Chromatin immunoprecipitation and library preparation: a powerful tool to unravel the epigenome

Chromatin immunoprecipitation (IP) followed by sequencing (ChIP-seq) is a powerful tool used in genome-wide DNA binding sequences for specific proteins, such as transcription factors and histone marks. ChIP-seq helps decipher cell identity by revealing interactions between transcription factors and regulatory DNA elements, as well as the regulation of gene expression during various stages of development, regeneration and disease [1–3]. Chromatin structures are influenced by intra- and extracellular signal transductions that control several processes like gene transcription, DNA replication, histone modifications, DNA repair and other transcription coregulators [2,4,5]. Understanding genome-wide DNA protein interactions and histone modifications is critical for deciphering the chromatin regulatory landscape. ChIP-seq provides a powerful tool to understand and determine the processes regulated by these specific interactions [2,6–8].

ChIP-seq method overview

The ChIP-seq protocol requires the optimization of several critical steps to produce high-quality data. Briefly, it involves fixing the desired cells or tissues with formaldehyde to crosslink the protein–DNA, followed by cell lysis and fragmentation of chromatin. Subsequently, fragmented chromatin is immunoprecipitated, using an antibody specific to the protein of interest, followed by purification of the antibody-bound target DNA. Finally, the protein–DNA complex is reverse-crosslinked to release enriched DNA for downstream assays. The DNA can be analyzed using either next-generation sequencing (NGS) for genome-wide analysis (ChIP-seq) or standard PCR/quantitative real-time PCR (qPCR) for assessment of targeted binding sites. This article focuses on best practices for ChIP and NGS library preparation for ChIP-seq.

Chromatin immunoprecipitation

Several factors can influence ChIP and, due to the diversity of target proteins, it is difficult to define all parameters. Some key points are discussed next.

Cell number

The starting material, whether it be the cell number or the amount of tissue required, is crucial for ChIP pull-down, lysis and sonication, as the signal-to-noise ratio is directly influenced by the cell number. Increasing the number of cells used results in a higher signal-to-noise ratio. Therefore, it is imperative to standardize the required number of cells needed for specific ChIP-seq experiments in order to minimize the signal-to-noise ratio. For instance, in ChIP-seq experiments, typically 1–10 million cells are sufficient to yield 10–100 ng of ChIP-seq DNA. Moreover, these cell numbers can be optimized by employing different cell numbers for pull-down followed by qPCR to assess the signal-to-noise ratio. If the protein of interest, such as H3K27me3 or RNA polII, is abundantly present, 1 million cells are generally adequate. However, if studying a less abundant transcription factor, at least 10 million cells are required, depending on the antibody used. Different cell numbers are employed for various experimental end points, such as 10⁴–10⁵ cells for qPCR, 10⁶ cells for ChIP-seq and 10⁸ cells for ChIP-chip, respectively. The minimum cell number is also critical for cell lysis and can be achieved by diluting cells in a lysis buffer. When performing sonication in a microcentrifuge tube, 1 million cells should be used in 100–150 ul of sonication buffer. If using a 15-ml falcon tube, 10 million or more cells should be suspended in 1 ml of sonication buffer. However, these numbers can be adjusted based on the experimental setup, but standardization is always necessary.

The cell number is also crucial for pull-down using specific antibodies, as certain antibodies yield excellent results with 1 million cells, while others may require 3–4 million cells, depending on the specific experimental conditions and antibodies used. Standardization can be accomplished prior to performing the actual experiments by employing positive and negative controls.

Crosslinking

Crosslinking is a process in which proteins form crosslinks with DNA. This is achieved by using formaldehyde fixation of the cells. The efficiencies of chromatin shearing and IP depend on the formaldehyde concentration, duration of formaldehyde fixation and the temperature at which crosslinking is performed [9]. Standardizing the formaldehyde concentration, fixation duration and temperature is crucial before initiating experiments because these parameters vary for different cell or tissue types. Generally, a formaldehyde concentration of 1–2%, incubation at 37°C and duration of 8–10 min are sufficient. Immediately after fixation, quenching the reaction with 125 mM glycine or 4.5M (final concentration 1.5M) tris pH 8.0 should be essential. Quenching is another important factor in this step. Over-crosslinking or under-crosslinking can impact fragmentation and the ability of proteins to crosslink to DNA [9]. Insufficient crosslinking can lead to the loss of protein–DNA interactions during fragmentation, resulting in successful IP. On the other hand, excessive crosslinking can affect chromatin shearing and cell lysis.

Cell lysis

The primary objective of this step is to isolate nuclei while removing the plasma membrane and cytoplasm. Cell or tissue lysis should be performed carefully on ice to minimize protein degradation, including epitopes. Additionally, the lysis buffer should contain protease inhibitors and, if necessary, phosphatase inhibitors. Several methods are available for cell lysis to enrich the nuclei. Based on our experience, lysis in a microcentrifuge tube is recommended. This can be achieved by gently pipetting the suspension up and down followed by placing the microcentrifuge tube on ice for 4–5 min. The process of pipetting the suspension and incubating on ice can be repeated 5–10 times to improve the lysis and enhance the nuclear enrichment. However, when working with skeletal muscle cells or tissues, it may be necessary to perform additional cycles for optimal lysis.

Chromatin shearing

After cell lysis, chromatin is fragmented within the range of 150–400 base pairs (bp), depending on the specific experiments, such as ChIP-seq (150–250 bp) or ChIP-on-chip (150–400 bp) and so on. These fragment sizes are ideal for IP and downstream processes like NGS library preparation and sequencing. Prior to sonication, the lysed cells or tissues are centrifuged to remove cell debris. Cell lysate should be appropriately diluted with shearing buffer. In the shearing buffer, 1% SDS is required for effective shearing. While SDS aids in shearing, it can also damage epitopes and denature the proteins, hindering the recognition of the targeted antibody. Therefore, it is crucial to standardize the SDS concentration in the shearing buffer. Alternatively, other detergents such as NP-40 or deoxycholate can be used to balance the reduced SDS percentage and facilitate shearing. The recommended cell density is 1 million per 130–150 μl of shearing buffer, depending on the sonication machine and tube used. For instance, if a Diagenode Bioruptor system is utilized, a maximum volume of 130–150 μl in a microcentrifuge tube is appropriate. Sonication time should be optimized for different types of cells typically ranging 30–40 s ON and 60–80 s OFF in cycles. Sonication conditions for chromatin shearing may vary depending on cell type, with skeletal muscle cells often requiring longer sonication times compared with kidney cells. Enzymatic digestion such as micrococcal nuclease (MNase) treatment, can also be employed for chromatin shearing, particularly in nucleosome occupancy experiments. Following shearing, the size of the chromatin fragment should be assessed after reversing the crosslink. Various methods, such as phenol-chloroform extraction and isopropanol precipitation, can be used to isolate DNA for testing the size of the sheared chromatin. One common approach is running the reverse-crosslinked and purified sheared DNA on a 2% agarose gel or using a Bioanalyzer/TapeStation. These methods allow for the evaluation of chromatin fragmentation size before proceeding with IP. Although we typically used 2% agarose gel electrophoresis due to cost considerations, the Bioanalyzer provides more accurate and precise results regarding fragment size.

Immunoprecipitation

In this step, sheared chromatin is incubated with an antibody specific to the target protein. The availability of validated antibodies is crucial for successful IP. Various factors can influence the effectiveness of IP, including the amount of antibody used, choice of beads (magnetic and agarose), size and quantity of sheared chromatin and duration of antibody incubation. The binding of the antibody to the beads is a critical step, and the ratio of beads (protein A or protein G) to antibody concentration should be carefully titrated.

Each IP should be accompanied by a mock control using IgG and an input control sample to validate the IP results. Additionally, the antibody binding to the beads requires standardization, typically with an incubation time of 2–3 h (although this can be adjusted accordingly). All binding and subsequent steps should be carried out at 4°C or according to specific recommendations by the manufacturer if using a kit. Following antibody binding to the beads, chromatin is added to the tubes containing the antibody-bound beads and incubated overnight on a rotator (20–40 r.p.m.) at 4°C in a cold room to prevent bead settling, and protein and epitope degradation.

Washes, reverse-crosslinking & recovery of ChIP DNA

After completing the IP, wash the complex with washing buffer to remove nonspecific binding to the complex (beads, antibody and chromatin). This step removes the unwanted materials/molecules that can interfere with downstream processes. Depending on the experimental requirements, different buffers such as high salt, low salt, lithium chloride and stringent TE wash buffers may be used to minimize nonspecific binding. It is crucial to follow the same washing steps, buffers, temperature, incubation time and rotator speed for replicates to ensure reproducible results.

Reverse crosslinking is then performed to dissociate the complex (beads, antibody and protein of interest bound to DNA) and release the ChIP DNA. For reverse crosslinking, the pull-down complex in elution buffer is incubated at 65°C for 4–5 h, followed by treatment with RNase A followed by proteinase K at 37–42°C for 1 h each. The DNA can be eluted using either a magnetic beads-based method or by precipitating with ethanol or isopropanol. The choice of elution method depends on the downstream procedure; magnetic bead methods are typically used for ChIP-seq, while isopropanol or ethanol precipitation can be employed for ChIP qPCR.

ChIP pull-down validation

ChIP pull-down validation is crucial for assessing the performance of the antibody and confirming the success of the ChIP pull-down. To validate both antibody specificity and ChIP pull-down, it is recommended to use two positive control loci and one negative control locus. In case the control loci are unknown, one positive control locus and one negative control antibody should be employed for validation using qPCR.The ChIP pull-down should exhibit minimal enrichment for the positive control loci or antibodies compared with the negative control. If the pull-down demonstrates enrichment, then further downstream experiments can proceed. However, if inadequate enrichment is observed, it is advisable to repeat the pull-down to obtain more reliable results.

NGS library preparation overview

NGS is a widely adopted, massively parallel sequencing technology used for analyzing DNA or RNA sequences involved in various biological processes. Prior to sequencing DNA or RNA using the NGS platform, these molecules need to be converted into sequencing platform-compatible libraries that can be processed on different NGS platforms. For short-read sequencing, Illumina sequencing platforms are most commonly employed, utilizing sequencing by synthesis technology. The library preparation process consists of several steps including indexing, which allows for the pooling and sequencing of multiple samples on the same run.

Here, we provide a brief overview of Illumina ChIP-seq library preparation. The isolated and purified ChIP DNA obtained after IP is used for library preparation. In the first step, the ends of DNA fragments are repaired and an A base is added to the blunt ends of each strand. Subsequently, sequencing adapters are ligated to the end-repaired and A-tailed DNA fragments. Each adapter contains a T-base overhang that complements the A-tailed fragments. Following adapter ligation, libraries are amplified using PCR, sample indexes are added and the final library with complete adapters, including indexes, is purified for sequencing. Library preparation involves multiple steps, and there are various kits available for this purpose. The choice of library preparation kit depends on factors such as sample input and cost.

End repair

To convert DNA into a blunt end, an end-repair mix can be utilized, which includes T4 DNA polymerase and Klenow. These enzymes fill the 5′ overhang and remove the 3′ overhang. Additionally, the mix contains the T4 polynucleotide kinase, which phosphorylates the 5′ end and adds a hydroxyl group to the 3′ end of the DNA. Kits provide specific protocols that should be carefully followed. The reaction times may vary from 5 to 60 min, depending on the kit or individual enzymes and solution mix used. It is important to set up reactions accurately using standardized pipets and autoclaves or filter tips. For ChIP-seq library preparation, commonly used kits typically require 1–10 ng of immunoprecipitated DNA.

Important note

During the library preparation process, each step involves bead purifications, which must be performed in accordance with the kit's protocol requirements. Several bead cleanup steps are employed, including after-end repair, A-overhang, adapter ligation, size selection and PCR amplification. These bead cleanup steps are highly sensitive, requiring high-fidelity methods to purify small amounts of DNA during library preparation. The same beads are also used for the size selection of the final libraries suitable for sequencing.

A-overhang generation

It is necessary to add an A nucleotide at the 3′ end because blunt-ended DNA may self-ligate during adapter ligation. Additionally, the adapters themselves contain a T-overhang, which is used to ligate the adapter to A-overhang on the target DNA molecule. It is advisable to use fresh kits or reaction mixtures for A-overhang generation.

Adapter ligation

This step is crucial in library preparation as it involves ligating indexing adapters to the ends of the immunoprecipitated ChIP DNA fragments, preparing them for ChIP-seq. Careful consideration should be given to selecting unique indexes to assign samples and ensure that they enable the pooling of samples in the same run.

PCR amplification

PCR amplification is performed to enrich the adapter-ligated ChIP DNA using PCR primers designed against the ligated adapters. Only a few PCR cycles (∼15–20 or fewer) are necessary to preserve library representations. It is important to avoid using more than 20 PCR cycles, as it can compromise the uniformity of ChIP-seq data.

Confirmation of library construction

It is highly recommended to perform qPCR after ChIP pull-down to assess the enrichment of the target DNA, while including negative and positive controls for comparison. Similarly, after library preparation, the libraries can be selected by running agarose gel, followed by purification. Subsequently, validation of the library's correct size range and absence of adapter dimer/primer dimer can be done using the Bioanalyzer. Once the library is validated on the Bioanalyzer, it is advisable to check the library's enrichment and DNA quantitation through qPCR before proceeding to sequencing.

Conclusion

Based on our experience and the available literature, we provided a detailed description of each step involved in ChIP and library preparation. We also discussed standardization and precautions, along with helpful notes, to assist beginners and users. Although word limitations restrict the inclusion of extensive expert opinions, we have endeavored to provide comprehensive details to explain and enhance our understanding of these techniques.