Dissection of Enhancer Function Using Multiplex CRISPR-based Enhancer Interference in Cell Lines

1. Generation of Cell Lines Stably Expressing SID4X-dCas9-KRAB

Note: The transfection conditions and drug concentrations presented here have been optimized for Ishikawa cells, an endometrial cancer cell line, grown in RPMI 1640 media supplemented with 10% FBS and 1% penicillin/streptomycin (complete RPMI). Other cell lines may require different transfection conditions and drug concentrations. Users can also perform transient transfection experiments in wild-type cells, instead of generating a stable cell line, with a plasmid expressing SID4X-dCas9-KRAB along with guide RNA expressing plasmids; however, results from transient transfections may be difficult to reproduce as SID4X-dCas9-KRAB levels may vary by transfection.

1. Plate Ishikawa cells in at least 2 wells of a 6-well plate at 30 – 50% confluence (approximately 300,000 Ishikawa cells) in 3 mL of complete RPMI.
   1. Aspirate the media from the cells. Wash the cells once with 1x PBS (pH 7.4). Aspirate the PBS and add trypsin (4 mL for a 10 cm dish or 5 mL for a T-75 flask).
   2. Incubate the cells for ~5 min at 37 °C, checking every 2 min for detached cells and gently shaking the vessel.
   3. Once cells have detached, pipette trypsinized cells up and down a few times, and pipette gently down the side of the vessel to release any attached cells.
   4. Transfer the cells to a 15 mL conical tube and spin the cells down for 5 min at 250 x g.
   5. Aspirate the trypsin and resuspend the cells in 5 – 10 mL of media. Use a P1000 pipette to dissociate cell clumps if necessary.
   6. Count the cells and determine the volume needed to plate ~300,000 cells per well in a total volume of 3 mL. Add the cells to 2 separate wells of a 6-well plate. Fill each well to 3 mL with complete RPMI.
   7. Gently shake the plate every 5 min in the first 15 min after plating to ensure that the cells are evenly distributed on the plate. Use a microscope to ensure that the cells have dispersed from the middle of the well.
2. Within 24 h of plating, perform the following transfections using an appropriate transfection reagent for the cell line of interest. For Ishikawa cells, use the procedure as described below.
   Note: This protocol assumes the use of cationic liposome-based transfection reagents. Electroporation provides an alternative method for cell types that are very sensitive to these reagents or that exhibit low transfection efficiency with lipofection. Transfection conditions must be optimized for the cell line of interest prior to attempting Enhancer-i experiments.
   1. In a 1.7 mL Eppendorf tube, dilute 2.5 μg of SID4X-dCas9-KRAB plasmid and 800 ng of plasmid expressing a fluorescent protein into serum-free media such that the final volume in the tube is 155 µL and the final concentration of plasmid is 0.020 µg/µL.
   2. In another tube, dilute 3.3 µg of a plasmid that does not contain a neomycin resistance cassette, such as pCMV-GFP, in serum-free media such that the final volume in the tube is 155 µL and the final concentration of plasmid is 0.020 µg/µL.
   3. Vortex each tube briefly and spin down using a microfuge.
   4. Add 9.9 µL of the transfection reagent (Table of Materials) to each tube. Mix by vortexing briefly at a low speed. Spin the tubes down with a microfuge.
   5. Incubate the tubes at room temperature for at least 5 min, but no more than 20 min.
   6. In the biosafety cabinet, add 150 µL of the prepared DNA:reagent mix dropwise to one well on the 6-well plate. Repeat for the other tube of prepared DNA:reagent mix. Mix the plates by swirling gently and return the plate to the incubator.
3. At day 2 post transfection, change the media and supplement with G418 to a final concentration of 600 ng/μL. This concentration may need to be optimized for the cell type.
4. Change complete RPMI media and supplement with G418 every other day for 2 – 4 weeks until the control transfected cells are dead and the wells containing SID4X-dCas9-KRAB become confluent. The exact amount of time needed for cells to recover will depend on the doubling time of the cells.
5. When the cells become confluent, passage to a T-25 or T-75 vessel in complete RPMI with a lower dose of G418 (300 ng/μL for Ishikawa cells). During this passage, make 2 aliquots of ~100,000 cells each (roughly 1/10^th of a 6-well plate) into 2 separate 1.7 mL Eppendorf tubes for RNA and DNA isolation, respectively. Spin these tubes down (5 min, 250 x g), remove trypsin by pipetting, and freeze the tubes at -20 °C for future use.
6. Isolate genomic DNA using commercially available kits and perform PCR using the “pAC95_PCR” or “SID4X_PCR” primers (Table 1) to verify the presence of the fusion protein within in the cell line. Use genomic DNA extracted from the parental line as a negative control, and SID4x-dCas9-KRAB plasmid DNA as a positive control. Use a high-fidelity polymerase master mix with 50 – 100 ng of genomic DNA and the following cycling conditions: 98 °C for 30 s, 25 cycles of (98 °C for 10 s, 58 °C for 30 s, 72 °C for 2 min), 72 °C for 5 min, hold at 4 °C.
7. To verify the expression of the fusion protein at the RNA level, perform qPCR with RNA extracted from the cell line using commercially available kits. Use the “dCas9_qPCR” primers (Table 1), and the one-step qPCR protocol provided in step 6.3 of this protocol.
8. To verify the protein-level expression of the fusion, perform a Western blot on lysates from the cell line. Use either anti-FLAG or anti-HA antibodies to detect the fusion protein.

2. Guide RNA Design

Note: This protocol is designed for use with the U6 guide RNA cloning vector created by the Church lab and available on Addgene (Addgene 41824). To create a version of this vector containing puromycin resistance that allowed for the same cloning strategy as 41824, we moved the multiple cloning site from this vector into the pGL3-U6-sgRNA-PGK-puromycin vector (Addgene 51133). Either Addgene 41824 or our version with puromycin (Addgene 106404) are compatible with the cloning strategy outlined below.

1. Obtain 600 – 900 basepairs of DNA sequence for each regulatory region of interest. Use transcription factor binding sites and/or chromatin accessibility for guidance as to where to define the region of interest (Figure 2A).
   Note: While the example in Figure 2A features upstream and downstream enhancers, it is also possible to target regulatory elements located within introns.
2. Place all sequences obtained in a single text file using the FASTA format.
3. Identify at least one negative control region that is not expected to change over experimental conditions, such as a promoter of a gene that is not expressed in the cell line of interest. Obtain DNA sequence for this region and add it to the text file in FASTA format.
   Note: We use guide RNAs targeting the IL1RN promoter²⁵ as a negative control for all regions we target. Users can also select intergenic sequence near the region of interest that does not contain transcription factor binding sites as a negative control. However, if multiple loci are being targeted simultaneously, a single negative control region simplifies experimental design and interpretation of results. If the targeted enhancer is intronic, it may be useful to target an intronic region at the same locus that does not contain a putative regulatory element as an additional negative control, as the dCas9 fusion may interfere with transcription.
4. Identify positive control regions, such as the promoters that are the putative targets of the regulatory regions of interest, or promoters of genes that are highly transcribed in the cell line of interest. Obtain DNA sequence for these regions and add it to the text file in FASTA format.
5. Use a program such as e-crisp²⁶ (http://www.e-crisp.org/E-CRISP/) on the DNA sequences generated to find guide RNAs with low off-targets (ideally 0-3). Guide RNAs consist of 20 nucleotides upstream of a protospacer adjacent motif (PAM), which takes the form “NGG” for the dCas9 from S. pyogenes.
   1. On the e-crisp website, select the organism of interest using the drop-down menu. The genome assembly appears to the right of the species name.
   2. Select the Input is FASTA sequence radio button. Copy the FASTA sequences from above and paste them into the dialog box. Ensure that a FASTA header is included for each sequence.
      Note: Up to 50 sequences can be queried simultaneously.
   3. Select the Medium radio button and Single design in the drop-down menu.
   4. Click the button Start sgRNA search. A new browser tab will open, and results will be displayed. Download the candidate sequences by clicking the button Download an Excel formatted tabular report for all query sequences together.
   5. Open the tabular report using Excel or a text editing program.
6. Use the UCSC genome browser to BLAT candidate full-length gRNA sequences (23 basepairs) to the genome.
   1. In a browser, navigate to the UCSC genome browser website (http://genome.ucsc.edu). Under the section Our tools, locate the word BLAT and click on it. The BLAT Search Tool will open.
   2. Use the drop-down menus located under the BLAT Search Genome text to select the organism and genome assembly of interest.
   3. Copy the guide RNA sequences from the tabular report generated by e-crisp and paste them into the dialog box. Ensure that each sequence has a unique FASTA header, then click 제출 at the bottom of the dialog box.
      Note: Up to 25 sequences can be examined at once. On the BLAT Search Results page, alignments of each guide RNA sequence will appear, with each line representing an alignment. Ideally, there should be one alignment for each guide RNA, indicating the uniqueness of that guide RNA.
   4. Avoid the guides that align to multiple locations in the genome if possible. 
   5. To examine guide RNA localization and distribution within the region of interest, click on the Browser link under the ACTIONS section for one of the queried guide RNAs. The Genome Browser will appear and will be centered on the selected guide RNA. Use the Zoom out buttons at the top of the page to visualize the distribution of other guide RNAs identified by e-crisp within the region of interest.
7. Select 4 preferably non-overlapping guide RNA sequences that are distributed throughout the region of interest (Figure 2B). If the region of interest exceeds 600 bp, consider adding 1 – 2 additional guides. Avoid guide RNAs with homopolymeric stretches and extreme GC content, as these features can hinder the guide RNA cloning process and reduce guide RNA targeting efficiency.
8. Once the guides have been selected, create a file containing the full guide RNA sequence (23 nucleotides) for each desired guide, and then remove the 5’ nucleotide as well as the PAM (NGG) from the 3’ end. This step facilitates oligo ordering.
9. Add the following sequence to the 5’ end of the oligonucleotide sequence: GTGGAAAGGACGAAACACCG.
10. Add the following sequence to the 3’ end of the oligonucleotide sequence: GTTTTAGAGCTAGAAATAGC.
    Note: The final sequence should be 59 nucleotides long and look like this: GTGGAAAGGACGAAACACCG-target (19 nt)-GTTTTAGAGCTAGAAATAGC.
11. Ensure that each regulatory element to be targeted with Enhancer-i has at least 4 unique oligonucleotides designed for it. Order these sequences along with the “U6_internal” primers listed in Table 1.

3. Guide RNA Cloning

Note: Guide RNA cloning via Gibson assembly has proven to be highly efficient in our hands, yielding hundreds of colonies per plate, with few if any colonies present in the vector only control. Such efficiency is critical for maintaining complexity during pooled cloning. Another advantage of Gibson assembly cloning is that users do not have to worry about the presence of a restriction enzyme cut site in the guide RNA they are trying to insert into the U6 cloning vector. Nonetheless, this protocol can be adapted for traditional restriction enzyme based cloning if desired.

1. Reconstitute guide RNA oligos at a final concentration of 100 μM in ultrapure water (RNase-free, DNase-free). There should be at least 4 separate guide RNA oligos for each region of interest.
2. For each regulatory region of interest, create a pool of all the oligos corresponding to the region of interest. In an Eppendorf tube, combine 5 μL of each individual reconstituted guide RNA oligo for each region. Mix the pool well by vortexing, then remove 1 μL and dilute this aliquot 1:200 in ultrapure water.
   Note: If desired, these pools targeting individual regulatory regions can be further combined to generate a complex pool targeting multiple regions. Up to 50 regulatory regions may be targeted simultaneously in a single pool (Figure 3C).
3. Perform a short PCR with the U6 primers to attach homology regions to the oligos prior to Gibson assembly. About 40 bases will be added to each oligo, yielding a ~100 bp product that contains sufficient homology to the U6 vector on both ends.
   1. For each guide RNA pool, set up a 20 μL PCR with a high-fidelity polymerase master mix and the following components: 1 μL of diluted oligo pool from step 3.2, 1 μL of U6 forward primer (10 μM), 1 μL of U6 reverse primer (10 μM), and water up to 20 μL.
   2. Incubate in a thermal cycler with the following conditions: 98 °C for 30 s, 10 cycles of (98 °C for 10 s, 55 °C for 30 s, 72 °C for 2 min), 72 °C for 5 min, and hold at 4 °C.
   3. Run 5 μL of the reaction on a 1 – 2% agarose gel with a low molecular weight ladder. The final product should be ~100 basepairs (Figure 2C).
   4. Clean up the extension reaction with a column-based DNA purification kit, and elute in 20 μL of elution buffer provided in the kit.
      Note: As the product is short, avoid using bead-based clean-ups, which are designed to exclude small fragments less than 100 bp.
   5. Quantify the purified DNA using a fluorometer or spectrophotometer (expected yield is 10-20 ng/μL). Guide RNA inserts can be stored at -20 °C, or can be used immediately in Gibson assembly with a linearized U6 vector.
4. To prepare the recipient U6 cloning vector for Gibson assembly, set up a restriction enzyme digest. If many Gibson assembly reactions are to be performed, set up multiple digests to ensure sufficient yield of cut vector.
   1. Use 20 units of AflII enzyme and 1 μg of plasmid in a 20 μL reaction with the appropriate restriction enzyme buffer. Incubate at 37 °C for 1-2 h.
   2. Clean up the digest with beads or a column-based kit for DNA purification and elute in 20 μL of elution buffer. Quantify the purified DNA using a fluorometer or spectrophotometer. Samples can be frozen at -20 °C for later use.
5. Perform Gibson assembly on the prepared vector and insert.
   1. Set up Gibson assembly reactions on ice. Use 50 ng of the vector and 7 ng of the insert in a 20 μL reaction. Dilute the inserts 1:10 in ultrapure water to facilitate pipetting. Set up a vector only Gibson assembly reaction, using 50 ng of the vector and replacing the insert with water.
   2. Incubate the Gibson assembly reactions for 15 min at 50 °C, followed by a hold at 4 °C.
   3. Transfer the assembled products to ice. Dilute the assembled products 1:4 in ultrapure water on ice. For example, add 5 μL of Gibson assembly product to 15 μL of ultrapure water.
6. Transform the diluted Gibson assembly products.
   1. Thaw high efficiency competent cells on ice and make 25 μL aliquots for each transformation. If a complex pool targeting multiple sites is desired, thaw enough cells into different tubes to perform multiple independent transformations of the same complex gRNA pool.
   2. For each diluted product, add 1 μL of this dilution to a 1.7 mL Eppendorf tube containing 25 μL of competent cells. Mix by briefly flicking the tube. Incubate the tubes on ice for 30 min.
   3. Heat shock the cells for 30 s at 42 °C, then transfer immediately to ice for 2 min.
   4. Add 300 μL SOC media (2% tryptone, 0.5% yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl₂, 10 mM MgSO₄, and 20 mM glucose) and let the cells recover for 1 h at 37 °C with shaking (300 rpm). During this time, warm the agar plates with ampicillin/carbenicillin to 37 °C in an incubator. Use one plate for each transformation.
   5. Plate 50 μL of cells and place the plates in a 37 °C incubator overnight. For the pools targeting individual sites, place directly in 3 – 5 mL of LB broth (Table of Materials) containing ampicillin/carbenicillin (1 mg/mL) and incubate overnight with shaking at 250 rpm at 37 °C for minipreps.
7. Harvest the cells and isolate DNA.
   1. For large guide RNA libraries targeting multiple sites, use a plate scraper to collect all colonies from each individual plate into one maxiprep (150 mL liquid culture). This can be facilitated by pouring ~5 mL of LB with the appropriate antibiotic into a 50 mL falcon tube and scraping the colonies into the tube. For libraries targeting individual sites, scrape the plates into a miniprep (3 – 5 mL liquid culture).
   2. Incubate these cultures with the appropriate antibiotic for 3 – 5 h at 37 °C with shaking at 250 rpm.
   3. Perform DNA extraction using a kit that results in endotoxin-free preps.
   4. Quantify DNA using a fluorometer or spectrophotometer. Plasmids can be used immediately in transfection or stored at -20 °C for future use.
8. To confirm the presence of guide RNA sequence within the U6 vector for small pools targeting single sites, use Sanger sequencing on the prepared miniprep with the “U6_PCR_R” primer listed in Table 1. Due to pooling of guide RNAs, the 19 basepair gRNA target sequence will yield mixed bases, but the U6 promoter and guide RNA scaffold surrounding this sequence should be intact.

4. Transfection of Enhancer-i

Note: For the successful blockade of an estrogen response using Enhancer-i in Ishikawa cells, it is necessary to deprive the cells of estrogen for 5 – 7 days prior to transfection by maintaining them in phenol red free RPMI with 10% charcoal-stripped FBS and 1% penicillin/streptomycin. Cells should be cultured in this media during and after transfection if trying to block an estrogen response. We recommend the use of phenol red free trypsin for passage of cells in complete phenol red free RPMI.

1. The day before transfection, plate the cells (either wild-type or stably expressing SID4X-dCas9-KRAB) in a 24-well plate at 30 – 50% confluency (~60,000 cells per well for Ishikawa cells). Plate enough cells such that transfections can be performed in duplicate, and include wells to be transfected with control guide RNAs.  Ensure that the cells are evenly distributed across the well by gently shaking the plate after cell plating as in step 1.1.7.
   Note: This protocol assumes the use of cationic liposome-based transfection reagents. Electroporation provides an alternative method for cell types that are very sensitive to these reagents. Transfection conditions must be optimized for the cell line of interest prior to attempting Enhancer-i experiments.
2. The following day, prepare transfections as per instructions of the transfection reagent of choice. For Ishikawa cells, use 550 ng of total plasmid for each well of a 24-well plate. Dilute the plasmids to a final concentration of 0.020 μg/μL in serum-free media (1.1 μg of DNA in 52 μL of total volume for transfecting 2 wells). Use 3 μL of transfection reagent for every 1 μg of DNA, vortex and incubate as described in step 1.2. Add 25 μL of the final mixture to each well.
   Note: To target combinations of sites, use the same weight of plasmid for each individual site, and then fill the leftover weight with a control plasmid (empty guide RNA cloning vector or guide RNAs targeting a negative control region such as the IL1RN promoter). For transient transfections, use a ratio of 3:2 Cas9 fusion:guide RNA plasmid. Plasmids containing fluorescent reporters may be added to monitor transfection efficiency.
3. At 36 h post transfection, change the media using phenol red free RPMI with 10% charcoal-stripped FBS and 1% penicillin/streptomycin (for Ishikawa cells) and supply puromycin (final concentration: 1 μg/mL) and neomycin (final concentration: 300 ng/mL). If cells are sensitive to transfection reagent, the media can be changed earlier, but antibiotics should be added no earlier than 24 h post transfection.
   Note: Wait at least 24 h after adding antibiotic before harvesting cells. Expression changes due to Enhancer-i can be detected as early as 48 h post transfection and up to 5 days post transfection. If working with Ishikawa cells that have been deprived of estrogen, perform an 8-h 10 nM 17β-estradiol (E2) induction the day after antibiotic treatment and then harvest cells immediately.

5. Cell Harvest and RNA Extraction

1. Prepare lysis buffer with 1% β-mercaptoethanol (BME). Ensure that there is enough lysis-BME mixture (300 μL for each well being harvested).
2. Aspirate the media using a vacuum aspirator.
3. Wash the cells once with an equal volume of 1x PBS (500 μL) and aspirate to remove as much PBS as possible.
4. Add 300 μL of lysis-BME solution to each well using a multichannel pipette. Pipette the lysis solution up and down 8 – 10 times, and transfer to a deep-well plate or 1.7 mL Eppendorf tubes on ice. RNA can be extracted immediately, or lysates can be frozen at -80 °C for future processing.
5. To extract RNA from lysates, use a commercially available kit that includes a DNase treatment. Elute in the smallest recommended volume of ultrapure water (RNase-free, DNase-free) or elution buffer and quantify RNA. For small numbers of samples, use a fluorometer or spectrophotometer. For large numbers of samples, use a fluorescent probe that detects RNA and measure on a plate reader. Samples can be frozen at -80 °C before or after quantification.

6. Quantifying Gene Expression Changes using One-step qPCR and RNA-seq

1. Obtain qPCR primers for the genes of interest and for at least one housekeeping gene that is expressed near the level of targeted genes and does not change across experimental conditions. Ideally, these primers will span an exon-exon junction to avoid amplification of genomic DNA.
   1. Test these primers on RNA obtained from the cell line of interest. Use melt curve analysis to verify the production of a single product. If a single product is not produced, test additional primer pairs.
2. For each Enhancer-i and control guide RNA treated sample, identify how many genes must be assayed in that sample. This set of genes should include housekeeping genes, such as CTCF or GAPDH.
3. Set up qPCR reactions.
   1. Dilute all samples to the same concentration in water, such that 50 ng of total RNA is easily pipetted, and there is enough diluted RNA for each reaction. For example, dilute RNA to ~16.6 ng/μL and use 3 μL of RNA in each reaction. Keep RNA on ice while setting up master mixes.
   2. Prepare separate master mixes for each gene to be measured using commercially available one-step qPCR kits. Use a 20 μL reaction volume with 1 μL of each primer (10 μM stock solution). Set up these reactions on ice.
   3. In a reaction plate that is appropriate for the thermal cycler, add RNA samples followed by master mixes. Seal with a plate sealer and mix gently by vortexing or pipetting. Briefly centrifuge the plate (140 x g for 60 s) to ensure that liquid is at the bottom of the wells.
   4. Incubate the plate in a thermal cycler as follows (or as kit instructs): 48 °C for 30 min, 95 °C for 10 min, 40 cycles of (95 °C for 15 s, 60 °C for 1 min).
4. Obtain Ct values for each gene measured within each sample. Use the comparative Ct method to identify changes in gene expression.
   1. Subtract the Ct of the housekeeping gene from the Ct of each gene of interest for each sample to generate normalized Ct values.
   2. For the control treated samples, take an average of the normalized Ct values for each gene. Log base 2-scale fold repression can then be calculated by subtracting the normalized Enhancer-i treated sample Ct for each gene from the same value for the control treated sample for the corresponding gene.
5. To determine global changes in gene expression following Enhancer-i treatment, prepare the samples for RNA sequencing using a commercially available kit compatible with the user’s sequencing technology. Use ~500 ng of RNA for starting material and prepare libraries for at least 2 biological replicates.

7. Verification of Specific Genomic Targeting by SID4X-dCas9-KRAB Using ChIP-seq

Note: The SID4X-dCas9-KRAB fusion protein contains both a FLAG epitope tag and an HA epitope tag, but best results for ChIP-seq were obtained with anti-FLAG antibodies. If desired, the user can perform additional ChIP-seq experiments for transcription factors potentially affected by Enhancer-i, or for H3K27ac, a mark of enhancer activity that is decreased by Enhancer-i. However, each ChIP-seq experiment requires 10 x 10⁶ cells, so plan accordingly.

1. Transfect cells with Enhancer-i pools.
   1. Plate 10 x 10⁶ cells in a 15 cm tissue culture dish. Each dish represents 1 ChIP-seq experiment for 1 factor of interest.
   2. The following day, transfect the cells using 20 μg of total DNA per dish. For transfections in cell lines stably expressing SID4X-dCas9-KRAB, the DNA should be a plasmid pool of guide RNAs targeting all sites of interest, and optionally a plasmid expressing fluorescent protein. For transient transfections, use a ratio of 3:2 dCas9 fusion protein:guide RNA pool. For ChIP-seq of other TFs or histone modifications, perform at least one additional control guide RNA transfection on another dish.  
   3. Treat the dishes with puromycin (1 μg/mL) and neomycin (300 ng/mL) at 24 – 48 h post transfection. Wait at least 24 h before harvesting chromatin.
2. Harvest chromatin from dishes.
   Note: To study the effects of Enhancer-i on ER genomic binding in Ishikawa cells, perform a 1 h 10 nM E2 treatment on dishes transfected with control guide RNAs and Enhancer-i prior to harvest. To study the effects of Enhancer-i on H3K27ac, perform an 8 h 10 nM E2 induction on dishes transfected with control guide RNAs and Enhancer-i prior to harvest.
   1. Apply 500 μL of 37% formaldehyde to each dish (final concentration of 1%). Swirl the plates briefly. Let the plates sit at room temperature for 10 min.
   2. Add 1 mL of 2.5 M glycine (final concentration of 125 mM). Swirl the plates briefly.
   3. Pour off the media with formaldehyde and glycine. Add an equal volume (~20 mL) of cold 1x PBS.
   4. Pour off the PBS. Aspirate with a vacuum aspirator to remove as much PBS as possible. Place the dishes on ice.
   5. Add 3 – 5 mL of cold 1x PBS or Farnham lysis buffer (5mM PIPES pH 8.0, 85 mM KCl, 0.5% NP-40) with 1x protease inhibitor (added just before use) to each plate. Scrape the dish with a plate scraper and transfer the solution to a 15 mL conical tube on ice.
   6. Pellet chromatin by spinning down tubes in a centrifuge for 5 min at 4 °C at 1000 x g. Discard the supernatant and store the pellets at -80 °C for future use, or proceed with the ChIP-seq protocol of choice using an anti-FLAG antibody or antibodies targeting other transcription factors or histone modifications of interest (H3K27ac, H3K9me3).