微生物学

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.

Sclerotinia sclerotiorum causes white mold, leading to substantial losses on a wide variety of hosts around the world. Many genes encoding effector proteins play important roles in the pathogenesis of S. sclerotiorum. Therefore, establishment of a transformation system for the exploration of gene function is necessarily significant. Here, we introduce a modified protocol to acquire protoplasts for transformation and generate knockout strains, which completements the transformation system of S. sclerotiorum.

Babesiosis is a tick-borne disease caused by pathogens belonging to the genus Babesia. In humans, the disease presents as a malaria-like illness and can be fatal in immunocompromised and elderly people. In the past few years, human babesiosis has been a rising concern worldwide. The disease is transmitted through tick bite, blood transfusion, and transplacentally in rare cases, with several species of Babesia causing human infection. Babesia microti, Babesia duncani, and Babesia divergens are of particular interest because of their important health impact and amenability to research inquiries. B. microti, the most commonly reported Babesia pathogen infecting humans, can be propagated in immunocompetent and immunocompromised mice but so far has not been successfully continuously propagated in vitro in human red blood cells (hRBCs). Conversely, B. divergens can be propagated in vitro in hRBCs but lacks a mouse model to study its virulence. Recent studies have highlighted the uniqueness of B. duncani as an ideal model organism to study intraerythrocytic parasitism in vitro and in vivo. An optimized B. duncani in culture and in mouse (ICIM) model has recently been described, combining long-term continuous in vitro culture of the parasite in human red blood cells with an animal model of parasitemia (P) and lethal infection in C3H/HeJ mice. Here, we provide a detailed protocol for the use of the B. duncani ICIM model in research. This model provides a unique and sound foundation to gain further insights into the biology, pathogenesis, and virulence of Babesia and other intraerythrocytic parasites, and has been validated as an efficient system to evaluate novel strategies for the treatment of human babesiosis and possibly other parasitic diseases.

Graphical abstract:

ICIM model [Adapted and modified from Pal et al. (2022)]

Microorganisms have evolved adaptive strategies to respond to the autonomous degradation of their environment. Indeed, a growing culture progressively exhausts nutrients from its media and modifies its composition. Yet, how single cells react to these modifications remains difficult to study since it requires population-scale growth experiments to allow cell proliferation to have a collective impact on the environment, while monitoring the same individuals exposed to this environment for days. For this purpose, we have previously described an integrated microfluidic pipeline, based on continuous separation of the cells from the media and subsequent perfusion of the filtered media in an observation chamber containing isolated single cells. Here, we provide a detailed protocol to implement this methodology, including the setting up of the microfluidic system and the processing of timelapse images.

Dozens of Mycoplasma species belonging to the class Mollicutes bind to solid surfaces through the organelle formed at a cell pole and glide in its direction by a unique mechanism. In Mycoplasma mobile, the fastest gliding species in Mycoplasma, the force for gliding is generated by ATP hydrolysis on an internal structure. However, the spatial and temporal behaviors of the internal structures in living cells were unclear. High-speed atomic force microscopy (HS-AFM) is a powerful method to monitor the dynamic behaviors of biomolecules and cells that can be captured while maintaining their active state in aqueous solution. In this protocol, we describe a method to detect their movements using HS-AFM. This protocol should be useful for the studies of many kinds of microorganisms.

Graphic abstract:

Scannnig Mycoplasma cell

Recent advancements in 3D microscopy have enabled scientists to monitor signals of multiple cells in various animals/organs. However, segmenting and tracking the moving cells in three-dimensional time-lapse images (3D + T images), to extract their dynamic positions and activities, remains a considerable bottleneck in the field. We developed a deep learning-based software pipeline called 3DeeCellTracker, which precisely tracks cells with large movements in 3D + T images, obtained from different animals or organs, using highly divergent optical systems. In this protocol, we explain how to set up the computational environment, the required data, and the procedures to segment and track cells with 3DeeCellTracker. Our protocol will help scientists to analyze cell activities/movements in 3D + T image datasets that have been difficult to analyze.

Graphic abstract:

The flowchart illustrating how to use 3DeeCellTracker. See the Equipment and Procedure sections for detailed explanations.

Hydrogen peroxide (H₂O₂) is a toxic oxidant produced as a byproduct of several biological processes. At too high levels of hydrogen peroxide cells will experience oxidative stress, leading to a cellular response to decrease its levels and to protect the cells. Previously, methods used to study and quantify intracellular H₂O₂ have been limited by both sensitivity and specificity. However, an increasing number of genetically encoded fluorescent indicators (GEFIs) are becoming available, which can specifically detect low levels of intracellular hydrogen peroxide. In this study, we use such a biosensor designed to monitor cytosolic H₂O₂ levels in the budding yeast Saccharomyces cerevisiae during continuous cultivation and in the absence of a fluorescence microscope. The fluorescent biosensor contains a peroxiredoxin protein fused to an engineered GFP molecule expressed from a commonly used yeast plasmid (pRS416-TEF1). The peroxiredoxin-based fluorescent indicator reduces H₂O₂, ultimately resulting in a GFP signal being emitted by the sensor. Here, we apply this biosensor to study cytosolic H₂O₂ levels in S. cerevisiae strains with and without recombinant protein production.

Graphic abstract:

Schematic overview of experimental steps.