Microbiology

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.

The Plasmodium parasites that cause malaria undergo an obligate, asymptomatic developmental stage in the host liver before initiating the symptomatic blood-stage infection. The parasite liver stage is a key intervention point for antimalarial chemoprophylaxis: successful targeting of liver-stage parasites prevents disease development in individuals and can help to reduce parasite transmission in populations, as the gametocyte forms that transmit infection to mosquitos are exclusively found in the blood stage. Antimalarial drugs that can target multiple parasite stages are thus highly desirable, and one emerging cellular target for such multistage active compounds is the process of protein synthesis or translation. Quantitative study of liver stage translation, and thus mechanistic evaluation of translation inhibitors against liver stage parasites, is not amenable to the methods allowing quantification of asexual blood stage translation, such as radiolabeled amino acid incorporation or lysate-based translation of reporter transcripts. Here, we present a method using o-propargyl puromycin (OPP) labeling of host and parasite nascent proteomes in the P. berghei-HepG2 infection model, followed by automated confocal image acquisition and computational separation of P. berghei vs. H. sapiens nascent proteome signals to allow simultaneous readout of the effects of translation inhibitors on both host and parasite. This protocol details our HepG2 cell culture and infected monolayer handling optimized for microscopy, our OPP labeling workflow, and our approach to automated confocal imaging, image processing, and data analysis.

Key features

• Uses the o-propargyl puromycin labeling technique developed by Liu et al. to quantitatively analyze protein synthesis in Plasmodium berghei liver-stage parasites in actively translating hepatoma cells.

• This quantitative approach should be adaptable for other puromycin-sensitive intracellular pathogens residing in actively translating host cells.

• The P. berghei–infected HepG2 recovery and reseeding protocol presented here is of use in applications beyond nascent proteome labeling and quantification.

Graphical overview

Many organisms alternate the expression of genes from large gene sets or gene families to adapt to environmental cues or immune pressure. The single-celled protozoan pathogen Trypanosoma brucei spp. periodically changes its homogeneous surface coat of variant surface glycoproteins (VSGs) to evade host antibodies during infection. This pathogen expresses one out of ~2,500 VSG genes at a time from telomeric expression sites (ESs) and periodically changes their expression by transcriptional switching or recombination. Attempts to track VSG switching have previously relied on genetic modifications of ES sequences with drug-selectable markers or genes encoding fluorescent proteins. However, genetic modifications of the ESs can interfere with the binding of proteins that control VSG transcription and/or recombination, thus affecting VSG expression and switching. Other approaches include Illumina sequencing of the VSG repertoire, which shows VSGs expressed in the population rather than cell switching; the Illumina short reads often limit the distinction of the large set of VSG genes. Here, we describe a methodology to study antigenic switching without modifications of the ES sequences. Our protocol enables the detection of VSG switching at nucleotide resolution using multiplexed clonal cell barcoding to track cells and nanopore sequencing to identify cell-specific VSG expression. We also developed a computational pipeline that takes DNA sequences and outputs VSGs expressed by cell clones. This protocol can be adapted to study clonal cell expression of large gene families in prokaryotes or eukaryotes.

Key features

• This protocol enables the analysis of variant surface glycoproteins (VSG) switching in T. brucei without modifying the expression site sequences.

• It uses a streamlined computational pipeline that takes fastq DNA sequences and outputs expressed VSG genes by each parasite clone.

• The protocol leverages the long reads sequencing capacity of the Oxford nanopore sequencing technology, which enables accurate identification of the expressed VSGs.

• The protocol requires approximately eight to nine days to complete.

Graphical overview

The mitochondrial electron transport chain (ETC) is a multi-component pathway that mediates the transfer of electrons from metabolic reactions that occur in the mitochondrion to molecular oxygen (O₂). The ETC contributes to numerous cellular processes, including the generation of cellular ATP through oxidative phosphorylation, serving as an electron sink for metabolic pathways such as de novo pyrimidine biosynthesis and for maintaining mitochondrial membrane potential. Proper functioning of the mitochondrial ETC is necessary for the growth and survival of apicomplexan parasites including Plasmodium falciparum, a causative agent of malaria. The mitochondrial ETC of P. falciparum is an attractive target for antimalarial drugs, due to its essentiality and its differences from the mammalian ETC. To identify novel P. falciparum ETC inhibitors, we have established a real-time assay to assess ETC function, which we describe here. This approach measures the O₂ consumption rate (OCR) of permeabilized P. falciparum parasites using a Seahorse XFe96 flux analyzer and can be used to screen compound libraries for the identification of ETC inhibitors and, in part, to determine the targets of those inhibitors.

Key features

• With this protocol, the effects of candidate inhibitors on mitochondrial O₂ consumption in permeabilized asexual P. falciparum parasites can be tested in real time.

• Through the sequential injection of inhibitors and substrates into the assay, the molecular targets of candidate inhibitors in the ETC can, in part, be determined.

• The assay is applicable for both drug discovery approaches and enquiries into a fundamental aspect of parasite mitochondrial biology.

Graphical overview

Seahorse assay experimental workflow. Prior to the assay, coat the cell culture microplate with Cell-Tak to help adhere the parasites to the wells; hydrate the cartridge wells to ensure proper sensor functionality and design the assay template using the Agilent Seahorse Wave Desktop software (Analyze Seahorse data files, Seahorse Wave desktop software|Agilent). On the day of the assay, prepare the inhibitors/substrates that are to be injected into the ports. Then, separate 3 × 10⁸ trophozoite-stage parasites from the uninfected red blood cells (RBCs) and ring-stage parasites using a MACS^® magnetic column. Check the purity of the parasites with Giemsa-stained smears. Determine the concentration of infected RBCs in the sample using a hemocytometer and dilute to approximately 5 × 10⁷ parasites per milliliter. Treat infected RBCs with saponin to permeabilize the host cell membrane and seed approximately 5 × 10⁶ parasites (100 μL) per well in mitochondria assay solution (MAS) buffer. Supplement MAS buffer with digitonin to permeabilize the parasite plasma membrane. Load the ports with the prepared inhibitors/substrates and run the assay using a Seahorse XFe96 analyzer. Once the assay is completed, analyze the data using the Wave desktop software. Further data processing can be done using statistical analysis software.

Cellular sensitivity is an approach to inhibit the growth of certain cells in response to any non-permissible conditions, as the presence of a cytotoxic agent or due to changes in growth parameters such as temperature, salt, or media components. Sensitivity tests are easy and informative assays to get insight into essential gene functions in various cellular processes. For example, cells having any functionally defective genes involved in DNA replication exhibit sensitivity to non-permissive temperatures and to chemical agents that block DNA replication fork movement. Here, we describe a sensitivity test for multiple strains of Saccharomyces cerevisiae and Candida albicans of diverged genetic backgrounds subjected to several genotoxic chemicals simultaneously. We demonstrate it by testing the sensitivity of DNA polymerase defective yeast mutants by using spot analysis combined with colony forming unit (CFU) efficiency estimation. The method is very simple and inexpensive, does not require any sophisticated equipment, can be completed in 2–3 days, and provides both qualitative and quantitative data. We also recommend the use of this reliable methodology for assaying the sensitivity of these and other fungal species to antifungal drugs and xenobiotic factors.

The cell cycle is a vital process of cell division that is required to sustain life. Since faithful cell division is critical for the proper growth and development of an organism, the study of the cell cycle becomes a fundamental research objective. Saccharomyces cerevisiae has been an excellent unicellular system for unraveling the secrets of cell division, and the process of synchronization in budding yeast has been standardized. Cell synchronization is a crucial step of cell cycle analysis, where cells in a culture at different stages of the cell cycle are arrested to the same phase and, upon release, they progress synchronously. The cellular synchronization of S. cerevisiae is easily achieved by a pheromone or other chemicals like hydroxyurea treatment; however, such methodologies seem to be ineffective in synchronizing cells of multimorphic fungi such as Candida albicans. C. albicans is a human pathogen that can grow in yeast, pseudohyphal, and hyphal forms; these forms differ in morphology as well as cell cycle progression. More importantly, upon subjecting to DNA replication inhibitors for synchronization, C. albicans develops hyphal structures and grows asynchronously. Therefore, here we describe a simple and easy method to synchronize C. albicans cells in the G1 phase and the subsequent analysis of cell cycle progression by using flow cytometry.

The envelope of Gram-negative bacteria consists of an outer membrane (OM), a peptidoglycan cell wall, and an inner membrane (IM). The OM and IM have different components of proteins and lipids. Separating the IM and OM is a basic biochemical procedure to further study lipids and membrane proteins in different locations. Sucrose gradient ultracentrifugation of lysozyme/EDTA-treated total membrane is the most widely used method to separate the IM and OM of Gram-negative bacteria. However, EDTA is often harmful to protein structure and function. Here, we describe a relatively simple sucrose gradient ultracentrifugation method to separate the IM and OM of Escherichia coli. In this method, the cells are broken by a high-pressure microfluidizer, and the total cell membrane is collected by ultracentrifugation. The IM and OM are then separated on a sucrose gradient. Because EDTA is not used, this method is beneficial for subsequent membrane protein purification and functional study.