微生物学

Isogenic populations of Saccharomyces cerevisiae exhibit significant proliferative heterogeneity, with individual cells within a clonal culture displaying divergent growth rates and metabolic states. Investigating the origins of this variation requires a method to reconstruct the individual histories of cells within the population. This protocol describes a method for single-cell microencapsulation in alginate microspheres to create a physically stable, traceable, three-dimensional genealogical environment. By utilizing the alginate matrix to prevent daughter cell migration, the replicative history of a founder cell can be mathematically reconstructed. This is achieved by correlating the total cell count (N) within a developed microcolony with the total number of accumulated bud scars (n) visualized via confocal microscopy.

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.

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.

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.

Detection of live African swine fever virus (ASFV) has historically relied on the use of primary swine macrophages (PSM). PSM do not replicate and have to be isolated fresh from donor swine. We previously identified that a MA-104 cells (ATCC #CRL-2378.1), a commercially available cell line isolated from African green monkey (Cercopithecus aethiops) kidney epithelial cells, supports the detection of ASFV from field samples with a sensitivity comparable to that of primary swine macrophages. Collection of swine blood or lungs is time costing, which is often not readily available in most veterinary diagnostic laboratories. MA-104 cells could thus be used as substitute for primary swine macrophages to save significant lead time by avoiding the production of primary swine macrophages.

The formation of neutrophil extracellular traps (NETs) is thought to play a critical role in infections and propagating sterile inflammation. Histone citrullination is an essential and early step in NETs formation, detectable prior to the formation of the hallmark extracellular DNA-scaffolded strands. In addition to the classical microscopy method, new technologies are being developed for studies of NETs and their detection, both for research and clinical purposes. Classical microscopy studies of NETs are subjective, low throughput and semi-quantitative, and limited in their ability to capture the early steps. We have developed this novel Imaging Flow Cytometry (IFC) method that specifically identifies and quantifies citrullination of histone H4 as a NETs marker and its relationship with other alterations at nuclear and cellular level. These include nuclear decondensation and super-condensation, multi-lobulated nuclei versus 1-lobe nuclei and cell membrane damage. NETs markers can be quantified following variable periods of treatment with NETs inducers, prior to the formation of the specific extracellular DNA-scaffolded strands. Because these high throughput image-based cell analysis features can be performed with statistical rigor, this protocol is suited for both experimental and clinical applications as well as clinical evaluations of NETosis as a biomarker.