms, it can also be helpful to have one positive control with a few thousand worms in parallel, to ensure DNA is obtained from the positive control as expected.

CUT&RUN Library Preparation for Illumina Sequencing

During library preparation, carefully pipette and treat all samples the same. The use of a mastermix can be helpful here, although note that NEB does NOT recommend adding adaptor to a mastermix during the Adaptor Ligation step; rather, it should be added directly to the End Prep Reaction Mixture in Step 6 of Basic Protocol 4. Similarly, index primers during step 20 of library amplification cannot be added to a mastermix, as they must be unique for each sample that will be pooled together in one sequencing lane.

The number of PCR cycles used for amplification during library preparation will be variable depending on the starting DNA concentration. We typically use 14 cycles for library amplification, and find that anything over 14 can lead to excessive PCR duplicates in the final analysis. Similarly, adaptor dilution can be adjusted according to experimental results. We typically use a 25-fold dilution of adaptors, but with undetectable input DNA, a 50-fold dilution may be more appropriate. However, keep in mind that too little adaptor can limit library complexity if there is not enough adaptor present for the DNA fragments of interest, so adaptor should always be present in excess.

When cleaning DNA with Ampure XP beads, make sure beads are at room temperature before adding them to the DNA sample, ideally by taking them out ~30 minutes or more before use. When drying the beads, make sure to dry thoroughly, to prevent ethanol from contaminating the sample. However, do not over dry the beads to the point of cracking, as this can reduce DNA yield. The ratio of beads to DNA volume is critical in determining the size selection capability of the beads. A size selection of 1.2X (i.e. a ratio of beads to DNA solution of 1.2:1) will usually remove residual adaptor while keeping the library intact (See Figure 4C–D for Bioanalyzer traces before and after size selection). Increasing this ratio would lead to the inclusion of smaller DNA fragments, while decreasing the ratio leads to the exclusion of small DNA fragments.

While high-throughput sequencing is the preferred approach to test whether CUT&RUN is working as planned, users can also consider testing their CUT&RUN libraries with fluorescence-based quantitative real-time PCR (qPCR) to evaluate whether enrichment in predicted binding regions can be seen in the test condition compared to control (e.g. H3K4me3 vs H3). To test samples with qPCR, library-prepped samples should be used (after Basic Protocol 4) to avoid amplification of large genomic DNA fragments likely to be present in the DNA samples before library preparation (at the end of Basic Protocol 3) (Janssens & Henikoff, 2019). Additionally, primers must be carefully designed and tested, as DNA fragments produced by CUT&RUN are small (often less than 120 bp, especially for transcription factors) due to the precise cutting of the MNase (Janssens & Henikoff, 2019). Therefore, we prefer the approach of pooling all CUT&RUN libraries prepared for optimization purposes together in one sequencing lane, unless precise peak regions are already well characterized for the protein of interest.

Sample Basic Data Analysis Using Linux

As recommended by Zhu and colleagues, we do not typically remove PCR duplicates as would be done in ChIP-seq analysis, because it is believed that the precision of MNase cutting can lead to duplicate DNA fragments being released from independent cells (Fu et al., 2018; Zhu et al., 2019). Specifically, because MNase cuts in a stereotypical manner, and because the fragments produced by CUT&RUN are so small, there is a possibility that identical sequencing reads result from independent cleavage events of the same DNA segment in multiple cells. In this case, reads appearing as duplicated are actually real signal obtained simultaneously and amplified together, rather than PCR duplicates originating from a single DNA cleavage event. However, duplicated reads can be removed on an individual basis depending on the factor studied.

Depending on the target (transcription factor versus histone mark), it may also be preferable to bioinformatically separate DNA obtained by CUT&RUN based on size. This can be done in Linux by using samtools and the awk command to manipulate the bam files produced after genomic alignment. The small fragments (<120bp) will represent direct binding targets of transcription factors, while larger fragments (>120bp) will represent the surrounding chromatin environment (Skene & Henikoff, 2017).

TROUBLESHOOTING

See Table 1 for a list of common problems with the protocol, their causes, and potential solutions.

Table 1.

Troubleshooting Guide for C. elegans dissociation and CUT&RUN.

Problem	Possible Cause	Solution
Insufficient numbers of embryos are obtained in step 11 of Basic Protocol 1	Worm strain used gives fewer progeny than N2; Temperature used differs from 20 °C Too many embryos washed off the plate in steps 4–5 of Basic Protocol 1 along with the adult population to be discarded	Increase the number of parental worms picked per plate in step 1 of Basic Protocol 1; Increase the number of plates prepared in step 1 of Basic Protocol 1; For slow-growing strains, increase the incubation time in step 3 of Basic Protocol 1 until plates are full of gravid adults. If embryos are being washed off the plates prematurely in steps 4–5 of Basic Protocol 1, save the adult population in a sterile tube, and allow the adult worms to settle to the bottom of the tube for ~30 seconds. Transfer the top ~90% of the supernatant to a new tube – this will contain primarily embryos and L1s.
Embryos stick to the pipette tips during embryo collection in Basic Protocol 1	Insufficient bacteria on the plate in step 6; Use of M9 instead of M9 + 0.05% (v/v) Tween in step 8 of Basic Protocol 1	Ensure that when embryos being pipetted, they are coated in either bacteria or M9 + Tween to avoid sticking. Tween percentage can be increased up to 0.1% (v/v) if necessary.
Worms are not well synchronized after step 13 of Basic Protocol 1	Adults and larvae were present in the embryo suspension seeded on plates in step 12 of Basic Protocol 1	Repeat step 5 of Basic Protocol 1 to wash all adults and larvae off the plates. Check under the microscope to ensure that only embryos (and some L1s) are present on the plates before washing off embryos. Check 15-cm plates after seeding them to ensure that only embryos (and some L1s) were seeded. Pick off any stray adult worms or older larvae that were seeded.
Worms are not well dissociated after Basic Protocol 2	Genotype or age of worms are more difficult to dissociate than N2 Day 1 adults	Optimize worm dissociation conditions – check dissociation efficiency under the microscope every 25 dounces for your strain or age of interest or increase cuticle disruption time. Follow Basic Protocol 2 Steps 14–17 to find the minimum amount of dissociation that leads to uniform permeability.
Worms break apart too quickly during Basic Protocol 2	Genotype or age of worms are less difficult to dissociate than N2 Day 1 adults	Optimize worm dissociation conditions – check dissociation efficiency under the microscope every 25 dounces for your strain or age of interest or decrease cuticle disruption time. Follow Basic Protocol 2 Steps 14–17 to find the minimum amount of dissociation that leads to uniform permeability.
ConA Beads clump during CUT&RUN in Basic Protocol 3	Cells lyse during the reaction Excessive cellular material on the beads (>>3,000 worms)	If beads clump and background in sequencing is high, optimize worm dissociation conditions – consider douncing fewer times or reducing cuticle disruption time if cells become damaged. Reduce worms used for each reaction.
Worms visibly stick to pipette tips during steps involving mixing by pipetting in Basic Protocol 3	BSA was left out of Wash Buffer or is at too low a concentration	Ensure BSA was included in the Wash Buffer. If it was included, remake fresh 10% (w/v) BSA and try again, or increase BSA concentration in the Wash Buffer.
Worms are lost during CUT&RUN wash steps (as evidenced by worm chunks seen in the pipette tip before discarding liquid during wash steps) in Basic Protocol 3	Too much liquid being withdrawn during wash steps; Too many worms used per CUT&RUN reaction	Remove less liquid during wash steps (keep up to 20 μL or slightly more if necessary). Carefully watch the worms on the magnet stand during each wash step, to remove the buffer with minimal worm loss. If it is unclear whether worms are being lost, save the beads containing cells and worm chunks that would normally be discarded at step 26 of Basic Protocol 3. Freeze the beads at -20 °C and return when convenient to count the real number of worms used per reaction.
The ice/water bath does not reach temperatures as low as 0 °C in Basic Protocol 3	Not enough salty water present in the ice/water bath	Add more NaCl and water to the ice/water bath. Ensure that the final mixture of salt, ice, and water is very wet. Keep ice/water bath in the 4 °C room with the top covered.
No or low (undetectable by Qubit) DNA detected before library prep (although this does not necessarily indicate a problem)	Low protein abundance; Low worm number; procedural failure	Proceed to library prep anyways and run a positive control for a histone mark in parallel to ensure the procedure worked.
No or low (<1 ng/μL) DNA detected after library prep	Too few worms used for the protein of interest Too few PCR cycles in library prep Failed library prep; Failed CUT&RUN	If low cell numbers were used and DNA is detectable, perform Bioanalyzer to check for size distribution and continue to sequence if possible; Increase worm number and use a positive control. Increase PCR cycle number, if using a low number of cycles, to increase DNA yield. If positive control fails, carefully check reagents and protocol.
High adaptor or primer content after library prep	Low DNA input	Increase DNA input to library prep; Reduce primer or adaptor concentration when preparing the library; Clean libraries with 1.2X Ampure XP beads an additional time to remove residual primers and adaptors.
A large proportion of amplified CUT&RUN library fragments (>10%) contains genomic DNA	Unsuccessful size selection while cleaning up PCR-amplified library in steps 22–24 of Basic Protocol 4 Carryover of worms or worm chunks to the DNA purification stage in steps 26–27 of Basic Protocol 3 Cell damage leading to excessive DNA leakage from cells in Step 24 of Basic Protocol 3	Repeat 0.5X size selection as in Steps 22–24 of Basic Protocol 4. Ensure that worms are not improperly carried over with the released CUT&RUN DNA fragments in steps 26–27 of Basic Protocol 3. If problem occurs persistently and sequencing data are low quality, carefully check cell dissociation as in the troubleshooting section for high background in sequencing.
Excessive PCR duplicates (>65% duplicated sequences) in sequencing results	Too many PCR cycles used in library preparation	As long as sufficient DNA quantity is produced by library preparation, reduce PCR cycles used in library prep. If this is not possible, start Basic Protocol 2 with more worms to obtain more DNA and then lower the cycle number in library prep.
Low alignment rate to the C. elegans genome	Low worm cell number relative to contaminating gut bacteria Antibody also has targets in bacteria present in the C. elegans gut	Increase worm number; Increase washes to remove bacteria when initially washing worms from plates Ensure that the antibody does not target bacterial proteins. This can be done by using the immunofluorescent staining method described in Critical Parameters with worm chunks, and checking if any of the bacteria from the animal’s gut (which can be observed as tiny, moving particles) also stain with the antibody.
No or low signal in sequencing (i.e. no or few peaks called by MACS2)	Antibody does not specifically bind the target of interest in CUT&RUN conditions or requires a higher concentration Protein abundance is low Antibody is not easily bound by pAG-MNase	Optimize antibody binding conditions with immunofluorescence; Try additional antibodies for the same target. Add more worms. Add a secondary antibody for primary antibodies not easily bound by pAG-MNase.
High background in sequencing (i.e. lots of noise present between peak regions when viewed on IGV)	Cell lysis during the reaction Nonspecific antibody binding; Insufficient antibody Excessive cleavage by pAG-MNase	Carefully check cell dissociation. Check if cells survive well overnight by counting on a hemocytometer. If cells are lost or lyse by the next day, decrease the harshness of the dissociation protocol (less time in Cuticle Disruption Buffer, fewer dounces, etc.). Optimize antibody binding conditions by immunofluorescence; Ensure antibody incubation and pAG-MNase binding steps are at 4 °C continuously. Ensure pAG-MNase cleavage step is at 0 °C throughout the reaction.

UNDERSTANDING RESULTS

We have described here our method for generating binding profiles for histone marks or chromatin factors by CUT&RUN in C. elegans. In this section, we discuss example results generated by our lab with the protocol described for several histone marks and chromatin factors of interest to us.

Evaluation of libraries after library preparation is the first opportunity to determine if the CUT&RUN protocol was likely successful. If library preparation worked, you should see that the concentration of DNA measured by Qubit^™ is higher than before library prep, and the majority of the library should be between ~180bp-600bp (See Figure 4). If the library contains primarily large genomic DNA fragments (>1000bp), the process was likely unsuccessful, as you have primarily captured uncut DNA leaking out of the cells. If only a small fraction of genomic DNA is present (<10% of the total library), libraries can still be sequenced (See Figure 4A–B), or can be subjected to an additional 0.5X Ampure XP bead size selection to remove large genomic DNA. Often, if CUT&RUN worked well for histone marks, a nucleosomal laddering pattern will be present in the Bioanalyzer trace (Figure 4A); however, absence of this clear pattern should not be taken as the only sign that the technique was unsuccessful. Finally, high contaminating adaptor or primer dimer in libraries (more than 5–10%) should be removed with an additional 1.2X size selection (See Figure 4C–D), as adaptor dimers can bind to the sequencing flow cell and disrupt the sequencing of your sample.

When trying CUT&RUN for the first time, we recommend targeting a histone mark. It is also a good idea to process ChIP-seq data for the same target in parallel, using publicly available data, either from existing publications or from modENCODE. Visualize the tracks of ChIP and CUT&RUN together to make sure the majority of expected peaks are present in your CUT&RUN data. Here, we show an example of ChIP-seq data for larval stage 4 wild-type worms generated previously by our lab (Wang et al., 2018) compared to CUT&RUN data for H3K4me3 in Day 1 adults generated with the protocol outlined here (See Figure 5). High-resolution profiles with very similar regions of enrichment to ChIP-seq were obtained with CUT&RUN. Notably, the H3K4me3 profile generated by CUT&RUN is detectable with only 10 worms, while high-resolution profiles were obtained with 100 worms or 1000 worms.

It is also a good idea to test antibodies against several different histone marks to ensure that CUT&RUN is performing as expected. For example, we show binding profiles of H3K9me3, H3K27me3, H3K4me3, and H3K9ac (See Figure 6). As expected, we see that repressive marks (H3K9me3 and H3K27me3) typically occupy distinct chromatin domains from active marks (H3K4me3 and H3K9ac), indicating that with this protocol, one is able to successfully capture the binding profile of a variety of histone marks.

Figure 6. CUT&RUN in C. elegans can capture both active and repressive histone marks.

CUT&RUN binding profiles for the repressive marks H3K9me3 and H3K27me3 are plotted alongside the active marks H3K4me3 and H3K9ac. Each track displays the log2 fold change of the histone mark relative to H3 control levels across the genome. Blue indicates enrichment, while red indicates depletion relative to H3. In general, active and repressive marks can be seen to occupy distinct regions. For example, (A) shows regions of active and repressive marks trading off, while (B) shows a large region of chromatin overall enriched for repressive marks and depleted for active marks. Antibodies used were: H3K9me3 (abcam, cat #ab176916), H3K27me3 (EMD Millipore, cat #07-449), H3K4me3 (Epicypher, cat #13-0041), H3K9ac (Cell Signaling Technology, cat #9649), and H3 (abcam, cat #ab1791). Wild-type worms of the N2 strain were used for sample preparation on Day 1 of adulthood. CUT&RUN tracks represent data from one biological replicate.

When trying CUT&RUN against a chromatin or transcription factor for the first time, it is also preferable to try it against a target with a known chromatin binding profile, for example, a factor that has a publicly available ChIP-seq profile. If you are testing the binding profile of a protein target with an unknown binding profile, we recommend testing at least two antibodies for the protein of interest, to make sure that you obtain consistent results. Here, we show our lab’s CUT&RUN data for two of our chromatin factors of interest, the highly similar and paralog H3K4me3 readers SET-9 and SET-26 (Wang et al., 2018). Using a C. elegans strain containing both tagged SET-9 and SET-26, we obtained a CUT&RUN profile for our factors (See Figure 7) and compared the profile to ChIP-seq data previously generated by our lab (Wang et al., 2018). As expected, the majority of peaks for SET-9 and SET-26 coincide between the ChIP-seq and CUT&RUN data, indicating that the CUT&RUN protocol described here works well for both histone marks and chromatin factors. Importantly, the CUT&RUN data for SET-9 and SET-26 are actually of higher resolution than the ChIP-seq data previously generated from our lab, which can be observed by the clear cutoffs between regions of enrichment (blue) and depletion (red) in the CUT&RUN data, which are not as clearly distinguishable in the ChIP-seq data. We also obtain more peaks using CUT&RUN to determine the binding profile of SET-9 and SET-26 than if we process the ChIP-seq data in parallel, suggesting we are able to uncover a more complete range of SET-9 and SET-26 binding sites, some of which were probably masked due to the lower-resolution data previously obtained by ChIP-seq. Importantly, this suggests that CUT&RUN will allow for more complete binding profiles of transcription factors and chromatin factors that were previously difficult to obtain with ChIP-seq, and may allow researchers to obtain binding profiles for factors which previously did not provide enough signal to detect in ChIP (e.g. a transcription factor only expressed in a few cells).

When visualizing your data to determine if CUT&RUN worked, make sure to always visualize a negative control and/or normalize your samples to background antibody binding. Every antibody will have background binding, even if the epitope it binds is not present in the strain of interest. For example, to obtain a SET-9/26 binding profile, we use a SET-9/26-tagged strain (set-9::GFP set-26::GFP). We visualize the binding profile of SET-9/26 by comparing the binding of the GFP antibody in the SET-9/26-tagged strain to the binding of the GFP antibody in N2 worms. Since the GFP antibody should not bind specifically in N2, we get the antibody background. In Figure 8, you can see that even without GFP present, some signal is obtained from using the antibody in N2 worms, but by normalizing the SET-9/26 sample to N2, we can make sure we do not falsely include background peaks as SET-9/26 binding regions, thus obtaining the most biologically meaningful data.

TIME CONSIDERATIONS

For Basic Protocol 1, growing worms can take a variable amount of time depending on the strain, temperature, and age desired. Typically, we keep our strains actively growing and well fed at 20 °C, so that from the time of picking 3–7 worms (Basic Protocol 1 Step 1) to the first day of CUT&RUN, is 6–7 days.

Generally, we divide the CUT&RUN protocol into 3 days for convenience, and to provide for optimum antibody incubation time and DNA recovery. Day 1 consists of Basic Protocol 2, worm dissociation, and the beginning of Basic Protocol 3, including all steps up to the overnight antibody incubation. Typically, this requires ~1 hour of buffer preparation before starting, and when working with multiple strains, can take around 1–1.5 hours for sample dissociation, bead binding, and antibody addition. Samples are then incubated overnight at 4 °C. Day 2 of CUT&RUN consists of the remainder of Basic Protocol 3 until the overnight ethanol precipitation step (Step 35), which typically takes between 6–8 hours total, depending if a secondary antibody binding step is also needed. However, note that 3–4 hours of CUT&RUN Day 2 consist of incubation waiting time. Day 3 of CUT&RUN consists of the final ethanol precipitation steps, which typically take only ~1 hour, and Qubit^™ quantification, which takes only ~15 minutes to prepare. If desired, Day 3 could be combined with either Day 2 (by shortening the ethanol precipitation to a few hours) or with library preparation.

Library preparation typically takes 4–6 hours total, and depending on the availability of the Bioanalyzer, it can take another few days to analyze the libraries before pooling them for sequencing. Once the data are back, and depending on the number of samples, it can take 1–2 days to process the data to the point of generating bigwig files and calling peaks to determine if the procedure worked. Altogether, in our lab, it typically takes ~4 weeks from the time of experimental inception until the time of determining whether the experiment worked, though the hands-on working time is more like one week in total.

Supplementary Material

Video 1.

Demonstration of CUT&RUN wash steps. Example of any of the CUT&RUN wash steps in Basic Protocol 3. Worms are bound to beads and have already been allowed to separate for 5 minutes prior to start of video. Using a multichannel pipette, the majority of the supernatant is removed, leaving ~10-20 μL per tube on the magnet. The supernatant is inspected for accidental worm removal and then discarded. Then, 200 μL of Dig-Wash Buffer is added to each tube, and samples are thoroughly resuspended before being placed back on the magnet stand for an additional 5 minutes of separation time.

DATA AVAILABILITY STATEMENT

The data, tools and material (or their source) that support the protocol are available from the corresponding author upon reasonable request.