Capturing Common Fragile Site Breaks by Native γH2A.X ChIP

Summary

We present a rapid and efficient method to detect common fragile site breaks through native γH2A.X chromatin immunoprecipitation (ChIP). This approach significantly reduces both the time and labor associated with traditional γH2A.X ChIP assays while maintaining high reproducibility and reliability of results.

Abstract

Replication stress induced by exposure to extrinsic agents can lead to DNA breaks at common fragile sites, which are regions in the genome known to be prone to structural instability. The γH2A.X chromatin immunoprecipitation (ChIP) assay serves as a powerful tool in genotoxicity studies, as γH2A.X phosphorylation is a well-established marker for DNA double-strand breaks. Traditional γH2A.X ChIP assays, however, are often labor-intensive and involve multiple, time-consuming steps. In this study, we present a simplified yet effective method that combines subcellular fractionation with native ChIP to isolate γH2A.X-associated complexes. This approach is particularly suitable for analyzing γH2A.X-chromatin interactions with enhanced specificity and efficiency. Using subcellular fractionation, chromatin-unbound materials are effectively removed, resulting in a purified chromatin fraction. Subsequent micrococcal nuclease (MNase) digestion under mild conditions allows chromatin fragmentation while preserving physiological interactions between γH2A.X and its associated protein complexes. This preservation is essential for studying native interaction partners involved in DNA damage response pathways. This optimized native ChIP protocol substantially reduces the time and labor associated with conventional γH2A.X ChIP assays. The streamlined procedure not only simplifies the workflow but also yields highly reproducible results, making it particularly advantageous in settings where high-throughput processing of multiple samples is required. This method has broad applicability in studies focused on genome stability, DNA repair, and chromatin biology, where accurate and efficient detection of DNA damage sites is critical. By employing optimized protocols and streamlined steps, this method enables the detection of DNA damage at fragile sites with improved sensitivity and minimal sample handling, making it a valuable tool for studies on genome stability and DNA damage response.

Introduction

Common fragile sites (CFSs) are large chromosomal regions found on every human chromosome prone to breaking during metaphase. Under replication stress, replication at these regions is significantly delayed, preventing their complete duplication before mitotic entry¹, which ultimately results in site-specific gaps and breaks. CFSs are hotspots for chromosomal instability and are a major cause of chromosomal rearrangements during early cancer development. Replication stress, which is often present under tumorigenic conditions, can lead to the loss of tumor suppressor genes and amplification of oncogenes-collectively referred to as copy number variation (CNV)^2,3,4,5,6. Additionally, CFSs are highly prone to viral integration, further promoting cancer development^7,8,9,10. Multiple homozygous deletions of tumor suppressor genes have been detected in CFS regions during pan-cancer analyses of primary tumors. The most commonly affected CFSs in cancer include FRA2F, FRA3B, FRA4F, FRA5H, and FRA16D¹¹. CFSs are particularly vulnerable to breakage in the presence of extrinsic carcinogenic agents¹². To assess the detrimental carcinogenic effects of environmental contaminants, a fast and reliable method for quantifying CFS break occurrence is needed.

Phosphorylation of H2A.X at the serine residue 139 (γH2A.X) by Ataxia Telangiectasia and Rad3-Related Protein (ATR) or Ataxia Telangiectasia Mutated (ATM) is a key event in signaling replication fork stalling¹³. γH2A.X serves as an indicator of stalled replication forks prior to double-strand break (DSB) formation¹³, creating a favorable chromatin environment to facilitate the efficient recruitment of repair proteins to stalled sites. Additionally, γH2A.X can be recruited to break sites following fork collapse^14,15, consistent with its primary role in DSB repair. Since CFS breaks are closely associated with chromosomal aberrations that drive cancer progression, detecting these breaks can be instrumental in understanding the early stages of tumorigenesis. The presence of γH2A.X at CFSs can be used as a biomarker to detect early events of genomic instability. This information can help identify potential carcinogens and evaluate the risk associated with exposure to various extrinsic agents. By measuring DNA breaks at CFSs induced by extrinsic agents, γH2A.X chromatin IP (ChIP) can provide insight into how such agents contribute to the mechanisms underlying tumorigenesis.

In the conventional ChIP (i.e., Cross-linked ChIP, X-ChIP), the association of γH2A.X with its target DNA sequences is stabilized by reversible formaldehyde crosslinking. Chromatin is subsequently sheared to fragments of approximately 500 base pairs (bp) through sonication, and the resulting solution is cleared of debris by sedimentation^16,17,18. A ChIP-grade γH2A.X antibody is then added to the cleared chromatin fraction, followed by the addition of Protein A/G agarose beads to enrich for γH2A.X-bound chromatin regions^16,17,18. The immune complexes (i.e., beads-antibody-γH2A.X-targeted DNA complex) are washed multiple times with stringent washing buffers to remove nonspecifically bound DNA fragments^16,17,18. After washing, the specifically bound DNA is eluted from the immune complexes. The formaldehyde cross-links are then reversed, followed by protein digestion using proteinase K, after which the enriched DNA is purified and concentrated^16,17,18. To assess the γH2A.X-associated regions, PCR, quantitative PCR (qPCR), or direct sequencing is used^16,17,18. The occupancy of γH2A.X at specific regions, such as CFS, is determined by the intensity of the PCR or qPCR signal, which is proportional to the amount of γH2A.X bound at that location, providing insights into site-specific DNA damage and repair events^16,17,18.

Despite being a powerful experimental approach, the X-ChIP has several significant limitations: (i) it requires a large number of cells, typically in the range of 1 x 10⁷ to 5 x 10⁷, due to the inefficiency of antibody precipitation associated with fixation, which increases the overall cost of the experiment¹⁹; (ii) the process of reversing formaldehyde cross-links and subsequent DNA purification is time-consuming and labor-intensive, making it challenging to maintain consistency and reliability in results; and (iii) γH2A.X-DNA interactions with minor functional significance may not be distinguished from those with greater significance because the cross-linking step can stabilize transient interactions, leading to the detection of interactions that may not be biologically relevant¹⁹.

Native chromatin immunoprecipitation (Native ChIP or N-ChIP) is an essential biochemical technique used to study protein-DNA interactions within their native chromatin context under physiological salt conditions. It has been instrumental in elucidating the spatial and temporal organization of chromatin, transcription factor binding, and histone modifications. Native ChIP has a long-standing role in the broader field of chromatin biology and epigenetics, providing unique advantages and limitations compared to X-ChIP. This method, introduced in the late 1980s²⁰, involves the isolation of chromatin from cells by methods that preserve its native structure, such as digestion with micrococcal nuclease (MNase)²¹. This preserves the inherent protein-DNA and histone-DNA contacts, which makes Native ChIP particularly well-suited for studying histone modifications and nucleosome positioning in their natural chromatin setting²². High-resolution Native ChIP studies have demonstrated the use of MNase digestion to reduce chromatin to individual nucleosomes, which facilitates the mapping of histone modifications with greater accuracy²³. Furthermore, because no chemical cross-linking is involved, the risk of introducing biases or artifacts that might misrepresent the protein-DNA interactions is minimized²⁴.

In contrast to X-ChIP, where formaldehyde or other cross-linking agents are used to fix protein-DNA interactions, Native ChIP provides a more realistic view of chromatin by avoiding potential cross-linking artifacts. However, while X-ChIP is generally better suited for detecting transient or dynamic interactions between DNA and regulatory proteins²⁵, Native ChIP is ideal for stable protein-DNA interactions, such as histones or other chromatin-bound proteins^26,27. One of the limitations noted for Native ChIP is the inability to capture low-affinity or transient binding events, which are often stabilized through cross-linking in X-ChIP²⁵.

A significant body of work in epigenetics has leveraged Native ChIP to uncover histone modifications in diverse biological settings²⁸. These efforts have been crucial in defining the histone code - the pattern of histone modifications that regulate gene expression and chromatin dynamics²⁹. Although H2A.X is a less strongly associated linker histone, the native H2A.X ChIP method has been successfully applied in embryonic stem cells³⁰. In this study, we optimized a chromatin extraction procedure to perform Native ChIP of γH2A.X in human 293T cells (Figure 1). Hydroxyurea and aphidicolin are widely used in research to investigate DNA replication stress, damage, and genomic instability³¹. In this study, these agents were applied to cells to induce replication stress and generate DNA breaks at CFS.

Using starting material of approximately 1 x 10⁶ to 5 x 10⁶ cells, this method can be divided into four main stages: (i) subcellular fractionation to isolate chromatin, (ii) micrococcal nuclease (MNase) digestion to fragment chromatin, (iii) immunoprecipitation and elution, and (iv) DNA analysis by quantitative PCR (qPCR). Conducting ChIP following subcellular fractionation provides several benefits and has been well-documented in numerous studies^32,33,34,35. This approach allows for the removal of chromatin-unbound proteins and other cellular debris, resulting in a highly purified chromatin fraction. By isolating chromatin before immunoprecipitation, subcellular fractionation helps maintain native chromatin interactions and reduces background noise from non-chromatin-associated proteins, which leads to more specific and reliable results, as only chromatin-bound complexes are retained for analysis. Moreover, subcellular fractionation enables milder conditions for chromatin digestion, thereby preserving physiological protein-DNA interactions and offering a more accurate representation of chromatin dynamics within the native cellular environment.

Using native ChIP of γH2AX to measure the impact of extrinsic agents on common fragile site breakage holds significant potential for cancer research. This technique enables the detection of DNA damage induced by exposure to environmental carcinogens, providing insights into the molecular mechanisms by which pollutants contribute to genomic instability and cancer development. By preserving the native chromatin context, this method facilitates the accurate assessment of DNA damage patterns associated with carcinogenic exposure, aiding in the evaluation of environmental risks and the study of pollution-driven tumorigenesis.

Protocol

1. Cell harvesting

1. Seed about 5 x 10⁵ HEK 293T cells into each of the four 6 cm dishes, each containing 4 mL of complete DMEM medium.
2. After 24 h, treat one dish with 2 µL of 1 mM Aphidicolin (refer to Table of Material) stock solution (final concentration of 0.5 µM) and another dish with 20 µL of 1 M hydroxyurea (refer to Table of Material) stock solution (final concentration of 5 mM) to induce replication stress. Add DMSO to the remaining two dishes to serve as controls.
3. After 24 h of treatment, discard the culture media. One 6 cm plate typically yields approximately 2 x 10⁶ cells at 60%-70% confluence.
4. Rinse each dish 2x with 5 mL of ice-cold 1x PBS. Use cell scrapers to detach the cells and transfer the cell suspension to four individual 1.5 mL tubes. Gently pipette up and down with a P1000 pipette to dissociate any cell clumps.
5. Centrifuge the cells at 500 x g for 5 min at 4 °C, then discard the supernatant. Place the cells on ice.

2. Subcellular fractionation

1. Resuspend the cell pellet in 500 µL of freshly prepared cold Buffer A (refer to Table 1), ensuring the complete dissociation of cell clumps by gentle pipetting.
2. Incubate the lysates on ice for 5-10 min. Check lysis progression under a microscope to make sure complete cell lysis.
   1. Take a small aliquot of the lysate (around 5 - 10 µL) and place it on a clean microscope slide. Cover it with a coverslip to avoid contamination.
   2. Use a light microscope with an appropriate magnification (e.g., 20x - 40x) to visualize cells or debris. Compare with an un-lysed control sample to differentiate between intact cells and lysed material.
      NOTE: A properly lysed sample will have no distinct cell outlines, only diffuse chromatin or cellular material. Adjust the focus to clearly observe the lysate. If needed, apply mechanical force during the lysis step, such as using a Dounce homogenizer, when working with certain cell types.
3. Centrifuge at 500 x g for 5 min at 4 °C, once cells are completely lysed. Carefully discard the supernatant. Resuspend the nuclei pellet in 500 µL of cold Buffer A using wide-orifice pipette tips.
   NOTE: Wide-orifice tips help minimize shear forces and protect delicate samples like chromatin. Make wide-bore tips by cutting the end of standard tips with a sharp blade.
4. Centrifuge at 500 x g for 5 min at 4 °C. Carefully discard the supernatant.
5. Prewarm an incubator to 37 °C and prepare a stopping buffer (100 mM EDTA, pH 8.0; Table 1).
6. Optimize Micrococcal Nuclease (MNase, refer to Table of Material) concentrations and incubation times in advance.
   1. Divide 40 µL of testing chromatin sample into several equal aliquots to test different MNase concentrations and incubation times.
   2. Use a range of MNase concentrations (e.g., 0.0625 U, 0.125 U, 0.25 U, 0.5 U, 1 U, 2 U, 4 U, 8 U per reaction) and test multiple incubation times (e.g., 2, 5, 10, and 15 min).
   3. Add MNase Buffer (refer to Table 1) containing various concentrations of MNase to the chromatin aliquots and incubate samples at 37 °C for the specified times.
   4. Terminate the reaction by adding 1/4 volume of stopping buffer (final concentration: 20 mM EDTA) immediately after the desired incubation time.
   5. Isolate DNA from the digested chromatin samples using a phenol/chloroform/isoamyl alcohol extraction method.
   6. Run the extracted DNA on a 1.5% agarose gel to visualize digestion patterns: Under-digestion will show high molecular weight bands (Figure 2, lane 1-4); over-digestion will result in a smear or very short fragments (Figure 2, lane 6-8), and optimal digestion will yield a clear nucleosomal ladder pattern (Figure 2, lane 5, e.g., mono-, di-, tri-nucleosomes).
   7. Identify the conditions that produce the desired nucleosomal resolution without excessive over-digestion.
      NOTE: CaCl₂ acts as a cofactor for MNase activity. Optimize the digestion by adjusting the CaCl₂ concentration between 1 mM and 5 mM.
7. Gently resuspend the intact nuclei with 100 μL of MNase Buffer by pipetting 5 - 10 times with wide-orifice tips. Immediately add the pre-determined amount of MNase to the samples (1.25 U MNase/100 µL MNase Buffer).
   NOTE: When working with multiple samples, digest each one individually to avoid over-digestion.
8. Place the tubes on a rotator and incubate for 5 min at 37 °C. Immediately return the tubes to ice and terminate MNase digestion by adding EDTA to a final concentration of 20 mM and mix by vortexing.
9. Add 500 µL of Buffer B (refer to Table 1) to each sample and mix thoroughly by pipetting up and down 5x - 10x. Solubilize proteins by incubating on ice for 5 min.
   NOTE: The salt and detergent in Buffer B help dissociate weak chromatin-bound proteins and expose the epitopes for immunoprecipitation.
10. Pellet the insoluble material by centrifuging at maximum speed for 5 min at 4 °C. Transfer the clear supernatant to new 1.5 mL tubes labeled as the native chromatin fraction. The samples can either be stored at -80 °C or used to validate the efficiency of chromatin fragmentation.
    NOTE: Avoid frequent freeze-thaw cycles, as they may disrupt protein-DNA interactions of interest. Minimize freeze-thaw cycles whenever possible.

3. Verification of chromatin fragmentation

1. Aliquot 10 µL of the supernatant from each sample into a new 1.5 mL tube. Mix with 20 µL of distilled water and 30 µL of phenol/chloroform/isoamyl alcohol (25:24:1).
2. Close the tubes tightly and vortex vigorously for 15-30 s. Centrifuge at 20,000 x g (or the maximum speed of the centrifuge) for 10 min at 4 °C. After centrifugation, three distinct layers will be observed: a clear top layer, a white middle layer, and a yellow bottom layer.
3. Carefully transfer 20 µL of the upper aqueous phase (containing DNA) to a fresh tube. Separate the purified DNA in 1.5% agarose gel for 30 min at 100 V and visualize the digestion patterns. Ensure that the size of chromatin fragments is primarily between 200 and 1000 base pairs.
   NOTE: Proper chromatin fragment sizes are crucial for the success of native ChIP and depend on MNase treatment conditions, including enzyme units, incubation time, and CaCl₂ concentration. MNase digestion efficiency can also vary based on cell type and number. The chromatin fragmentation pattern shown in Figure 2 (lane 5) is recommended for this ChIP assay.

4. Immunoprecipitation

1. Aliquot 20 µL of digested chromatin from each sample into a fresh 1.5 mL tube and mix with 180 µL of Elution Buffer (refer to Table 1). Label these tubes as Input samples and store at -20 °C.
2. Transfer 400 µL of chromatin sample into another 1.5 mL tube for ChIP.
3. Add γH2A.X antibody (refer to Table of Material) to one DMSO-treated, one Aphidicolin-treated sample, and one hydroxyurea-treated sample. Add the same amount of normal IgG (refer to Table of Material) to another DMSO-treated sample as a negative control for the ChIP assay.
   NOTE: Here, 1 μg of primary antibody is typically used for 400 μL of chromatin (i.e., antibody final concentration is 2.5 µg/mL). However, the optimal amount should be empirically determined for different γH2A.X antibodies.
4. Place the ChIP tubes on a rotator at 4 °C and incubate for at least 5 h, or preferably overnight.
5. Meanwhile, aliquot 100 µL of ChIP-grade magnetic Protein A/G beads (refer to Table of Material) into a new 1.5 mL tube. Use wide-orifice tips and pipette slowly to ensure accurate measurement of the beads. Place the tube on a magnetic stand for at least 1 min, then carefully discard the liquid.
6. Resuspend the beads in 1 mL of 1x PBS containing 0.5% BSA. Rotate at 4 °C for approximately 4 h. Place the tube on a magnetic stand for at least 1 min and discard the supernatant.
7. Wash the beads again with 1 mL of 1x PBS containing 0.5% BSA. Place the tube on the magnetic stand for 1 min to pellet the magnetic beads, then discard the supernatant.
   NOTE: Steps 4.5 to 4.7 are the pre-coating of beads to reduce non-specific binding of antibodies to magnetic beads.
8. Resuspend the pre-coated beads in 100 µL of Buffer B using wide-orifice tips. Add 25 µL of the pre-coated magnetic bead suspension to each ChIP sample tube. Rotate at 4 °C for 2 h.
9. Place the ChIP tubes on the magnetic stand and wait until the beads are completely attached to the side of the tube and the solution becomes clear.
10. Discard the clear supernatant without disturbing the magnetic beads. Resuspend the beads with 1 mL of Wash Buffer (refer to Table 1) and rotate at 4 °C for 10 min.
11. Place the tubes back on the magnetic stand and wait until the solution becomes clear. Discard the wash buffer. Repeat washing for a total of four washes.
12. Discard the wash buffer after final wash and briefly centrifuge the tubes at 400 x g for 30 s at 4 °C to spin down any residual liquid. Place the tubes back on the magnetic stand and carefully remove any remaining liquid from the bottom of the tube.

5. Elution and DNA precipitation

NOTE: Antibody efficiency may vary among different batches. It is important to confirm the binding affinity of a new antibody by checking the immunoprecipitated samples through Western blot analysis.

1. Verify ChIP antibody pull-down efficiency using Western blot (WB) as described below.
   1. Take a small aliquot of the ChIP sample for analysis (i.e., usually 10% of the ChIP sample). Include Input chromatin (pre-immunoprecipitation) and negative control (e.g., IgG pull-down) for comparison.
   2. Elute proteins from the antibody-bound beads by heating in 20 µL of 1x SDS-PAGE loading buffer (refer to Table of Material) at 95 °C for 5 min.
   3. Load the IP samples, input, and controls onto a 15% SDS-PAGE gel. Run the gel.
   4. Transfer the proteins to a 0.2 µm nitrocellulose (refer to Table of Material) or PVDF membrane using a wet or semi-dry transfer system.
   5. Block the membrane with 5% non-fat milk or BSA in TBST (refer to Table 1) for 1 h at room temperature to prevent non-specific binding.
   6. Incubate the membrane with the primary antibody against the γH2A.X (refer to the Table of Material) diluted in blocking buffer for 1-2 h at room temperature or overnight at 4 °C.
   7. Wash the membrane 3x with TBST to remove unbound antibodies. Incubate the membrane with an HRP-conjugated secondary antibody (refer to the Table of Material) for 1 h at room temperature. Wash the membrane again to remove excess secondary antibodies.
   8. Develop the membrane using a chemiluminescent substrate and visualize the signal with an imager. Compare the signal intensity between the IP, input, and control lanes to assess the efficiency and specificity of the pull-down.
      NOTE: A band corresponding to the target protein in the IP lane confirms successful antibody pull-down. This approach ensures you can evaluate the efficacy of the antibody in capturing the target protein during the ChIP experiment.
2. Add 50 µL of Elution Buffer (refer to the Table 1) to each of the remaining ChIP samples. Place the tubes on a thermomixer and shake for 15 min at room temperature.
3. Place the tubes on the magnetic holder for at least 1 min. Collect the elute into new tubes. Repeat 1x and collect the elute in the same tubes.
4. Add additional 100 µL of Elution Buffer to each ChIP elution sample and 180 µL of Elution Buffer to each Input sample.
5. Add 200 µL of phenol/chloroform/isoamyl alcohol (25:24:1) to each sample and vortex vigorously. Centrifuge the samples at 20,000 x g (or maximum speed) for 10 min at 4 °C.
6. Add 19 µL of 3M sodium acetate (NaOAc, pH 5.2; refer to the Table 1) and 2 µL of glycogen solution (20 mg/mL, refer to the Table of Material) to each new 1.5 mL centrifuge tube.
7. After centrifugation, carefully transfer the upper aqueous layer (approximately 190 µL) to the tubes containing NaOAc and glycogen and mix by vortexing.
8. Add 500 µL of 100% ethanol and vortex. Precipitate the DNA by incubating the samples at -20 °C for at least 2 h or overnight.
9. Centrifuge the tubes at 20,000 x g (or maximum speed) for 10 min at 4 °C. Discard the supernatant, taking care not to disturb the white pellet. Resuspend the pellet in 1 mL of 70% ethanol and vortex thoroughly.
10. Centrifuge the tubes at 20,000 x g (or maximum speed) for 5 min at 4 °C. Carefully remove the supernatant. Briefly centrifuge the tubes again to spin down any residual ethanol. Carefully remove the ethanol using a P20 pipette. Air dry the DNA pellets for 2-3 min.
    NOTE: Avoid over-drying the pellet, as this can make the DNA difficult to re-dissolve.
11. For ChIP samples, resuspend the DNA in 400 µL of TE buffer (refer to Table 1). For Input DNA, resuspend in 1000 µL of TE buffer. The eluted samples can now be stored at -20°C.

6. qPCR quantification

1. Perform qPCR using a commercial kit (refer to Table of Material) with technical triplicates for each sample. Confirm the presence of a single specific PCR product by conducting melting curve analysis to ensure the specificity of amplification³⁶.
2. Data analysis
   NOTE: In relative quantification analysis, the test sample is expressed as a fold change relative to a control sample (immunoprecipitated using normal purified IgG or mock IP). DNA loci known to be unoccupied by the immunoprecipitated protein (negative locus) can be used in this manner as a reference gene compared to known, occupied, positive control DNA loci³⁶.
   1. Calculate the percent of input for each ChIP using the formula below
      %Input = 2(-ΔCt [normalized ChIP])
   2. Normalize the positive locus ΔCt values to negative locus (ΔΔCt) by subtracting the ΔCt value obtained for the positive locus from the ΔCt value for the negative locus using the formula below
      (ΔΔCt = ΔCtpositive - ΔCtnegative)
   3. Calculate the fold enrichment of the positive locus sequence in ChIP DNA over the negative locus using the formula below
      Fold enrichment =2ΔΔCt
      The sequences of the qPCR primers used for analysis are provided in Table 2. The genomic organization of FRA3B and FRA16D³⁷is depicted in Figure 3A,B.
3. Statistical analysis
   1. Analyze the results statistically using Student's paired t-test. A p-value of ≤0.05 is considered statistically significant, indicating that the observed differences are unlikely to be due to random variation³⁸.

Results

The size of chromatin fragments is crucial for the success of Native ChIP, as it directly impacts the accessibility of DNA regions for antibody binding. To determine the optimal MNase concentration for chromatin fragmentation, we prepared a series of microcentrifuge tubes containing varying concentrations of MNase (i.e., 0.0625 U, 0.125 U, 0.25 U, 0.5 U, 1 U, 2 U, 4 U, 8 U per reaction) and 40 µL of isolated nuclei. Each reaction was incubated at 37 °C for 5 min to achieve a range of chromatin fragment sizes. The results...

Discussion

Environmental pollution is a significant contributor to human cancers. Many pollutants are carcinogenic, meaning they can cause genetic damage that leads to the development of cancer^40,41. However, determining whether a particular substance is tumorigenic is a challenging task. A fast, reliable, and cost-efficient method for identifying carcinogenic potential would empower scientists to efficiently screen environmental pollutants and assess their impact on genomi...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.2 µm nitrocellulose membrane	Amersham	10600011
Actin B	proteintech	20536-1-AP
Aphidicolin	MedChemExpress	HY-N6733
ChIP-grade magnetic Protein A/G beads	ThermoFisher	26162
Clarity Western ECL Substrate	Bio-Rad	#1705061
Glycogen, molecular biology grade	ThermoFisher	Cat. No.  R0561
HRP-conjugated secondary antibody	proteintech	SA00001-2
hydroxyurea	MedChemExpress	HY-B0313
Micrococcal Nuclease	NEB	M0247S
normal IgG	Santa Cruz	sc-2025
Taq Universal SYBR Green Supermix	BioRad	1725120
γH2A.X antibody  (for ChIP)	Sigma-Aldrich	05-636
γH2A.X antibody (for WB)	Cell Signaling	#25955