Adipocyte-Specific ATAC-Seq with Adipose Tissues Using Fluorescence-Activated Nucleus Sorting

Animal care and experimentation were performed according to procedures approved by the Institutional Animal Care and Use Committee of Indiana University School of Medicine.

1. Preparations before beginning the experiment

1. Tissue preparation
   1. For adipocyte nucleus labeling, cross NuTRAP mice with adipocyte-specific adiponectin-Cre lines (Adipoq-Cre) to generate Adipoq-NuTRAP mice, which are hemizygous for both Adipoq-Cre and NuTRAP.
   2. Dissect the adipose tissues of interest from the Adipoq-NuTRAP mice as described previously¹⁴.
   3. Snap-freeze the tissues in liquid nitrogen, and store them at −80 °C until use.
      NOTE: Each fat pad can be stored as a whole, and the frozen tissues can be later cut on dry ice into smaller pieces (~50 mg) for experiments. Alternatively, the fat tissues can be cut, aliquoted, and frozen in tubes for storage (~50 mg per tube). Frozen samples are never allowed to thaw after freezing until use.
2. Buffer preparation
   NOTE: Keep the buffers on ice throughout the experiment.
   1. Prepare nucleus preparation buffer (NPB): 250 mM sucrose, 10 mM HEPES (pH 7.5), 10 mM KCl, 1.5 mM MgCl₂, and 0.1% NP40. Prepare 7 mL of NPB per sample. Add fresh 100x protease inhibitor cocktail (final 1x), DTT (final 1 mM), and Hoechst (final 1 µg/mL) to the NPB before use.
      NOTE: It is recommended to prepare fresh NPB, but it can be stored at 4 °C for up to 1 month.
   2. Prepare 1x phosphate-buffered saline with 0.1% NP-40 (PBS-N).

2. Nucleus isolation

1. Chill glass Dounce homogenizers, with one glass Dounce per sample, on ice. Then, add 7 mL of the NPB mix to each glass Dounce.
   NOTE: It is recommended to use up to eight samples in each experiment. Working with more than eight samples would significantly delay all the steps, and it may especially downgrade the nucleus quality during sorting.
2. Put the frozen adipose tissue (50-100 mg) in the glass Dounce, and immediately cut it into a few smaller pieces using a long pair of scissors. Stroke 10x with a loose pestle for all the samples, and then 10x with a tight pestle.
   NOTE: Adipose tissues are soft, so cutting them into a few small pieces should be sufficient. For rare samples, smaller pieces (10-20 mg) can be used, although the collected adipocyte nucleus number may vary.
3. Filter through a 100 µm strainer into a new 50 mL tube. Move the filtered homogenates to a new 15 mL tube by pouring. Spin at 200 × g for 10 min at 4 °C in a swing-bucket centrifuge. Remove the supernatant as much as possible.
   NOTE: Aspirate first using a vacuum, and then remove the remaining volume by using a micropipette.
4. Resuspend the pellet in 500 µL of PBS-N by gentle but thorough finger tapping.
   NOTE: Avoid pipetting, as this may damage the nuclei.
5. Filter through a 40 µm strainer into a new 50 mL tube using gentle pipetting. Rinse the strainer with 250 µL of PBS-N. Transfer the filtered nuclear resuspension to a fluorescence-activated cell sorting (FACS) tube using a micropipette.
   ​NOTE: If available, tubes and cell strainers can be used for smaller volumes to minimize sample loss.

3. Nucleus sorting

1. Collection tube preparation
   1. Add 500 µL of PBS-N to new 1.5 mL tubes, and invert a few times to wet all inner surfaces with the buffer.
   2. Briefly spin the tubes to bring down all the liquid, and then chill them on ice until sorting.
2. FACS (Figure 2)
   1. Use the FSC-A/FSC-H gating for singlets and FSC-A/SSC-A for small or large debris removal.
   2. Gate for the mCherry/GFP-positive adipocyte nucleus population.
   3. Collect 10,000 nuclei in each of the collection tubes prepared in step 3.1.
3. Post-sorting nucleus preparation
   1. Add 500 µL of PBS-N to each collection tube, and invert a few times to mix.
   2. Spin the collection tubes at 200 × g for 10 min at 4 °C in a swing-bucket centrifuge. Completely remove the supernatant.
      NOTE: Use a vacuum first to remove bubbles, pour off the supernatant, and use paper wipes to remove the remaining liquid completely. The pellet is invisible at this point. Prepare the Tn5 master mixture during the centrifugation step as described below.

4. Tn5 Tagmentation (Table 1)

1. To prepare 25 µL of Tn5 master mixture, mix 12.5 µL of 2x tagmentation DNA buffer (TD buffer), 1.25 µL of Tn5 transposase (TDE I enzyme), and 11.25 µL of nuclease-free water.
2. Add 25 µL of Tn5 master mixture to each nucleus pellet, and resuspend by gentle pipetting. Incubate at 600 rpm for 30 min at 37 °C using a thermomixer.

5. DNA purification

NOTE: The following procedure uses the PCR purification kit mentioned in the Table of Materials. Any other similar DNA purification methods can be used.

1. Add 25 µL of nuclease-free water to the 25 µL of the mixture of Tn5 nucleus resuspension obtained after step 4.2. The final volume is 50 µL per sample.
2. Add 250 µL of buffer PB.
3. Add 5 µL of 3 M sodium acetate (NaOAc) (pH 5.2).
4. Transfer the mixture to a column (provided with the referenced kit), and centrifuge at 17,900 × g for 1 min at room temperature (RT).
5. Discard the flowthrough, and place the spin column back in the same collection tube. Add 750 µL of buffer PE containing absolute ethanol (EtOH), and centrifuge at 17,900 × g for 1 min at RT.
6. Discard the flowthrough, and place the spin column back into the same collection tube. Centrifuge at 17,900 × g for 1 min at RT, and discard the collection tube with the flowthrough.
7. To elute the DNA fragments, equip the column with a new 1.5 mL tube, add 10 µL of buffer EB, and leave at RT for 3 min. Centrifuge at 17,900 × g for 1 min at RT.
   ​NOTE: The samples can be stored at 4 °C for days or at −20 °C for weeks.

6. PCR amplification (Table 2)

NOTE: The primers used in this study are listed in Table 3.

1. To amplify transposed DNA fragments, prepare PCR master mixture without adding a unique Ad2.n barcoding primer (primer #2). Use 38 µL of the PCR master mixture per sample: 11 µL of nuclease-free water, 2 µL of 25 µM Ad1_noMX primer (primer #1), and 25 µL of 2x PCR Master Mix.
2. Add 38 µL of the PCR master mixture into a 0.2 mL PCR tube.
3. Add 10 µL of the eluted DNA fragments from step 5.7.
4. Add 2 µL of 25 µM primer #2, which needs to be specific for each sample.
5. Run the PCR according to the cycling conditions indicated in Table 2.

7. Real-time quantitative PCR (qPCR) test (Table 4)

NOTE: This step aims to determine the additional cycles needed to amplify the DNA. It is optional but highly recommended, especially for new experiments.

1. Prepare qPCR master mixture without adding primer #2. Use 9.975 µL of the qPCR master mixture per sample: 4.41 µL of nuclease-free water, 0.25 µL of 25 µM primer #1, 5 µL of 2x PCR Master Mix, and 0.09 µL 100x SYBR Green I.
   NOTE: To prepare 100x SYBR Green I, dilute the 10,000x stock with nuclease-free water. As the volume of 100x SYBR Green I used for the qPCR master mixture is negligible, it is easier to make an additional master mixture of diluted 100x SYBR Green I (e.g., for 8-10 samples).
2. Add 9.975 µL of the qPCR master mixture into each well of a qPCR plate.
3. Add 0.25 µL of 25 µM primer #2 into each well.
   NOTE: Be sure to add the same primer #2 to match what was added to each specific sample in step 6.4.
4. Add 5 µL of the PCR reaction obtained after the PCR amplification in step 6.5, and mix by pipetting.
   NOTE: The remaining 45 µL of the PCR reaction will be used for the second PCR amplification.
5. Run the qPCR according to the cycling conditions indicated in Table 5.
6. Check the amplification curves, and identify the number of additional PCR cycles needed by estimating the number of cycles that reached ~35% of the maximum (Figure 3A).
   NOTE: Typically, 10,000 nuclei require five to eight additional PCR cycles.

8. Additional PCR amplification

1. Run the PCR using all of the remaining 45 µL of the PCR reaction according to the calculations from step 7.6 (Table 6).
   ​NOTE: Each sample could require a different number of amplification cycles.

9. Second DNA purification using the PCR purification kit

1. Use the same procedures as in section 5, but elute the amplified DNA fragments with 20 µL of buffer EB.

10. DNA fragment size selection

NOTE: Use solid phase reversible immobilization beads, as described below. Any other similar DNA purification beads can be used according to the manufacturer's instructions. The SPRI bead suspension should be completely resuspended and equilibrated at RT with rotation before use.

1. Transfer the eluted DNA to a new PCR tube, and add 80 µL of buffer EB to make a final volume of 100 µL.
2. Add 55 µL of SPRI beads (0.55x sample volume), and mix by pipetting. Incubate for 5 min at RT, and then separate on a mini magnetic stand for PCR 8-tube strips for 5 min.
3. Transfer 150 µL of the supernatant to a new PCR tube. Add 95 µL of SPRI beads, and mix by pipetting.
   NOTE: The supernatant contains DNA of approximately 500 bp or smaller.
4. Incubate for 5 min at RT, and then separate on the mini magnetic stand for 5 min. Discard the supernatant carefully.
5. Wash the beads for 1 min with 200 µL of 70% EtOH. Repeat twice for three washes in total.
   NOTE: Do not move PCR tubes from the magnetic stand during the washes.
6. After the final wash, spin down the tubes at 1,000 × g for 1 min, and pipet out the remaining EtOH. Dry the pellet with the lid open at 37 °C for 2 min on a PCR machine.
   NOTE: The bead pellets should display only some minor cracks once dried. Avoid over-drying.
7. Resuspend the pellet with 20 µL of buffer EB by pipetting, and incubate for 5 min at RT. Separate on the mini magnetic stand for 5 min, and then transfer 18 µL of the supernatant containing the final library to a new 1.5 mL tube.
   ​NOTE The library can be stored at −20 °C while performing a quality check.

11. DNA quantification using a fluorometer

1. To prepare the master mixture, dilute 200x dsDNA High Sensitivity (HS) reagent with dsDNA HS buffer to a final concentration of 1x.
2. For standards, add 190 µL of the 1x master mixture and 10 µL of standard 1 or standard 2 to a fluorometer tube.
   NOTE: Equilibrate the standards at RT in the dark before starting the quantification.
3. For the library samples, add 198 µL of the 1x master mixture and 2 µL of each sample to a separate fluorometer tube.
   NOTE: If the sample amounts are limited, the sample volume can be reduced to 1 µL, and the 1x master mixture volume can be increased to 199 µL.
4. Vortex or shake the fluorometer tubes, and let them sit for 5 min at RT in the dark. Measure the DNA concentration using the dsDNA HS assay of the fluorometer.

12. Library quality check by high-sensitivity electrophoresis systems

1. Take the ATAC-seq library, and dilute with nuclease-free water to a final concentration of 1-5 ng/µL.
2. Analyze the size distribution of the ATAC-seq libraries using high-sensitivity, automated electrophoresis systems following the manufacturer's protocols.

13. Quality check of the ATAC-seq library by using targeted qPCR

1. Take 5-10 ng of the ATAC-seq library, and dilute with 50 µL of nuclease-free water.
2. Run the qPCR using primers targeting the promoters and enhancers known to be active in adipocytes according to the cycling conditions indicated in Table 7.
3. Use negative control primers targeting closed/silent genomic regions (Table 7).
4. Calculate the enrichment of the promoter/enhancers over the negative controls (Table 8).
   ​NOTE: It is expected to see ≥10-20-fold enrichments with successful samples.

14. Sequencing

1. Submit the ATAC-seq libraries for sequencing. Use paired-end sequencing (2 x 34 bp, 2 x 50 bp or longer) with average read numbers of 10-30 million reads per sample.