In-Nucleus Hi-C in Drosophila Cells

1. Fixation

1. Start with 10 million Schneider's line 2 plus (S2R+) cells to prepare 17.5 mL of a cell suspension in Schneider medium containing 10% fetal bovine serum (FBS) at room temperature (RT).
2. Add methanol-free formaldehyde to obtain a final concentration of 2%. Mix and incubate for 10 min at RT, taking care to mix every minute.
   NOTE: Formaldehyde is a hazardous chemical. Follow the appropriate health and safety regulations, and work in the fume hood.
3. Quench the reaction by adding glycine to achieve a final concentration of 0.125 M and mix. Incubate for 5 min at RT, followed by 15 min on ice.
4. Centrifuge for 400 × g at RT for 5 min and then for 10 min at 4 °C; discard the supernatant. Resuspend the pellet carefully in 25 mL of cold 1x phosphate-buffered saline.
5. Centrifuge at 400 × g for 10 min at 4 °C, then discard the supernatant.
   NOTE: If continuing with the protocol, go to step 2.1 for lysis; otherwise, flash-freeze the pellet in liquid N₂ and store the pellet at -80 °C.

2. Lysis

1. Resuspend the cells in 1 mL of ice-cold lysis buffer (10 mM Tris-HCl, pH 8; 0.2% of non-ionic surfactant (see the Table of Materials); 10 mM NaCl; 1x protease inhibitors), and adjust the volume to 10 mL with ice-cold lysis buffer. Adjust the volume to obtain a concentration of 1 × 10⁶ cells/mL. Incubate on ice for 30 min, mixing every 2 min by inverting the tubes.
2. Centrifuge the nuclei at 300 × g for 5 min at 4 °C, and then carefully discard the supernatant. Wash the pellet 1x with 1 mL of cold lysis buffer, and transfer it to a microcentrifuge tube. Wash the pellet 1x with 1 mL of cold 1.25x restriction buffer, and resuspend each cell pellet in 360 µL of 1.25x restriction buffer.
3. Add 11 µL of 10% sodium dodecyl sulfate (SDS) per tube (0.3% final concentration), mix carefully by pipetting, and incubate at 37 °C for 45 min, shaking at 700-950 rpm. Pipet up and down to disrupt clumps a few times during incubation.
4. Quench the SDS by adding 75 µL of non-ionic surfactant (10% solution, see the Table of Materials) per tube (1.6% final concentration), and incubate at 37 °C for 45 min, shaking at 950 rpm. Pipet up and down a few times to disrupt clumps during incubation.
   NOTE: If clumps are large and difficult to disrupt, decrease the rotating speed to 400 rpm during SDS and surfactant treatments. If the clumps are difficult to disaggregate by pipetting, split the sample into two; adjust the volumes of restriction buffer, SDS, and surfactant; and proceed with the permeabilization. Next, spin the nuclei at minimum speed (200 × g), carefully discard the supernatant, pool the samples together in 1X restriction buffer, and proceed with digestion. Take a 10 µL aliquot as the undigested sample (UD).

3. Enzymatic digestion

1. Digest the chromatin by adding 200 units (U) of Mbo I per tube, and incubate at 37 °C for a period ranging from 4 h to overnight while rotating (950 rpm).
2. On the next day, add an additional 50 U of Mbo I per tube, and incubate at 37 °C for 2 h while rotating (950 rpm).
3. Inactivate the enzyme by incubating the tubes at 60 °C for 20 min. Place the tubes on ice.
   NOTE: Take a 10 µL aliquot as the digested sample (D).

4. Biotinylation of DNA ends

1. To fill in the restriction fragment overhangs and label the DNA ends with biotin, add 1.5 µL each of 10 mM dCTP, dGTP, dTTP, 20 µL of 0.4 mM biotin dATP, 17.5 µL of Tris low-EDTA (TLE) buffer [10 mM Tris-HCl, 0.1 mM EDTA, pH 8.0], and 10 µL of 5 U/µL of Klenow (DNA polymerase I large fragment) to all the tubes. Mix carefully and incubate for 75 min at 37 °C. Shake at 700 rpm every 10 s for 30 s. Place all the tubes on ice while preparing the ligation mix.

5. Ligation

1. Transfer each digested chromatin mixture to a separate 1 mL tube with ligation mix (100 µL of 10x ligation buffer, 10 µL of 10 mg/mL bovine serum albumin, 15 U of T4 DNA ligase, and 425 µL of double-distilled water [ddH₂O]). Mix thoroughly by gentle pipetting, and incubate overnight at 16 °C.

6. Crosslink reversal and DNA purification

1. Degrade proteins by adding 50 µL of 10 mg/mL Proteinase K per tube, and incubate at 37 °C for 2 h. Reverse crosslinks by increasing the temperature to 65 °C and incubate overnight.
2. Degrade RNA by adding 20 µL of 10 mg/mL RNase A, and incubate at 37 °C for 1 h.
3. Perform phenol:chloroform extraction followed by ethanol precipitation.
   1. Add 1 volume of phenol-chloroform, and mix thoroughly by inversion to obtain a homogeneous white phase.
      NOTE: Phenol is a hazardous chemical. Follow the appropriate health and safety regulations. Work in the fume hood.
   2. Centrifuge at 15,000 × g for 15 min. Transfer the aqueous phase into a fresh 2 mL microcentrifuge tube. Perform a back-extraction of the lower layer with 100 µL of TLE buffer, and transfer the aqueous phase into the same 2 mL tube.
   3. Precipitate DNA by adding 2 volumes of 100% ethyl alcohol (EtOH), 0.1 volumes of 3 M sodium acetate, and 2 µL of 20 mg/mL glycogen. Incubate at -20 °C for a period ranging from 2 h to overnight.
   4. Spin at 15,000 × g for 30 min at 4 °C, and wash the pellets 2x with ice-cold 70% EtOH. Dry the pellets at RT, and resuspend in 100 µL of TLE buffer. Quantify the DNA using a fluorogenic dye that binds selectively to DNA and a fluorometer according to the manufacturer's instructions (Table of Materials).
      NOTE: The protocol can be paused here. Store ligation products at 4 °C for the short term or at -20 °C for the long term. Use an aliquot (100 ng) of the material as the ligated sample (L) for quality control.

7. Assess Hi-C template quality

1. Digestion and ligation qualitative controls
   1. Purify DNA from the UD and D aliquots by reversing the crosslinks, and perform phenol:chloroform extraction and ethanol precipitation as described above.
   2. Load 100 ng of UD, D, and L samples in a 1.5% agarose gel. Look for a smear centered around 500 bp in the D sample versus a high molecular weight band for the L sample (see representative results).
2. Digestion efficiency quantitative control
   NOTE: To assess the digestion efficiency more accurately, use the UD and D samples as templates to perform quantitative polymerase chain reactions (qPCR) using primers designed as follows.
   1. Design a primer pair that amplifies a DNA fragment containing the DNA restriction site for the enzyme used for digestion (Mbo I in the present protocol), called R in the formula in step 7.2.3.
   2. Design a primer pair that amplifies a control DNA fragment that does not contain the restriction site for the enzyme used for digestion (Mbo I for the present protocol), called C in the formula in step 7.2.3.
   3. Use the cycle threshold values (Ct values) of the amplification to calculate restriction efficiency according to the formula shown below:
      % Restriction = 100 – 100/2^{{(CtR – CtC)D – (CtR – CtC)UD}}
      Where CtR refers to the Ct value of fragment R, and CtC refers to the Ct value of the fragment C for sample D and sample UD.
      NOTE: The restriction percentage reflects the efficiency of the restriction enzyme cleaving the restricted (R) DNA fragment compared to a control (C) DNA fragment that does not contain the restriction DNA site. A restriction efficiency of ≥ 80% is recommended.
3. Detection of known interactions
   1. Perform PCR to amplify an internal ligation control to examine short-range and/or medium- or long-range interactions (see representative results).
   2. Alternatively, design primers to amplify a ligation product in which the primers are in forward-forward or reverse-reverse orientation in adjacent restriction fragments.
4. Fill-in and biotin-labeling control
   1. Verify Hi-C marking and ligation efficiency by amplification and digestion of a known interaction or a ligation product between adjacent restriction fragments in the genome, as described above.
      NOTE: Successful fill-in and ligation of the Mbo I site (GATC) generates a new site for the restriction enzyme Cla I (ATCGAT) at the ligation junction and regenerates the Mbo I site.
   2. Digest the PCR product with Mbo I, Cla I, or both. After running the samples on a 1.5-2% gel, estimate the relative number of 3C and Hi-C ligation junctions by quantifying the intensity of the cut and uncut bands²⁶.
      NOTE: An efficiency of > 70% is desired (see representative results).

8. Sonication

1. Sonicate the samples to obtain 200-500 bp DNA fragments. For the instrument used in this protocol (see the Table of Materials), dilute the sample (from 5 to 10 µg) in 130 µL of ddH₂O per tube, and set the instrument to sonicate to 400 bp: fill level: 10; duty factor: 10%; peak incident power (w): 140; cycles per burst: 200; time (s): 80.

9. Biotin removal/end repair

NOTE: The steps shown below are adjusted for 5 µg of Hi-C DNA.

1. To perform biotin removal, transfer the sample (130 µL) into a fresh microcentrifuge tube. Add 16 µL of 10x ligation buffer, 2 µL of 10 mM dATP, 5 µL of T4 DNA Polymerase (15U), and 7 µL of ddH₂O (160 µL of total volume). Incubate at 20 °C for 30 min.
2. Add 5 µL of 10 mM dNTPs, 4 µL of 10x ligation buffer, 5 µL of T4 polynucleotide kinase (10 U/µL), 1 µL of Klenow, and 25 µL of ddH₂O (200 µL of total volume). Incubate at 20 °C for 30 min.

10. Size selection

1. To select fragments mostly in the 250-550 bp size range, perform sequential solid phase reversible immobilization (SPRI) size selection first with 0.6x, followed by 0.90x according to the manufacturer's instructions, and elute the DNA using 100 µL of TLE.

11. Biotin pulldown/A-tailing/adapter ligation

NOTE: Perform the washes by resuspending the magnetic beads by vortexing, rotate the samples for 3 min on a rotating wheel, and then briefly spin down the sample and place it on the magnetic stand. Allow the beads to stick to the magnet, discard the supernatant, and proceed with the following wash step. Perform the washes at 55 °C on a thermo-block with rotation instead of the rotating wheel.

1. Make up the final volume to 300 µL per sample with TLE for pull-down. Prepare the bead-washing buffers: 1x Tween Buffer (TB) (TB: 5 mM Tris-HCl pH 8.0, 0.5 mM EDTA, 1 M NaCl, 0.05% Tween), 0.5x TB, 1x No-Tween buffer (NTB) (5 mM Tris-HCl pH 8.0, 0.5 mM EDTA, 1 M NaCl), 2x NTB.
2. Use 150 µL of streptavidin-linked magnetic beads (see the Table of Materials) per library. Wash the beads 2x with 400 µL of 1x TB. Then resuspend beads in 300 µL 2x NTB.
3. Mix the beads with the 300 µL Hi-C material and incubate at RT for 30 minutes on a rotating wheel to allow biotin binding to streptavidin beads.
4. Wash the beads with 400 µL of 0.5x TB, and incubate at 55 °C for 3 min, rotating at 750 rpm. Wash the beads in 200 µL of 1x restriction buffer.
5. Resuspend the beads in 100 µL of dATP tailing mix (5 µL of 10 mM dATP, 10 µL of 10x restriction buffer, 5 µL of Klenow exo-, and 80 µL of ddH₂O). Incubate at 37 °C for 30 min.
6. Remove the supernatant, wash the beads 2x with 400 µL of 0.5x TB by incubating at 55 °C for 3 min and rotating at 750 rpm.
7. Wash the beads with 400 µL of 1x NTB and then with 100 µL of 1x ligation buffer.
8. Resuspend the beads in 50 µL of 1x ligation buffer, and transfer the suspension to a new tube. Add 4 µL of pre-annealed PE adapters (15 µM stock) and 2 µL of T4 ligase (400 U/µL, i.e., 800 U/tube); incubate at RT for 2 h.
   NOTE: Pre-anneal the adapters by adding equal volumes of both PE 1.0 and PE 2.0 adapters (30 µM stock) and incubating for 10 min at RT (see the Table of Materials).
9. Recapture the beads by removing the supernatant and washing the beads 2x with 400 µL of TB. Wash the beads with 200 µL of 1x NTB, then with 100 µL of 1x restriction buffer, and resuspend the beads in 40 µL of 1x restriction buffer.

12. PCR amplification

1. Set up PCRs of 25 µL volume with 5, 6, 7, and 8 cycles. For each PCR, use:

   Reaction Recipe
   Hi-C beads	2.5 µL
   10 µM PE PCR primer 1 (Table of Materials)	0.75 µL
   10 µM PE PCR primer 2 (Table of Materials)	0.75 µL
   10 mM dNTP	0.6 µL
   5x reaction buffer	5 µL
   DNA polymerase	0.3 µL
   ddH2O	14.65 µL
   Total	25 µL

   Cycles	Temperature	Time
   1	98 °C	30 s
   n cycles	98 °C	10 s
   	65 °C	30 s
   	72 °C	30 s
   1	72 °C	7 min

13. Final PCR amplification

1. Perform final PCR amplification using the same conditions as described above and the number of cycles selected. Split the sample using 5 µL of Hi-C beads as template in 50 µL reactions.
2. Collect all PCR reactions and transfer them to a fresh tube. Use the magnet to remove the streptavidin beads and recover the supernatant (PCR products). Transfer the beads to a fresh tube, wash the beads as indicated in step 11.2.9, and store in 1x restriction buffer at 4 °C as a backup.
3. Purify the PCR products using 0.85x the volume of the SPRI beads according to the manufacturer´s instructions. Elute with 30 µL of TLE buffer.
4. Quantify the Hi-C library using a fluorometric instrument, and confirm the quality of the library by chip-based capillary electrophoresis.
5. As a last quality checkpoint, use 1 µL of the Hi-C library as a template to perform a PCR reaction using 10 cycles using the same conditions described in step 12.1. Divide the PCR product into two microcentrifuge tubes: digest one with Cla I and leave the other one undigested as a control. Run the products in a 1.5-2% agarose gel (see representative results).
6. Proceed to 50 bp or 75 bp paired-end sequencing on a suitable sequencing platform.