Microbiology

The ribosome, a complex macromolecular machine, plays a vital role in cellular translation. To investigate its structure and conduct in vitro experiments, isolating the ribosomes from cells is the first step. While isolating ribosomes from bacterial cells is routine, obtaining them from mycobacteria proves challenging due to the protective mycolic acid layer, which hinders cell lysis. In this study, we present a straightforward and efficient protocol for isolating ribosomes from Mycobacterium smegmatis. Additionally, we introduce a co-sedimentation assay using density gradient ultracentrifugation, providing a simple yet powerful method for studying ribosome–protein interactions. The re-association assay also offers a practical approach for obtaining tRNA-free 70S ribosomes and evaluating the anti-association properties of potential ligands. While these assays are commonly used, our protocol stands out for its simplicity, requiring limited specialized instruments. These methods can also be scaled up or down per requirement. By employing sonication for cell rupture and utilizing basic lab equipment for ultracentrifugation-based assays, our method greatly simplifies ribosome isolation and related research.

Orthoflavivirus is an enveloped, positive-stranded RNA virus that buds into the endoplasmic reticulum (ER) lumen. The budded virus particles are subsequently transported to the Golgi apparatus and secreted into the extracellular environment via the conventional secretion pathway. In this protocol, we describe a method for monitoring the secretion of Orthoflavivirus particles from the ER. To visualize intracellular membrane trafficking, we combine two distinct imaging techniques: the retention using selective hooks (RUSH) system and the split green fluorescent protein (GFP) system. In this approach, GFP11, a peptide tag fused to prME, the outer coat structural protein of Japanese encephalitis virus particles, was co-expressed in HeLa cells along with two additional components: GFP1-10 fused to a streptavidin-binding peptide and a hook construct consisting of streptavidin fused to the ER retention sequence KDEL. Time-lapse imaging was performed after the addition of biotin, which releases the captured GFP-labeled subviral particles from the ER. This method enables synchronized visualization of intracellular subviral particle trafficking and serves as a valuable tool for analyzing the maturation process of Orthoflavivirus particles within cells.

Microbial biofilms are structured communities of microorganisms embedded in a self-produced extracellular matrix, adhering to surfaces. These biofilms enhance bacterial resistance to antibiotics, immune responses, and environmental stress. Current microscopy techniques, such as scanning electron microscopy (SEM), confocal laser scanning microscopy (CLSM), and fluorescence microscopy, are commonly used to visualize and differentiate biofilms. However, their high cost and complexity often render them impractical. In contrast, simpler methods like crystal violet and Congo red staining are limited in distinguishing bacterial cells from the biofilm matrix. This study introduces a cost-effective, dual-staining method using Maneval’s stain to visualize and differentiate microbial biofilms. It requires only basic equipment and minimal reagents, making it ideal for routine use in clinical diagnosis and microbial research.

Cricket paralysis virus (CrPV), a member of the family Dicistroviridae, is a single-stranded positive-sense RNA virus that primarily infects arthropods. Some members of the dicistrovirus family, including the honey bee viruses Israeli acute paralysis virus and Acute bee paralysis virus and the shrimp-infecting Taura syndrome virus, pose significant threats to agricultural ecosystems and economies worldwide. Dicistrovirus infection in Drosophila is used as a model system to study fundamental insect–virus–host interactions. The availability of a CrPV infectious clone allows controlled manipulation of the viral genome at a molecular level. Effective viral propagation and titration techniques are crucial for understanding the pathogenesis and epidemiology of dicistrovirus infections. Traditional methods for assessing viral titers, such as plaque assays, are unsuitable for CrPV, since Drosophila tissue culture cells like Schneider 2 cells cannot readily form adherent plaques. Here, we present a streamlined protocol for generating a recombinant virus from a CrPV infectious clone, propagating the virus in S2 cells and titering the virus by an immunofluorescence-based focus-forming assay (FFA). This protocol offers a rapid and reliable approach for generating recombinant viruses, viral amplification, and determining CrPV titers, enabling efficient investigation into viral biology and facilitating the development of antiviral strategies.

Cryo-electron microscopy (cryo-EM) is a powerful technique capable of investigating samples in a hydrated state, compared to conventional high-vacuum electron microscopy that requires samples to be completely dry. During the drying process, numerous features and details may be lost due to damage caused by dehydration. Cryo-EM circumvents these problems by cryo-fixing the samples, thereby retaining the intact and original features of hydrated samples. This protocol describes a step-by-step cryo-scanning electron microscopy (cryo-SEM) experimental procedure with Chlorella sorokiniana as the subject. By employing filter paper as the sample substrate, we propose a simple and reliable method for cryo-fixation and freeze-fracture of Chlorella sorokiniana in water suspension. The advantage of using filter paper as a substrate lies in its ability to support a thin film of sample, enabling a cold knife to make a cut effortlessly and produce a clean freeze-fractured surface for SEM investigation. By following the approach described in this protocol, both the internal structure and surface morphology of Chlorella sorokiniana can be easily resolved with high quality. This protocol is highly versatile and can be applied to samples dispersed in water or solvents, including cyanobacterial cells, algal cells, and any kind of sample that can be adsorbed onto filter paper.

In modern science, the exchange of scientific material between different institutions and collaborating working groups constitutes an indispensable endeavor. For this purpose, bacterial strains are frequently shipped to collaborators to advance joint research projects. Bacterial strains are usually safely shipped as cultures on solid medium, whereas the shipment of liquid cultures requires specific safety measures due to the risk of leakage. Cyanobacterial cultures are frequently maintained as liquid stock cultures, and this problem typically arises. This protocol describes a new method for the shipment of liquid cyanobacterial stock cultures by agarose gel embedding (SCAGE). More specifically, a cyanobacterial culture is mixed with low-melting agarose and cast into sterile plastic bags, resulting in a thin, solid cyanobacterial agarose gel (cyanogel) that can be easily shipped. After delivery, subsequent regeneration of the cyanogel material in liquid media results in full recovery of the examined bacterial strains. Thus, the packaging method devised in the present study comprises an innovative technique to facilitate the shipment of bacterial strains, whilst eliminating previously encountered issues like cell culture leakage.

Candida albicans is the most common human fungal pathogen, able to reside in a broad range of niches within the human body. Even though C. albicans systemic infection is associated with high mortality, the fungus has historically received relatively little attention, resulting in a lack of optimized molecular and fluorescent tools. Over the last decade, some extra focus has been put on the optimization of fluorescent proteins (FPs) of C. albicans. However, as the FPs are GFP-type, they require an aerobic environment and a relatively long period to fully mature. Recently, we have shown the application of a novel type of fluorogen-based FP, with an improved version of fluorescence activating and absorption shifting tag (iFAST), in C. albicans. Due to the dynamic relation between iFAST and its fluorogens, the system has the advantage of being reversible in terms of fluorescence. Furthermore, the combination of iFAST with different fluorogens results in different spectral and cellular properties, allowing customization of the system.

The sensing of and response to ambient chemical gradients by microorganisms via chemotaxis regulates many microbial processes fundamental to ecosystem function, human health, and disease. Microfluidics has emerged as an indispensable tool for the study of microbial chemotaxis, enabling precise, robust, and reproducible control of spatiotemporal chemical conditions. Previous techniques include combining laminar flow patterning and stop-flow diffusion to produce quasi-steady chemical gradients to directly probe single-cell responses or loading micro-wells to entice and ensnare chemotactic bacteria in quasi-steady chemical conditions. Such microfluidic approaches exemplify a trade-off between high spatiotemporal resolution of cell behavior and high-throughput screening of concentration-specific chemotactic responses. However, both aspects are necessary to disentangle how a diverse range of chemical compounds and concentrations mediate microbial processes such as nutrient uptake, reproduction, and chemorepulsion from toxins. Here, we present a protocol for the multiplexed chemotaxis device (MCD), a parallelized microfluidic platform for efficient, high-throughput, and high-resolution chemotaxis screening of swimming microbes across a range of chemical concentrations. The first layer of the two-layer polydimethylsiloxane (PDMS) device comprises a serial dilution network designed to produce five logarithmically diluted chemostimulus concentrations plus a control from a single chemical solution input. Laminar flow in the second device layer brings a cell suspension and buffer solution into contact with the chemostimuli solutions in each of six separate chemotaxis assays, in which microbial responses are imaged simultaneously over time. The MCD is produced via standard photography and soft lithography techniques and provides robust, repeatable chemostimulus concentrations across each assay in the device. This microfluidic platform provides a chemotaxis assay that blends high-throughput screening approaches with single-cell resolution to achieve a more comprehensive understanding of chemotaxis-mediated microbial processes.

The bacterial membrane vesicles (MVs) are non-replicative, nanoscale structures that carry specific cargos and play multiple roles in microbe–host interactions. An appropriate MV isolation method that mimics complex pathogen infections in vivo is needed. After bacterial MVs extraction, flagella or pili can be frequently observed along with MVs by transmission electron microscope (TEM). Recently, MVs from Pseudomonas aeruginosa were found to coexist with Pf4 phages, and this MV–phages complex exhibited a different impact on host cell innate immunity compared with MVs or phages solely. The presence of this MVs–phages complex simulates the real condition of complex pathogen infections within the host. This protocol outlines the extraction of the MVs and Pf4 phages complex of P. aeruginosa PAO1, including the respective isolation and qualification approaches. Our step-by-step bacterial MVs–phages complex extraction protocol provides valuable insights for further studying microbe–host cell interactions and the development of novel phage therapies.

Diseases caused by trypanosomatid parasites remain a significant unmet medical need for millions of people globally. Trypanosomatid parasites such as Trypanosoma cruzi and subspecies of Trypanosoma brucei cause Chagas disease and human African trypanosomiasis (HAT), respectively. Although efforts to find novel treatments have been successful for HAT, Chagas disease is still treated with decades-old therapies that suffer from long treatment durations and severe safety concerns. We recently described the identification and characterization of the cyanotriazole compound class that kills trypanosomes, in vitro and in vivo, by selective inhibition of the trypanosome nuclear topoisomerase II enzyme. To evaluate whether inhibition of the topoisomerase II enzyme led to parasite death due to lethal double-strand DNA breaks, we developed assays for detecting DNA damage in both intracellular amastigotes of T. cruzi and bloodstream-form T. brucei by using the canonical DNA damage marker γH2A. Herein, this article describes the protocols for detecting DNA damage using an immunofluorescence assessment of γH2A by microscopy in trypanosome parasites.