Molecular 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

Nucleotide excision repair (NER) removes a wide variety of structurally unrelated lesions from the genome, including UV-induced photolesions such as 6–4 pyrimidine–pyrimidone photoproducts (6-4PPs) and cyclobutane pyrimidine dimers (CPDs). NER removes lesions by excising a short stretch of single-stranded DNA containing the damaged DNA, leaving a single-stranded gap that is resynthesized in a process called unscheduled DNA synthesis (UDS). Measuring UDS after UV irradiation in non-dividing cells provides a measure of the overall NER activity, of which approximately 90% is carried out by the global genome repair (GGR) sub pathway. Here, we present a protocol for the microscopy-based analysis and quantification of UDS as a measurement for GGR activity. Following local UV-C irradiation, serum-starved human cells are supplemented with the thymidine analogue 5-ethynyl-2'-deoxyuridine (EdU), which is incorporated into repair patches following NER-dependent dual incision. The incorporated nucleotide analogue is coupled to a fluorophore using Click-iT chemistry, followed by immunodetection of CPD photolesions to simultaneously visualize both signals by fluorescence microscopy. Accompanying this protocol is a custom-built ImageJ plug-in to analyze and quantify UDS signals at sites of CPD-marked local damage. The local UDS assay enables an effective and sensitive fluorescence-based quantification of GGR activity in single cells with application in basic research to better understand the regulatory mechanism in NER, as well as in diagnostics to characterize fibroblasts from individuals with NER-deficiency disorder.

Graphical abstract

8-oxo-7,8-dihydro-2′-deoxyguanosine (8-oxodG) is considered to be a premutagenic DNA lesion generated by 2'-deoxyguanosine (dG) oxidation due to reactive oxygen species (ROS). In recent years, the 8-oxodG distribution in human, mouse, and yeast genomes has been underlined using various next-generation sequencing (NGS)–based strategies. The present study reports the OxiDIP-Seq protocol, which combines specific 8-oxodG immuno-precipitation of single-stranded DNA with NGS, and the pipeline analysis that allows the genome-wide 8-oxodG distribution in mammalian cells. The development of this OxiDIP-Seq method increases knowledge on the oxidative DNA damage/repair field, providing a high-resolution map of 8-oxodG in human cells.

Analysis of DNA double strand breaks (DSBs) is important for understanding dyshomeostasis within the nucleus, impaired DNA repair mechanisms, and cell death. In the C. elegans germline, DSBs are important indicators of all three above-mentioned conditions. Although multiple methods exist to assess apoptosis in the germline of C. elegans, direct assessment of DSBs without the need for a reporter allele or protein-specific antibody is useful. As such, unbiased immunofluorescent approaches can be favorable. This protocol details a method for using terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) to assess DNA DSBs in dissected C. elegans germlines. Germlines are co-labeled with DAPI to allow for easy assessment of DNA DSBs. This approach allows for qualitative or quantitative measures of DNA DSBs.

Graphic abstract:

Schematic for TUNEL labeling of C. elegans germlines.

Directed evolution is a powerful approach to obtain genetically-encoded sought-for traits. Compared to the prolonged adaptation regimes to mutations occurring under natural selection, directed evolution unlocks rapid screening and selection of mutants with improved traits from vast mutated sequence spaces. Many systems have been developed to search variant landscapes based on ex vivo or in vivo mutagenesis, to identify and select new-to-nature and optimized properties in biomolecules. Yet, the majority of such systems rely on tedious iterations of library preparation, propagation, and selection steps. Furthermore, among the relatively few in vivo directed evolution systems developed to mitigate handling of repetitive ex vivo steps, directed evolution of DNA is the standard approach. Here, we present the protocol for designing the transfer of genetic information from evolving RNA donors to DNA in baker’s yeast, using CRISPR- and RNA-assisted in vivo directed evolution (CRAIDE). We use mutant T7 RNA polymerase to introduce mutations in RNA donors, while incorporation into DNA is directed by CRISPR/Cas9. As such, CRAIDE offers an opportunity to study fundamental questions, such as RNA’s contribution to the evolution of DNA-based life on Earth.

Graphic abstract:

CRISPR- and RNA-assisted in vivo directed evolution (CRAIDE).

Double-strand breaks (DSBs) are lesions in DNA that, if not properly repaired, can cause genomic instability, oncogenesis, and cell death. Multiple chromatin posttranslational modifications (PTMs) play a role in the DNA damage response to DSBs. Among these, RNF168-mediated ubiquitination of lysines 13 or 15 at the N-terminal tail of histone H2A (H2AK13/15Ub) is essential for the recruitment of effectors of both the non-homologous end joining (NHEJ) and the homologous recombination (HR) repair pathways. Thus, tools and techniques to track the spatiotemporal dynamics of H2AK13/15 ubiquitination at DNA DSBs are important to facilitate studies of DNA repair. Previous work from other groups used the minimal focus-forming region (FFR) of the NHEJ effector 53BP1 to detect H2AK15Ub generated upon damage induced by gamma or laser irradiation in live cells. However, 53BP1-FFR only binds nucleosomes modified with both H2AK15Ub and dimethylation of lysine 20 on histone H4 (H4K20me2); thus, 53BP1-FFR does not recognize H2AK13Ub–nucleosomes or nucleosomes that contain H2AK15Ub but lack methylation of H4K20 (H4K20me0). To overcome this limitation, we developed an avidity-based sensor that binds H2AK13/15Ub without dependence on the methylation status of histone H4K20. This sensor, called Reader1.0, detects DNA damage-associated H2AK13/15Ub in live cells with high sensitivity and selectivity. Here, we present a protocol to detect the formation of H2AK13/15Ub at laser-induced DSBs using Reader1.0 as a live-cell reporter for this histone PTM.

Graphic abstract:

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.

In vertebrates, hematopoietic stem cells (HSCs) regulate the supply of blood cells throughout the lifetime and help to maintain homeostasis. Due to their long lifespan, genetic integrity is paramount for these cells, and accordingly, a number of stem cell-specific mechanisms are employed. However, HSCs tend to show more DNA damage with increasing age due to an imbalance between proliferation rates and DNA damage responses. The comet assay is the most common and reliable method to study DNA strand breaks at the single-cell level. This procedure is based on the electrophoresis of agarose-embedded lysed cells. Following the electrophoretic mobilization of DNA, it is stained with fluorescent DNA-binding dye. Broken DNA strands migrate based on fragment size and form a tail-like structure called “the comet,” whereas intact nuclear DNA remains a part of the head of the comet. Since the alkaline comet assay fails to differentiate between single and double-strand breaks (DSBs), we used a neutral comet assay to quantitate the DSBs in HSCs upon aging and other physiological stresses. The protocol presented here provides procedural details on this highly sensitive, rapid, and cost-effective assay, which can be used for rare populations of cells such as HSCs.

Graphical abstract:

The neutral comet assay is an extremely useful tool that allows the detection and quantitation of double-strand DNA breaks at the single-cell level. The graphical abstract represents a flowchart for the neutral comet assay procedure.

Hepatitis B virus (HBV) is the major cause of liver diseases and liver cancer worldwide. After infecting hepatocytes, the virus establishes a stable episome (covalently closed circular DNA, or cccDNA) that serves as the template for all viral transcripts. Specific and accurate quantification of cccDNA is difficult because infected cells contain abundant replicative intermediates of HBV DNA that share overlapping sequences but arranged in slightly different forms. HBV cccDNA can be detected by Southern blot or qPCR methods which involve enzymatic digestion. These assays are laborious, have limited sensitivity, or require degradation of cellular DNA (which precludes simple normalization). The method described in this protocol, cccDNA inversion quantitative (cinq)PCR, instead uses a series of restriction enzyme-mediated hydrolysis and ligation reactions that convert cccDNA into an inverted linear amplicon, which is not amplified or detected from other forms of HBV DNA. Importantly, cellular DNA remains quantifiable during sample preparation, allowing normalization and markedly improving precision. Further, a second linear fragment (derived from enzymatic digestion of a separate region of the HBV DNA genome and is present in all forms of HBV DNA) can be used to simultaneously quantify total HBV levels.

Graphic abstract:

Selective detection of HBV cccDNA and total HBV DNA using cinqPCR (Reproduced from Tu et al., 2020a).

7,8-dihydro-8-oxoguanine (8-oxoG) is one of the most common and mutagenic oxidative DNA damages induced by reactive oxygen species (ROS). Since ROS is mainly produced in the inner membranes of the mitochondria, these organelles and especially the mitochondrial DNA (mtDNA) contained therein are particularly affected by this damage. Insufficient elimination of 8-oxoG can lead to mutations and thus to severe mitochondrial dysfunctions. To eliminate 8-oxoG, the human body uses the enzyme 8-oxoguanine DNA glycosylase 1 (OGG1), which is the main antagonist to oxidative damage to DNA. However, previous work suggests that the activity of the human OGG1 (hOGG1) decreases with age, leading to an age-related accumulation of 8-oxoG. A better understanding of the exact mechanisms of hOGG1 could lead to the discovery of new targets and thus be of great importance for the development of preventive therapies. Because of this, we developed a real-time base excision repair assay with a specially designed double-stranded reporter oligonucleotides to measure the activity of hOGG1 in lysates of isolated mitochondria. This system presented here differs from the classical assays, in which an endpoint determination is performed via a denaturing acrylamide gel, by the possibility to measure the hOGG1 activity in real-time. In addition, to determine the activity of each enzymatic step (N-glycosylase and AP-lyase activity) of this bifunctional enzyme, a melting curve analysis can also be performed. After isolation of mitochondria from human fibroblasts using various centrifugation steps, they are lysed and then incubated with specially designed reporter oligonucleotides. The subsequent measurement of hOGG1 activity is performed in a conventional real-time PCR system.