微生物学

Riboswitches are structured non-coding RNA elements that regulate gene expression in response to small molecules; they serve as valuable systems in both public health and biophysical research by elucidating principles around RNA–ligand interactions, structure, and cellular function. Traditional approaches to studying riboswitches have relied on low-throughput techniques such as reporter assays or gel electrophoresis analysis of transcriptional products, which are limited in scalability. In this study, we present a high-throughput protocol to characterize the transcriptional activity of nearly 2,000 natural variants of the fluoride riboswitch in in vitro transcription. Starting with bioinformatics, we compiled a comprehensive dataset of riboswitch variants and then employed massive parallel oligonucleotide synthesis to generate an oligo pool of the riboswitch library. This pool was transcribed in vitro, converted into an Illumina-compatible next-generation sequencing (NGS) library, and analyzed to identify transcriptionally active riboswitch candidates. The workflow integrates natural riboswitch bioinformatic acquisition into a quantitative readout in a single streamlined pipeline, enabling large-scale exploration of transcriptional riboswitch function. This protocol offers a scalable method for mapping genotype-to-function relationships across transcriptional riboswitch families, accelerating the identification of functional variants for desired applications.

RNA sequencing (RNA-seq) has revolutionized transcriptomics, ribosome footprinting, and polysome profiling, providing a wealth of data. Many RNA-based omics typically remove ribosomal RNA (rRNA) or select for messenger RNA (mRNA) prior to sequencing, thereby enriching reads that map to the translationally active part of the transcriptome. Prokaryotic mRNA lacks the 3′ polyadenylated tail, which excludes the use of poly(A)-based selection methods. While commercial rRNA depletion products exist for prokaryotes, their proprietary nature and potential inefficiency with non-model organisms are factors that may limit broad-scale application. To mitigate this issue, we designed DepStep, a consolidated workflow for one-step rRNA depletion using species-specific biotinylated antisense probes for selective hybridization and removal of the target rRNA molecules. As a proof-of-concept, RNA-seq libraries of the psychrophilic gram-negative bacterium Shewanella glacialimarina TZS-4_T were prepared using both DepStep and a commercial rRNA depletion kit for gram-negative bacteria, to which DepStep was benchmarked. DepStep compares favorably to the commercial depletion kit; it removes >98.6% of the rRNA content in the sample, resulting in sequencing libraries where the coding DNA sequence (CDS) reads account for >80% of the total read count. Importantly, DepStep’s cost-per-sample is three times lower than the commercial kit, establishing DepStep as a simple yet cost-effective alternative to commercial solutions.

The genomes of RNA viruses can fold into dynamic structures that regulate their own infection and immune evasion processes. Proximity ligation methods (e.g., SPLASH) enable genome-wide interaction mapping but lack specificity when dealing with low-abundance targets in complex samples. Here, we describe HiCapR, a protocol integrating in vivo psoralen crosslinking, RNA fragmentation, proximity ligation, and hybridization capture to specifically enrich viral RNA–RNA interactions. Captured libraries are sequenced, and chimeric reads are analyzed via a customized computational pipeline to generate constrained secondary structures. HiCapR generates high-resolution RNA interaction maps for viral genomes. We applied it to resolve the in vivo structure of the complete HIV-1 RNA genome, identifying functional domains, homodimers, and long-range interactions. The protocol's robustness has been previously validated on the SARS-CoV-2 genome. HiCapR combines proximity ligation with targeted enrichment, providing an efficient and specific tool for studying RNA architecture in viruses, with broad applications in virology and antiviral development.

Pseudoalteromonas haloplanktis TAC125 is a psychrophilic marine bacterium widely used to study cold adaptation and increasingly exploited as a non-conventional platform for biotechnological applications. The strain harbors the endogenous megaplasmid pMEGA (64.7 kb), whose presence may limit its exploitation as a cell factory, making its elimination advantageous to strain engineering. Traditional plasmid-curing approaches based on chemical and physical agents are often inefficient and unsuitable for stable endogenous replicons, such as pMEGA. Here, we describe a targeted protocol for pMEGA curing in P. haloplanktis TAC125 that combines homologous recombination with paired-termini antisense RNA (PTasRNA) gene silencing. First, a selectable marker cassette is inserted into pMEGA by homologous recombination using a suicide vector, enabling selective discrimination between plasmid-positive and plasmid-cured bacteria. Next, PTasRNA gene silencing technology is applied to target a gene essential for the replication of pMEGA, thereby transiently interfering with its replication and promoting its loss. This approach provides a specific method to cure a highly stable endogenous megaplasmid in a psychrophilic non-conventional bacterium, enabling improved functional studies and strain optimization, establishing a broadly applicable framework for targeted curing across diverse bacterial systems.

Engineering of microbial cells, including E. coli, is essential in prototyping genetic designs used in numerous applications throughout synthetic biology. While many advanced genome editing tools, such as CRISPR-based tools, offer new capabilities with genetically recalcitrant organisms, these tools often do not offer an immediate advantage in readily manipulated microbes, such as E. coli, especially when scarless modifications are not critical. We describe a comprehensive recombineering tutorial that we commonly use for multiplex engineering of E. coli using antibiotic markers. We leverage a group of 15 antibiotic resistance cassettes, most of which can be readily included when designing double-stranded DNA donors intended for recombineering and purchased from several vendors. Using these methods, 10–15 defined modifications to a single host strain can be achieved in less than three weeks, using two-day editing cycles. We discuss sequences and protocols as well as the optimal design of genetic modifications and the associated DNA.

Cyanobacteria have been widely used as model organisms in photobiochemical research and have recently been exploited as hosts in numerous pilot studies to produce valuable biochemicals via genetic and metabolic modifications. Analyzing cellular RNA is a suitable method for studying genetic changes in cells. Several methods have previously been reported for cyanobacterial RNA extraction. However, the majority of these methods rely heavily on phenol and chloroform, which are hazardous. Additionally, these methods are time-consuming and difficult to perform. Using Synechocystis sp. PCC 6803 as a model, this study developed a novel method for extracting total ribonucleic acid (RNA) using standard centrifugation techniques and laboratory chemicals such as citric acid, ethylenediaminetetraacetic acid, sodium dodecyl sulfate, sodium chloride, and tri-sodium citrate dihydrate to extract RNA from cyanobacterial cells. This method does not necessitate the use of hazardous chemicals, especially phenol and chloroform. Furthermore, it is cost-effective since it does not require expensive chemicals. The results of the quantification, purity, and integrity checks show the effectiveness of this method for extracting good-quality RNA. Furthermore, RT-qPCR results demonstrate that the quality of the extracted RNA is suitable for downstream applications.

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.

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.

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.