ATAC-seq library prep

Documentation

Assay for Transposase Accessible Chromatin (ATAC-seq) has been shown to be compatible with many methods for cell collection and has also worked effectively across many cell types and species. However, the following protocol has been optimized for human lymphoblastoid cells. Minor variations (i.e. cell number, centrifugation speeds, and lysis conditions) may be required to optimize for your particular application. We have seen that crosslinking greatly reduces library efficiency, and therefore we recommend starting with fresh unfixed cells.

What you’ll need

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All of the items for this protocol need to be prepared fresh or purchased

  • 2x TD Buffer (recipe below)
  • lysis buffer (10 mM Tris-HCl, pH7.4, 10 mM NaCl, 3 mM MgCl, 0.1% IGEPAL CA-630)
  • Tn5 Transposes (Illumina Cat #FC-121-1030)
  • Qiagen PCR Cleanup Kit
  • 10,000x SYBR Green I (Invitrogen Cat #S-7563)
  • NEBNext High-Fidelity 2x PCR Master Mix (New England Labs Cat #M0541)
  • 1.5mL DNA LoBind tubes (to be used for all steps in this protocol

Cell Preparation

  • Harvest your cells (no fixation). How exactly you do this will depend on your particular tissue, cell type of interest, and experimental system.
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Make sure you are very accurate about cell number and remember to use the LoBind tubes

  • Spin down 50,000 cells at 750 ×g for 5 min, 4°C.
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Pellets will be very small, so keep an eye on the orientation of your tubes in the centrifuge and be careful to recognize this as you remove the tubes, since it will help you identify the pellet.

  • Very slowly remove the liquid. Wash once with 50 μL of cold 1x PBS buffer. DO NOT PIPET UP AND DOWN.
  • Spin down at 750 ×g for 5 min, 4°C.
  • Very slowly remove the liquid. Gently pipette to resuspend (just 3 times up and down) the cell pellet in 50 μL of cold lysis buffer.
  • Incubate on ice for 2 minutes.
  • Spin down immediately at 750 ×g for 10 min, 4°C.
  • Discard the supernatant, and immediately continue to transposition reaction.
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Make sure you remove all the buffer before proceeding

Transposition Reaction and Purification

  • Make sure your cell pellet is on ice, and make the following transposition reaction mix

Transposition reaction mix

Namevolume (uL)
2x TD Buffer
25
Tn5 Transpose
2.5
Nuclease-free water
22.5
50 uL total
  • Gently pipette to resuspend nuclei in the transposition reaction mix.
 Pipet up and down 10 times.
  • Incubate the transposition reaction at 37°C for 45 min (mouse) or 30 min (human).
  • Immediately following transposition, purify using a Qiagen MinElute Kit.
 Follow kit instructions —check pH!
  • Elute transposed DNA in 10 μL Elution Buffer (10mM Tris buffer, pH 8).
 Allow EB buffer to incubate for 5min on column, then spin.
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Purified DNA can be stored at -20°C until you are ready to carry out PCR

PCR Amplification

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Be sure to record which Illumina primers you use in the reaction below

  • To amplify transposed DNA fragments, combine the following:

PCR reaction master mix

NameVolume (uL)Comments
Transposed DNA
10
Nuclease-free water
12.7
Illumina Primer 1
1
Illumina Primer 2
1
100x SYBR Green I
0.3
10,000x SYBR Green I is diluted in EB Buffer to make a 100x working solution.
NEBNext High-Fidelity 2x PCR Master Mix
25
5o uL total

PCR cycling condition

Temp (C)TagsFiles
72
5:00
1
98
0:30
1
98
0:10
4
63
0:30
4
72
1:00
4
4
hold
NA
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The number of PCR cycles corresponds to the starting material. Do not perform more than 16 cycles as it can lead to amplification of background material. Use the following table below to choose appropriate cycles.

Untitled

Input of total RNA (ng)Typical number of PCR cycles Regular RNA
50
9-10
10
12
1
14-15
0.5
16
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Safe stopping point, samples can be left overnight at 4C. If not processed within the next day, freeze the PCR products at -20C for up to 2 weeks.

Purification of Final RNA-Seq Library Using AMPure Beads

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Move samples to a 96 well plate before beginning this step. We do not have a magnetic stand that holds PCR strip tubes. You may recombine your split samples from the previous step here, or keep them split and do the bead cleanup separately on each.

Allow AMPure XP beads to come to room temperature for about 30 minutes before use. Vortex beads for 2 minutes to mix well. Make sure samples are in a 96 well plate that will fit on our magnetic stand. Prepare fresh 80% ethanol, you will about 400 uL per sample.
Add 100 uL of AMPure beads to each sample and pipette 10 times to mix. [This is a 1:1 ratio; if you kept each sample separated into 2 wells, add 50 uL of beads. If all 100uL of PCR product is one a single well, add 100uL of beads very carefully because this will completely fill the wells.]
Incubate at room temperature for 8 minutes to allow the DNA to bind to the beads.
Place plate on magnetic stand for 5 minutes or longer until the solution is completely clear.
While samples are on magnetic stand, remove and discard supernatant. Do not disrupt beads.
Keeping samples on magnetic stand, add 180 uL (use 180 uL per well regardless of whether samples were split) of freshly made 80% ethanol to each sample without disturbing beads. Wait 30 seconds and carefully pipette out and discard supernatant. cDNA will remain bound to beads during the washing process.
Repeat wash step above 1 more time.
Let samples air dry for about 1 minute on magnetic stand then remove any excess ethanol with a P20.
Air dry samples on magnetic stand for 3-5 minutes at room temperature until pellets appear dry and matte. Once you start to see pellets crack, take off magnetic stand to ensure you do not overdry your samples.
Once beads are dry, remove from magentic stand and add 20 uL (10uL per well if samples still split) of Tris Buffer to cover beads, pipette to mix thoroughly until the beads are resuspended.
Incubate at room temperature for 5 minutes.
Place plate back on magnetic stand for 1 minute or longer until the supernatant is clear.
Pipette 18 uL (9 uL per well if samples still split) supernatant to LoBind Tubes. This is the final product. If samples were split, recombine supernatant from both halves into a single tube with 18uL total.

Quality Check of Final Product

Run samples on HSD1000 Tapestation Assay.
Measure concentration with HS dsDNA Qubit assay.

Sequencing Guidelines

Normalize and Pool

  1. If not done already, quantify each of your libraries on Qubit. For most libraries, using the HS dsDNA Qubit assay with 2uL of input will yield a reading. Record the concentration in ng/uL for each library.
  2. Run your samples on Tapestation with either the D1000 or the HSD1000 assay. Remember to allow Tapestation reagents to sit at room temperature for at least 30 minutes before use. Save your Tapestation results by going to File -> Create Report -> Save as pdf. This file can then be emailed or uploaded to Asana. For the base pair length, we usually use the value of the peak identified by the Tapestation analysis software. This value is shown both on the tracing itself and in the Peak Table for each sample.
  3. Download our nM Conversion Calculator here. Enter the concentration (from Qubit) and the base pair length (from Tapestation) in the appropriate cells and it will give you the nM concentration for each library. Normalize and pool all your libraries to 4, 2, 1, or 0.5 nM in a LoBind microcentrifuge tube. If you need to dilute your libraries, we recommend using at least 2uL to minimize pipetting errors. The example sheet of the calculator provides further detail.
  4. Quantify your pool on Qubit and enter into the calculator sheet to check that your pool is close to the nM concentration you normalized to.

Setup Run in Basespace

  1. Sign into Basespace, then go to the Prep tab, Biological Samples, and select Import Samples on the upper right. Use Illumina’s Sample Import Template to enter information about your samples. The SampleID and Name can be the same, but make sure they are unique for each sample. Species can be left blank. Upload the completed .csv to import your samples.
  2. Continue to Prep Libraries. Select the library prep kit “TruSeqHT” If you used another index format, you will need to use a different entry for library prep kit. The your project name as the Plate ID. For each sample, check the box next to it on the left, then drag the sample name to the appropriate index well.
  3. Proceed to Pool Libraries. Select all your samples on the left, then drag and drop in the pool on the right. Name the pool your project name.
  4. Continue to Plan Run. Select NextSeq and name your run your project name. Select Single Read or Paired End Read, then enter the cycle numbers based on your selected kit. For example, if you were doing a run using a High Output 75 Cycle kit, you would select Single Read and enter 76 for Read 1 Cycles and 0 for Read 2 Cycles.
  5. Press Sequence to complete planning the run. The run will now be available for selection on the sequencer.

Loading the Sequencer

  • The next step is to dilute and denature the prepared libraries. Illumina’s general guidelines for this on the NextSeq can be found here.
  • Illumina’s system guide for the NextSeq, which covers the sequencing workflow, can be found here.
  • Your final loading concentration should be 1.3 - 1.8 pM, with most pools loaded at 1.4 or 1.5 pM.