微生物学

Mal de Río Cuarto disease, caused by a Fijivirus, is a major constraint for maize production in Argentina. The traditional evaluation of resistant hybrids is limited by the low efficiency of natural virus transmission and the lack of standardized field inoculation methods. We developed a protocol that combines laboratory mass-rearing of the planthopper vector Delphacodes kuscheli with a controlled field transmission system. The method involves the synchronized production of large insect populations, acquisition of viruliferous vectors under controlled conditions, and their safe transport to the field using specialized containers. Transmission is achieved through individual cages placed on maize seedlings, ensuring high inoculation pressure under field-like conditions. This protocol enables reliable and reproducible virus transmission, facilitating large-scale screening of maize hybrids and other cereals. Its main advantages are the high throughput of vector production, improved transmission efficiency, and adaptability to diverse experimental designs.

Single-cell RNA sequencing (scRNA-seq) is a powerful technique for exploring cellular heterogeneity and host–pathogen interactions. This protocol details the Zika virus (ZIKV)-targeted scRNA-seq workflow for preparing high-quality single-cell suspensions from the whole brain tissues of neonatal mice, high-quality single-cell sorting, cDNA reverse transcription, amplification, ZIKV enrichment and host transcriptome library preparation, and sequencing dataset integration in downstream analysis to complete the quantification of ZIKV RNA in individual cells.

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.

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.