Molecular Biology

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.

Transient transfection is commonly used for the commercial production of adeno-associated viral particles for gene therapy. In this process, packaging cells such as HEK293 cells are transfected with three plasmids, including the Rep/Cap plasmid, the Helper plasmid, and the gene-of-interest plasmid containing the transgene/gene therapy product. The combination of these plasmids allows for the robust production of recombinant adeno-associated viral particles. As a result, the concentration of these plasmids plays a critical role in viral production and must be accurately assessed. Typically, A₂₆₀/A₂₈₀ readings are utilized to measure plasmid titer; however, this approach lacks accuracy and specificity and is susceptible to matrix interference. To address these shortcomings, a digital droplet PCR method was developed to titer plasmids. This method uses a combined restriction digest/PCR protocol to linearize the plasmid template and evaluate copy numbers of a plasmid-specific gene. Qualification demonstrated that the method is highly accurate, specific to plasmid DNA, and impervious to matrix interference.

CRISPR/Cas9-based genome editing is a powerful approach for functional genomics and bioenergy research in woody plants. However, conventional single guide RNA (gRNA) strategies predominantly generate small insertions or deletions that may not fully disrupt gene function and often require extensive sequencing for mutation identification. Here, we present an optimized protocol for the efficient generation of large-fragment deletion mutants in Populus tremula × P. alba clone INRA 717-1B4 using a dual-gRNA CRISPR/Cas9 system. Co-expression of two gRNAs flanking the target region induces double-strand breaks at both sites, enabling the deletion of the intervening genomic fragment, typically larger than 50 bp. This protocol describes step-by-step procedures for gRNA design, vector construction, Agrobacterium-mediated transformation, plant regeneration, and molecular validation. Using the PtFBX230 gene as a representative target, large deletions are readily identified by conventional PCR and agarose gel electrophoresis, enabling rapid and cost-effective genotyping. This protocol can be readily adopted to other loci in poplar and related woody species and provides a robust framework for generating null alleles to support functional genomics and bioenergy-related trait engineering in woody plants.

Nanobodies are recombinant single-domain antibodies (VHHs) derived from the heavy chain–only subset of camelid immunoglobulins that can be reverse-engineered into bivalent antibodies by fusion to immunoglobulin Fc constant regions. Mammalian cells are the system of choice to produce VHH-Fcs to ensure authentic folding and post-translation glycosylation of the expressed VHH-Fcs. In a recent project to find neutralising VHH-Fc binders to the spike proteins of SARS-CoV-2 viruses, we identified a need for rapid expression and purification of multiple VHH-Fc fusions from nanobodies selected by phage display. Here, we present a protocol for the construction of expression vectors by parallel ligase-independent cloning, transient small-scale expression in mammalian cells (4 mL culture volume), screening antigen-binding activity, and midi-scale purification (30 mL culture volume) for downstream activity assays. The workflow is completely transferable between different vector formats, of which three are described herein: Fc fusion dimers, monomeric CD4 fusions, and His-tagged monomers.