Molecular Biology

Real-time quantitative PCR (qPCR) is a pivotal technique for analyzing gene expression and DNA copy number variations. However, the limited availability of user-friendly software tools for qPCR data analysis presents a significant challenge for experimental biologists with limited computational skills. To address this issue, we developed Click-qPCR, a user-friendly and web-based Shiny application for qPCR data analysis. Click-qPCR streamlines ΔCq and ΔΔCq calculations using user-uploaded CSV data files. The interactive interface of the application allows users to select genes and experimental groups and perform Welch’s t tests and one-way analysis of variance with Dunnett’s post-hoc test for pairwise and multi-group comparisons, respectively. Results are visualized via interactive bar plots (mean ± standard deviation with individual data points) and can be downloaded as publication-quality images, along with summary statistics. Click-qPCR empowers researchers to efficiently process, interpret, and visualize qPCR data regardless of their programming experience, thereby facilitating routine analysis tasks. Click-qPCR Shiny application is available at https://kubo-azu.shinyapps.io/Click-qPCR/, while its source code and user guide are available at https://github.com/kubo-azu/Click-qPCR.

Preserving biological samples in the field is essential for ensuring high-quality nucleic acid extraction and reliable downstream molecular analyses. Broadly, two main preservation strategies are available: physical preservation, such as flash freezing in liquid nitrogen, which halts enzymatic activity by rapid cooling, and chemical preservation, using stabilizing reagents that inactivate nucleases and protect nucleic acids even at ambient temperatures. This protocol presents a comparative approach using liquid nitrogen and a commercial stabilizing reagent (DNA/RNA Shield, Zymo Research) to preserve tissue from five marine invertebrate species: two cold-water corals, two sponges, and one bivalve. Samples preserved by each method were processed with the AllPrep DNA/RNA Mini kit (Qiagen) to extract both RNA and DNA. RNA quality was assessed using RNA Integrity Number (RIN) scores. The stabilizing reagent preserved high-quality RNA in sponge and bivalve samples but did not prevent RNA degradation in coral tissues, which showed lower RIN scores compared to those preserved in liquid nitrogen. DNA yields were also consistently lower in tissues preserved with DNA/RNA Shield across all species. These findings suggest that DNA/RNA Shield can be a viable alternative to liquid nitrogen for some marine invertebrates, particularly in field conditions where cryopreservation is impractical. However, for cold-water corals, liquid nitrogen remains essential to ensure RNA integrity for transcriptomic analyses and other sensitive molecular applications (e.g., RT-qPCR).

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.

DNA methylation is a fundamental epigenetic mark with critical roles in epigenetic regulation, development, and genome stability across diverse organisms. Whole genome bisulfite sequencing (WGBS) enables single-base resolution mapping of cytosine methylation patterns and has become a standard method in epigenomics. This protocol provides a detailed, step-by-step workflow for WGBS library construction starting from genomic DNA. It includes steps of RNaseA treatment, DNA shearing, end-repair and A-tailing, adapter ligation, bisulfite conversion, library amplification, and quantification. Notably, the method uses self-prepared reagents and customizable index systems, avoiding the constraints of commercial library preparation kits. This flexibility supports cost-effective, scalable methylome profiling, suitable for diverse experimental designs, including high-throughput multiplexed sequencing.

The RNA-guided Cas enzyme specifically cuts chromosomes and introduces a targeted double-strand break, facilitating multiple kinds of genome editing, including gene deletion, insertion, and replacement. Caulobacter crescentus and its relatives, such as Agrobacterium fabrum and Sinorhizobium meliloti, have been widely studied for industrial, agricultural, and biomedical applications; however, their genetic manipulations are usually characterized as time-consuming and labor-intensive. C. crescentus and its relatives are known to be CRISPR/Cas-recalcitrant organisms due to intrinsic limitations of SpCas9 expression and possible CRISPR escapes. By fusing a reporting gene to the C terminus of SpCas9M and precisely manipulating the expression of SpCas9M, we developed a CRISPR/SpCas9M-reporting system and achieved efficient genome editing in C. crescentus and relatives. Here, we describe a protocol for rapid, marker-less, and convenient gene deletion by using the CRISPR/SpCas9M-reporting system in C. crescentus, as an example.

The RNA-guided CRISPR-Cas9 endonuclease has been a transformative tool for laboratory biochemistry with huge potential as a precision therapeutic. This tool site-specifically cleaves double-stranded DNA following the recognition of a unique protospacer-adjacent motif (PAM). Activation of the protein–nucleic acid Cas complex has also been widely recognized to feature an allosteric mechanism dependent on structural remodeling and interdomain crosstalk. Biophysical methods have probed the impact of allosteric perturbations on cleavage and specificity of Cas9, with the aim of engineering enhanced Cas effectors. These studies include Cas9 from thermophilic organisms that edit at higher temperatures and are active in human plasma. Validation of biophysical insights has necessitated the quantitation of DNA cleavage in vitro and, subsequently, the adaptation of established protocols to encompass temperature-dependent function that is evident in extremophilic Cas systems, such as Cas9 from Geobacillus stearothermophilus and the mesophilic SpCas9. This protocol is advantageous for probing functional temperature ranges of DNA cleavage that can theoretically be applied to any Cas-RNP system.

No specific ecological niche has been identified for Serratia proteamaculans. Different strains of the bacterium have been described as opportunistic pathogens of plants, animals, and humans, as plant symbionts, and as free-living bacteria. This makes S. proteamaculans and its particular strains promising models for research, particularly aimed at studying the role of various genes in interspecific interactions. Genome editing is one of the most significant approaches used to study gene function. However, as each bacterial species has its own characteristics, editing methods often need to be adapted. In this study, we adapted a conventional approach based on homologous recombination—the allelic exchange method—to edit the genome of S. proteamaculans, with the aim of examining the biological role of protealysin. Plasmids for recombination were created using the suicidal vector pRE118, and then an auxotrophic Escherichia coli ST18 strain was used to deliver these plasmids to S. proteamaculans through conjugation. This method is valid and can potentially be used to create knockouts, knockins, and point mutations in the S. proteamaculans genome, without the need to insert a selective marker into the genome.

The clustered regularly interspaced short palindromic repeat (CRISPR)/CRISPR-associated protein 9 (Cas9) system is a widely used programmable nuclease system for gene modification in many organisms, including Physcomitrium patens. P. patens is a model species of moss plants, a basal land plant group, which has been extensively studied from the viewpoint of evolution and diversity of green plant lineages. So far, gene modifications by CRISPR/Cas9 in P. patens have been carried out exclusively by the polyethylene glycol (PEG)-mediated DNA transfer method, in which a transgene (or transgenes) is introduced into protoplast cells prepared from protonemal tissues. However, this PEG-mediated method requires a relatively large amount of transgene DNA (typically 30 µg for a single transformation), consists of many steps, and is time-consuming. Additionally, this PEG-mediated method has only been established in a few species of moss. In the current protocol, we succeeded in CRISPR/Cas9-induced targeted mutagenesis of P. patens genes by making good use of the biolistic method, which i) requires amounts of transgene DNA as low as 5 μg for each vector, ii) consists of fewer steps and is time-saving, and iii) is known to be applicable to a wide variety of species of plants.

Transposon-based genetic transformation enables stable transgene integration in avian genomes and is increasingly used in the development of transgenic chickens for enhanced disease resistance, productivity, and biopharmaceutical applications. Conventional transformation techniques in avian biotechnology, including viral vectors and primordial germ cell (PGC) manipulation, are limited by biosafety risks, low efficiency, and technical complexity. This protocol outlines a two-step cloning approach for generating transposon-compatible gene constructs suitable for chicken embryo microinjection. Topoisomerase-based (TOPO) cloning is used as the first step due to its ability to directly clone PCR-amplified products without the need for restriction site-engineered primers while simultaneously producing an insert flanked with EcoRI restriction sites. The insert is subsequently transferred into the transposon vector through EcoRI-mediated restriction digestion and ligation. This approach simplifies construct generation by integrating the speed of TOPO cloning with the precision of restriction cloning, while ensuring compatibility with transposon-mediated integration systems. The protocol is efficient, reproducible, and does not require specialized equipment, providing a practical and scalable tool for gene construct assembly in avian transgenesis research.

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.