Microbiology

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.

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.

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.

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.

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.

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.

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.

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.

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.

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.