分子生物学

The release and updating of whole-genome sequences of several representatives of the human opportunistic pathogen Pseudomonas aeruginosa have laid the ground for investigating the mechanisms of antibiotic resistance, biofilm formation, and virulence, while also offering opportunities for researchers to find new therapeutic targets to control this bacterium using a functional genomics approach. However, there is still a lack of detailed protocols describing gene inactivation methods in P. aeruginosa, resulting in failures and extra time spent designing in-house protocols. Here, we introduce a rapid, efficient, and unmarked deletion mutagenesis method combining overlap extension PCR, efficient conjugation, and the traditionally used sacB-based counter-selection procedure. Efficient generation of deletion mutants using this detailed protocol can be easily completed in one week using standard lab reagents. Importantly, this method may be adaptable to other bacteria where the sacB-based counter-selection system works.

Reliable DNA extraction is essential for genetic research on marine species; however, obtaining sufficient DNA from single fish eggs remains challenging. Existing protocols often require optimization to achieve high PCR efficiency. The optimized phenol–chloroform–isoamyl extraction protocol presented in this paper improves DNA yield and quality from individual eggs of Atlantic bluefin tuna (Thunnus thynnus), bogue (Boops bops), saddled seabream (Oblada melanura), and painted comber (Serranus scriba) by modifying buffer volumes, incubation times, and washing steps, following prior micropuncturing of eggs on a glass slide. DNA quality is confirmed by spectrophotometry, PCR amplification of the mitochondrial COI gene, electrophoresis, and Sanger sequencing. This method provides a low-cost and effective approach for species identification from individual fish eggs.

ADP ribosylation (ADPr) is a crucial post-translational modification that plays a vital role in DNA damage repair. Catalyzed by ADP ribose polymerases using NAD⁺ as a substrate, ADPr activates DNA repair pathways rapidly, thereby maintaining genomic integrity. The involvement of ADP ribose hydrolases in this process is significant, as they hydrolyze PAR chains, facilitating the release of ADPr-modified proteins from DNA or other proteins, which is essential for subsequent DNA repair steps. This protocol outlines a high-throughput screening method for identifying inhibitors of ADP ribose hydrolases, utilizing His-Tb-conjugated and ADPr-modified His-ADP ribose polymerase as the signal donor, and GST-d2-conjugated GST-XRCC1 as the signal receptor. The detection of time-resolved fluorescence signals enables efficient evaluation of compounds with potential therapeutic activity against cancer.

Gastric cancer remains a major global health challenge, and reliable prognostic biomarkers are urgently needed to guide treatment decisions. Here, we present a simple and efficient protocol for a novel liquid biopsy approach based on quantifying gastric fluid DNA (gfDNA) collected during routine esophagogastroduodenoscopy (EGD). We have previously shown that gfDNA carries gastric cancer–derived mutations; moreover, its concentration increases with tumor progression and varies according to cancer prognosis. This empirically observed increase in gfDNA may mechanistically stem from enhanced cellular turnover, tissue disorganization, dysbiosis of the local microbiota, and/or fluctuations in immune cell infiltrates. Surprisingly, however, in patients diagnosed with gastric cancer, elevated gfDNA levels were also associated with improved survival. This paradoxical finding may be reconciled by an increased anti-tumor immune cell response in treatment-responsive gastric cancers, as well as by the contribution of non-tumoral DNA from inflammatory processes within the microenvironment of the stomach. Here, we detail a standardized protocol for gastric fluid collection and processing, designed to support downstream gfDNA quantification among other potential molecular applications.

Chromatin modifications regulate genome function by recruiting proteins that control transcription, genome organization, and DNA repair. Identifying the proteins associated with specific chromatin modifications is therefore essential for understanding how these regulatory processes operate. Traditional approaches, including chromatin immunoprecipitation and affinity purification coupled to mass spectrometry, have uncovered many chromatin-associated proteins. However, they often rely on crosslinking and chromatin fragmentation, which can disrupt native chromatin architecture and limit the detection of transient interactions. Here, we describe a proximity-labeling protocol for identifying the chromatin-dependent protein interactome associated with specific chromatin marks, termed ChromID. ChromID uses engineered chromatin readers (eCRs) fused to a promiscuous biotin ligase, which labels proteins in the immediate vicinity of the targeted chromatin mark. The protocol includes in vivo biotin labeling, nuclear extract preparation, streptavidin-based enrichment, and tryptic digestion for downstream LC-MS/MS analysis. The protocol has been validated across multiple cell types and chromatin contexts and can be extended to other chromatin-associated proteins, providing a versatile approach to profile chromatin-associated proteomes within their native cellular environment.

Chemotherapy-resistant persister cells are a major driver of cancer recurrence, yet their epigenetic basis remains poorly characterized. This protocol describes a computational pipeline for identifying DNA-binding factors (DBFs) that are enriched in accessible chromatin that collectively define a persister cell signature (PCS). Starting from single-nucleus ATAC-seq (snATAC-seq) data processed through the 10x Genomics CellRanger ARC pipeline, this protocol covers (1) the creation of a Seurat/Signac object with ATAC peaks, (2) the optional integration of DNA-binding data from the ReMap2022 database as a per-cell chromatin module assay, (3) differential accessibility analysis across clinically defined comparison groups, and (4) identifying and defining the top enriched DBFs as the PCS. This approach is applicable to any snATAC-seq dataset in which cells can be grouped by clinical response, treatment status, or resistance phenotype.

Site-directed mutagenesis is an indispensable molecular biology tool, but traditional methods often suffer from extended reaction time, structural limitations, and variable success rates. This article details three optimized protocols: P3a (primer pairs with 3′-overhangs, version a), P3b, and QuickChange 2.0, which rely on two highly processive DNA polymerases (Platinum SuperFi II and Q5) to accelerate and standardize plasmid engineering. The P3a method utilizes partially complementary primer pairs with distinct 3′-overhangs, achieving ~100% efficiency and enabling seamless cassette mutagenesis (insertion, deletion, and replacement). Building on this, the P3b method introduces specific thermal cycling modifications and a pre-denaturation step to overcome structural barriers resulting from GC-rich sequences. QuickChange 2.0 applies these two advanced polymerases to completely complementary primer pairs, even though the average efficiency decreases to 50%–60%. Replacing Pfu with the highly processive DNA polymerases also reduces PCR time to approximately 2 h. Thus, these new methods are more efficient and rapid than classical QuickChange mutagenesis based on Pfu polymerase.

Mosaic animals are highly valuable for investigating complex biological processes and cell lineages in vivo. Traditional mosaic techniques in Drosophila, such as the FRT/Flp system, rely on exogenous site-specific recombination sequences, preventing their application to unmodified mutant chromosomes or wild-derived strains. Mosaic analysis by gRNA-induced crossing-over (MAGIC) overcomes this limitation by utilizing the CRISPR/Cas9 system to generate targeted double-strand breaks (DSBs) that induce somatic homologous recombination in precursor cells. Here, we describe a comprehensive protocol for applying MAGIC with a newly developed, genome-wide MAGIC kit. This protocol utilizes optimized gRNA-markers with the Qtg2.1 scaffold for high-efficiency clone induction, alongside improved fluorescent labeling strategies for both positive MAGIC (pMAGIC) and negative MAGIC (nMAGIC). The procedure details the genetic crossing schemes, temporal induction of clones, and tissue processing for diverse Drosophila cell types. This method enables convenient mosaic analysis across all chromosomes and allows for the study of pericentromeric genes, deficiency chromosomes, and species-specific alleles in interspecific hybrids.

Expressing large DNA constructs in the native three-dimensional brain microenvironment remains technically challenging. Although viral vectors provide high transduction efficiency and cell-type selectivity, their genetic payload capacity is limited. Various non-viral approaches have been used in brain tissue, but they may compromise tissue viability or require specialised equipment, such as biolistic delivery or electroporation. We present an adapted protocol for delivering the large DNA vector encoding the optical PIEZO1 sensor GenEPi into brain tissue to enable sensor expression in pyramidal neurons. By applying DNA–Lipofectamine liposomes directly to the slice surface, we achieved efficient, minimally invasive transfection of pyramidal neurons in the CA1 and CA3 regions of organotypic hippocampal slices. PIEZO1 sensor expression was detectable as early as 7 days after transfection, increased with longer tissue maintenance, and was sustained for 3–4 weeks in vitro. This protocol describes a cost-effective, non-invasive approach that preserves cell viability and enables investigation of PIEZO1-mediated mechanotransduction in a native brain microenvironment.

Careful dissection of insect gut tissues is essential for microbiome studies to ensure accurate characterization of internal microbial communities and preservation of DNA integrity. Because insect-associated microbiomes are highly sensitive to contamination, effective removal of external microbes prior to dissection is critical to minimize bias in downstream analyses. While ethanol- and bleach-based surface sterilization methods are commonly used, standardized workflows integrating field collection, sterilization, and dissection remain limited. Here, we present a step-by-step protocol for the field collection, surface sterilization, and dissection of gut tissues from the agricultural pest Diaprepes abbreviatus (Coleoptera: Curculionidae), optimized for genomic DNA extraction and microbiome analyses. Using wild-caught specimens, this workflow incorporates a rigorous surface sterilization and dissection strategy that minimizes external contamination while preserving biologically relevant microbial signatures and DNA integrity for downstream microbiome analyses. The protocol provides a standardized framework for insect gut microbiome studies and can be broadly adapted to other wild-caught insect species requiring careful collection, disinfection, and sterile dissection prior to molecular analysis. The protocol integrates field collection and laboratory processing steps into a streamlined workflow that minimizes contamination while preserving tissue integrity for downstream applications.