Hi-C is a high-throughput genomic and epigenomic technique to capture chromatin conformation (3C). In general, Hi-C is considered as a derivative of a series of chromosome conformation capture technologies, including but not limited to 3C (chromosome conformation capture), 4C (chromosome conformation capture-on-chip/circular chromosome conformation capture), and 5C (chromosome conformation capture carbon copy). Hi-C comprehensively detects genome-wide chromatin interactions in the cell nucleus by combining 3C and next-generation sequencing (NGS) approaches and has been considered as a qualitative leap in C-technology (chromosome conformation capture-based technologies) development and the beginning of 3D genomics.
Similar to the classic 3C technique, Hi-C measures the frequency (as an average over a cell population) at which two DNA fragments physically associate in 3D space, linking chromosomal structure directly to the genomic sequence. The general procedure of Hi-C involves first crosslinking chromatin material using formaldehyde. Then, the chromatin is solubilized and fragmented, and interacting loci are re-ligated together to create a genomic library of chimeric DNA molecules. The relative abundance of these chimeras, or ligation products, is correlated to the probability that the respective chromatin fragments interact in 3D space across the cell population. While 3C focuses on the analysis of a set of predetermined genomic loci to offer "one-versus-some" investigations of the conformation of the chromosome regions of interest, Hi-C enables "all-versus-all" interaction profiling by labeling all fragmented chromatin with a biotinylated nucleotide before ligation. As a result, biotin-marked ligation junctions can be purified more efficiently by streptavidin-coated magnetic beads, and chromatin interaction data can be obtained by direct sequencing of the Hi-C library.
Analyses of Hi-C data not only reveal the overall genomic structure of chromosomes, but also offer insights into the biophysical properties of chromatin as well as more specific, long-range contacts between distant genomic elements (e.g. between genes and regulatory elements), including how these change over time in response to stimuli. In recent years, Hi-C has found its application in a wide variety of biological fields, including cell growth and division, transcription regulation, fate determination, development, autoimmune disease, and genome evolution. By combining Hi-C data with other datasets such as genome-wide maps of chromatin modifications and gene expression profiles, the functional roles of chromatin conformation in genome regulation and stability can also be delineated.
History At its inception, Hi-C was a low-resolution, high-noise technology that was only capable of describing chromatin interaction regions within a bin size of 1 million base pairs (Mb). The Hi-C library also required several days to construct, and the datasets themselves were low in both output and reproducibility. Nevertheless, Hi-C data offered new insights for chromatin conformation as well as nuclear and genomic architectures, and these prospects motivated scientists to put efforts to modify the technique over the past decade.
Between 2012 and 2015, several modifications to the Hi-C protocol have taken place, with 4-cutter digestion or adapted deeper sequencing depth to obtain higher resolution. The use of restriction endonucleases that cut more frequently, or DNaseI and Micrococcal nucleases also significantly increased the resolution of the method. More recently (2017), Belaghzal et al. described a Hi-C 2.0 protocol that was able to achieve kilobase (kb) resolution. The key adaptation to the base protocol was the removal of the SDS solubilization step after digestion to preserve nuclear structure and prevent random ligation between fragmented chromatin by ligation within the intact nuclei, which formed the basis of in situ Hi-C. In 2021, Hi-C 3.0 was described by Lafontaine et al., with higher resolution achieved by enhancing crosslinking with formaldehyde followed by disuccinimidyl glutarate (DSG). While formaldehyde captures the amino and imino groups of both proteins and DNA, the NHS-esters in DSG react with primary amines on proteins and can capture amine-amine interactions. These updates to the base protocol allowed the scientists to look at more detailed conformational structures such as chromosomal compartment and topologically associating domains (TADs), as well as high-resolution conformational features such as DNA loops.
To date, a variety of derivatives of Hi-C have already emerged, including in situ Hi-C, low Hi-C, SAFE Hi-C, and Micro-C, with distinctive features related to different aspects of standard Hi-C, but the basic principle has remained the same.
Traditional Hi-C The outline of the classical Hi-C workflow is as follows: cells are cross-linked with formaldehyde; chromatin is digested with a restriction enzyme that generates a 5' overhang; the 5' overhang is filled with biotinylated bases and the resulting blunt-ended DNA is ligated. The ligation products, with biotin at the junction, are selected for using streptavidin and further processed to prepare a library ready for subsequent sequencing efforts.
The pairwise interactions that Hi-C can capture across the genome are immense and so it is important to analyze an appropriately large sample size, in order to capture unique interactions that may only be observed in a minority of the general population. To obtain a high complexity library of ligation products that will ensure high resolution and depth of data, a sample of 20–25 million cells is required as input for Hi-C. Primary human samples, which may be available only in fewer cell numbers, could be used for standard Hi-C library preparation with as low as 1–5 million cells. However, using such a low input of cells may be associated with low library complexity which results in a high percentage of duplicate reads during library preparation.
Standard Hi-C gives data on pairwise interactions at the resolution of 1 to 10 Mb, requires high sequencing depth and the protocol takes around 7 days to complete.
Formaldehyde cross-linking
Cell and nuclear membranes are highly permeable to formaldehyde. Formaldehyde cross-linking is frequently employed for the detection and quantification of DNA-protein and protein-protein interactions. Of interest in the context of Hi-C, and all 3C-based methods, is the ability of formaldehyde to capture cis chromosomal interactions between distal segments of chromatin. It does so by forming covalent links between spatially adjacent chromatin segments. Formaldehyde can react with macromolecules in two steps: first it reacts with a nucleophilic group on a DNA base for example, and forms a methylol adduct, which is then converted to a Schiff base. In the second step, the Schiff base, which can decompose rapidly, forms a methylene bridge with another functional group on another molecule. It can also make this methylene bridge with a small molecule in solution such as glycine, which is used in excess to quench formaldehyde in Hi-C. Quenchers can typically exert an effect on formaldehyde from outside the cell. A key feature of this two-step formaldehyde crosslinking reaction is that all the reactions are reversible, which is vital for chromatin capture.
Crosslinking is a pivotal step of the chromatin capture workflow as the functional readout of the technique is the frequency at which two genomic regions are crosslinked to each other. Thus, the standardization of this step is important and for that, one must consider potential sources of variation. Presence of serum, which contains a high concentration of protein, in culture media can decrease the effective concentration of formaldehyde available for chromatin crosslinking, by sequestering it in the culture media. Therefore, in cases where serum is used in culture, it should be removed for the crosslinking step. The nature of cells, i.e., whether they are suspension or adherent, is also a pertinent consideration for the crosslinking step. Adherent cells bind to surfaces with the help of molecular mechanisms of cytoskeletons. It has been shown that there is a link between cytoskeleton-maintained nuclear and cellular morphology which, if altered, may negatively impact global nuclear organization. Adherent cells therefore, should be crosslinked while still attached to their culture surface.
Lysis, restriction digest and biotinylation Cells are lysed on ice with cold hypotonic buffer containing sodium chloride, Tris-HCl at pH 8.0, and non-ionic detergent IGEPAL CA-630, supplemented with protease inhibitors. The protease inhibitors and incubation on ice help preserve the integrity of crosslinked chromatin complexes from endogenous proteases. The lysis step helps to release the nucleic material from the cells.
Following cell lysis, chromatin is solubilized with dilute SDS in order to remove proteins that have not been crosslinked and to open chromatin and make it more accessible for subsequent restriction endonuclease-mediated digestion. If the incubation with SDS exceeds the recommended 10 minutes, the formaldehyde crosslinks can be reversed and so the incubation with SDS must be immediately followed by an incubation on ice. A non-ionic detergent called Triton X-100 is used to quench SDS in order to prevent enzyme denaturation in the next step.
Any restriction enzyme that generates a 5' overhang, such as HindIII can be used to digest the now accessible chromatin overnight. This 5' overhang provides the template required by the Klenow fragment of DNA Polymerase I to add biotinylated CTP or ATP to the digested ends of chromatin. This step allows for the selection of Hi-C ligation products for library preparation.
Proximity ligation A dilution ligation is performed on DNA fragments that are still crosslinked to one another in order to favor the intramolecular ligation of fragments within the same chromatin complex instead of ligation events between fragments across different complexes. Since this ligation step occurs between blunt-ended DNA fragments (since the sticky ends have been filled in with biotin-labeled bases), the reaction is allowed to go on for up to 4 hours to make up for its inherent inefficiency. As a result of proximity ligation, the terminal HindIII sites are lost and an NheI site is generated.
Biotin removal, DNA shearing, size selection and end repair The biotin-labeled ligation products can be purified using phenol-chloroform DNA extraction. To remove any fragments with biotin-labeled ends that have not been ligated, T4 DNA Polymerase with 3' to 5' exonuclease activity is used to remove nucleotides from the ends of such fragments. This step ensures that none of these unligated fragments are selected for library preparation. The reaction is stopped with EDTA and the DNA is purified once again using phenol-chloroform DNA extraction.
The ideal size of DNA fragments for the sequencing library depends on the sequencing platform that will be used. DNA can first be sheared to fragments around 300–500 bp long using sonication. Fragments of this size are suitable for high-throughput sequencing. Following sonication, fragments can be size selected using AMPure XP beads from Beckman Coulter to obtain ligation products with a size distribution between 150 and 300 bp. This is the optimal fragment size window for HiSeq cluster formation.
DNA shearing causes asymmetric DNA breaks and must be repaired before biotin pulldown and sequencing adaptor ligation. This is achieved by using a combination of enzymes that fill in 5' overhangs, and add 5' phosphate groups and adenylate to the 3' ends of fragments to allow for ligation of sequencing adaptors.
Biotin pull-down