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

Investigating the three-dimensional architecture of chromatin offers invaluable insight into the mechanisms of gene regulation. Here, we describe a protocol for performing the chromatin conformation capture technique in situ Hi-C on staged Drosophila melanogaster embryo populations. The result is a sequencing library that allows the mapping of all chromatin interactions that occur in the nucleus in a single experiment. Embryo sorting is done manually using a fluorescent stereo microscope and a transgenic fly line containing a nuclear marker. Using this technique, embryo populations from each nuclear division cycle, and with defined cell cycle status, can be obtained with very high purity. The protocol may also be adapted to sort older embryos beyond gastrulation. Sorted embryos are used as inputs for in situ Hi-C. All experiments, including sequencing library preparation, can be completed in five days. The protocol has low input requirements and works reliably using 20 blastoderm stage embryos as input material. The end result is a sequencing library for next generation sequencing. After sequencing, the data can be processed into genome-wide chromatin interaction maps that can be analyzed using a wide range of available tools to gain information about topologically associating domain (TAD) structure, chromatin loops, and chromatin compartments during Drosophila development.


Introduction
Chromatin conformation capture (3C) has emerged as an exceptionally useful method to study the topology of chromatin in the nucleus 1 . The 3C variant Hi-C allows measuring the contact frequencies of all chromatin interactions that occur in the nucleus in a single experiment 2 . Application of Hi-C has played an important role in the discovery and characterization of many fundamental principles of chromatin organization, such as TADs, compartments, and loops 3,4,5 .
Studies of chromatin architecture in the context of developmental transitions and cell differentiation are increasingly used to unravel the mechanisms of gene regulation during these processes 6,7,8,9 . One of the model organisms of great interest is Drosophila melanogaster, whose development and genome are well characterized. However, few studies that investigate chromatin architecture in Drosophila outside of in vitro tissue culture settings have been conducted 10,11 . In embryos 16-18 h post fertilization, TADs and compartments reminiscent of similar structures in mammals were identified 10 , which raises the question of which role they are playing in gene regulation during Drosophila embryo development. Especially in the early stages of development, prior to gastrulation, such studies are technically challenging. Before gastrulation, Drosophila embryos undergo 13 synchronous nuclear divisions that proceed at an extremely rapid pace of 8-60 min per cycle 12,13 . In addition to this, the lack of visual features to distinguish the different stages make it difficult to obtain tightly staged embryo material in sufficient quantities.
In order to develop a protocol that allows studying chromatin architecture in early Drosophila development at nuclear cycle resolution, we combined two existing techniques: in situ Hi-C, which allows the generation of high resolution whole genome contact maps 5 , and embryo staging using a transgenic Drosophila line expressing a eGFP-PCNA transgene 13,14 . This transgene localizes to the nucleus during interphase and disperses throughout the syncytial blastoderm during mitosis. Using this property, it is possible to easily distinguish different stages by their nuclear density and mitotic embryos by the dispersion of the GFP signal.
Together, these techniques enable studying the three-dimensional structure of chromatin in high resolution from as few as 20 Drosophila embryos. This protocol includes the instructions for harvesting and sorting Drosophila embryos to obtain populations of embryos from a single nuclear division cycle. It further describes how the obtained embryos are used to perform in situ Hi-C. The end result is a nucleotide library suitable for sequencing on next generation sequencing machines. The resulting sequencing reads can then be processed into detailed chromatin interaction maps covering the entire Drosophila genome.
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 baskets 17 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.

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.

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.
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. 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.

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.

Representative Results
Sorted embryo populations at nuclear cycle 12, 13, and 14 (corresponding to 1:30, 1:45, and 2:10 hours post fertilization, respectively 12 ) and 3-4 h post fertilization (hpf) were obtained according to the procedures described in the protocol. By taking pictures of the eGFP-PCNA signal of each sorted embryo batch, it is possible to document the precise stage and cell cycle status of every single embryo that is used in downstream experiments. Example pictures of embryos from sorted populations are shown in Figure 1B-E. The output of the in situ Hi-C protocol is a nucleotide library ready to be sequenced on next generation sequencing machines. For this purpose, a final library concentration of at least 2-4 nM is usually required. Using the recommended amounts of input material, this concentration is reliably achieved ( Table 1).
The expected size distribution of DNA fragments after size selection is between 300-600 bp, with a maximum at around 500 bp (Figure 2A), depending on the exact shearing and size selection parameters. For sequencing, we recommend paired-end reads of at least 75 bp length to minimize the number of unmappable restriction fragments in the genome. High-resolution maps with 1-2 kb bin size can be obtained from 400 million reads. We recommend sequencing multiple biological replicates at a lower depth of ~150 million reads each, instead of sequencing a single replicate at very high depth. This allows assessment of the biological variation and leads to a lower number of discarded reads due to PCR duplication. For visual representation, the replicates can be combined. Before committing to sequencing a sample at high depth, we recommend running samples using shallow sequencing (a few million reads per sample) to determine basic library quality parameters as in Figure 2B. Analysis of Hi-C data requires significant computational resources and bioinformatics expertise. As a rough overview, the paired reads are mapped independently to the reference genome, the resulting alignments are filtered for quality and orientation, then a matrix of contacts at a given bin resolution or fragment level can be generated from the filtered alignments. The contact matrix is the basis for all further downstream analysis exploring TADs, loops, and compartments. For the initial analysis of the sequencing reads, several bioinformatics pipelines are available that enable processing of raw reads into contact matrices without much specialized bioinformatics knowledge 18,19,20,21,22,23 . How further analysis is carried out depends largely on the exact biological question under study and might require significant experience in programming and scripting in R or Python. However, several tools and algorithms to call TADs are available 5,24,25,26,27,28 , as well as software to analyze and explore Hi-C data in the web browser and as stand-alone desktop applications 29,30,31,32 .
Once processed, the quality of the library can be determined using different metrics ( Figure 2B). First, the rate of PCR duplicates, which is the number of sequenced read pairs arising from the same original molecule, should be as low as possible to limit the amount of wasted sequence reads. However, even libraries with >40% PCR duplication can be processed into high-quality contact maps if the duplicates are filtered. Second, the rate of filtered reads due to their orientation, as described in 4 , should consistently be lower than 10% of aligned read pairs.
During pre-gastrular development of Drosophila between nuclear cycle 12 and 14, the nuclear architecture is drastically remodeled 33 (Figure   3). At nuclear cycle 12, few TADs are detected, and the overall distribution of contacts is very smooth without many discernable features. This is dramatically changed at nuclear cycle 13 and 14, when TADs are increasingly prominent and unspecific long-range contacts are depleted.   Table 1: List of representative sequencing library statistics. For each library in the list, the number of embryos that were used for its generation, the amount of total DNA before biotin pulldown and shearing measured by Qubit, the number of PCR cycles used for amplification, and the final concentration of the sequencing library after purification and size selection are indicated.

Discussion
The protocol presented here is very effective at generating high-quality maps of the chromatin architecture in early Drosophila embryos.
Compared to an earlier protocol 34 , the approach described here uses an up-to-date in situ Hi-C procedure 5 , resulting in quicker processing, higher resolution, and less reagent usage. The overall procedure including the in situ Hi-C protocol is expected to work on a wide range of stages and experimental systems besides Drosophila. Since the protocol has a low input requirement, it could also be used on isolated cell populations. In Drosophila, when using the protocol for embryos outside the range described here, some parameters, in particular the fixation of the material, might need to be adjusted. Since older embryos develop a highly impermeable cuticle, raising the concentration of formaldehyde and prolonging fixation may be appropriate. For collection of embryos at stages other than nuclear cycle 14, the incubation times of embryos at 25 °C in step 1.4 need to be adjusted as follows: nuclear cycle 12, 70 min; nuclear cycle 13, 90 min; 3-4 hpf, 3:30 h.
During the 13 cleavage divisions (stage 1-4), the nuclei density roughly doubles with each division. The nuclei can easily be identified by their bright GFP fluorescence. During mitosis, eGFP-PCNA is not located in the nucleus, and its signal is dispersed throughout the embryo. This feature makes identifying embryos that are undergoing a synchronous cleavage division possible. For studying chromatin conformation, these mitotic embryos are usually not desirable, since the mitotic organization of chromatin is drastically different than the interphase organization 35 .
It is possible to adapt the protocol to specifically select embryos undergoing a synchronous mitotic division. In this case, only embryos with dispersed, non-nuclear distribution of eGFP-PCNA should be kept, and all other embryos should be discarded. Since the nuclear density cannot be determined, alternative methods to stage embryos by their morphology viewed in transmitted light microscopy must be employed. Presence of pole cells and nuclei at the embryo periphery indicate that the embryo has completed at least nuclear cycle 9, whereas visible cellularization at the periphery indicates nuclear cycle 14 12 .
Hi-C experiments can be successfully performed using a wide selection of restriction enzymes 5 . Current approaches typically use enzymes that recognize either a 4-base sequence, such as MboI, or a 6-base recognition site, such as HindIII. The advantage of 4-base cutters over 6-base cutters is that they offer higher potential resolution, given enough sequencing depth, and a more even coverage of restriction sites across the genome. There is no clear advantage in choosing one 4-base cutter over another 5,23,36,37 . The two most commonly used enzymes, MboI and DpnII, both recognize the same GATC recognition site. DpnII is less sensitive to CpG methylation, which is of no concern in Drosophila. The protocol presented here can also be successfully completed using DpnII as a restriction enzyme. In section 4.2. restriction enzyme and buffer have to be adjusted for DpnII compatibility, according to the manufacturer's recommendations.
If the fragment size of the sequencing library deviates significantly from the range shown in Figure 2A, cluster formation during sequencing may be less efficient or fail completely. In this case, the size distribution after shearing should be checked and shearing parameters adjusted accordingly. Peaks in the distribution of DNA fragments of very small (<100 bp) or very large (>1,000 bp) sizes indicates problems with size selection, such as carry over of beads or supernatant that are supposed to be discarded. Often these libraries with small peaks at these undesirable sizes, such as the one pictured, are still sequenced successfully with only a minor decrease in clustering efficiency.
High rates of PCR duplication should be avoided because this drastically reduces the number of usable sequence reads. The rate of PCR duplicates is directly related to the amount of input material. Using more input therefore usually alleviates problems with PCR duplication.