Microbiology

RNA-binding proteins (RBPs) have pleiotropic roles in modulating the physiology of both eukaryotic and prokaryotic cells, enabling them to adapt to environmental variations. The importance of RBPs has led to the development of a variety of methods aiming to identify them. However, most of these approaches have primarily been implemented and optimized in eukaryotic systems. To both uncover novel RBPs involved in Bacillus subtilis sporulation and capture their RNA-binding ability dynamically, we adapted the orthogonal organic phase separation technique (OOPS), which had previously been used in Escherichia coli to reveal its RNA-binding proteome (RBPome). We optimized the UV cross-linking process used to stabilize RNA–protein interactions in vivo and the bacterial lysis process to overcome the robust cell wall of Gram-positive sporulating cells. RNA–protein complexes are then recovered after phase separation steps using guanidinium thiocyanate–phenol–chloroform, and RNA-associated proteins are identified and label-free-quantified by liquid chromatography–mass spectrometry. Collecting samples at various time points during sporulation further enables tracking the dynamics of the RBPome. In addition to being applicable to bacteria and requiring minimal starting material, this method has provided a comprehensive map of the RBPome during sporulation, refining the roles of known factors and revealing new players.

The discovery of broad-spectrum antiparasitic agents relies on the ability to evaluate drug efficacy under harmonized in vitro conditions across related species. However, current drug screening pipelines for intraerythrocytic parasites are constrained by the use of species-specific media with distinct nutrient compositions and serum sources, which hinder direct comparison of compound potency. To address this gap, we describe a unified erythrocytic culture system based on DMEM/F12 supplemented with 20% fetal bovine serum (DFS20), which supports robust asexual growth of multiple Plasmodium falciparum strains (3D7, Dd2, HB3, V1/S), Babesia duncani, Babesia divergens (Rouen 87), and Babesia MO1. Parasite proliferation and morphology in DFS20 are comparable to those observed in established species-specific media such as RPMI-1640 for P. falciparum and B. divergens and HL-1/Claycomb/DMEM/F12/SFM for B. duncani, while eliminating reliance on undefined or discontinued proprietary components. Importantly, this standardized medium enables cross-species growth inhibition assays for direct comparison of drug efficacy under identical conditions. Using this platform, we recently screened dihydrotriazines and biguanides targeting the conserved DHFR-TS enzymes and identified potent antifolate candidates with broad-spectrum activity against Babesia and Plasmodium species. For B. duncani, which is uniquely supported by both a continuous in vitro human erythrocyte culture system and a lethal in vivo mouse infection model, integration with the in-culture and in-mouse (ICIM) pipeline enables systematic validation of pharmacodynamics, pharmacokinetics, resistance, and toxicity. This unified DFS20-based system establishes a scalable and reproducible protocol for harmonized drug efficacy evaluation across intraerythrocytic parasites and provides a foundation for the development and prioritization of pan-antiparasitic therapies.

The emergence of antimicrobial resistance and the persistence of Klebsiella pneumoniae biofilms represent significant challenges to public health. Hermetia illucens (HI) larvae are considered a sustainable reservoir of novel bioactive compounds. This protocol details a method for extracting fatty acids from HI larvae fat (AWME3 fraction) and studying their effects on multidrug-resistant and hypervirulent Klebsiella pneumoniae strains. Effects are evaluated by crystal violet and ethidium bromide uptake assays, motility assays (swimming, swarming, and twitching), minimal biofilm inhibitory and eradication concentration tests (MBIC/MBEC) for single, mixed, and mature biofilms, light, fluorescence, and scanning electron microscopy imaging, and microbial adhesion to solvents (MATS). This protocol offers a reliable methodology for evaluating the anti-biofilm and anti-virulence properties of natural compounds.

This protocol describes an easy, quick, cheap, and effective method for the purification and concentration of bacteriophages (phages) produced in rich culture media, meeting the quality criteria required for structural analyses. It is based on a tube dialysis system that replaces the classical but expensive and tedious density gradient ultracentrifugation step. We developed this protocol for the Oenococcus oeni bacteriophage OE33PA from its amplification to imaging by negative stain electron microscopy (NS-EM). The host bacterium, O. oeni, is a lactic acid bacterium that lives in harsh oenological ecosystems and grows only in rich and complex media such as Man–Rogosa–Sharpe (MRS) or fruit juice-based media in laboratory conditions. This raises experimental challenges in pure and concentrated phage preparations for further uses such as structure-function studies.

A prompt and accurate diagnosis of respiratory viral diseases in intensive poultry production is essential to safeguard animal health and ensure the economic sustainability of farms. Currently, much effort is being devoted to preventing the spread of the avian influenza virus in farms. However, the diagnosis of other relevant respiratory viruses, as infectious laryngotracheitis virus (ILTV), is also crucial. Indeed, infection by ILTV does lead to substantial economic losses due to high morbidity, reduced growth, and decreased productivity, making rapid detection a critical aspect of disease control. Conventional diagnostics, including PCR and qPCR, while sensitive and specific, require expensive laboratory infrastructure and well-trained personnel, limiting their deployment in field settings where immediate intervention is most valuable. To address these limitations, this protocol describes a portable molecular diagnostic workflow based on loop-mediated isothermal amplification (LAMP) combined with gold nanoparticle–DNA nanoprobes for specific and visual detection of ILTV directly at the point of need. Gold nanoparticles synthesized via the Turkevich method are functionalized with thiolated DNA probes, which undergo full-length, sequence-specific hybridization to LAMP amplicons, enabling a naked-eye colorimetric readout. The procedure integrates streamlined steps for DNA probe preparation, nanoparticle synthesis and assembly, and minimal sample processing, compatible with on-farm deployment. Results obtained with this workflow on field samples demonstrated 100% sensitivity and specificity, matching the performance of gold-standard assays. This approach offers a rapid, cost-effective, and equipment-free detection system of viral pathogens, enabling timely decision-making for disease containment and biosecurity. By overcoming the barriers of conventional diagnostics, this protocol enables producers with powerful tools for efficient monitoring and response to respiratory outbreaks in poultry farms.

This article presents an efficient protocol for refolding recombinant proteins that are prone to aggregation and form inclusion bodies during expression in Escherichia coli. As a model system, the homolog of CRISPR-associated effector protein CasV-M was investigated. The key element of the developed approach is refolding directly on a metal-affinity Ni-TED (N,N,N´-tris(carboxymethyl)ethylendiamine) resin using a dual-gradient system: a stepwise reduction in the concentration of the chaotropic agent combined with a simultaneous increase in the concentration of a mild nonionic detergent. This combination ensures spatial separation of protein molecules, minimizes aggregation, and promotes the recovery of the native conformation. The resulting method appears to be an alternative to conventional refolding strategies, with potential improvements in the reproducibility and yield of soluble protein compared to dialysis or dilution. The proposed approach can be extended to a broad range of aggregation-prone proteins and is considered a promising strategy for obtaining otherwise insoluble recombinant proteins.

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.

Accurate profiling of soil and root-associated bacterial communities is essential for understanding ecosystem functions and improving sustainable agricultural practices. Here, a comprehensive, modular workflow is presented for the analysis of full-length 16S rRNA gene amplicons generated with Oxford Nanopore long-read sequencing. The protocol integrates four standardized steps: (i) quality assessment and filtering of raw reads with NanoPlot and NanoFilt, (ii) removal of plant organelle contamination using a curated Viridiplantae Kraken2 database, (iii) species-level taxonomic assignment with Emu, and (iv) downstream ecological analyses, including rarefaction, diversity metrics, and functional inference. Leveraging high-performance computing resources, the workflow enables parallel processing of large datasets, rigorous contamination control, and reproducible execution across environments. The pipeline’s efficiency is demonstrated on full-length 16S rRNA gene datasets from yellow pea rhizosphere and root samples, with high post-filter read retention and high-resolution community profiles. Automated SLURM scripts and detailed documentation are provided in a public GitHub repository (https://github.com/henrimdias/emu-microbiome-HPC; release v1.0.2, emu-pipeline-revised) and archived on Zenodo (DOI: 10.5281/zenodo.17764933).

Toxoplasma gondii is an apicomplexan parasite that infects a wide variety of eukaryotic hosts and causes toxoplasmosis. The cell cycle of T. gondii exhibits a distinct architecture and regulation that differ significantly from those observed in well-studied eukaryotic models. To better understand the tachyzoite cell cycle, we developed a fluorescent ubiquitination-based cell cycle indicator (FUCCI) system that enables real-time visualization and quantitative assessment of the different cell cycle phases via immunofluorescence microscopy. Quantitative immunofluorescence and live-cell imaging of the ToxoFUCCI^S probe with specific cell cycle markers revealed substantial overlap between cell cycle phases S, G₂, mitosis, and cytokinesis, further confirming the intricacy of the apicomplexan cell cycle.

During herpesvirus replication, capsids are assembled inside the nucleus and translocated into the cytosol by a non-canonical nucleocytoplasmic export process termed nuclear egress. Traditional methods of measuring nuclear egress rely on imaging-based technologies such as confocal and electron microscopy. These techniques are labor-intensive, limited by the number of cells that can be examined, and may not accurately represent the entire population, generating a potential bias during data interpretation. To overcome these problems, we have developed a flow cytometry–based method to measure HSV-1 nuclear egress that we termed FLARE (FLow cytometry–based Assay of nucleaR Egress). This assay uses a double fluorescent reporter system, utilizing HSV-1-tdTomato to identify infected cells and an Alexa Fluor-488-conjugated, capsid-specific antibody to detect cytosolic capsids, thereby distinguishing infected cells with nuclear egress from those without it. This assay provides more quantitative results than traditional methods, enables large-scale high throughput, and can be adapted for use with other herpesviruses.