Microbiology

Genetic modification is essential for understanding parasite biology, yet it remains challenging in Plasmodium. This is partially due to the parasite’s low genetic tractability and reliance on homologous recombination, since the parasites lack the canonical non-homologous end-joining pathway. Existing approaches, such as the PlasmoGEM project, enable genome-wide knockouts but remain limited in coverage and flexibility. Here, we present the Plasmodium berghei high-throughput (PbHiT) system, a scalable CRISPR-Cas9 protocol for efficient genome editing in rodent malaria parasites. The PbHiT method uses a single cloning step to generate vectors in which a guide RNA (gRNA) is physically linked to short (100 bp) homology arms, enabling precise integration at the target locus upon transfection. The gRNA also serves as a unique barcode, allowing pooled vector transfections and identification of mutants by downstream gRNA sequencing. The PbHiT system reliably recapitulates known mutant growth phenotypes and supports both knockout and tagging strategies. This protocol provides a reproducible and scalable tool for genome editing in P. berghei, enabling both targeted functional studies and high-throughput genetic screens. Additionally, we provide an online resource covering the entire P. berghei protein-coding genome and describe a step-by-step pooled ligation approach for large-scale vector production.

Since its introduction, the CRISPR/Cas9 system has been used in many organisms for precise and rapid genome editing, as well as for editing multiple genes at once. This targeted mutagenesis makes it easy to analyze the function of a gene of interest (goi). The standard method for genetic manipulation of the model organism Neurospora crassa has been homologous recombination. It is well established and widely used to create knock-out or overexpression mutants. The recently developed CRISPR/Cas9 system is an addition to the toolkit for genetically manipulating N. crassa. For this protocol, a strain stably expressing the Cas9 endonuclease is required. After designing the gRNA with the online tool CHOP-CHOP, a synthetic gRNA is used to transform macroconidia via electroporation. Combining the goi-gRNA with a gRNA targeting the csr-1 gene as a selection marker allows for easy identification of colonies with mutations at the target site of the goi, since the obtained resistance to Cyclosporin A (CsA) allows for selecting editing events. The mutation type can be detected by PCR of the edited gene region followed by Sanger sequencing. This system is fast and easy to handle, offering an attractive alternative to homologous recombination, especially for targeting multiple genes simultaneously.

This protocol presents a modified version of the Filterprep method originally reported in New Biotechnology, adding an optional step to reduce endotoxin levels. Filterprep is a simple, rapid, and cost-effective approach to plasmid DNA purification that couples ethanol precipitation with a single spin-column filtration step, eliminating chaotropic salts and silica binding. The formulations and parameters are fully transparent and do not rely on proprietary buffers, using only standard laboratory reagents and widely available miniprep columns. Under matched conditions, the method recovers high-purity plasmid DNA with yields up to fivefold higher than those obtained with representative commercial midiprep kits. The workflow is readily adoptable in most molecular biology laboratories and, under routine conditions, can be completed in approximately 40 min. The resulting DNA is suitable for molecular cloning, PCR, sequencing, and other downstream biochemical applications. Endotoxin is a lipopolysaccharide (LPS) found in the outer membrane of Gram-negative bacteria and may carry over during plasmid preparation. For experiments requiring lower endotoxin input, an optional modification resuspends the DNA pellet in a Triton X-114 wash buffer before column loading to decrease lipopolysaccharide carryover. The method is modular and extensible, allowing adjustment of precipitation and wash conditions, variation in the number of washes, selection of alternative column formats, and integration of endotoxin-reduction modules without altering the core principle. These features facilitate troubleshooting and quality control, enable scaling from routine batches to larger culture volumes and higher throughput, and allow seamless integration with existing workflows.

The exploration of microbial genomes through next-generation sequencing (NGS) and genome mining has transformed the discovery of natural products, revealing an immense reservoir of previously untapped chemical diversity. Bacteria remain a prolific source of specialized metabolites with potential applications in medicine and biotechnology. Here, we present a protocol to access novel biosynthetic gene clusters (BGCs) that encode natural products from soil bacteria. The protocol uses a combination of Oxford Nanopore Technology (ONT) sequencing, de novo genome assembly, antiSMASH for BGC identification, and transformation-associated recombination (TAR) for cloning the BGCs. We used this protocol to allow the detection of large BGCs at a relatively fast and low-cost DNA sequencing. The protocol can be applied to diverse bacteria, provided that sufficient high-molecular-weight DNA can be obtained for long-read sequencing. Moreover, this protocol enables subsequent cloning of uncharacterized BGCs into a genome engineering-ready vector, illustrating the capabilities of this powerful and cost-effective strategy.

Genome-walking protocols have been extensively used to clone unknown genomic sequences next to known DNAs. Existing genome-walking protocols need further improvement in methodological specificity or operation. Here, we describe a novel genome-walking protocol based on fusion primer–driven racket PCR (FPR-PCR). FPR-PCR involves four sequence-specific oligos (SSO), SSO1, SSO2, SSO3, and SSO4, which are sequentially chosen from known DNA in the direction 5’→3’. The fusion primer, mediating primary FPR-PCR, is generated by attaching SSO3 to the 5’ end of SSO1. The SSO3 encourages the target DNA of primary PCR to form a racket-like structure by mediating intra-strand annealing. SSO2 and SSO4 are directly used as sequence-specific primers (SSP) in secondary FPR-PCR, which selectively amplifies this racket-like DNA. This protocol was verified by cloning several unknown genomic sequences. Compared to traditional PCRs, FPR-PCR offers the advantages of higher specificity and fewer rounds, primarily attributed to the omission of arbitrary walking primers typically required in traditional methods.

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.

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.

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.

Endophytic actinomycetes, particularly Streptomyces species, have gained significant attention due to their potential to produce novel bioactive compounds. In this study, we isolated and characterized an endophytic Streptomyces sp. VITGV100 from the tomato plant (Lycopersicon esculentum), employing the direct streak method and whole-genome sequencing. A genome analysis was done to uncover its biosynthetic potential and identify indole-type compounds. The strain's secondary metabolite production was evaluated through GC–MS analysis, and its antimicrobial activity was tested against selected human pathogenic bacteria. Our protocol outlines a comprehensive approach, describing the isolation and extraction of metabolites and genome mining for indole-type compounds. This isolate has potential pharmaceutical applications, accelerating the discovery of novel indole-type bioactive compounds.

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.