Generation of Genome-wide Chromatin Conformation Capture Libraries from Tightly Staged Early Drosophila Embryos

1. Drosophila Embryo Collection

NOTE: An equivalent embryo collection can be performed as shown in a previous publication¹⁵.

1. Transfer young eGFP-PCNA flies (<1 week old) into egg collection cages with yeasted collection plates¹⁶ (1% ethanol, 1% acetic acid, and 4% agar).
2. Move collection cages to an incubator set at 25 °C. Incubation for 1–2 days prior to egg collection improves the egg yield significantly. Change collection plates twice a day.
3. Remove plates containing embryos from the collection chamber in 30–60 min intervals. Smaller intervals result in fewer embryos, but tighter distribution of developmental stages. Collect from multiple cages in parallel so that ideally >200 eggs are laid every 30–60 min.
4. Store the plates at 25 °C until the embryos reach the desired age. For blastoderm stage embryos (nuclear cycle 14), incubate for approximately 2 h.
5. After 2 h of incubation, add tap water from a squirt bottle to the collection plate so that the entire surface is covered with water. Suspend embryos and yeast using a soft brush.
6. Pour resuspended embryos from the collection plate into an embryo collection basket (commercial cell strainers with 100 µm pore size or homemade baskets¹⁷ work well), adding additional tap water from a squirt bottle, if necessary. At this stage, combine embryos from all plates that were collected in parallel. The pooled sample represents a single batch.
7. Wash embryos well by rinsing the basket with tap water from a squirt bottle for 30 s until all yeast residue is washed away.
8. Dechorionate embryos by placing the collection basket into a 2.5% sodium hypochlorite solution in water. Light agitation by swirling facilitates removal of the chorion. Continue until embryos are sufficiently hydrophobic so that they float at the surface of the solution when the basket is lifted out and submerged again, which should take ~1.75–2 min.
   Caution: Sodium hypochlorite is corrosive. Wear appropriate personal protective equipment. Solutions containing <10% sodium hypochlorite can usually be disposed in the sink, make sure to check the regulations of the host institute.
9. Remove the basket from the solution and rinse thoroughly with tap water from a squirt bottle until the bleach smell is no longer noticeable.

2. Embryo Fixation

NOTE: Optimal fixation conditions, primarily the concentration of detergent, formaldehyde and the duration of fixation, need to be empirically determined to fit the stage of the embryos. For stages around the syncytial blastoderm, a final concentration of 0.5% Triton X-100 and 1.8% formaldehyde in the aqueous phase work well. For later stages beyond embryo stage 9, further optimization of these parameters may be necessary. All solutions used during fixation and sorting should contain protease inhibitors.

1. Invert collection basket and place it over a 15 mL conical centrifugation tube. Flush embryos from the basket into the tube using a Pasteur pipette dispensing PBS-T (PBS, 0.5% Triton X-100).
2. Let embryos settle at the bottom and adjust the total volume to 2 mL with PBS-T.
3. Add 6 mL of heptane and 100 µL of 37% formaldehyde in water.
   Caution: Heptane and formaldehyde are toxic when inhaled or after skin contact. Wear appropriate personal protective equipment and work in a fume hood. Waste containing heptane or formaldehyde has to be disposed separately according to the host institute's regulations.
4. After the addition of the formaldehyde, start a 15 min timer and vigorously shake the tube up and down for 1 min by hand. The aqueous and organic phase will combine to form a shampoo-like consistency.
5. Agitate on a rotatory mixer until 10 min after the addition of formaldehyde.
6. Centrifuge at 500 x g for 1 min at room temperature to collect embryos at the bottom of the tube.
7. Aspirate the entire shampoo-like liquid and discard it, taking care not to aspirate any embryos. Small remaining quantities of shampoo-like supernatant do not cause problems.
8. 15 min after the addition of formaldehyde, resuspend the embryos in 5 mL of PBS-T with 125 mM glycine to quench the formaldehyde. Mix vigorously by shaking up and down for 1 min.
9. Centrifuge at 500 x g at room temperature for 1 min and aspirate supernatant.
10. Wash embryos by resuspending them in 5 mL of ice-cold PBS-T. Let embryos settle and aspirate all supernatant.
11. Repeat the wash in step 2.10 two more times.
12. Keep embryos on ice until sorting. Usually, it is a good idea to collect 3–4 batches of fly embryos before proceeding to sorting. However, embryos should be sorted the same day. Extended storage on ice or in the fridge leads to altered embryo morphology.

3. Embryo Sorting

NOTE: Sorting can be done on any fluorescent stereo microscope equipped with a GFP filter at 60–80X magnification.

1. Using a 1,000 µL pipette, transfer a batch of approximately 100 embryos to a small glass vessel suitable for sorting, preferably of a dark color, and place it on ice.
2. Sort embryos by nuclear density and cell cycle status (Figure 1) by pushing desirable embryos into a separate pile using a needle or syringe tip.
   1. Remove all embryos with dispersed, non-nuclear distribution of eGFP-PCNA (Figure 1E). Also, embryos that partially show a non-nuclear GFP signal should be removed.
   2. To aid in the sorting, assemble a line-up of reference embryos at nuclear cycle 12, 13, and 14 in each batch using the pictures in Figure 1 as a guide. Use this line-up to match embryos of an unknown stage with one of the reference embryos in order to determine their stage.
   3. To verify the developmental stage for reference embryos, measure the nuclear density by imaging the embryo and counting the number of nuclei at the surface of the embryo in an area of 2,500 µm² using imaging software that provides distance information.
      NOTE: The expected number of nuclei for an area of 2,500 µm² is 12 to 16 nuclei at nuclear cycle 12, and 20 to 30 nuclei at nuclear cycle 13¹³.
3. Once all embryos at the appropriate stage are separated, take pictures of the embryos for documentation and quality control. If the stereo microscope is not itself equipped with a camera module, any epifluorescence microscope with GFP filters may be used.
4. Pipette up the desired embryos using a 1,000 µL pipette, transfer to a fresh tube, and place on ice.
5. Continue until enough embryos are sorted for the planned experiment. For embryos older than stage 9, generally 20 embryos are sufficient for one in situ Hi-C experiment. At nuclear cycle 12, 80 embryos are a good starting point. In earlier cycles, the number of embryos should approximately be doubled for every cycle.
6. Pool and split embryos into 1.5 mL tubes in such a way that one tube contains enough embryos for a single in situ Hi-C experiment. It is advisable to use tubes with low DNA binding characteristics, since the same tube will be used for the entire protocol and adsorption of DNA can lead to significant losses at low DNA concentrations.
7. Spin tubes briefly at 100 x g at room temperature and remove supernatant. The embryos should be as dry as possible for freezing.
8. Flash freeze embryos by submerging the tubes in liquid nitrogen and store at -80 °C.

4. In Situ Hi-C

1. Lysis
   1. Place tubes with frozen embryos on ice.
   2. Resuspend embryos in 500 µL of ice-cold lysis buffer (10 mM Tris-Cl pH 8.0, 10 mM NaCl, 0.2% IGEPAL CA-630, protease inhibitors; dissolved in water). Then wait 1 min to let embryos settle at the bottom of the tube.
   3. Grind embryos using a metal micro pestle, pre-cooled on ice, that is designed to tightly fit a 1.5 mL microcentrifuge tube.
      1. To avoid agitating the embryos, insert the pestle slowly until it touches the bottom of the tube, push down, and then grind by rotating the pestle twice in both directions.
      2. Lift the pestle very slightly, push to the bottom of the tube again, and repeat grinding.
      3. Repeat 4.1.3.2 10 times, or until the embryos are completely lysed. The solution should be homogenous, and no residual large pieces of embryos should remain.
   4. Incubate the homogenized suspension on ice for 15 min. Spin at 1,000 x g, 4 °C for 5 min, and discard supernatant.
   5. Wash pellet by resuspending in 500 µL ice-cold lysis buffer, pipetting up and down.
   6. Spin again as in 4.1.4, and discard supernatant.
   7. Resuspend washed pellet in 100 µL of 0.5% sodium dodecyl sulfate (SDS), pipetting up and down. Permeabilize nuclei by incubating for 10 min at 65 °C in a heating block. Quench SDS by adding 50 µL of 10% Triton X-100 and 120 µL water. Mix by flicking the tube.
   8. Incubate at 37 °C for 15 min in heat block.
2. Restriction enzyme digestion
   1. Add 25 µL of 10x restriction enzyme buffer and 20 U of 5 U/µL MboI. Mix by flicking the tube.
   2. Digest DNA by incubating for 90 min at 37 °C in heat block under slight agitation (750 rpm).
   3. Add another 20 U of MboI and continue incubation for 90 min.
   4. Heat-inactivate MboI by incubating at 62 °C for 20 min.
3. Overhang fill-in
   NOTE: Filling in the overhang with biotinylated dATP allows selection of specific ligated fragments. Biotin-dATP at ligation junctions is protected from the exonuclease activity of T4 DNA Polymerase (section 4.6), whereas biotin-dATP at unligated blunt ends is efficiently removed. The pulldown using streptavidin-coated beads in section 4.7 therefore specifically enriches for ligated, chimeric DNA fragments.
   1. Add 18 µL of 0.4 mM biotin-14-dATP, 2.25 µL of an unmodified dCTP/dGTP/dTTP mix (3.3 mM each), and 8 µL of 5 U/µL DNA Polymerase I Klenow Fragment.
   2. Mix by flicking the tube and incubate at 37 °C for 90 min in heat block.
4. Ligation
   1. Add 657 µL of water, 120 µL of 10x T4 DNA Ligase Buffer, 100 µL of 10% Triton X-100, 6 µL of 20 mg/mL bovine serum albumin (BSA), and mix by flicking the tube. Finally add 5 µL of 5 U/µL T4 DNA Ligase and mix by flicking the tube.
   2. Rotate tube gently (20 rpm) at room temperature for 2 h.
   3. Add a second installment of 5 µL of 5 U/µL T4 DNA Ligase and continue rotating for 2 more h.
   4. Spin down nuclei at 2,500 x g for 5 min and discard supernatant.
5. DNA extraction
   1. Resuspend pellet in 500 µL of extraction buffer (50 mM Tris-Cl pH 8.0, 50 mM NaCl, 1 mM Ethylenediaminetetraacetic acid (EDTA), 1% SDS; dissolved in water) and add 20 µL of 20 mg/mL proteinase K. Mix by flicking the tube.
   2. Digest protein by incubating at 55 °C for 30 min, shaking at 1,000 rpm.
   3. To de-crosslink, add 130 µL of 5 M NaCl and incubate overnight at 68 °C, shaking at 1,000 rpm.
   4. Pipette sample into a new 2 mL tube, preferentially with low DNA binding characteristics.
   5. Add 0.1x volumes (63 µL) of 3 M sodium acetate pH 5.2 and 2 µL of 15 mg/mL GlycoBlue. Mix well by inverting. Add 1.6x volumes (1,008 µL) of pure absolute ethanol and mix by inverting.
   6. Incubate at -80 °C for 15 min. Centrifuge at 20,000 x g at 4 °C for at least 30 min. The DNA pellet is often very small, almost invisible, and can only be spotted due to the blue color of GlycoBlue.
   7. Remove supernatant very carefully, moving the pipette tip into the tube along the opposite wall from where the DNA pellet is located. Small remaining droplets are often easily removed during this step and the following washes by pushing them out of the tubes using a P10 tip rather than pipetting them out.
   8. Wash pellet by adding 800 µL of 70% ethanol. Mix by inverting and centrifuge at 20,000 x g at room temperature for 5 min. Repeat this wash at least once.
   9. Remove all traces of ethanol and leave the tube standing with the lid open for up to 5 min to air-dry. Once no liquid is remaining, add 50 µL of 10 mM Tris-Cl pH 8.0. Repeatedly pipette the solution over the area on the wall of the tube where the pellet was located to solubilize the DNA.
   10. Add 1 µL of 20 mg/mL RNase A, mix by flicking the tube, and incubate at 37 °C for 15 min to digest RNA. The sample can now be stored in the fridge overnight or frozen at -20 °C indefinitely.
   11. Check the concentration of DNA using a fluorescent dye based assay according to the manufacturer's instructions. The total amount of DNA in the sample should be at least 10 ng, otherwise too little material is available for amplification and library complexity will likely be low. When this happens, the amount of starting material was probably not sufficient, or material was lost along the way, perhaps during lysis and precipitation.
6. Biotin removal and DNA shearing
   1. Add together 12 µL of 10x T4 DNA Polymerase buffer, 3 µL of 1 mM dATP, 3 µL of 1 mM dGTP, and 46 µL of water. Mix by flicking the tube. Add 5 µL of 3 U/mL T4 DNA Polymerase, mix by flicking the tube and incubate at 20 °C for 30 min.
   2. Add 3 µL of 0.5 M EDTA to stop the reaction, and use water to bring the sample to a volume of approximately 120 µL.
   3. Shear the DNA to a size of 200–400 bp using a sonication device according to the manufacturer's instructions. Using the sonicator mentioned in the Table of Materials, the following program is appropriate: 2 cycles each of 50 s, 10% duty, intensity 5, 200 cycles/burst.
7. Biotin pulldown
   1. Pipette 30 µL of 10 mg/mL streptavidin coated magnetic beads into a new tube, separate them on a magnetic stand, and discard supernatant.
   2. Resuspend beads in 1x B&W buffer (5 mM Tris-Cl pH 7.4, 0.5 mM EDTA, 1 M NaCl; dissolved in water) + 0.1% Triton X-100 and mix by vortexing. Place tube on a magnetic stand and wait for 1–5 min until the beads are separated, depending on the make and model.
   3. Aspirate and discard supernatant while sliding the pipette tip along the wall opposite of where the beads are located. Resuspend beads in 120 µL of 2x B&W buffer (10 mM Tris-Cl pH 7.4, 1 mM EDTA, and 2 M NaCl). Mix by vortexing.
   4. Transfer sheared DNA to a new low DNA binding tube, and mix with 120 µL of the bead suspension in 2X B&W buffer by vortexing. Rotate beads with the DNA sample at 20 rpm for 15 min.
   5. Separate beads on a magnetic stand and discard supernatant.
   6. Resuspend beads in 600 µL of 1x B&W + 0.1% Triton X-100, and incubate at 55 °C for 2 min, shaking at 1,000 rpm. After separation, discard supernatant. Repeat this wash once.
   7. Wash beads once with 600 µL of 10 mM Tris-Cl pH 8.0, and discard supernatant after separation.
   8. Resuspend beads in 50 µL of 10 mM Tris-Cl pH 8.0.

5. Sequencing Library Preparation

NOTE: All library steps are done using components from a commercial DNA library preparation kit (see Table of Materials). However, alternative kits or other reagents may be substituted. Precipitation tends to form in the library preparation agents during freezer storage. It is therefore important to make sure that all precipitation is dissolved before using the reagents.

1. End repair
   1. Transfer the bead suspension in 50 µL of 10 mM Tris-Cl pH 8.0 into a new PCR tube.
   2. Add 3 µL of End Prep Enzyme Mix and 7 µL of End Prep Reaction Buffer. Mix by pipetting up and down.
   3. Transfer tube to a thermal cycler and run the following program: 20 °C for 30 min, 65 °C for 30 min, and hold at 4 °C.
2. Adapter ligation
   1. Add 30 µL of Ligation Master Mix, 2.5 µL of 1.5 µM Sequencing Adaptor (dilute to 1.5 µM from stock), and 1 µL of Ligation Enhancer to the bead suspension. Mix by pipetting up and down.
   2. Incubate at 20 °C for 15 min in a thermal cycler.
   3. Add 3 µL of USER enzyme. Mix by pipetting up and down.
   4. Incubate at 37 °C for 15 min in a thermal cycler.
   5. Separate beads on a magnetic stand and remove supernatant.
   6. To wash beads, resuspend beads in 100 µL of 1x B&W buffer + 0.1% Triton X-100. Mix by vortexing, and transfer to a new microcentrifuge tube. Separate beads on a magnetic stand and remove supernatant.
   7. Repeat this wash once using 600 µL of the same buffer.
   8. Resuspend beads in 600 µL of 10 mM Tris-Cl pH 8.0, mix by vortexing, and transfer beads to a new tube.
   9. Separate beads on a magnetic stand, discard supernatant, and resuspend beads in 50 µL of 10 mM Tris-Cl pH 8.0.
3. PCR amplification
   1. Prepare two PCR tubes and in each, mix 25 µL of Polymerase Master Mix, 1.5 µL of 10 µM Forward (unindexed) PCR primer, and 1.5 µL of 10 µM Reverse (indexed) PCR primer.
      NOTE: Forward (unindexed) PCR primer:
      5'-AATGATACGGCGACCACCGAGATCTACACTCTTTCCCTACACGACGCTCTTCCGATC*T-3´.
      Reverse (indexed) PCR primer:
      5'-CAAGCAGAAGACGGCATACGAGATNNNNNNGTGACTGGAGTTCAGACGTGTGCTCTTCCGATC*T-3´. * indicates phosphorothioate bonds and Ns in the indexed PCR primer.
   2. In each tube, add 22 µL of bead suspension and mix by pipetting up and down.
   3. Run PCR using the following program: 98 °C for 1 min, (98 °C for 15 s, 65 °C for 75 s, ramping 1.5 °C/s) repeated 9-12 times, 65 °C for 5 min, and hold at 4 °C.
      NOTE: The number of amplification cycles has to be determined empirically. However, we found that libraries that required more than 12 cycles were generally of low complexity and did not result in high quality Hi-C maps. On the other hand, libraries that required less than 12 cycles were not negatively affected by amplifying for a full 12 cycles. Therefore, it is possible to default to 12 cycles of amplification.
   4. Pool the two PCR reactions in a single microcentrifuge tube, separate beads on a magnetic stand, and transfer the supernatant containing the library to a new tube.
4. Size selection
   1. Bring Ampure XP bead suspension to room temperature and mix well by shaking.
   2. Bring volume of the pooled PCR reaction to exactly 200 µL with water. During PCR and the magnetic separation, some of the original volume is usually lost. Verify volume by setting the pipette to 200 µL and aspirate the entire volume of the reaction. If air is aspirated, more water needs to be added. If the volume exceeds 200 µL, adjust the volume of beads added in steps 5.4.3 and 5.4.6 proportionally.
      NOTE: The volumes in parentheses are valid if the total volume of the pooled PCR reactions is exactly 200 µL.
   3. Add 0.55x volumes (110 µL) of Ampure XP bead suspension and mix by pipetting up and down at least 10 times.
   4. Incubate at room temperature for 5 min, separate beads on a magnetic stand for 5 min.
   5. Move supernatant to a new tube. Discard the tube containing the beads. The beads have bound DNA >700 bp, which is too large to be sequenced.
   6. To the supernatant, add 0.2x volumes (40 µL, resulting in a total of 0.75x Ampure buffer in the sample) of Ampure XP bead suspension and mix by pipetting up and down 10 times.
   7. Incubate at room temperature for 5 min, separate beads on a magnetic stand for 5 min.
   8. Discard supernatant which contains DNA <200 bp, which includes free primers, primer dimers, and fragments too small to be sequenced.
   9. Leave the tube on the magnetic stand. To wash beads, add 700 µL of 80% ethanol, taking care not to disturb the bead pellet, and incubate for 30 s.
   10. Discard supernatant, then take the tube off the magnetic stand and resuspend beads in 100 µL of 10 mM Tris-Cl pH 8.0. Mix by pipetting up and down 10 times, and incubate at room temperature for 1 min.
   11. Add 0.8x volumes (80 µL) of Ampure XP bead suspension. Mix by pipetting up and down 10 times and incubate at room temperature for 5 min. This second round of lower bound size selection ensures that the final library is completely free of primers and primer dimers.
   12. Separate beads on a magnetic stand for 5 min and discard supernatant.
   13. Wash the bead pellet twice with 700 µL of 80% ethanol for 30 s each, while leaving the tube on the magnetic stand, as above.
   14. With the tube still on the magnetic stand, remove all traces of ethanol. It helps to push droplets of ethanol out of the tube using a P10 pipette. Let residual ethanol evaporate for a maximum of 5 min.
   15. Take tube off the magnetic stand and resuspend beads in 50 µL of 10 mM Tris-Cl pH 8.0. Mix by pipetting up and down 10 times.
   16. Incubate at room temperature for 5 min, then separate beads on a magnetic stand.
   17. Transfer supernatant to a fresh tube. This is the final Hi-C library, ready to be quantified and sequenced on next generation sequencing machines, according to the manufacturer's instructions.