分子生物学

N⁶-methyladenosine (m⁶A) is the most abundant internal modification in mRNA and is regulated primarily by the balance between the METTL3 methylase complex and two demethylases, FTO (fat mass and obesity-associated protein) and ALKBH5 (α-ketoglutarate-dependent dioxygenase alkB homolog). Reflecting this prevalence, m⁶A participates in virtually every step of RNA metabolism, influencing a wide range of physiological and pathological processes. The first step in studying m⁶A is genome-wide mapping, typically performed by m⁶A-seq, which sequences RNA fragments immunoprecipitated with an m⁶A-specific antibody. This is followed by identification of RRACH motifs (R = A or G; H = A, C, or U) within these sequences, with m⁶A being located at the third nucleotide. The second step involves mutating the putative m⁶A sites to establish a causal link between the modification and downstream biological effects. Since the mapping step has been covered in several detailed protocols, this article focuses on the second step—mutagenesis of RRACH motifs and subsequent functional analysis of the mutations by ectopic expression. The 3′ untranslated region (UTR) of the mouse Runx2 gene is used as an example. The mutant and wild-type sequences are inserted into a luciferase reporter vector and transfected into 293FT cells to evaluate how loss of m⁶A affects luciferase protein levels. The same reporter plasmids are also used in an RNA stability assay with a transcription inhibitor. Although site-specific demethylation of endogenous mRNA would be preferable, it remains technically challenging despite many attempts. Thus, ectopic expression of the mutated target gene remains a widely used and practical alternative.

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.

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.

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.

This protocol describes a standardized and economically accessible method for synthesizing mRNA-encapsulated lipid nanoparticles using routine laboratory equipment, including precision syringe pumps, Y-shaped glass microfluidic chips, and silicone tubing. Designed to address the cost and accessibility limitations of commercial microfluidic platforms, the system achieves performance metrics comparable to high-end devices while reducing equipment costs by 90%. By systematically optimizing hydrodynamic parameters (total flow rate: 12 mL/min; lipid-to-aqueous phase ratio: 3:1), the protocol enables consistent production of lipid nanoparticles with key quality attributes: high mRNA encapsulation efficiency (≥ 80%), narrow particle size distribution (100–120 nm, polydispersity index ≤ 0.2), and excellent storage performance (≥ 7 days at 4 °C ).

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.

Lipid nanoparticles (LNPs) are powerful carriers for nucleic acid delivery, but plasmid DNA-loaded LNPs (pDNA-LNPs) have been limited by inflammation and toxicity. We showed that standard pDNA-LNPs activate the cGAS–STING pathway, leading to severe immune responses and mortality in mice. To overcome this, we co-loaded nitro-oleic acid (NOA), an endogenous STING inhibitor, into pDNA-LNPs. NOA-pDNA-LNPs mitigated inflammation, enabled safe in vivo delivery, and supported sustained gene expression for months. Here, we present a detailed protocol for producing and characterizing NOA-pDNA-LNPs to facilitate safer, long-term gene delivery applications.

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.

Chromatin-associated RNAs (caRNAs) have been increasingly recognized as key regulators of gene expression and genome architecture. A few technologies, such as ChRD-PET and RedChIP, have emerged to assess protein-mediated RNA–chromatin interactions, but each has limitations. Here, we describe the TaDRIM-seq (targeted DNA-associated RNA and RNA–RNA interaction mapping by sequencing) technique, which combines Protein G (PG)-Tn5-targeted DNA tagmentation with in situ proximity ligation to simultaneously profile caRNAs across genomic regions and capture global RNA–RNA interactions within intact nuclei. This approach reduces the required cell input, shortens the experimental duration compared to existing protocols, and is applicable to both mammalian and plant systems.

Human coronavirus OC43 (HCoV-OC43) is an endemic “common cold” coronavirus widely used to study fundamental aspects of coronavirus biology and to test therapeutic interventions. Recently, we used a yeast-based reverse genetics strategy to create recombinant HCoV-OC43 and fluorescent reporter viruses. We assembled a DNA copy of the HCoV-OC43 genome from six linear dsDNA fragments and a linearized yeast centromeric plasmid/bacterial artificial chromosome (YCpBAC) vector in Saccharomyces cerevisiae using transformation-associated recombination (TAR). Reporter genes encoding mCardinal fluorescent protein or histone H2B fused to mClover3 (mClover-H2B) or mRuby3 (mRuby-H2B) were inserted into an intergenic region between the HCoV-OC43 M and N genes. Assembled full-length HCoV-OC43-encoding plasmids were delivered into permissive mammalian cells to initiate viral gene expression, genome replication, and production of infectious progeny. This technique allows for the precise mutagenesis of any area of the HCoV-OC43 genome using homologous recombination, yielding genetically defined reference plasmids for the future generation of HCoV-OC43 virus stocks.