Cancer Biology

Quantification of DNA double-strand breaks (DSBs) is critical for assessing genomic damage and cellular response to stress. γH2AX is a well-established marker for DNA double-strand breaks, but its quantification is often performed manually or semi-quantitatively, lacking standardization and reproducibility. Here, we present a standardized and automated workflow for γH2AX foci quantification in irradiated cells using immunofluorescence and a custom Fiji macro. The protocol includes steps for cell irradiation, immunostaining, image acquisition, and automated foci counting. The protocol is also adaptable to colony-like formations in multi-well plates, extending its utility to clonogenic assays. This protocol enables high-throughput, reproducible quantification of DNA damage with minimal user bias and can be readily implemented in routine laboratory settings.

Homologous recombination (HR) is a major pathway to repair DNA double-strand breaks. Hereditary breast and ovarian cancer syndrome (HBOC) is caused by germline pathogenic variants of HR-related genes, such as BRCA1 and BRCA2 (BRCA1/2). Cancer cells with HR deficiency are sensitive to poly(ADP-ribose) polymerase (PARP) inhibitors. Therefore, accurate evaluation of HR activity is helpful to diagnose HBOC and predict the effects of PARP inhibitors. The direct-repeat GFP (DR-GFP) assay has been utilized to evaluate cellular HR activity. However, evaluation by the DR-GFP assay tends to be qualitative and requires the establishment of stable cell lines. Therefore, we developed an assay to quantitatively measure HR activity called Assay for Site-Specific HR Activity (ASHRA), which can be performed by transiently transfecting two plasmids. In ASHRA, we use Cas9 endonuclease to create DNA double-strand breaks at specific sites in the genome, enabling the targeting of any endogenous loci. Quantification of HR products by real-time PCR using genomic DNA allows HR activity evaluated at the DNA level. Thus, ASHRA is an easy and quantitative method to evaluate HR activity at any genomic locus in various samples. Here, we present the protocols for adherent cells, suspension cells, and tumor tissues.

In response to DNA-damaging physical or chemical agents, the DNA damage repair (DDR) pathway is activated in eukaryotic cells. In the radiobiology field, it is important to assess the DNA damage effect of a certain irradiation regime on cancer cells and compare it to the effect on non-transformed cells exposed to identical conditions. The first step in the DNA repair mechanism consists of the attachment of proteins such as the phosphorylated histone γ-H2AX (p-γ-H2AX) to DNA double-strand breaks (DSB) in the nucleus, which leads to the formation of repairing foci. Therefore, imaging methods were established to evaluate the presence of foci inside the nucleus after exposure to DNA-damaging agents. This approach is superior in sensitivity to other methods, such as the comet assay or the pulsed-field gel electrophoresis (PFGE), that allow direct detection of cleaved DNA fragments. These electrophoresis-based methods require high ionizing radiation dosages and are difficult to reproduce compared to imaging-based assays. Conventionally, the number of foci is determined visually, with limited accuracy and throughput. Here, by exploring the effect of laser-plasma accelerated electrons FLASH irradiation on cancer cells, we describe an image cytometry protocol for the quantification of foci with increased throughput, upon large areas, with increased precision and sample-to-sample consistency. It consists of the automatic scanning of fluorescently labeled cells and using a gating strategy similar to flow cytometry to discriminate cells in co-culture based on nuclei elongation properties, followed by automatic quantification of foci number and statistical analysis. The protocol can be used to monitor the kinetics of DNA repair by quantification of p-γ-H2AX at different time points post-exposure or by quantification of other DNA repair proteins that form foci at the DNA DSB sites. Also, the protocol can be used for quantifying the response to chemical agents targeting DNA. This protocol can be performed on any type of cancer cells, and our gating strategy to discriminate cells in co-culture can also be used in other research applications.

Maintenance of genome integrity requires efficient and faithful resolution of DNA breaks and DNA replication obstacles. Dysfunctions in any of the processes orchestrating such resolution can lead to chromosomal instability, which appears as numerical and structural chromosome aberrations. Conventional cytogenetics remains as the golden standard method to detect naturally occurring chromosomal aberrations or those resulting from the treatment with genotoxic drugs. However, the success of cytogenetic studies depends on having high-quality chromosome spreads, which has been proven to be particularly challenging. Moreover, a lack of scoring guidelines and standardized methods for treating cells with genotoxic agents contribute to significant variability amongst different studies. Here, we report a simple and effective method for obtaining well-spread chromosomes from mammalian cells for the analysis of chromosomal aberrations. In this method, cells are (1) arrested in metaphase (when chromosome morphology is clearest), (2) swollen in hypotonic solution, (3) fixed before being dropped onto microscope slides, and (4) stained with DNA dyes to visualize the chromosomes. Metaphase chromosomes are then analyzed using high-resolution microscopy. We also provide examples, representative images, and useful guidelines to facilitate the scoring of the different chromosomal aberrations. This method can be used for the diagnosis of genetic diseases, as well as for cancer studies, by identifying chromosomal defects and providing insight into the cellular processes that influence chromosome integrity.

Graphical overview

In the human cell cycle, complete replication of DNA is a fundamental process for the maintenance of genome integrity. Replication stress interfering with the progression of replication forks causes difficult-to-replicate regions to remain under-replicated until the onset of mitosis. In early mitosis, a homology-directed repair DNA synthesis, called mitotic DNA synthesis (MiDAS), is triggered to complete DNA replication. Here, we present a method to detect MiDAS in human U2OS 40-2-6 cells, in which repetitive lacO sequences integrated into the human chromosome evoke replication stress and concomitant incomplete replication of the lacO array. Immunostaining of BrdU and LacI proteins is applied for visualization of DNA synthesis in early mitosis and the lacO array, respectively. This protocol has been established to easily detect MiDAS at specific loci using only common immunostaining methods and may be optimized for the investigation of other difficult-to-replicate regions marked with site-specific binding proteins.

DNA double strand breaks (DSBs) constantly arise in cells during normal cellular processes or upon exposure to genotoxic agents, and are repaired mostly by homologous recombination (HR) and non-homologous end joining (NHEJ). One key determinant of DNA DSB repair pathway choice is the processing of broken DNA ends to generate single strand DNA (ssDNA) overhangs, a process termed DNA resection. The generation of ssDNA overhangs commits DSB repair through HR and inhibits NHEJ. Therefore, DNA resection must be carefully regulated to avoid mis-repaired or persistent DSBs. Accordingly, many approaches have been developed to monitor ssDNA generation in cells to investigate genes and pathways that regulate DNA resection. Here we describe a flow cytometric approach measuring the levels of replication protein A (RPA) complex, a high affinity ssDNA binding complex composed of three subunits (RPA70, RPA32, and RPA14 in mammals), on chromatin after DNA DSB induction to assay DNA resection. This flow cytometric assay requires only conventional flow cytometers and can easily be scaled up to analyze a large number of samples or even for genetic screens of pooled mutants on a genome-wide scale. We adopt this assay in G₀- and G₁- phase synchronized cells where DNA resection needs to be kept in check to allow normal NHEJ.

Maintenance of DNA integrity is of pivotal importance for cells to circumvent detrimental processes that can ultimately lead to the development of various diseases. In the face of a plethora of endogenous and exogenous DNA damaging agents, cells have evolved a variety of DNA repair mechanisms that are responsible for safeguarding genetic integrity. Given the relevance of DNA damage and its repair for disease pathogenesis, measuring them is of considerable interest, and the comet assay is a widely used method for this. Cells treated with DNA damaging agents are embedded into a thin layer of agarose on top of a microscope slide. Subsequent lysis removes all protein and lipid components to leave ‘nucleoids’ consisting of naked DNA remaining in the agarose. These nucleoids are then subjected to electrophoresis, whereby the negatively charged DNA migrates towards the anode depending on its degree of fragmentation, creating shapes resembling comets, which can be visualized and analysed by fluorescence microscopy. The comet assay can be adapted to assess a wide variety of genotoxins and repair kinetics, and both DNA single-strand and double-strand breaks. In this protocol, we describe in detail how to perform the neutral comet assay to assess double-strand breaks and their repair using cultured human cell lines. We describe the workflow for assessing the amount of DNA damage generated by ionizing radiation or present endogenously in the cells, and how to assess the repair kinetics after such an insult. The procedure described herein is easy to follow and cost-effective.

Maintenance of DNA integrity is of pivotal importance for cells to circumvent detrimental processes that can ultimately lead to the development of various diseases. In the face of a plethora of endogenous and exogenous DNA-damaging agents, cells have evolved a variety of DNA repair mechanisms that are responsible for safeguarding genetic integrity. Given the relevance of DNA damage and its repair in disease, measuring the amount of both aspects is of considerable interest. The comet assay is a widely used method that allows the measurement of both DNA damage and its repair in cells. For this, cells are treated with DNA-damaging agents and embedded into a thin layer of agarose on top of a microscope slide. Subsequent lysis removes all protein and lipid components to leave so-called ‘nucleoids’ consisting of naked DNA remaining in the agarose. These nucleoids are then subjected to electrophoresis, whereby the negatively charged DNA migrates toward the anode depending on its degree of fragmentation and creates shapes resembling comets, which can be subsequently visualized and analyzed by fluorescence microscopy. The comet assay can be adapted to assess a wide variety of genotoxins and repair kinetics, in addition to both DNA single-strand and double-strand breaks. In this protocol, we describe in detail how to perform the alkaline comet assay to assess single-strand breaks and their repair using cultured human cell lines. We describe the workflow for assessing the amount of DNA damage generated by agents such as hydrogen peroxide (H₂O₂) and methyl-methanesulfonate (MMS) or present endogenously in cells, and how to assess the repair kinetics after such an insult. The procedure described herein is easy to follow and allows the cost-effective assessment of single-strand breaks and their repair kinetics in cultured cells.

Recent studies from multiple labs including ours have demonstrated the importance of extrachromosomal circular DNA (eccDNA) from yeast to humans (Shibata et al., 2012; Dillon et al., 2015; Møller et al., 2016; Kumar et al., 2017; Turner et al., 2017; Kim et al., 2020). More recently, it has been found that cancer cells obtain a selective advantage by amplifying oncogenes on eccDNA, which drives genomic instability (Wu et al., 2019; Kim et al., 2020). Previously, we have purified circular DNA and enriched the population using rolling circle amplification followed by high-throughput sequencing for the identification of eccDNA based on the unique junctional sequence. However, eccDNA identification by rolling circle amplification is biased toward small circles. Here, we report a rolling circle-independent method to detect eccDNA in human cancer cells. We demonstrate a sensitive and robust step-by-step workflow for finding novel eccDNAs using ATAC-seq (Assay for Transposase-Accessible Chromatin using sequencing) combined with a Circle_finder bioinformatics algorithm to predict the eccDNAs, followed by its validation using two independent methods, inverse PCR and metaphase FISH (Fluorescence in situ Hybridization).

In the last decade, genome editing has been the center of attention as a novel tool for mechanistic investigations and for potential clinical applications. Various genome editing tools like meganucleases, zinc finger nucleases (ZFNs), transcription activator-like effector-based nucleases (TALEN), and the clustered regularly interspaced short palindromic repeats (CRISPR)-associated genes (Cas), have been developed in recent years. For the optimal use as well as continued developments of these genome editing tools, the evaluation of their efficiencies and accuracies is vital. Here, we present a protocol for a reporter based on frameshift fluorescence protein which we recently developed to evaluate the efficiency and accuracy of genome editing tools. In this method, a ~20 bp target sequence containing frame-shifting is inserted after the start codon of a cerulean fluorescence protein (CFP) to inactivate its fluorescence, and only a new insertion/deletion event in the target sequence will reactivate the CFP fluorescence. To increase the traceability, an internal ribosome entry site and a red fluorescence protein, mCherryFP, are placed downstream of the reporter. The percentage of CFP-positive cells resulted from in/del mediated fluorescence restoration can be quantified by fluorescence measuring devices as the readout for genome editing frequency. As a demonstration, we present the usage for CRISPR-Cas9 technique here with flow cytometer as the readout for fluorescence changes.