Adapted ATAC-seq protocol from Monahan et al eLife 2017

Assay for Transposase Accessible Chromatin

Adapted by Kevin Monahan (2017)

Original protocol by: Beijing Wu, Jason Buenrostro, 

Greenleaf & Chang Lab Stanford University

November 2013

Notes/changes are bolded and underlined.

I. Cell Preparation 

1. Harvest cells (no fixation), protocol to be defined by the user. 

2. Spin down 50,000 cells at 500 ×g for 5 min, 4°C. 

   (Use very clear microcentrifuge tubes to facilitate pellet identification – DNA LoBind work well)

3. Wash once with 50 μL of cold 1x PBS buffer. Spin down at 800 ×g for 5 min, 4°C. 

4. Gently pipette to resuspend the cell pellet in 50 μL of cold lysis buffer (10 mM Tris-HCl, pH 7.4, 10 mM NaCl, 3 mM MgCl2, 0.1% IGEPAL CA-630). Spin down immediately at 1000 ×g for 10 min, 4°C. 

5. Discard the supernatant, and immediately continue to transposition reaction. 

II. Transposition Reaction and Purification 

1. Make sure the cell pellet is set on ice. 

2. To make the transposition reaction mix, combine the following: 

            25 μL 2x TD Buffer (Illumina Cat #FC-121-1030)

            2.5 μL Tn5 Transposes (Illumina Cat #FC-121-1030)

            22.5 μL Nuclease Free H2O

            50 μl Total

3. Gently pipette to resuspend nuclei in the transposition reaction mix. 

4. Incubate the transposition reaction at 37°C for 30 min 

    (Test different transposition times to improve fragment size distribution) . 

5. Immediately following transposition, purify using a Qiagen MinElute Kit (Zymo 5 also works). 

    6. Elute transposed DNA in 10 μL Elution Buffer (10mM Tris buffer, pH 8). 

7. Purified DNA can be stored at -20°C. 

III. PCR Amplification 

1. To amplify transposed DNA fragments, combine the following in a PCR tube: 

            10 μL Transposed DNA 

            10 μL Nuclease Free H2O 

            2.5 μL 25μM Customized Nextera PCR Primer 1* 

            2.5 μL 25μM Customized Nextera PCR Primer 2* [Barcode] 

            25 μL NEBNext High-Fidelity 2x PCR Master Mix (New England Labs Cat #M0541) 

            50 μL Total

2. Cycle as follows:

            (1) 72°C, 5 min 

            (2) 98°C, 30 sec 

            (3) 98°C, 10 sec 

            (4) 63°C, 30 sec 

            (5) 72°C, 1 min 

            (6) Repeat steps 3-5, 4x 

            (7) Hold at 4°C 

3. In order to reduce GC and size bias in PCR, the PCR reaction is monitored using qPCR to stop amplification prior to saturation. To run a qPCR side reaction, combine the following: 

            5 μL 5 cycles PCR amplified DNA 

            4.41 μL Nuclease Free H₂O 

            0.25 μL 25μM Customized Nextera PCR Primer 1* 

            0.25 μL 25μM Customized Nextera PCR Primer 2* 

            0.09 μL 100x SYBR Green I **

            5 μL NEBNext High-Fidelity 2x PCR Master Mix 

            15 μL Total 

            * Complete list of primers available in Section VI of this protocol 

            **10,000x SYBR Green I is diluted in 10mM Tris buffer, pH 8 to make a 100x working solution. 

4. qPCR cycle as follows: (I use white-walled PCR strips for this)

           (1) 98°C, 30 sec 

           (2) 98°C, 10 sec 

           (3) 63°C, 30 sec 

           (4) 72°C, 1 min 

           (5) Repeat steps 2-4, 19x 

           (6) Hold at 4°C 

5. The additional number of cycles needed for the remaining 45 μL PCR reaction is determined as following: 

For each sample, determine maximum fluorescent intensity – then set threshold where fluorescent intensity reached 1/5^th of this maximum value. Round down the CT for this threshold, this is the number of additional cycles to run (e.g. a 1/5^th max CT of 5.7 = 5 more cycles)

6. Run the remaining 45 μL PCR reaction to the correct # of cycle. Cycle as follows: 

          (1) 98°C, 30 sec 

          (2) 98°C, 10 sec 

          (3) 63°C, 30 sec 

          (4) 72°C, 1 min 

          (5) Repeat steps 2-4, x times 

          (6) Hold at 4°C 

7. Purify amplified library using ampure XP beads

1. Transfer PCR product to Lo-Bind tube. 
2. Add 72uL room temperature (RT) Ampure XP beads to 45uL. Pipet to mix. Incubate 10 minutes at RT.
3. Place on magnet for 5 minutes
4. Remove 105uL of supernatant (leaving ~12uL)
5. Wash twice for 30 seconds with 200uL FRESH 70% EtOH.
6. Pipet out last wash. Wait until beads dry completely (10-15 minutes)
7. Add 33uL pH 8 Low-EDTA TE (10mM Tris pH 8, 0.1mM EDTA). Resuspend beads by pipetting. Incubate 3 minutes at RT.
8. Place on magnet for 3 minutes.
9. Collect 30uL of eluate containing finished library. 

8. QC finished library:

      -run on gel/bioanalyzer to examine fragment size distribution

      -run KAPA, usually 10-40nM

N.B.: This protocol is inferior to newer protocols, such OMNI-ATAC (https://doi.org/10.1038/nmeth.4396)