Molecular Biology

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.

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.

Transposon mutagenesis is a powerful tool for investigating gene function in bacteria, particularly in newly discovered species. In this study, we applied the hyperactive EZ-Tn5 transposase system to Pseudomonas argentinensis SA190, an endophytic bacterium known for enhancing plant resilience under drought stress. By leveraging the random amplification of transposon ends (RATE)-PCR method, we successfully mapped the insertion sites of the transposon within the SA190 genome. This approach enabled the precise identification of disrupted genes, offering insights into their roles in bacterial function and interaction with host plants. Our comprehensive protocol, including competent cell preparation, transformation, and insertion site mapping, provides a reliable framework for future studies aiming to explore gene function through mutagenesis.

We recently developed an approach for cell type–specific CRISPR/Cas9 editing and transgene expression using a single viral vector. Here, we present a protocol describing how to design and generate plasmids and adeno-associated viruses (AAVs) compatible with this single-vector gene editing approach. This protocol has four components: (1) guide RNA (gRNA) design to target specific genes of interest, (2) ligation and cloning of CRISPR-competent AAV vectors, (3) production of vector-containing AAVs, and (4) viral titer quantification. The resultant vectors are compatible for use with mouse lines expressing the Cas9 protein from Streptococcus pyogenes (SpCas9) and Cre recombinase to enable selective co-expression of standard neuroscience tools in edited cells. This protocol can produce AAVs of any serotype, and the resulting AAVs can be used in the central and peripheral nervous systems. This flexible approach could help identify and test the function of novel genes affecting synaptic transmission, circuit activity, or morphology with a single viral injection.

This protocol provides a step-by-step approach for generating single-gene knockout in hard-to-transfect suspension immune cell lines like THP1, specifically demonstrated by knocking out the GSDMD gene. By employing CRISPR-Cas9 system delivered via lentivirus, this protocol enables precise gene disruption through targeted single-guide RNAs (sgRNAs). Key steps include designing specific sgRNAs, cloning them into a CRISPR vector, viral packaging, and transducing the target cells, followed by selection and validation. This optimized protocol is particularly useful for functional studies in immune cells, allowing researchers to reliably explore gene function in complex cellular pathways.

Malaria remains a major public health threat, especially in tropical and subtropical regions. Accurate and rapid diagnosis is essential for effective disease management and control, yet conventional malaria diagnostics, including blood smear microscopy using Giemsa staining, PCR, and rapid diagnostic tests (RDTs), are limited by the need for trained personnel, reliance on laboratory infrastructure, and reduced sensitivity at low parasite densities, respectively. This protocol details an innovative, rapid, and economical diagnostic platform combining a simplified Chelex-100 resin-based nucleic acid extraction method with a multiplex loop-mediated isothermal amplification microscanner (LAMP-MS) assay. The malaria diagnostic platform enables simultaneous detection of Plasmodium falciparum (Pf), Plasmodium vivax (Pv), pan-malaria (Pan), and an internal control (IC) within 40 min, from DNA extraction to result interpretation. It demonstrates sensitivity and specificity comparable to traditional PCR-based diagnostics, making it a practical and scalable solution for use in resource-constrained environments.

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.

In many plant species, self-incompatibility (SI) is a mechanism that inhibits inbreeding. SI is gametophytic in the Solanaceae, with specificity determined by S-ribonucleases (S-RNases) in the pistil and S-locus F-box proteins (SLFs) in the pollen. The role of these proteins has been studied extensively in the Solanaceae, often using Petunia as a model. Using degenerate PCR and Sanger sequencing, this protocol identified three SLF sequences from self-incompatible diploid potato (Solanum okadae). While SLFs are well-characterized in model species like Petunia, there is limited sequence data and no standardized protocols for identifying SLFs in non-model species such as S. okadae, hindering broader insights into SI across the Solanaceae. This protocol fills that gap by using degenerate PCR and Sanger sequencing with primers designed from conserved Petunia SLF regions to identify SLF sequences in S. okadae. SLF sequences from 10 distinct Solanaceae members sharing maximum identity with the S2-haplotype of Petunia were used to design two pairs of primers targeting different regions of the target sequence. PCR amplification using designed degenerate primers yielded amplicons that were directly sequenced and joined together to get the partial SLF sequence. It was observed that the S. okadae shared an orthologous relation with the Petunia SLF according to the phylogenetic analysis. These SLFs could be used in future SI breakdown experiments via the competitive interaction route. This protocol, including the primer design, is novel for detecting SLF sequences in S. okadae.

The standard protocols for allelic exchange using homologous recombination deploy suicide vectors with negative selection markers. However, the use of multiple restriction enzymes to generate sticky ends in the vector and the insert for cloning is time-consuming, resource-intensive, and challenging. The advent of next-generation proofreading enzymes is enabling researchers to routinely carry out long-range PCR. Hence, amplifying 5–6 kb of complete low-complex DNA cloning vectors and 2–3 kb of complex genomic regions is much easier. Here, we report a simple, accurate, rapid, and unidirectional approach for chromosomal in-frame gene deletion and complementation by reconstitution of the full-length gene without using any restriction enzymes. The method requires long-range PCR using Phusion polymerase to linearize the vector and amplify the target gene to create a recombinant vector (pRM1) and further inverse PCR amplification of pRM1 to create a recombinant vector (pRM4) with a deleted version of the gene. The cloning steps involve the use of kinase and ligase for phosphorylation and ligation steps, respectively. The recombinant plasmid, pRM4, is finally transformed into electrocompetent cells of Xanthomonas sontii, a gram-negative phytobacterium, for final genomic integration/excision to obtain an in-frame gene deletion mutant (PPL1RM15). Gene reconstitution for complementation is carried out by electroporating the deletion mutant with the recombinant plasmid (pRM1) carrying the wild-type allele. Clean gene mutation, allele restoration, and plasmid excision are confirmed using whole-genome sequencing.