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14689
MLL1 (D2M7U) Rabbit mAb (Amino-terminal Antigen)
Primary Antibodies
Monoclonal Antibody

MLL1 (D2M7U) Rabbit mAb (Amino-terminal Antigen) #14689

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  1. WB
  2. IP
  3. C&R
Western Blotting Image 1: MLL1 (D2M7U) Rabbit mAb (Amino-terminal Antigen)
Western blot analysis of extracts from various cell lines using MLL1 (D2M7U) Rabbit mAb (Amino-terminal Antigen).
No image available
CUT and RUN Image 1: MLL1 (D2M7U) Rabbit mAb (Amino-terminal Antigen)
CUT&RUN was performed with NCCIT cells and MLL1 (D2M7U) Rabbit mAb (Amino-terminal Antigen), using CUT&RUN Assay Kit #86652. DNA Library was prepared using SimpleChIP® ChIP-seq DNA Library Prep Kit for Illumina® #56795. The figure shows binding across ELOC, a known target gene of MLL1 (see additional figure containing CUT&RUN-qPCR data).
CUT and RUN Image 2: MLL1 (D2M7U) Rabbit mAb (Amino-terminal Antigen)
CUT&RUN was performed with NCCIT cells and MLL1 (D2M7U) Rabbit mAb (Amino-terminal Antigen), using CUT&RUN Assay Kit #86652. DNA Libraries were prepared using SimpleChIP® ChIP-seq DNA Library Prep Kit for Illumina® #56795. The figures show binding across chromosome 8 (upper), including ELOC (lower), a known target gene of MLL1 (see additional figure containing CUT&RUN-qPCR data).
CUT and RUN Image 3: MLL1 (D2M7U) Rabbit mAb (Amino-terminal Antigen)
CUT&RUN was performed with NCCIT cells and either MLL1 (D2M7U) Rabbit mAb (Amino-terminal Antigen) or Rabbit (DA1E) mAb IgG XP® Isotype Control (CUT&RUN) #66362, using CUT&RUN Assay Kit #86652. The enriched DNA was quantified by real-time PCR using human RTN4 promoter, human ELOC promoter, and SimpleChIP® Human α Satellite Repeat Primers #4486. The amount of immunoprecipitated DNA in each sample is represented as signal relative to the total amount of input chromatin, which is equivalent to one.
To Purchase # 14689
Cat. # Size Qty. Price
14689S
100 µl $ 276

Supporting Data

REACTIVITY H M R Mk
SENSITIVITY Endogenous
MW (kDa) 300
Source/Isotype Rabbit IgG

Application Key:

  • WB-Western Blot
  • IP-Immunoprecipitation
  • IHC-Immunohistochemistry
  • ChIP-Chromatin Immunoprecipitation
  • IF-Immunofluorescence
  • F-Flow Cytometry
  • E-P-ELISA-Peptide

Species Cross-Reactivity Key:

  • H-Human
  • M-Mouse
  • R-Rat
  • Hm-Hamster
  • Mk-Monkey
  • Vir-Virus
  • Mi-Mink
  • C-Chicken
  • Dm-D. melanogaster
  • X-Xenopus
  • Z-Zebrafish
  • B-Bovine
  • Dg-Dog
  • Pg-Pig
  • Sc-S. cerevisiae
  • Ce-C. elegans
  • Hr-Horse
  • All-All Species Expected

Product Usage Information

The CUT&RUN dilution was determined using the CUT&RUN Assay Kit #86652.
Application Dilution
Western Blotting 1:1000
Immunoprecipitation 1:50
CUT&RUN 1:50

Storage

Supplied in 10 mM sodium HEPES (pH 7.5), 150 mM NaCl, 100 µg/ml BSA, 50% glycerol and less than 0.02% sodium azide. Store at –20°C. Do not aliquot the antibody.

Protocol

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Western Blotting Protocol

For western blots, incubate membrane with diluted primary antibody in 5% w/v BSA, 1X TBS, 0.1% Tween® 20 at 4°C with gentle shaking, overnight.

NOTE: Please refer to primary antibody product webpage for recommended antibody dilution.

A. Solutions and Reagents

From sample preparation to detection, the reagents you need for your Western Blot are now in one convenient kit: #12957 Western Blotting Application Solutions Kit

NOTE: Prepare solutions with reverse osmosis deionized (RODI) or equivalent grade water.

  1. 20X Phosphate Buffered Saline (PBS): (#9808) To prepare 1 L 1X PBS: add 50 ml 20X PBS to 950 ml dH2O, mix.
  2. 10X Tris Buffered Saline (TBS): (#12498) To prepare 1 L 1X TBS: add 100 ml 10X to 900 ml dH2O, mix.
  3. 1X SDS Sample Buffer: Blue Loading Pack (#7722) or Red Loading Pack (#7723) Prepare fresh 3X reducing loading buffer by adding 1/10 volume 30X DTT to 1 volume of 3X SDS loading buffer. Dilute to 1X with dH2O.
  4. 10X Tris-Glycine SDS Running Buffer: (#4050) To prepare 1 L 1X running buffer: add 100 ml 10X running buffer to 900 ml dH2O, mix.
  5. 10X Tris-Glycine Transfer Buffer: (#12539) To prepare 1 L 1X Transfer Buffer: add 100 ml 10X Transfer Buffer to 200 ml methanol + 700 ml dH2O, mix.
  6. 10X Tris Buffered Saline with Tween® 20 (TBST): (#9997) To prepare 1 L 1X TBST: add 100 ml 10X TBST to 900 ml dH2O, mix.
  7. Nonfat Dry Milk: (#9999).
  8. Blocking Buffer: 1X TBST with 5% w/v nonfat dry milk; for 150 ml, add 7.5 g nonfat dry milk to 150 ml 1X TBST and mix well.
  9. Wash Buffer: (#9997) 1X TBST.
  10. Bovine Serum Albumin (BSA): (#9998).
  11. Primary Antibody Dilution Buffer: 1X TBST with 5% BSA; for 20 ml, add 1.0 g BSA to 20 ml 1X TBST and mix well.
  12. Biotinylated Protein Ladder Detection Pack: (#7727).
  13. Blue Prestained Protein Marker, Broad Range (11-250 kDa): (#59329).
  14. Blotting Membrane and Paper: (#12369) This protocol has been optimized for nitrocellulose membranes. Pore size 0.2 µm is generally recommended.
  15. Secondary Antibody Conjugated to HRP: Anti-rabbit IgG, HRP-linked Antibody (#7074).
  16. Detection Reagent: SignalFire™ ECL Reagent (#6883).

B. Protein Blotting

A general protocol for sample preparation.

  1. Treat cells by adding fresh media containing regulator for desired time.
  2. Aspirate media from cultures; wash cells with 1X PBS; aspirate.
  3. Lyse cells by adding 1X SDS sample buffer (100 µl per well of 6-well plate or 500 µl for a 10 cm diameter plate). Immediately scrape the cells off the plate and transfer the extract to a microcentrifuge tube. Keep on ice.
  4. Sonicate for 10–15 sec to complete cell lysis and shear DNA (to reduce sample viscosity).
  5. Heat a 20 µl sample to 95–100°C for 5 min; cool on ice.
  6. Microcentrifuge for 5 min.
  7. Load 20 µl onto SDS-PAGE gel (10 cm x 10 cm).

    NOTE: Loading of prestained molecular weight markers (#59329, 10 µl/lane) to verify electrotransfer and biotinylated protein ladder (#7727, 10 µl/lane) to determine molecular weights are recommended.

  8. Electrotransfer to nitrocellulose membrane (#12369).

C. Membrane Blocking and Antibody Incubations

NOTE: Volumes are for 10 cm x 10 cm (100 cm2) of membrane; for different sized membranes, adjust volumes accordingly.

I. Membrane Blocking

  1. (Optional) After transfer, wash nitrocellulose membrane with 25 ml TBS for 5 min at room temperature.
  2. Incubate membrane in 25 ml of blocking buffer for 1 hr at room temperature.
  3. Wash three times for 5 min each with 15 ml of TBST.

II. Primary Antibody Incubation

  1. Incubate membrane and primary antibody (at the appropriate dilution and diluent as recommended in the product webpage) in 10 ml primary antibody dilution buffer with gentle agitation overnight at 4°C.
  2. Wash three times for 5 min each with 15 ml of TBST.
  3. Incubate membrane with Anti-rabbit IgG, HRP-linked Antibody (#7074 at 1:2000) and anti-biotin, HRP-linked Antibody (#7075 at 1:1000–1:3000) to detect biotinylated protein markers in 10 ml of blocking buffer with gentle agitation for 1 hr at room temperature.
  4. Wash three times for 5 min each with 15 ml of TBST.
  5. Proceed with detection (Section D).

D. Detection of Proteins

Directions for Use:

  1. Wash membrane-bound HRP (antibody conjugate) three times for 5 minutes in TBST.
  2. Prepare 1X SignalFire™ ECL Reagent (#6883) by diluting one part 2X Reagent A and one part 2X Reagent B (e.g. for 10 ml, add 5 ml Reagent A and 5 ml Reagent B). Mix well.
  3. Incubate substrate with membrane for 1 minute, remove excess solution (membrane remains wet), wrap in plastic and expose to X-ray film.

* Avoid repeated exposure to skin.

posted June 2005

revised June 2020

Protocol Id: 10

Immunoprecipitation for Native Proteins

This protocol is intended for immunoprecipitation of native proteins for analysis by western immunoblot or kinase activity utilizing Protein A magnetic separation.

A. Solutions and Reagents

NOTE: Prepare solutions with reverse osmosis deionized (RODI) or equivalent grade water.

  1. 20X Phosphate Buffered Saline (PBS): (#9808) To prepare 1 L of 1X PBS, add 50 ml 20X PBS to 950 ml dH2O, mix.
  2. 10X Cell Lysis Buffer: (#9803) To prepare 10 ml of 1X cell lysis buffer, add 1 ml cell lysis buffer to 9 ml dH2O, mix.

    NOTE: Add 1 mM PMSF (#8553) immediately prior to use.

  3. 3X SDS Sample Buffer: Blue Loading Pack (#7722) or Red Loading Pack (#7723) Prepare fresh 3X reducing loading buffer by adding 1/10 volume 30X DTT to 1 volume of 3X SDS loading buffer.
  4. Protein A Magnetic Beads: (#73778).
  5. Magnetic Separation Rack: (#7017) or (#14654).
  6. 10X Kinase Buffer (for kinase assays): (#9802) To Prepare 1 ml of 1X kinase buffer, add 100 µl 10X kinase buffer to 900 µl dH2O, mix.
  7. ATP (10 mM) (for kinase assays): (#9804) To prepare 0.5 ml of ATP (200 µM), add 10 µl ATP (10 mM) to 490 µl 1X kinase buffer.

B. Preparing Cell Lysates

  1. Aspirate media. Treat cells by adding fresh media containing regulator for desired time.
  2. To harvest cells under nondenaturing conditions, remove media and rinse cells once with ice-cold 1X PBS.
  3. Remove PBS and add 0.5 ml ice-cold 1X cell lysis buffer to each plate (10 cm) and incubate on ice for 5 min.
  4. Scrape cells off the plate and transfer to microcentrifuge tubes. Keep on ice.
  5. Sonicate on ice three times for 5 sec each.
  6. Microcentrifuge for 10 min at 4°C, 14,000 x g and transfer the supernatant to a new tube. The supernatant is the cell lysate. If necessary, lysate can be stored at -80°C.

C. Immunoprecipitation

Cell Lysate Pre-Clearing (Optional)

A cell lysate pre-clearing step is highly recommended to reduce non-specific protein binding to the Protein A Magnetic beads. Pre-clear enough lysate for test samples and isotype controls.

  1. Briefly vortex the stock tube to resuspend the magnetic beads.

    IMPORTANT: Pre-wash #73778 magnetic beads just prior to use:

  2. Transfer 20 μl of bead slurry to a clean tube. Place the tube in a magnetic separation rack for 10-15 seconds.

    Carefully remove the buffer once the solution is clear. Add 500 μl of 1X cell lysis buffer to the magnetic bead pellet, briefly vortex to wash the beads. Place tube back in magnetic separation rack. Remove buffer once solution is clear. Repeat washing step once more.

  3. Add 200 μl cell lysate to 20 μl of pre-washed magnetic beads.

    IMPORTANT: The optimal lysate concentration will depend on the expression level of the protein of interest. A starting concentration between 250 μg/ml-1.0 mg/ml is recommended.

  4. Incubate with rotation for 20 minutes at room temperature.
  5. Separate the beads from the lysate using a magnetic separation rack, transfer the pre-cleared lysate to a clean tube, and discard the magnetic bead pellet.
  6. Proceed to immunoprecipitation section.

Immunoprecipitation

IMPORTANT: Appropriate isotype controls are highly recommended in order to show specific binding in your primary antibody immunoprecipitation. Use Normal Rabbit IgG #2729 for rabbit polyclonal primary antibodies, Rabbit (DA1E) mAb IgG XP® Isotype Control #3900 for rabbit monoclonal primary antibodies, Mouse (G3A1) mAb IgG1 Isotype Control #5415 for mouse monoclonal IgG1 primary antibodies, Mouse (E5Y6Q) mAb IgG2a Isotype Control #61656 for mouse monoclonal IgG2a primary antibodies, Mouse (E7Q5L) mAb IgG2b Isotype Control #53484 for mouse monoclonal IgG2b primary antibodies, and Mouse (E1D5H) mAb IgG3 Isotype Control #37988 for mouse monoclonal IgG3 primary antibodies. Isotype controls should be concentration matched and run alongside the primary antibody samples.

  1. Add primary antibody (at the appropriate dilution as recommended in the product datasheet) to 200 µl cell lysate. Incubate with rotation overnight at 4°C. to form the immunocomplex.
  2. Pre-wash magnetic beads (see Cell Lysate Pre-Clearing section, steps 1 and 2).
  3. Transfer the lysate and antibody (immunocomplex) solution to the tube containing the pre-washed magnetic bead pellet.
  4. Incubate with rotation for 20 min at room temperature.
  5. Pellet beads using magnetic separation rack. Wash pellets five times with 500 μl of 1X cell lysis buffer. Keep on ice between washes.
  6. Proceed to analyze by western immunoblotting or kinase activity (section D).

D. Sample Analysis

Proceed to one of the following specific set of steps.

For Analysis by Western Immunoblotting

  1. Resuspend the pellet with 20-40 µl 3X SDS sample buffer, briefly vortex to mix, and briefly microcentrifuge to pellet the sample.
  2. Heat the sample to 95-100°C for 5 min.
  3. Pellet beads using magnetic separation rack. Transfer the supernatant to a new tube. The supernatant is the sample.
  4. Analyze sample by western blot (see Western Immunoblotting Protocol).

NOTE: To minimize masking caused by denatured IgG heavy chains (~50 kDa), we recommend using Mouse Anti-Rabbit IgG (Light-Chain Specific) (D4W3E) mAb (#45262) or Mouse Anti-Rabbit IgG (Conformation Specific) (L27A9) mAb (#3678) (or HRP conjugate #5127). To minimize masking caused by denatured IgG light chains (~25 kDa), we recommend using Mouse Anti-Rabbit IgG (Conformation Specific) (L27A9) mAb (#3678) (or HRP conjugate #5127).

For Analysis by Kinase Assay

  1. Wash pellet twice with 500 µl 1X kinase buffer. Keep on ice.
  2. Suspend pellet in 40 µl 1X kinase buffer supplemented with 200 µM ATP and appropriate substrate.
  3. Incubate for 30 min at 30°C.
  4. Terminate reaction with 20 µl 3X SDS sample buffer. Vortex, then microcentrifuge for 30 sec.
  5. Transfer supernatant containing phosphorylated substrate to another tube.
  6. Heat the sample to 95-100°C for 2-5 min and microcentrifuge for 1 min at 14,000 x g.
  7. Load the sample (15-30 µl) on SDS-PAGE gel.

posted December 2008

revised April 2021

Protocol Id: 410

CUT&RUN Protocol

! This ! signifies an important step in the protocol regarding volume changes based on the number of CUT&RUN reactions being performed.
!! This !! signifies an important step to dilute a buffer before proceeding.
SAFE STOP This is a safe stopping point in the protocol, if stopping is necessary.

I. Cell and Tissue Preparation

For most cell types, live cells can be used in the CUT&RUN assay to generate robust enrichment of histones, transcription factors, and cofactors. For certain cell types that are fragile or sensitive to Conconavalin A, a light cell fixation helps to preserve the cells and keep them intact. In addition, fixation may help to boost enrichment of low abundance and/or weak binding transcription factors and cofactors if robust signal is not observed using fresh cells. Please note that over-fixation of cells will inhibit the CUT&RUN assay.

Our CUT&RUN assay works with a wide range of cell or tissue inputs. As defined in the protocol, one CUT&RUN reaction can contain between 5,000 to 250,000 cells or 1 to 5 mg of tissue. Buffer volumes used throughout the protocol do not need to be adjusted based on the amount of cells or tissue per reaction, as long as it is within this range. When indicated, buffer volumes do need to be increased proportionally based on the number of reactions being performed. If possible, we recommend using 100,000 cells or 1 mg of tissue per reaction. If cells are limiting, we recommend using at least 5,000 to 10,000 cells per reaction for histone modifications and 10,000 to 20,000 cells per reaction for transcription factors or cofactors.

NOTE: The amount of digitonin recommended for cell permeabilization is in excess and should be sufficient for permeabilization of most cell lines and tissue types. However, not all cell lines and tissues exhibit the same sensitivity to digitonin. If your specific cell line or tissue does not work with the recommended digitonin concentration, you can optimize conditions by following the protocol provided in Appendix A. Digitonin treatment should result in permeabilization of >90% of the cell population.

A. Live Cell Preparation

Before Starting:

! All buffer volumes should be increased proportionally based on the number of CUT&RUN reactions being performed.

  • Remove and warm 200X Protease Inhibitor Cocktail #7012 and 100X Spermidine #27287. Make sure both are completely thawed.
  • Prepare 1X Wash Buffer (2 ml for each cell line and additional 100 µl for each reaction or input sample). For example, to prepare 2.5 ml of 1X Wash Buffer, add 250 µl 10X Wash Buffer #31415 + 25 µl 100X Spermidine #27287 + 12.5 µl 200X Protease Inhibitor Cocktail #7012 + 2212.5 µl Nuclease-free Water #12931. Equilibrate it to room temperature to minimize stress on the cells.

    NOTE: Steps for live cell (no fixation) preparation should be performed in succession at room temperature to minimize stress on the cells. To minimize DNA fragmentation, avoid vigorous vortexing and cavitation of samples during resuspension. When preparing live cells for CUT&RUN, we recommend preparing the Concanavalin A Beads (Section II, Steps 1 to 5) prior to preparing the cells as to minimize the amount of time the cells sit around during bead preparation. Activated beads can be stored on ice until ready to use.

  1. Harvest fresh cell cultures at room temperature to minimize stress on the cells. Collect 5,000 to 100,000 cells for each reaction and an additional 5,000 to 100,000 cells for the input sample. Be sure to include reactions for the positive control Tri-Methyl-Histone H3 (Lys4) (C42D8) Rabbit mAb #9751 and the negative control Rabbit (DA1E) mAb IgG XP® Isotype Control (CUT&RUN) #66362.

    NOTE: For adherent cells, the cells first need to be detached from the dish using Trypsin and neutralized with at least 3 volumes of tissue culture medium. We do not recommend scraping the cells from the dish because this can stress and even lyse the cells. Cells should be counted using a hemocytometer or other cell counter to ensure the proper number of cells are being used for the experiment.

  2. Centrifuge cell suspension for 3 min at 600 x g at room temperature and remove the liquid.

    NOTE: The challenge of working with low cell numbers (<100,000 total cells) is that the centrifuged cell pellet is not always visible by eye, making it easy to lose cells during the wash steps. Therefore, when working with low cell numbers, we recommend skipping the wash steps 3 to 5 below. The binding of the Concanavalin A beads to cells is tolerant to having 40% cell medium in the binding reaction. Therefore, after the initial centrifugation of the cell suspension in Step 2, one can remove most of the supernatant, leaving behind ≤40 µl cell medium per reaction. Then in Step 6 add enough 1X Wash Buffer (+ spermidine + PIC) to the cell suspension to achieve a total volume of 100 µl per reaction.

  3. Resuspend cell pellet in 1 ml of 1X Wash Buffer (+ spermidine + PIC) at room temperature by gently pipetting up and down.
  4. Centrifuge for 3 min at 600 x g at room temperature and remove the liquid.
  5. Wash the cell pellet a second time by repeating steps 3 and 4 one time.
  6. For each reaction or input sample, add 100 µl of 1X Wash Buffer (+ spermidine + PIC) and resuspend the cell pellet by gently pipetting up and down.
  7. Transfer 100 µl of cells to a new tube and store at 4°C until Section V. This is the input sample.

    NOTE: The input sample will be incubated at 55°C later in the protocol, so it is recommended to use a safe-lock 1.5 ml tube to reduce evaporation during the incubation.

  8. Immediately proceed to Section II.

B. Fixed Cell Preparation

NOTE: The following reagents are required for fixed cell preparation and are not included in this kit: 37% formaldehyde or 16% Formaldehyde Methanol-Free #12606, Glycine Solution (10X) #7005, and 10% SDS Solution #20533.

Before Starting:

! All buffer volumes should be increased proportionally based on the number of CUT&RUN reactions being performed.

  • Remove and warm 200X Protease Inhibitor Cocktail #7012 and 100X Spermidine #27287. Make sure both are completely thawed.
  • Prepare 1X Wash Buffer (2 ml for each cell line and additional 100 µl for each reaction or input sample). For example, to prepare 2.5 ml of 1X Wash Buffer, add 250 µl 10X Wash Buffer #31415 + 25 µl 100X Spermidine #27287 + 12.5 µl 200X Protease Inhibitor Cocktail #7012 + 2212.5 µl Nuclease-free Water #12931. Equilibrate it to room temperature to minimize stress on the cells.
  • Prepare 2.7 µl of 37% formaldehyde or 6.25 µl of 16% Formaldehyde Methanol-Free #12606 per 1 ml of cell suspension to be processed and keep at room temperature. Use fresh formaldehyde that is not past the manufacturer’s expiration date.
  1. Collect 5,000 to 100,000 cells for each antibody/MNase reaction and an additional 5,000 to 100,000 cells for the input sample. Be sure to include reactions for the positive control Tri-Methyl-Histone H3 (Lys4) (C42D8) Rabbit mAb #9751 and the negative control Rabbit (DA1E) mAb IgG XP® Isotype Control (CUT&RUN) #66362.

    NOTE: With adherent cell lines, cells first need to be detached from the dish using Trypsin and neutralized with at least 3 volumes of medium. We don’t recommend scraping the cells from the dish because this can stress and even lyse the cells. Cells should be counted using a hemocytometer or other cell counter to ensure the proper number of cells are being used for the experiment.

  2. Add 2.7 µl 37% formaldehyde or 6.25 µl 16% Formaldehyde Methanol-Free #12606 per 1 ml of cell suspension to achieve a final concentration of 0.1% formaldehyde. Swirl tube to mix and incubate at room temperature for 2 min.
  3. Stop cross-linking by adding 100 µl of Glycine Solution (10X) #7005 per 1 ml of fixed cell suspension. Swirl the tube to mix and incubate at room temperature for 5 min.
  4. Centrifuge cell suspension for 3 min at 3,000 x g at 4°C and remove the liquid. Immediately proceed to Step 5. (SAFE STOP) Alternatively, fixed cell pellets may be stored at -80°C before using for up to 6 months.

    NOTE: The challenge of working with low cell numbers (<100,000 total cells) is that the centrifuged cell pellet is not always visible by eye, making it easy to lose cells during the wash steps. In this case we do NOT recommend freezing down cell pellets. In addition, when working with these low cell numbers, we recommend skipping the wash steps 5 to 7 below. The binding of the Concanavalin A beads to cells is tolerant to having 40% cell medium in the binding reaction. Therefore, after the initial centrifugation of the cell suspension in Step 4, one can remove most of the supernatant, leaving behind ≤40 µl cell medium per reaction. Then in Step 8 add enough 1X Wash Buffer (+ spermidine + PIC) to the cell suspension to achieve a total volume of 100 µl per reaction.

  5. Resuspend cell pellet in 1 ml of 1X Wash Buffer (+ spermidine + PIC) by gently pipetting up and down.
  6. Centrifuge for 3 min at 3,000 x g at 4°C and remove the liquid.
  7. Wash the cell pellet a second time by repeating steps 5 and 6 one time.
  8. For each reaction or input sample, add 100 µl of 1X Wash Buffer (+ spermidine + PIC) and resuspend the cell pellet by gently pipetting up and down.
  9. Transfer 100 µl of cells to a new tube and store at 4°C until Section V. This is the input sample.

    NOTE: The input sample will be incubated at 55°C later in the protocol, so it is recommended to use a safe-lock 1.5 ml tube to reduce evaporation during the incubation.

  10. Immediately proceed to Section II.

C. Tissue Sample Preparation

For most tissue types, 1 mg of lightly fixed tissue (0.1% formaldehyde for 2 min) can generate robust enrichment of histones, transcription factors and cofactors. Formaldehyde fixation is not required for enrichment of histone modifications. However, many transcription factors and cofactors do require light fixation of the tissue for optimal results. Some low abundance and/or weak binding transcription factors and cofactors may require a medium fixation (0.1% formaldehyde for 10 min) for optimal results. In addition, medium fixation may improve results when using difficult tissue types, like fibrous tissues. Please note that over-fixation will inhibit the CUT&RUN assay. Fixed tissue samples can be frozen at -80°C for up to 6 months before using.

NOTE: When preparing fresh tissue (no fixation) for CUT&RUN, we recommend preparing the Concanavalin A Beads (Section II, Steps 1 to 5) prior to preparing the tissue as to minimize the amount of time the cells sit around during bead preparation. Activated beads can be stored on ice until ready to use.

NOTE: The following reagents are required for fixed tissue preparation and are not included in this kit: 37% formaldehyde or 16% Formaldehyde Methanol-Free #12606, Phosphate Buffered Saline (PBS) #9872, Glycine Solution (10X) #7005, and 10% SDS Solution #20533.

Before Starting:

! All buffer volumes should be increased proportionally based on the number of CUT&RUN reactions being performed.

  • Remove and warm 200X Protease Inhibitor Cocktail #7012 and 100X Spermidine #27287. Make sure both are completely thawed.
  • Prepare 1X Wash Buffer (3 ml for each tissue type and additional 100 µl for each reaction or input sample). For example, to prepare 3.5 ml of 1X Wash Buffer, add 350 µl 10X Wash Buffer #31415 + 35 µl 100X Spermidine #27287 + 17.5 µl 200X Protease Inhibitor Cocktail #7012 + 3097.5 µl Nuclease-free Water #12931. Equilibrate it to room temperature to minimize stress on the cells.
  • Prepare the following buffers if tissue fixation is needed:
    • Prepare 1 ml fixation buffer for each tissue type by adding 2.7 µl of 37% formaldehyde or 6.25 µl of 16% Formaldehyde Methanol-Free #12606 and 5 µl 200X Protease Inhibitor Cocktail (PIC) #7012 into 1 ml of Phosphate Buffered Saline (PBS) #9872. Use fresh formaldehyde that is not past the manufacturer’s expiration date.
    • Prepare 1 ml of PBS #9872 + 5 µl Protease Inhibitor Cocktail (PIC) #7012 for each tissue type and place on ice.
    • Prepare 100 µl of Glycine Solution (10X) #7005 per 1 ml of fixation buffer.
  1. Weigh 1 mg fresh tissues for each antibody/MNase reaction and an additional 1 mg of tissue for the input sample. Be sure to include reactions for the positive control Tri-Methyl-Histone H3 (Lys4) (C42D8) Rabbit mAb #9751 and the negative control Rabbit (DA1E) mAb IgG XP® Isotype Control (CUT&RUN) #66362.

    NOTE: For some transcription factors or cofactors, or for difficult tissue types like fibrous tissues, up to 5 mg tissue per reaction can be used without scaling up reagents.

  2. Place tissue sample in a dish and finely mince using a clean scalpel or razor blade. Keep dish on ice. It is important to keep the tissue cold to avoid protein degradation.

    NOTE: We recommend light fixation of tissues because this condition works optimally for most tissue types and protein targets. However, if fresh tissues are desired, skip Steps 3 to 8 and immediately proceed to Step 9.

  3. Immediately transfer minced tissue to 1 ml of fixation solution and swirl tube to mix.

    NOTE: This volume of fixation solution is sufficient for up to 50 mg of tissue. If processing >50 mg, scale up fixation solution and 1X PBS + PIC solution in Step 7 accordingly.

  4. Incubate at room temperature for 2 min.

    NOTE: For difficult tissue types (like fibrous tissues) or low abundance and/or weak binding transcription factors or cofactors, extending the formaldehyde fixation to 10 min may improve results.

  5. Stop cross-linking by adding 100 µl of Glycine Solution (10X) #7005 per 1 ml of fixation buffer. Swirl the tube to mix and incubate at room temperature for 5 min.
  6. Centrifuge tissue for 5 min at 2,000 x g at 4°C and remove the liquid.
  7. Resuspend tissue with 1 ml of 1X PBS + PIC.
  8. Centrifuge for 5 min at 2,000 x g at 4°C and remove the liquid and proceed to step 9. (SAFE STOP) Alternatively, fixed tissue pellets may be stored at -80°C before disaggregation for up to 6 months.
  9. Resuspend tissue in 1 ml of 1X Wash Buffer (+ spermidine + PIC) and transfer the sample to a Dounce homogenizer.
  10. Disaggregate tissue pieces into single-cell suspension with 20-25 strokes until no tissue chunks are observed.
  11. Transfer cell suspension to a 1.5 ml tube and centrifuge at 3,000 x g for 3 min at room temperature, remove supernatant from cells.
  12. Resuspend cell pellet in 1 ml of 1X Wash Buffer (+ spermidine + PIC).
  13. Centrifuge cell suspension for 3 min at 3,000 x g at room temperature and remove the liquid.
  14. Wash the cell pellet a second time by repeating steps 12 and 13 one time.
  15. For each reaction, add 100 µl of 1X Wash Buffer (+ spermidine + PIC) and resuspend the cell pellet by gently pipetting up and down.
  16. Transfer 100 µl of cells to a new tube and store at 4°C until Section V. This is the input sample.

    NOTE: The input sample will be incubated at 55°C later in the protocol, so it is recommended to use a safe-lock 1.5 ml tube to reduce evaporation during the incubation.

  17. Immediately proceed to Section II.

II. Binding of Concanavalin A Beads and Primary Antibody

Before Starting:

! All buffer volumes should be increased proportionally based on the number of CUT&RUN reactions being performed.

  • Place Concanavalin A Bead Activation Buffer on ice.
  • For each reaction, prepare 1 µl 100X Spermidine #27287 + 0.5 µl 200X Protease Inhibitor Cocktail #7012 + 2.5 µl Digitonin Solution #16359 + 96 µl Antibody Binding Buffer #15338 and place on ice (100 µl per reaction).
  1. Carefully resuspend Concanavalin A Magnetic Beads by gently pipetting up and down, making sure not to splash any bead suspension out of the tube. Transfer 10 µl of the bead suspension per each CUT&RUN reaction to a new 1.5 ml microcentrifuge tube.

    NOTE: Avoid vortexing the Concanavalin A Magnetic Bead suspension as repeated vortexing may displace the Concanavalin A from the beads.

  2. Add 100 µl Concanavalin A Bead Activation Buffer per 10 µl beads. Gently mix beads by pipetting up and down.
  3. Place tube on a magnetic rack until solution becomes clear (30 sec to 2 min) and then remove the liquid.

    NOTE: To avoid loss of beads, remove liquid using a pipet. Do not aspirate using a vacuum.

  4. Remove tubes from the magnetic rack. Wash the beads a second time by repeating steps 2 and 3 one time.
  5. Add a volume of Concanavalin A Bead Activation Buffer equal to the initial volume of bead suspension added (10 µl per sample) and resuspend by pipetting up and down.

    NOTE: If Concanavalin A Beads are prepared prior to cell or tissue preparation, as recommended for live cells and fresh tissue, the activated beads can be stored on ice until use.

  6. Make sure Concanavalin A Beads are mixed well into solution. Add 10 µl of activated bead suspension per reaction to the washed cell suspension prepared in Section I-A Step 8, Section I-B Step 10, or Section I-C Step 17.
  7. Rotate the sample for 5 min at room temperature.

    NOTE: Concanavalin A Magnetic Beads may clump or stick to the sides of the tube. Rocking instead of rotating the tubes may help to mitigate this issue. Beads can be resuspended by pipetting up and down.

  8. Briefly centrifuge the sample at 100 x g for 2 sec to remove cell:bead suspension from the cap of the tube. Place the tube on the magnetic rack until the solution turns clear (30 sec to 2 min), then remove and discard the liquid.
  9. Remove tube from the stand. Add 100 µl of Antibody Binding Buffer (+ spermidine + PIC + digitonin) per reaction and place on ice.
  10. Aliquot 100 µl of the cell:bead suspension into separate 1.5 ml tubes for each reaction and place on ice.
  11. Add the appropriate amount of antibody to each reaction and mix gently by pipetting up and down.

    NOTE: The amount of antibody required for CUT&RUN varies and should be determined by the user. For the positive control Tri-Methyl-Histone H3 (Lys4) (C42D8) Rabbit mAb #9751, add 2 µl of antibody to the sample. For the negative control Rabbit (DA1E) mAb IgG XP® Isotype Control (CUT&RUN) #66362, add 5 µl to the sample. We strongly recommend using the negative control antibody and NOT a no-antibody control, because the latter results in high levels of non-specific MNase digestion and high background signal. We recommend using the input sample for comparison with both qPCR and NG-seq analysis, when possible.

  12. Rotate tubes at 4°C for 2 hr. This step can be extended to overnight.

III. Binding of pAG-MNase Enzyme

Before Starting:

! All buffer volumes should be increased proportionally based on the number of CUT&RUN reactions being performed.

  • Remove and warm Digitonin Solution #16359 to 90-100°C for 5 min and make sure it is completely thawed and in solution. Immediately place the thawed Digitonin Solution #16359 on ice.

    NOTE: Digitonin Solution #16359 should be stored at -20°C. Please keep on ice during use and store at -20°C when finished for the day.

  • For each reaction, prepare 1.05 ml of Digitonin Buffer (105 µl 10X Wash Buffer #31415 + 10.5 µl 100X Spermidine #27287 + 5.25 µl 200X Protease Inhibitor Cocktail #7012 + 26.25 µl Digitonin Solution #16359 + 903 µl Nuclease-free Water #12931).
  • For each reaction, make a pAG-MNase pre-mix by adding 50 µl of Digitonin Buffer (described above) and 1.5 µl of pAG-MNase Enzyme to a new tube. For example, for 10 reactions, transfer 500 µl of Digitonin Buffer to a new tube and add 15 µl of pAG-MNase Enzyme. Mix by gently pipetting up and down and place on ice.
  1. Briefly centrifuge samples from Section II, Step 12 at 100 x g for 2 sec to remove cell:bead suspension from the caps of the tubes.
  2. Place the tubes on the magnetic rack until the solution turns clear (30 sec to 2 min) and then remove the liquid.
  3. Remove tubes from the magnetic rack and add 1 ml of Digitonin Buffer (+ spermidine + PIC + digitonin). Resuspend beads by gently pipetting up and down, make sure to collect all beads that are stuck to the tube wall.
  4. Place the tubes on the magnetic rack until the solution turns clear (30 sec to 2 min) and then remove the liquid.
  5. Remove tubes from magnetic rack. Add 50 µl of pAG-MNase pre-mix to each tube and gently mix the sample by pipetting up and down.
  6. Rotate tubes at 4°C for 1 hr.

    NOTE: Concanavalin A Magnetic Beads may clump or stick to the sides of the tube. Rocking instead of rotating the tubes may help to mitigate this issue. Beads can be resuspended by pipetting up and down.

  7. Immediately proceed to Section IV.

IV. DNA Digestion and Diffusion

Before Starting:

! All buffer volumes should be increased proportionally based on the number of CUT&RUN reactions being performed.

  • Remove and warm Digitonin Solution #16359 to 90-100°C for 5 min and make sure it is completely thawed and in solution. Immediately place the thawed Digitonin Solution #16359 on ice.

    NOTE: Digitonin Solution #16359 should be stored at -20°C. Please keep on ice during use and store at -20°C when finished for the day.

  • For each reaction, prepare 2.15 ml Digitonin Buffer (215 µl 10X Wash Buffer #31415 + 21.5 µl 100X Spermidine #27287 + 10.75 µl 200X Protease Inhibitor Cocktail #7012 + 53.75 µl Digitonin Solution #16359 + 1.849 ml Nuclease-free Water #12931).
  • Place Calcium Chloride on ice.
  • Make sure the 10% SDS Solution #20533 is completely in solution. Warming it up to 37°C will help to dissolve the SDS precipitates.
  • For each reaction, prepare 150 µl of 1X Stop Buffer (37.5 µl 4X Stop Buffer #48105 + 3.75 µl Digitonin Solution #16359 + 0.75 µl RNAse A #7013 + 108 µl Nuclease-free Water #12931).

    Optional: Sample Normalization Spike-In DNA can be added into the 1X Stop Buffer if sample normalization is desired (for example, see Figure 8 in Section VII). For qPCR analysis, we recommend adding 5 µl (5 ng) of Spike-In DNA to each reaction. For NG-seq analysis, we recommend diluting the Sample Normalization Spike-In DNA 100-fold into Nuclease-free Water #12931 and then adding 5 µl (50 pg) of Spike-In DNA to each reaction. When using 100,000 cells or 1 mg of tissue per reaction this ensures that the normalization reads are around 0.5% of the total sequencing reads. If more or less than 100,000 cells or 1 mg of tissue are used per reaction, proportionally scale the volume of Sample Normalization Spike-In DNA up or down to adjust normalization reads to around 0.5% of total reads.

  1. Briefly centrifuge samples from Section III, Step 6 at 100 x g for 2 sec to remove cell:bead suspension from the caps of the tubes.
  2. Place the tubes on the magnetic separation rack until the solution turns clear (30 sec to 2 min) and then remove the liquid.
  3. Remove tubes from the magnetic separation rack. Add 1 ml of Digitonin Buffer (+ spermidine + PIC + digitonin) and resuspend beads by gently pipetting up and down.
  4. Repeat steps 2 and 3 one time.
  5. Place the tubes on the magnetic rack until the solution turns clear (30 sec to 2 min) and then remove the liquid.
  6. Remove tubes from magnetic rack. Add 150 µl of Digitonin Buffer (+ spermidine + PIC + digitonin) to each tube and mix by pipetting up and down.
  7. Place tubes on ice for 5 min to cool before digestion.
  8. Activate pAG-MNase by adding 3 µl cold Calcium Chloride to each tube and mix by pipetting up and down.
  9. Incubate samples at 4°C for 30 min.

    NOTE: Digestion should be performed in a 4°C cooling block or refrigerator. The temperature of ice can get as low as 0°C, which can limit digestion and decrease signal.

  10. Add 150 µl of 1X Stop Buffer (+ digitonin + RNAse A + spike-in DNA [optional]) to each sample and mix by pipetting up and down.
  11. Incubate the tubes at 37°C for 10 min without shaking to release DNA fragments into the solution.
  12. Centrifuge at 4°C for 2 min at 16,000x g and place the tubes on a magnetic rack until the solution is clear (30 sec to 2 min).
  13. Transfer the supernatants to new 2 ml microcentrifuge tubes. These are your enriched chromatin samples.

    NOTE: If live cells or fresh tissues (not fixed) are used for the CUT&RUN assay, skip Steps 14-15 and immediately proceed to Step 16.

    NOTE: Fixed samples will be incubated at 65°C later in the protocol, so it is recommended to use a safe-lock 2 ml tube to reduce evaporation during the incubation.

  14. To reverse the crosslinks in fixed cell or tissue samples, allow samples to warm to room temperature and add 3 µl of 10% SDS Solution #20533 (0.1% final concentration) and 2 µl of proteinase K (20 mg/ml) #10012 to each sample.

    NOTE: SDS may precipitate out of solution if samples are not pre-warmed to room temperature.

  15. Vortex each sample and incubate at 65°C for at least 2 hr. This incubation can be extended overnight. After incubation, quickly spin samples at 10,000 x g for 1 sec to collect evaporation from the cap of tubes.
  16. Equilibrate samples to room temperature and proceed to Section VI. (SAFE STOP) Alternatively, samples can be stored at -20°C for up to 1 week. However, be sure to warm samples to room temperature before DNA purification (Section VI).

V. Preparation of the Input Sample

Fragmentation of input DNA is required for compatibility with downstream NG-Sequencing but is not necessary for downstream qPCR analysis. If a sonicator is not available, we recommend using the unfragmented input DNA for qPCR analysis; however, the input DNA should be purified using phenol/chloroform extraction and ethanol precipitation because the size of unfragmented input DNA is too large to be purified using DNA spin columns. If a sonicator is not available and downstream NG-Sequencing analysis is desired, one can use the CUT&RUN normal IgG antibody sample as the negative control, although this is not ideal because the normal IgG-enriched sample may show non-specific DNA enrichment. Alternatively, an input DNA fragmentation protocol using MNase is available at https://cst-science.com/CUT-RUN-input-digestion.

! All buffer volumes should be increased proportionally based on the number of input samples being prepared.

Before Starting:

  • Remove and warm DNA Extraction Buffer #42015. Make sure it is completely thawed.
  • For each input sample, prepare 2 µl Proteinase K #10012 + 0.5 µl RNAse A #7013 + 197.5 µl DNA Extraction Buffer #42015 (200 µl total per input sample).
  1. Add 200 µl of DNA Extraction Buffer (+ Proteinase K + RNAse A) to the 100 µl input sample from Section I-A Step7, Section I-B Step 9, or Section I-C Step 16. Mix by pipetting up and down.
  2. Place the tube at 55°C for 1 hr with shaking.
  3. Place the tubes on ice for 5 min to completely cool the sample.
  4. Lyse the cells and fragment the chromatin by sonicating the input samples. Incubate samples on ice for 30 seconds between pulses.

    NOTE: Sonication conditions may need to be determined empirically by testing different sonicator power settings and/or durations of sonication, following the protocol in Appendix B. Optimal sonication conditions will generate chromatin fragments ranging in size from 100-600 bp. Sonication for 5 sets of 15-sec pulses using a VirTis Virsonic 100 Ultrasonic Homogenizer/Sonicator at setting 6 with a 1/8-inch probe sufficiently fragments the input chromatin.

  5. Clarify lysates by centrifugation at 18,500 x g for 10 min at 4°C. Transfer supernatant to a new 2 ml microcentrifuge tube.
  6. Immediately proceed to Section VI DNA Purification. (SAFE STOP) Alternatively, samples can be stored at -20°C for up to 1 week. However, be sure to warm samples to room temperature before DNA purification procedures (Section VI).

VI. DNA Purification

DNA can be purified from input and enriched chromatin samples using DNA spin columns, as described in Section VI - A, or phenol/chloroform extraction followed by ethanol precipitation as described in Section VI - B. Purification using DNA spin columns is simple and fast, providing good recovery of DNA fragments above 35 bp (Figure 7A, Lane 2). Phenol/chloroform extraction followed by ethanol precipitation is more difficult, but provides better recovery of DNA fragments below 35 bp (Figure 7A, Lane 3); however, as shown in Figure 7B, the majority of DNA fragments generated in the CUT&RUN assay are larger than 35 bp. Therefore, DNA spin columns provide a quick and simple method for purification of > 98% of the total CUT&RUN DNA fragments.

Purified DNA can be quantified prior to NG-seq analysis using a picogreen-based DNA quantification assay. For CUT&RUN reactions containing 100,000 cells, the expected DNA yield for a CUT&RUN reaction ranges from 0.5 to 10 ng per reaction for transcription factors and cofactors, and 1 to 20 ng per reaction for histone modifications.

FIGURE 7

FIGURE 7. Comparison of DNA purification using spin columns or phenol/chloroform extraction followed by ethanol precipitation. (A) A low range DNA ladder mix (lane 1, unpurified) was purified using either DNA Purification Buffers and Spin Columns (ChIP, CUT&RUN) #14209 (lane 2) or phenol/chloroform extraction followed by ethanol precipitation (lane 3) and separated by electrophoresis on a 4% agarose gel. As shown, phenol/chloroform followed by ethanol precipitation efficiently recovers all DNA fragment sizes, while DNA spin columns recover DNA fragments ≥ 35 bp. (B) DNA was purified using phenol/chloroform extraction followed by ethanol precipitation from a CUT&RUN assay performed using TCF4/TCF7L2 (C48H11) Rabbit mAb #2569. The size of the DNA fragments in the library was analyzed using a Bioanalyzer (Agilent Technologies). The adaptor and barcode sequences added to the library during construction account for 140 bp in fragment length. Therefore, starting 35 bp DNA fragments would be 175 bp in length after library preparation (indicated with blue vertical line in figure). As shown, less than 2% of the total CUT&RUN enriched DNA fragments are less than 175 bp (starting length of 35 bp), suggesting that DNA purification spin columns are sufficient for capture of > 98% of the total CUT&RUN DNA fragments.

A. DNA Purification Using Spin Columns

NOTE: DNA can be purified from input and enriched chromatin samples using the DNA Purification Buffers and Spin Columns (ChIP, CUT&RUN) #14209 (not included in this kit) and the modified protocol below. Steps 1 through 5 are modified to reflect the requirement for adding 5 volumes (1.5 ml) of DNA Binding Buffer to the 300 µl of input and enriched chromatin samples.

Before starting:

  • !! Add 24 ml of ethanol (96-100%) to DNA Wash Buffer before use. This step only has to be performed once prior to the first set of DNA purifications.
  • Remove one DNA Purification collection tube for each enriched chromatin sample or input sample to be purified.
  1. Add 1.5 ml DNA Binding Buffer to each input and enriched chromatin sample and mix by pipetting up and down.

    NOTE: 5 volumes of DNA Binding Buffer should be used for every 1 volume of sample.

  2. Transfer 600 µl of each sample from Step 1 to a DNA spin column in collection tube.
  3. Centrifuge at 18,500 x g in a microcentrifuge for 30 sec.
  4. Remove the spin column from the collection tube and discard the liquid. Replace the spin column in the empty collection tube.
  5. Repeat steps 2-4 until the entire sample from Step 1 has been spun through the spin column. Replace the spin column in the empty collection tube.
  6. Add 750 µl of DNA Wash Buffer to the spin column in collection tube.
  7. Centrifuge at 18,500 x g in a microcentrifuge for 30 sec.
  8. Remove the spin column from the collection tube and discard the liquid. Replace spin column in the empty collection tube.
  9. Centrifuge at 18,500 x g in a microcentrifuge for 30 sec.
  10. Discard collection tube and liquid. Retain spin column.
  11. Add 50 µl of DNA Elution Buffer to each spin column and place into a clean 1.5 ml tube.
  12. Centrifuge at 18,500 x g in a microcentrifuge for 30 sec to elute DNA.
  13. Remove and discard DNA spin column. Eluate is now purified DNA. (SAFE STOP) Samples can be stored at -20°C for up to 6 months.

B. DNA Purification Using Phenol/Chloroform Extraction and Ethanol Precipitation

NOTE: The following reagents are required for the phenol/chloroform extraction and ethanol precipitation and are not included in this kit: phenol/chloroform/isoamyl alcohol (25:24:1), chloroform/isoamyl alcohol (24:1), 3M Sodium Acetate (pH 5.2), 20mg/ml glycogen, 100% ethanol, 70% ethanol, and 1X TE buffer or Nuclease-free Water #12931.

  1. Add 300 µl of phenol/chloroform/isoamyl alcohol (25:24:1) to each input and enriched chromatin sample and mix thoroughly by vortexing for 30 sec.
  2. Separate layers by centrifugation at 16,000 x g for 5 min in a microcentrifuge. Carefully transfer most of the top aqueous layer (avoiding the interphase) to a new tube.
  3. Add 300 µl of chloroform/isoamyl alcohol (24:1) to the aqueous sample and mix thoroughly by vortexing for 30 sec.
  4. Separate layers by centrifugation at 16,000 x g for 5 min in a microcentrifuge. Carefully transfer most of the top aqueous layer (avoiding the interphase) to a new tube.
  5. Add 25 µl of 3M Sodium Acetate (pH 5.2), 1 µl 20mg/ml glycogen, and 600 µl of 100% ethanol to each aqueous sample and mix by vortexing for 30 sec.
  6. Incubate samples at -80°C for 1 hr or -20°C overnight to precipitate DNA.
  7. Pellet DNA by centrifugation at 16,000 x g for 5 min at 4°C in a microcentrifuge.
  8. Carefully remove supernatant and wash pellet with 70% ethanol.
  9. Pellet DNA by centrifugation at 16,000 x g for 5 min at 4°C in a microcentrifuge.
  10. Decant supernatant and air dry pellet.
  11. Resuspend pellet in 50 µl of 1X TE buffer or Nuclease-free Water #12931. This is the purified DNA. (SAFE STOP) Samples can be stored at -20°C for up to 6 months.

VII. Quantification of DNA by qPCR

Recommendations:

  • The Sample Normalization Primer Set included in the kit is specific for the S. cerevisiae ACT1 gene and can be used to quantify the signal from the Sample Normalization Spike-In yeast DNA for sample normalization (optional).
  • The additional control primers included in the kit are specific for the human or mouse RPL30 gene (#7014 or #7015) and can be used for quantitative real-time PCR analysis of the Tri-Methyl-Histone H3 (Lys4) (C42D8) Rabbit mAb #9751 sample. If the user is performing CUT&RUN on another species, the user needs to design the appropriate control primers and determine the optimal PCR conditions for that species.
  • PCR primer selection is critical. For CUT&RUN, PCR amplicon sizes should be approximately 60 to 80 bp in length. Primers should be designed with optimum melting temperature around 60°C and GC content around 50%.
  • 2 µl of purified DNA is sufficient for qPCR-mediated quantification of target genes for histones, transcription factors, and cofactors.
  • A Hot-Start Taq polymerase is recommended to minimize the risk of nonspecific PCR products.
  • Use Filter-tip pipette tips to minimize risk of contamination.
  1. Label the appropriate number of PCR tubes or PCR plates compatible with the model of PCR machine to be used. PCR reactions should include the positive control tri-methyl-histone H3 Lys4 sample, the negative control rabbit IgG sample, a tube with no DNA to control for DNA contamination, and a serial dilution of the input DNA (undiluted, 1:5, 1:25, 1:125) to create a standard curve and determine the efficiency of amplification and quantify the amount of DNA in each immune-enriched sample.

    NOTE: If sample normalization is performed, only the CUT&RUN samples are to be analyzed using the Sample Normalization Primer Set. The input DNA does not contain the Normalization Spike-In DNA.

  2. Add 2 µl of the appropriate DNA sample to each tube or well of the PCR plate.
  3. Prepare a master reaction mix as described below. Set up 2-3 replicates for each PCR reaction. Add enough reagents to account for loss of volume (1-2 extra reactions). Add 18 µl of reaction mix to each PCR reaction tube or well.
Reagent Volume for 1 PCR Reaction (18 µl)
Nuclease-free H2O #12931 6 µl
5 µM Primers 2 µl
SimpleChIP® Universal qPCR Master Mix #88989 10 µl
  1. Start the following PCR reaction program:
a. Initial Denaturation 95°C for 3 min
b. Denature 95°C for 15 sec
c. Anneal and Extension 60°C for 60 sec
d. Repeat steps b and c for a total of 40 cycles.
  1. Analyze quantitative PCR results using the software provided with the real-time PCR machine. Alternatively, one can calculate the IP efficiency manually using the Percent Input Method and the equation shown below. With this method, signals obtained from each immunoprecipitation are expressed as a percent of the total input chromatin.
    • Percent Input = 100% x 2(C[T] 100%Input Sample - C[T] IP Sample)
    • C[T] = CT = Average threshold cycle of PCR reaction
  2. For sample normalization, choose the sample that has the lowest C[T] value for the Sample Normalization Primer Set as the selected sample (e.g. Sample 1 in the example table below) and calculate the normalization factor of other samples using the below equation. Adjust the signals from the test primer sets using the respective normalization factors.
An Example of Sample Normalization for qPCR Assay (see Figure 8)
C[T] value of Sample Normalization Primer Set **Normalization Factor for qPCR Signal Before Normalization (% Input Calc'd from Step 5) Signal After Normalization
Sample 1 23.31 2(23.31-23.31)=1.00 24.4% 24.4%/1.00=24.4%
Sample 2 24.24 2(23.31-24.24)=0.52 12.0% 12.0%/0.52=23.1%
Sample 3 25.08 2(23.31-25.08)=0.29 6.28% 6.28%/0.29=21.7%
Sample 4 26.30 2(23.31-26.30)=0.13 2.72% 2.72%/0.13=20.9%

**Normalization Factor for qPCR = 2(C[T] Selected Sample - C[T] the Other Sample)

FIGURE 8

FIGURE 8. Normalization of CUT&RUN signals using spike in DNA for qPCR analysis. CUT&RUN was performed with a decreasing number of HCT116 cells and either Tri-Methyl-Histone H3 (Lys4) (C42D8) Rabbit mAb #9751 (upper panels) or Phospho-Rpb1 CTD (Ser2) (E1Z3G) Rabbit mAb #13499 (lower panels). Enriched DNA was quantified by real-time PCR using SimpleChIP® Human GAPDH Exon 1 Primers #5516, SimpleChIP® Human β-Actin Promoter Primers #13653, SimpleChIP® Human β-Actin 3' UTR Primers #13669, and SimpleChIP® Human MyoD1 Exon 1 Primers #4490. The amount of immunoprecipitated DNA in each sample is represented as signal relative to the total amount of input chromatin for 100,000 cells. Non-normalized enrichments are depicted in the left panels. The Sample Normalization Spike-In DNA was added into each reaction proportionally to the starting cell number. Based on the difference of qPCR signals from spike in DNA in each sample, CUT&RUN signals were normalized to the sample containing 100,000 cells. Normalized enrichments are depicted in the right panels.

VIII. NG-Sequencing Library Construction

The immuno-enriched DNA samples prepared with this kit are directly compatible with NG-seq. For downstream NG-seq DNA library construction, use a DNA library preparation protocol or kit compatible with your downstream sequencing platform. For sequencing on Illumina® platforms, we recommend using the SimpleChIP® ChIP-seq DNA Library Prep Kit for Illumina® #56795 with SimpleChIP® ChIP-seq Multiplex Oligos for Illumina® #29580 or #47538.

Additional Recommendations for DNA Library Preparation:

  • During DNA End Preparation (Section I, Step 4 of the SimpleChIP® ChIP-seq DNA Library Prep Kit for Illumina® #56795), change the thermocycler program to incubate at 50°C for 30 min instead of 65°C for 30 min to avoid denaturation of small DNA fragments. Denatured DNA fragments are not compatible with adaptor ligation.
  • We recommend diluting the Adaptor for Illumina® in 10 mM Tris-HCl (pH 8.0-8.5) to avoid adaptor contamination in the library, based on the actual amount of starting CUT&RUN DNA (Section II, Step 1 of the SimpleChIP® ChIP-seq DNA Library Prep Kit for Illumina® #56795). If starting DNA is 1.25 ng to 2.5 ng, dilute the adaptor 1:50 to generate a working concentration of 0.3 µM. If starting DNA is less than 1.25 ng, dilute the adaptor 1:125 to generate a working concentration of 0.12 µM.
  • When purifying adaptor-ligated DNA (Section III, Step 1 of the SimpleChIP® ChIP-seq DNA Library Prep Kit for Illumina® #56795), add 1.1X volume instead of 0.9X volume of AMPure® XP beads or SPRIselect® beads to samples to increase the capture of smaller DNA fragments.
  • During PCR Enrichment of adaptor-ligated DNA (Section IV, Step 3 of the SimpleChIP® ChIP-seq DNA Library Prep Kit for Illumina® #56795), reduce the Anneal and Extension time from 75 sec to 13 sec to exclude amplification of large library DNA fragments (>1,000 bp).
  • The DNA yield from the CUT&RUN assay may be lower than the 5 ng of starting DNA recommended for ChIP-seq library preparation, especially when low cell numbers are used. Based on the actual CUT&RUN DNA yield, we recommend increasing the number of PCR amplification cycles up to 20 cycles in order to generate a library with DNA concentration of 10-30 ng/µl. We recommend 13 cycles for 3-5 ng starting DNA, 16 cycles for 0.75-1 ng starting DNA, or 20 cycles if there is less than 0.2 ng starting DNA. When 17-20 cycles are used, it’s better to split the total cycles into two PCR programs to ensure the optimal performance of Q5® PCR Master Mix due to the limited amount of components in the PCR Master Mix (eg. dNTP). For example, if 17 cycles are needed, run a first round of PCR program with 9 cycles, purify the PCR product following the Section V of the SimpleChIP® ChIP-seq DNA Library Prep Kit for Illumina® #56795, then set up a second round of PCR with the same Index Primers (containing specific barcodes) used in the first round of PCR and amplify for another 8 cycles. Continue with Section V of the SimpleChIP® ChIP-seq DNA Library Prep Kit for Illumina® #56795 to complete the library preparation.
  • When purifying library DNA (Section V, Step 1 of the SimpleChIP® ChIP-seq DNA Library Prep Kit for Illumina® #56795), add 1.0X volume instead of 0.9X volume of AMPure® XP beads or SPRIselect® beads to samples to increase the capture of smaller DNA fragments.
  • If a higher library concentration is desired, decrease the volume of 10 mM Tris-HCl (pH 8.0-8.5) to elute DNA from AMPure® XP beads or SPRIselect® beads (Section V, Steps 8 of the SimpleChIP® ChIP-seq DNA Library Prep Kit for Illumina® #56795). For example, use 17 µl of 10 mM Tris-HCl and collect 15 µl of supernatant as the final library DNA sample.
  • Because of the very low background signal generated in CUT&RUN, a sequencing depth of 5 million reads per sample is usually sufficient for histone modifications and transcription factors. The duplication rate of reads significantly increases if the sequencing depth is greater than fifteen million per sample. The signal to noise ratio decreases if the sequencing depth is lower than two million per sample.
  • For less than 20,000 starting number of cells, it is common to obtain lower mapping rates or higher duplication rates in the NGS reads. If this happens, we recommend increasing the sequencing depth to obtain a sufficient amount of unique mapped reads for downstream data analysis.
  • When performing sample normalization, map CUT&RUN sequencing data for all samples to both the test reference genome (e.g. human) and the sample normalization S. cerevisiae yeast genome. Choose the sample that has the least number of unique yeast reads as the selected sample (e.g. Sample 1 in table below) and calculate the normalization factor of other samples using the equation below. Downsize the number of unique reads aligned to test reference genome for each sample using the respective normalization factors. Use the downsized dataset for further NGS analysis.
An Example of Sample Normalization for NGS Assay
The Number of Unique Reads Aligned to Yeast Normalization Factor for NGS The Number of Unique Reads Aligned to Test Reference Genome Before Normalization The Number of Unique Reads Aligned to Test Reference Genome After Normalization
Sample 1 219,275 219,275/219,275 = 1.00 5,077,747 5,077,747 X 1.00 = 5,077,747
Sample 2 411,915 219,275/411,915 = 0.53 9,896,671 9,896,671 X 0.53 = 5,268,306
Sample 3 816,235 219,275/816,235 = 0.27 17,842,773 17,842,773 X 0.27 = 4,793,320
Sample 4 1,120,826 219,275/1,120,826 = 0.20 23,836,679 23,836,679 X 0.20 = 4,663,339

Normalization Factor for NGS = the number of unique yeast reads from Selected Sample / the number of unique yeast reads from the other sample

APPENDIX A: Determination of Cell Sensitivity to Digitonin

In the CUT&RUN protocol, the addition of digitonin to the buffers facilitates the permeabilization of cell membranes and entry of the primary antibody and pAG-MNase enzyme into the cells and nuclei. Therefore, having an adequate amount of digitonin in the buffers is critical to the success of antibody and enzyme binding and digestion of targeted genomic loci. Different cell lines show differing sensitivities to digitonin cell permeabilization. While the amount of digitonin recommended in this protocol should be sufficient for permeabilization of most cell lines or tissues, you can test your specific cell line or tissue using this protocol. We have found that the addition of excess digitonin is not deleterious to the assay, so there is no need to perform a concentration curve. Rather, a quick test to determine if the recommended amount of digitonin works for your cell line is sufficient.

Before starting:

  • Remove and warm Digitonin Solution #16359 to 90-100°C for 5 min. Make sure it is completely thawed. Immediately place the thawed Digitonin Solution #16359 on ice.

    NOTE: Digitonin Solution #16359 should be stored at -20°C. Please keep on ice during use and store at -20°C when finished for the day.

  • For each cell or tissue sample, prepare 100 µl of Wash Buffer (10 µl 10X Wash Buffer #31415 + 90 µl Nuclease-free Water #12931). It is not necessary to add spermidine or Protease Inhibitors for this test.
  1. In a 1.5 ml tube, collect 10,000 - 100,000 cells. For tissue, collect disaggregated cells from 1 mg of tissue (Section I-C Step 1-13). If you are using fixed cells or tissue in your CUT&RUN experiment, be sure to fix the cells or tissue the same way for this test.
  2. Centrifuge for 3 min at 600 x g at room temperature and withdraw the liquid.

    NOTE: If the cell pellet is not visible by eye, we recommend removing as much cell medium as possible without disturbing the cell pellet after the initial centrifugation of the cell suspension in Step 2 and leave behind some cell medium per reaction. Then in Step 3 add enough 1X Wash Buffer to the cell suspension to achieve a total volume of 100 µl.

  3. Resuspend cell pellet in 100 µl of Wash Buffer.
  4. Add 2.5 µl Digitonin Solution #16359 to each reaction and incubate for 10 min at room temperature.
  5. Mix 10 µl of cell suspension with 10 µl of 0.4% Trypan Blue Stain.
  6. Use a hemocytometer or cell counter to count the number of stained cells and the total number of cells. Sufficient permeabilization results in > 90% of cells staining with Trypan blue.
  7. If less than 90% of cells stain with Trypan blue, then increase the amount of Digitonin Solution #16359 added to the Digitonin Buffer and repeat steps 1-5 until > 90% cells are permeabilized and stained. Use this amount of Digitonin Solution #16359 in Sections I - IV.

APPENDIX B: Sonication Optimization for the Input Sample

Sonication of the input DNA sample is recommended because only fragmented genomic DNA (<10 kb) can be purified using DNA purification spin columns. Additionally, the fragmented genomic DNA (<1kb) may be used as the negative control in NG-seq analysis. Sonication should be optimized so that the input DNA is 100-600 bp in length.

We recommend using the input sample for NG-seq because it provides a convenient and unbiased representation of the cell genome. While the IgG sample can also be used as a negative control for NG-seq, it may show enrichment of specific regions of the genome due to non-specific binding. Unfragmented input DNA can be used for qPCR analysis. However, unfragmented DNA must be purified using phenol/chloroform extraction followed by ethanol precipitation.

Before starting:

! All buffer volumes should be increased proportionally based on the number of input samples being prepared.

  • Remove and warm DNA Extraction Buffer #42015 at room temperature, making sure it's completely thawed and in solution.
  • For each input sample, prepare 2.1 ml 1X Wash Buffer (210 µl 10X Wash Buffer #31415 + 1.89 ml Nuclease-free Water #12931) and equilibrate it to room temperature to minimize stress on the cells. It is not necessary to add spermidine or Protease Inhibitor Cocktail #7012 to this Wash Buffer.
  • For each input sample, prepare 2 µl Proteinase K #10012 + 0.5 µl RNAse A #7013 to 197.5 µl DNA Extraction Buffer #42015 (200 µl per input sample).
  1. In a 1.5 ml tube, collect the same number of cells you use for the input in your CUT&RUN experiment (5,000 to 100,000 cells) for each sonication condition being tested. For tissue, collect disaggregated cells from the same amount of tissue you use for the input in your CUT&RUN experiment (Section I-C Step 1-13) for each sonication condition being tested. If you are using fixed cells or tissue in your experiment, be sure to fix the cells or tissue the same way for this test.
  2. Centrifuge for 3 min at 600 x g at room temperature and remove the liquid.

    NOTE: If the centrifuged cell pellet is not visible by eye when working with low cell numbers (<100,000 cells), we recommend skipping the wash steps 3-5 below. Remove as much cell medium as possible without disturbing the cell pellet after the initial centrifugation of the cell suspension in Step 2 and leave behind some cell medium per reaction. Then in Step 6 add enough 1X Wash Buffer (+ spermidine + PIC) to the cell suspension to achieve a volume of 100 µl per sonication condition being tested.

  3. Resuspend cell pellet in 1 ml of 1X Wash Buffer by gently pipetting up and down.
  4. Centrifuge for 3 min at 600 x g at room temperature and remove the liquid.
  5. Wash the cell pellet again by repeating steps 3 and 4 one time.
  6. Add 100 µl of 1X Wash Buffer per sonication condition being tested and resuspend the cell pellet by gently pipetting up and down.
  7. Aliquot 100 µl of the cell suspension into a new tube for each sonication condition.

    NOTE: Samples will be incubated at 55°C in Step 9, so it is recommended to use a safe-lock 1.5 ml tube to reduce evaporation during the incubation.

  8. Add 200 µl DNA Extraction Buffer (+ Proteinase K + RNAse A) to each sample and mix by pipetting up and down.
  9. Place the tubes at 55°C for 1 hr with shaking.
  10. Place the tubes on ice for 5 min to completely cool down the samples.
  11. Determine optimal sonication conditions for your sonicator by setting up a time-course experiment with increasing numbers of 15 sec pulse sonication cycles. Be sure to incubate samples on ice for 30 sec between pulses.
  12. Clarify lysates by centrifugation at 18,500 x g in a microcentrifuge for 10 min at 4°C. Transfer supernatant to a new 2 ml microcentrifuge tube.
  13. Purify the DNA samples with DNA Purification Spin Columns or phenol/chloroform extraction followed by ethanol precipitation, following the directions in Section VI.
  14. Elute the DNA from the column or resuspend DNA pellet in 30 µl of 1X TE buffer or Nuclease-free Water #12931.
  15. Determine DNA fragment sizes by electrophoresis. Load > 15 µl sample on a 1% agarose gel with a 100 bp DNA marker. A dye-free loading buffer (30% glycerol) is recommended to better observe the DNA smear on gel.
  16. Choose the sonication conditions that generate the optimal DNA fragment size of 100-600 bp and use for Preparation of the Input Sample in Section V, Step 4. If optimal sonication conditions are not achieved, increase or decrease the power setting of the sonicator or number of sonication cycles and repeat the sonication time course experiment.

APPENDIX C: Troubleshooting Guide

For a detailed troubleshooting guide, please go to https://cst-science.com/troubleshooting-CUT-RUN

Protocol Id: 1884

Specificity / Sensitivity

MLL1 (D2M7U) Rabbit mAb (Amino-terminal Antigen) recognizes endogenous levels of total MLL1 protein.

Species Reactivity:

Human, Mouse, Rat, Monkey

Source / Purification

Monoclonal antibody is produced by immunizing animals with recombinant protein specific to the amino terminus of human MLL1 protein.

Background

The Set1 histone methyltransferase protein was first identified in yeast as part of the Set1/COMPASS histone methyltransferase complex, which methylates histone H3 at Lys4 and functions as a transcriptional co-activator (1). While yeast contain only one known Set1 protein, mammals contain six Set1-related proteins: SET1A, SET1B, MLL1, MLL2, MLL3, and MLL4, all of which assemble into COMPASS-like complexes and methylate histone H3 at Lys4 (2,3). These Set1-related proteins are each found in distinct protein complexes, all of which share the common subunits WDR5, RBBP5, ASH2L, CXXC1 and DPY30, which are required for proper complex assembly and modulation of histone methyltransferase activity (2-6). MLL1 and MLL2 complexes contain the additional protein subunit, menin (6).MLL1 functions as a master regulator of both embryogenesis and hematopoiesis, and is required for proper expression of Hox genes (7,8). MLL1 is a large, approximately 4000 amino acid, protein that is cleaved by the taspase 1 threonine endopeptidase to form N-terminal (MLL1-N) and C-terminal MLL1 (MLL1-C) fragments, both of which are subunits of the functional MLL1/COMPASS complex (9,10). MLL1-N, MLL1-C, WDR5, RBBP5 and ASH2L define the core catalytic component of the MLL1/COMPASS complex, which is recruited to target genes and methylates histone H3 lysine 4 to regulate transcriptional initiation (11). At least 60 different MLL1 translocation partners have been molecularly characterized and associated with various hematological malignancies. The most common translocation partners include AF4, AF9, ENL, AF10, ELL and AF6 (8,12,13). With the exception of AF6, all of these partners are nuclear proteins that function to positively regulate transcriptional elongation. AF4, AF9 and ENL are all components of the super elongation complex (SEC), while AF4, AF9, AF10 and ENL all interact with the histone H3 lysine 79 methyltransferase DOT1L. Many MLL1 target genes are normally regulated by promoter-proximal pausing, with the release of RNA polymerase and transcriptional elongation occurring in response to proper stimuli (14). The association of MLL1 translocation partners with SEC and DOT1L suggest that MLL1-fusion proteins may function to sustain specific gene expression programs by constitutively activating transcriptional elongation.
  1. Miller, T. et al. (2001) Proc Natl Acad Sci U S A 98, 12902-7.
  2. Shilatifard, A. (2008) Curr Opin Cell Biol 20, 341-8.
  3. Tenney, K. and Shilatifard, A. (2005) J Cell Biochem 95, 429-36.
  4. Lee, J.H. and Skalnik, D.G. (2005) J Biol Chem 280, 41725-31.
  5. Lee, J.H. et al. (2007) J Biol Chem 282, 13419-28.
  6. Hughes, C.M. et al. (2004) Mol Cell 13, 587-97.
  7. Eissenberg, J.C. and Shilatifard, A. (2010) Dev Biol 339, 240-9.
  8. Smith, E. et al. (2011) Genes Dev 25, 661-72.
  9. Takeda, S. et al. (2006) Genes Dev 20, 2397-409.
  10. Yokoyama, A. et al. (2002) Blood 100, 3710-8.
  11. Dou, Y. et al. (2006) Nat Struct Mol Biol 13, 713-9.
  12. Yip, B.H. and So, C.W. (2013) Exp Biol Med (Maywood) 238, 315-23.
  13. Neff, T. and Armstrong, S.A. (2013) Blood 121, 4847-53.
  14. Wang, P. et al. (2009) Mol Cell Biol 29, 6074-85.

Pathways & Proteins

Explore pathways + proteins related to this product.

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