Molecular Biology

Our genome is duplicated during every round of cell division through the process of DNA replication, but this fundamental process is subjected to various stresses arising from endogenous or exogenous sources. Thus, studying replication dynamics is crucial for understanding the mechanisms underlying genome duplication in physiological and replication stress conditions. Earlier, radioisotope-based autoradiography and density-labeling methods were used to study replication dynamics, which were limited in spatial resolution, representing only average estimates from many DNA samples. Here, we describe a DNA fiber assay that utilizes different thymidine analog incorporation, like 5-chloro-2’-deoxyuridine (CldU) and 5-iodo-2’-deoxyuridine (IdU), into replicating DNA. Such labeled DNA can be stretched and fixed on silanized glass slides, which are denatured with mild acidic treatment to expose the labeled nascent DNA. This DNA can then be visualized by using primary antibodies against CldU and IdU, followed by fluorophore-conjugated secondary antibodies, and observing them using a fluorescence microscope. The DNA fiber assay allows the visualization of individually replicating DNA at a single-molecular resolution and is highly quantitative, high-throughput, and easily reproducible. This technique offers insights into different replication parameters, like rate of DNA synthesis, extent of reversed fork protection, restart of stalled forks, and fork asymmetry under untreated or replication stress conditions at a single-molecule level.

Telomere length maintenance is strongly linked to cellular aging, as telomeres progressively shorten with each cell division. This phenomenon is well-documented in mitotic, or dividing, cells. However, neurons are post-mitotic and do not undergo mitosis, meaning they lack the classical mechanisms through which telomere shortening occurs. Despite this, neurons retain telomeres that protect chromosomal ends. The role of telomeres in neurons has gained interest, particularly in the context of neurodegenerative diseases such as amyotrophic lateral sclerosis (ALS), where aging is a major risk factor. This has sparked interest in investigating telomere maintenance mechanisms in post-mitotic neurons. Nevertheless, most existing telomere analysis techniques were developed for and optimized using mitotic cells, posing challenges for studying telomeres in non-dividing neuronal cells. Thus, this protocol adapts an already established technique, the combined immunofluorescence and telomere fluorescent in situ hybridization (IF-FISH) on mitotic cells to study the processes occurring at telomeres in cortical neurons of the mouse ALS transgenic model, TDP-43 rNLS. Specifically, it determines the occurrence of DNA damage and the alternative lengthening of telomeres (ALT) mechanism through simultaneous labeling of the DNA damage marker, γH2AX, or the ALT marker, promyelocytic leukemia (PML) protein, together with telomeres. Therefore, the protocol enables the visualization of DNA damage (γH2AX) or the ALT marker (PML) concurrently with telomeres. This technique can be successfully applied to brain tissue and enables the investigation of telomeres specifically in cortical neurons, rather than in bulk tissue, offering a significant advantage over Southern blot or qPCR-based techniques.

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.

The DNA double-strand breaks (DSBs) generated by exogenous and endogenous factors are repaired by two pathways: homologous recombination (HR) and non-homologous end-joining (NHEJ). These two pathways compete for DSB repair, and the choice of pathway depends on the context of the DNA lesion, the stage of the cell cycle, and the ploidy in the yeast Saccharomyces cerevisiae. However, the mechanistic details of the DSB repair pathway choice and its consequences for S. cerevisiae genome stability remain unclear. Here, we present PCR-based and cell-based assays as well as data analysis methods to quantitatively measure the efficiency of HR and NHEJ at DSBs in S. cerevisiae. An intermolecular recombination assay between plasmid and chromosomal DNA involving G-quadruplex DNA and a “suicide-deletion” assay have been utilized to evaluate the efficiency of HR and NHEJ, respectively. These streamlined protocols and optimized growth conditions can be used to identify the NHEJ- and HR-deficient S. cerevisiae mutant strains.

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: