Mapping Genome-wide Accessible Chromatin in Primary Human T Lymphocytes by ATAC-Seq

All the procedures were approved by institutional review board of Bar Ilan University and the protocol follows guidelines provided by the committee approving the experiments.

1. Purification of Naïve Human CD4+ Cells and Polarization to T Helper 1 (Th1) and Th2 Cells

Note: Here we describe the procedure starting from frozen human peripheral blood mononuclear cells (PBMCs). The first step consists of isolating CD4+ cells using microbeads and columns that usually give us more than 95% of CD4+ cells. However, this step may vary according to the preferred protocol in each lab. The protocol for T cell activation and polarization was modified from Jenner et al. (2009)¹⁸. Isolation of CD4+ cells from 10 million PBMCs gives rise to 4 – 6 million of CD4+ cells. They are split into two flasks and grown under Th1 and Th2 polarizing conditions yielding 3-5 million Th1 and Th2 cells within just a week.

Note: Cool down the centrifuge to 4 °C before starting.

1. Thaw 1 mL of human PBMCs (10⁷ cells) in a 50 mL tube containing 10 mL of RPMI medium supplemented with 1% Penicillin-Streptomycin, 2 mM L-glutamine and 10% heat-inactivated fetal bovine serum. Centrifuge at 500 x g for 5 min. Remove the supernatant and resuspend the cells with a sterile 25 mL pipette in 15 mL of supplemented RPMI medium. Transfer the cells (with sterile 25 mL pipette) to a T75 culture flask.
2. Leave the cells overnight in a humidified incubator (37 °C, 5% CO₂).
3. Transfer the floating cells with a sterile 25 mL pipette to 50 mL tube. Determine cell number and viability by trypan blue exclusion.
4. Isolate CD4+ cells from 10 million live non-adherent PBMCs by positive selection using CD4+ microbeads and columns according to the manufacturer's recommendations (see Table of Materials/Equipment) with the following modifications: 10⁷ PBMCs are labeled with 30 µL of CD4 microbeads in 120 µL of 0.5% BSA in PBS.
5. Activate the CD4+ T cells for 72 h by rhIL-2 (10 ng/mL), plate-bound anti-CD3 (5 µg/mL) and soluble anti-CD28 (2 µg/mL). For Th1 polarization, add rhIL-12 (20 ng/mL) and anti-IL-4 (10 µg/mL). For Th2 polarization, add rhIL-4 (40 ng/mL) and anti-IFN-γ (10 µg/mL).
6. Culture the cells for additional 7 days in the presence of rhIL-2 (10 ng/mL) and the same polarizing cytokines (rhIL-12 for Th1 and rhIL-4 for Th2).

2. Nuclei Isolation

NOTE: ATAC-seq is performed with intact nuclei. Lysis buffer containing 0.05% nonylphenyl polyethylene glycol (see Table of Materials/Equipment) was calibrated for isolating nuclei from primary human Th1 and Th2 cells. We recommend calibrating this step with the laboratory reagents and cells. An excess of intact cells from insufficient detergent decreases the efficiency of the transposition reaction. Cell lysis efficiency is determined by the number of nuclei (trypan blue positive cells) relative to the total number of cells.
NOTE: Prepare the lysis buffer (10 mM Tris-HCl, pH 7.5, 10 mM NaCl, 3 mM MgCl₂). Cool down a centrifuge with a swing-bucket rotor to 4 °C. Pelleting the cells in a swing-bucket centrifuge instead of a fixed angle centrifuge reduces cell/nuclei loss. To avoid nuclei or cell loss, pipet carefully when discarding the supernatant.

1. Add fresh nonylphenyl polyethylene glycol (to a final concentration of 0.05%) and 100x protease inhibitors (to a final concentration of 1x) to the cold lysis buffer immediately before use. Keep the buffer on ice.
2. Count T cells to determine their amount and viability using trypan blue method. Viability that is lower than 90% results in higher non-specific digestion.
3. Transfer 0.5 x 10⁶ T cells (Th1 or Th2) to 1.5 mL microtubes. Spin down at 500 x g for 5 min at 4 °C.
4. Resuspend the cell pellet in 1 mL of cold phosphate-buffered saline (PBS) solution. Spin down at 500 x g for 5 min at 4 °C.
5. Resuspend the cell pellet in 1 mL of cold lysis buffer (containing nonylphenyl polyethylene glycol and protease inhibitors). Keep the tube on ice. Pipette gently to avoid disrupting the nuclei.
6. Quickly take 10 µL and count the cells with an automated cell counter while the microtube with lysed cells is on ice. This step should not exceed five minutes to avoid damaging the nuclei. At least 80% of the cells should be lysed.
7. Continue immediately with the transposition reaction. Keep the prepared nuclei on the ice.

3. Transposition reaction

NOTE: In this step, isolated nuclei are incubated with prokaryotic Tn5 transposase (TDE1) loaded with adapters for NGS sequencing. Hyperactive Tn5 simultaneously fragments DNA and ligates adapters into accessible regions of the genome (tagmentation process). The ratio between nuclei and Tn5 transposase is critical for preferential cleavage at accessible chromatin. This protocol is calibrated for 100,000 nuclei in a 100 μL reaction volume. However, the reaction can be scaled down.

1. Set temperature in a thermal shaker to 37 °C.
2. Transfer 100,000 nuclei to a 1.5 mL microtube.
3. Centrifuge at 500 x g for 10 min at 4 °C and gently remove the supernatant.
4. Add the transposition reaction components to the nuclei as specified in Table 1.
5. Resuspend by gentle pipetting.
6. Incubate the transposition reaction in a thermal shaker at 37 °C for 30 min with gentle shaking (500 rpm).
   Note: DNA Cleanup is performed by solid-phase reversible immobilization beads¹⁹ (see Table of Materials/Equipment) or PCR purification columns. At the end of the cleanup, elute the DNA fragments in 20 µL of 10 mM Tris-HCl, pH 8. Avoid EDTA in the elution buffer.

4. PCR Enrichment of ATAC-seq Libraries

NOTE: This step is aimed to amplify the ATAC-seq library i.e., DNA fragments with inserted adapters. To allow mixing of several ATAC-seq libraries in the same next-generation sequencing lane ("multiplexing") use unindexed Primer 1 (Ad1_noMx)¹ for all samples, and a different indexed (barcoded) Primer 2 (Ad 2.1 – 2.24)¹ for each sample. The primer sequences are provided in the supplemented Table of Materials/Equipment.

1. Initial PCR amplification
   NOTE: The working concentration of Primer 1 (Ad1_noMx) and Primer 2 is 25 µM. All the primers are diluted from original stock of 100 µM to 25 µM. In all of the PCR reactions, use Primer 1 (Ad1_noMx) and only one of the indexed Primer 2.
   1. Add the components of the PCR reaction as specified in Table 2 to a sterile PCR tube.
   2. Place PCR tube into a thermal cycler and perform PCR amplification using the cycling conditions detailed in Table 3.
2. Assessment of number of additional amplification cycles
   NOTE: The number of additional PCR cycles should yield sufficient amount of library fragments for a successful next-generation sequencing run, while minimized to avoid GC and size bias²⁰. The determination of the number of PCR cycles (N) required for optimal library fragment amplification is done by quantitative PCR (qPCR).
   1. Dilute Primers 1 (Ad1_noMx) and 2 (used for initial library amplification) from 25 µM to 6.25 µM.
   2. Add the components to optical PCR tubes or a plate as stated in Table 4.
   3. Place in a qPCR instrument and cycle as specified in Table 5.
   4. To estimate the required number of additional amplification cycles (N), plot cycle number on the x-axis and relative fluorescence (RFU) on the y-axis.
   5. The number of additional amplification cycles (N) is 1/3 of the number of cycles at which the qPCR reaction reached the plateau. Figure 2 provides examples for three ATAC-seq libraries that reached plateau at ~2,350 relative fluorescence units, RFU (thick green line). The number of PCR cycles in which one third of the maximal amount (783 RFU, marked on y-axis) is amplified corresponds to 8 cycles for two of the libraries (red and blue amplification curves) and 9 PCR cycles for the third library (pink).
3. Final PCR amplification
   1. Amplify the remaining 45 µL of the PCR reaction. Place a PCR tube containing amplification reaction from step 4.1.2 in a thermal cycler. Run the PCR program described in Table 6. Use the previously determined (step 4.2.5) number of amplification cycles (N).

5. Size Selection of ATAC-Seq Libraries

NOTE: In our experience, size selection of amplified ATAC-seq libraries improves next-generation sequencing results because it eliminates high molecular weight library fragments from the final ATAC-seq library.
NOTE: Allow the magnetic beads to warm to room temperature 30 min before use.
Prepare fresh 70% ethanol in nuclease-free water.

1. Resuspend the magnetic beads by mixing.
2. Add nuclease-free water to the ATAC-seq libraries (obtained in step 4.3.1.) and bring up to 100 µL.
3. Add 50 µL (0.5x) of resuspended DNA-binding magnetic beads to 100 µL of amplified libraries. Mix by pipetting up and down at least 10 times. Incubate samples for 5 min at room temperature. If necessary, quickly spin down the microtubes.
4. Place the tube on an appropriate magnetic stand for 2 min to separate the magnetic beads from the supernatant. After 2 min, transfer the supernatant to a new microtube.
5. Measure the volume of the supernatant by pipetting and add 0.7x of magnetic beads. Mix by pipetting up and down at least 10 times.
6. Incubate 5 min at room temperature. Place on a magnetic stand for 2 min.
7. After the 2 min incubation, discard the supernatant. Add 200 µL freshly made 70% ethanol to wash the beads while the tubes are on the magnetic stand.
8. Keep the microtube on the magnet for 30 s and then discard the ethanol.Repeat step 5.7.for two final ethanol washes.
9. Completely remove the remaining ethanol and let the beads air-dry for 5 min while the tube is on the magnet. If necessary, briefly spin the microtube. Remove the traces of ethanol with a p10 pipette tip.
10. Remove the microtube from the magnet and add 22 µL of 10 mM Tris-HCl, pH 8. Do not elute the ATAC-seq libraries in buffer containing EDTA.
11. Incubate the tube for 2 min at room temperature and then place on the magnetic stand.
12. When the solution is clear, transfer 20 µL of eluted libraries to a new sterile microtube.
13. Store the size selected ATAC-seq libraries at -20 °C.

6. Quality Analysis of the ATAC-Seq Libraries

1. Validation of the quality of ATAC-seq libraries by Real-Time PCR
   NOTE: It is important to assess the signal to noise ratio of the ATAC-seq libraries prior to next-generation sequencing. This is done by determining the relative amount of DNA fragments from accessible and inaccessible loci using quantitative PCR (qPCR). The inaccessible loci (negative control, chr1:48,137,860-48,137,934 and chr1:193,093,748-193,093,827) are amplified by primer pairs 1 and 2. The accessible loci (positive control, chr19:30,336,166-30,336,253 and chr19:11546154-11546237) are amplified by primer pairs 3 and 4. Positive and negative loci were defined from chromatin accessibility (DHS-seq) profiles of human CD4+ cells (ENCODE accessions ENCSR000EQE and ENCSR000EQG). Negative primer 1 is located in a large heterochromatic intergenic region (230 kb from TRADB2 and 88 kb from FOXD2). Negative control region 2 is in within the first intron of CDC73 gene. Positive primer pair 3 is within an open chromatin region downstream of the Cyclin E (CCNE1) gene while positive primer pair 4 is centered within the promoter of protein kinase C substrate 80K-H (PRKCSH). Importantly, these control loci show a similar pattern of accessibility in other human cell types from the ENCODE project³, suggesting that they can be applied to monitor ATAC-seq libraries from a broad spectrum of human cell types. Primer efficiency and specificity of all primer pairs was verified by qPCR on a serial dilution of genomic DNA (from human Th cells) and a melting curve analysis of obtained amplified products.
   1. Isolate genomic DNA using a commercially available kit (see Table of Materials/Equipment).
   2. Dilute the amplified ATAC-seq library 1:10 (1 µL library + 9 µL of nuclease-free water) and genomic DNA to ~5 ng/µL.
   3. Prepare reaction mixture (Table 7) for each positive and negative control primer pair, taking into account that the reactions are performed in triplicate.
   4. Incubate in qPCR thermal cycler according to the protocol recommended by qPCR master mix supplier.
   5. Analyze the results in qPCR instrument's software (see Table of Materials/Equipment). Choose genomic DNA as a control sample. The obtained values represent an enrichment of accessible regions (amplified by positive control primers). An example is shown in Table 8.
      NOTE: Estimation of the average library size and concentration: the size distribution of DNA fragments from ATAC-seq libraries is determined by high-sensitivity automated electrophoresis systems according to the manufacturer's instructions. It is advisable to measure the sample concentration on a fluorometer using dsDNA high-sensitivity kit and at least 2 µL of each DNA sample.
      NOTE: Next-generation sequencing – prior to multiplexing the libraries, calculate the molarity of each ATAC-seq library by using the formula: (ng/µL x 10⁶)/(660 x average library fragment length). Aim for > 30 million reads of each ATAC-seq library to assess open chromatin regions of human samples. If you wish to determine if the library is good enough for NGS sequencing, initially aim for ~10 million reads (5% of the sequencing lane on DNA sequencing instrument in rapid run mode). Keep in mind that to infer nucleosome positioning, paired-end sequencing is needed¹.

7. Analysis of the Obtained Next-Generation Sequencing Results

1. Infer the quality of the sequencing reads by inspecting the FastQC files, separately for every library.
2. Align the reads to human reference genome (hg19 assembly) using Bowtie²¹ software in the Unix/Linux environment. The command is 'bowtie -m 1 -q -S genome directory reads.fastq output_aligned.sam'. Genome directory stands for the folder where the genome indexes of Bowtie are stored. The parameter -m 1 is for not allowing alignment of reads to more than one locus in the genome, -q is for the input file that should be in fastq format, -S is for the output that is in SAM format.
3. Remove duplicate reads using SAMtools²² rmdup option in the Unix/Linux environment. The commands are 'samtools view -S output_aligned.sam -b | samtools sort -o -output_aligned > output_aligned.bam',
   samtools rmdup -s output_aligned.bam output_aligned _rmdup.bam'. The first command, view, changes the SAM format into BAM format which is then sorted. The option of rmdup is then applied on the sorted BAM file. Optionally one can adjust the reads for the transposon insertion site, as described in the original ATAC-seq paper¹.This is done using BEDtools commands²³ in the Unix/Linux environment. The commands are 'bamToBed -i output_aligned _rmdup.bam > output_aligned _rmdup.bed ', 'shiftBed -i output_aligned _rmdup.bed -p 4 -m -5 -g genome > output_aligned _rmdup_adjusted.bed'. The first command, bamTobed, changes the BAM format into BED format which can then be used for the shiftBed command. The genome file is a tab delimited file that contains the length of each chromosome in the genome. The file is usually added in the BEDtools directory.
4. Perform peak calling using model-based analysis of ChIP-seq (MACS2)²⁴ software in the Unix/Linux environment on the shifted BED file with the following parameters: –nomodel –extsize 75 –shift -30. These parameters are used to adjust the reads so that the transposon insertion site is in the middle of each sequencing read.