Cancer Biology

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

Errors in chromosome segregation during mitosis lead to chromosome instability, resulting in an unbalanced number of chromosomes in the daughter cells. Light microscopy has been used extensively to study chromosome missegregation by visualizing errors of the mitotic spindle. However, less attention has been paid to understanding spindle function in the broader context of intracellular structures and organelles during mitosis. Here, we outline a protocol to visualize chromosomes and endomembranes in mitosis, combining light microscopy and 3D volume electron microscopy, serial block-face scanning electron microscopy (SBF-SEM). SBF-SEM provides high-resolution imaging of large volumes and subcellular structures, followed by image analysis and 3D reconstruction. This protocol allows scientists to visualize the whole subcellular context of the spindle during mitosis.

Mitochondria play decisive roles in bioenergetics and intracellular communication. These organelles contain a circular mitochondrial DNA (mtDNA) genome that is duplicated within one to two hours by a mitochondrial replisome, independently from the nuclear replisome. mtDNA stability is regulated in part at the level of mtDNA replication. Consequently, mutations in mitochondrial replisome components result in mtDNA instability and are associated with diverse disease phenotypes, including premature aging, aberrant cellular energetics, and developmental defects. The mechanisms ensuring mtDNA replication stability are not completely understood. Thus, there remains a need to develop tools to specifically and quantifiably examine mtDNA replication. To date, methods for labeling mtDNA have relied on prolonged exposures of 5′-bromo-2′-deoxyuridine (BrdU) or 5′-ethynyl-2′-deoxyuridine (EdU). However, labeling with these nucleoside analogs for a sufficiently short time in order to monitor nascent mtDNA replication, such as under two hours, does not produce signals suited for efficient or accurate quantitative analysis. The assay system described here, termed Mitochondrial Replication Assay (MIRA), utilizes proximity ligation assay (PLA) combined with EdU-coupled Click-IT chemistry to address this limitation, thereby enabling sensitive and quantitative analysis of nascent in situ mtDNA replication with single-cell resolution. This method can be further paired with conventional immunofluorescence (IF) for multi-parameter cell analysis. By enabling monitoring nascent mtDNA prior to the complete replication of the entire mtDNA genome, this new assay system allowed the discovery of a new mitochondrial stability pathway, mtDNA fork protection. Moreover, a modification in primary antibodies application allows the adaptation of our previously described in situ protein Interactions with nascent DNA Replication Forks (SIRF) for the detection of proteins of interest to nascent mtDNA replication forks on a single molecule level (mitoSIRF).

Graphical overview

Schematic overview of Mitochondrial Replication Assay (MIRA). 5′-ethynyl-2′-deoxyuridine (EdU; green) incorporated in DNA is tagged with biotin (blue) using Click-IT chemistry. Subsequent proximity ligation assay (PLA, pink circles) using antibodies against biotin allows the fluorescent tagging of the nascent EdU and amplification of the signal sufficient for visualization by standard immunofluorescence. PLA signals outside the nucleus denote mitochondrial DNA (mtDNA) signals. Ab, antibody. In in situ protein interactions with nascent DNA replication forks (mitoSIRF), one of the primary antibodies is directed against a protein of interest, while the other detects nascent biotinylated EdU, thus enabling in situ protein interactions with nascent mtDNA.

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.

The duplication of DNA is a fundamental process that is required for the transfer of the genetic information from parent to daughter cells. Aberrant DNA replication processes are associated with diverse disease phenotypes, including developmental defects, ageing disorders, blood disorders such as Fanconi Anemia, increased inflammation and cancer. Therefore, the development of tools to study proteins associated with error-free DNA replication processes is of paramount importance. So far, methods to study proteins associated with nascent replication forks relied on conventional immunofluorescence and immunoprecipitation assays of 5′-ethylene-2′-deoxyuridine (EdU) labeled DNA (iPOND). While greatly informative and important, these methods lack specificities for nascent fork interactions (e.g., IF) or assay an average change of millions of cells without single-cell resolution (e.g., iPOND). The assay system described here combines proximity ligation assay (PLA) with EdU coupled click-iT chemistry, which we termed “in situ Protein Interaction with Nascent DNA Replication Forks (SIRF)”. This method enables sensitive and quantitative analysis of protein interactions with nascent DNA replication forks with single-cell resolution, and can further be paired with conventional immunofluorescence marker analysis for added multi-parameter analysis.