Microbiology

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.

The mitochondrial electron transport chain (ETC) performs several critical biological functions, including maintaining mitochondrial membrane potential, serving as an electron sink for important metabolic pathways, and contributing to the generation of ATP via oxidative phosphorylation. The ETC is important for the survival of many eukaryotic organisms, including intracellular parasites such as the apicomplexan Toxoplasma gondii. The ETC of T. gondii and related parasites differs in several ways from the ETC of the mammalian host cells they infect, and can be targeted by anti-parasitic drugs, including the clinically used compound atovaquone. To characterize the function of novel ETC proteins found in the parasite and to identify new ETC inhibitors, a scalable assay that assesses both ETC function and non-mitochondrial parasite metabolism (e.g., glycolysis) is desirable. Here, we describe methods to measure the oxygen consumption rate (OCR) of intact T. gondii parasites and thereby assess ETC function, while simultaneously measuring the extracellular acidification rate (ECAR) as a measure of general parasite metabolism, using a Seahorse XFe96 extracellular flux analyzer. We also describe a method to pinpoint the location of ETC defects and/or the targets of inhibitors, using permeabilized T. gondii parasites. We have successfully used these methods to investigate the function of T. gondii proteins, including the apicomplexan parasite-specific protein subunit TgQCR11 of the coenzyme Q:cytochrome c oxidoreductase complex (ETC Complex III). We note that these methods are also amenable to screening compound libraries to identify candidate ETC inhibitors.

Experimental results in fungal biology research are usually obtained as average measurements across whole populations of cells, whilst ignoring what is happening at the single cell level. Microscopy has allowed us to study single-cell behavior, but it has low throughput and cannot be used to select individual cells for downstream experiments. Here we present a method that allows for the analysis and selection of single fungal cells in high throughput by flow cytometry and fluorescence activated cell sorting (FACS), respectively. This protocol can be adapted for every fungal species that produces cells of up to 70 microns in diameter. After initial setting of the flow cytometry gates, which takes a single day, accurate single cell analysis and sorting can be performed. This method yields a throughput of thousands of cells per second. Selected cells can be subjected to downstream experiments to study single-cell behavior.

The genus Flavivirus within the family Flaviviridae includes many viral species of medical importance, such as yellow fever virus (YFV), Zika virus (ZIKV), and dengue virus (DENV), among others. Presently, the identification of flavivirus-infected cells is based on either the immunolabeling of viral proteins, the application of recombinant reporter replicons and viral genomes, or the use of cell-based molecular reporters of the flaviviral protease NS2B-NS3 activity. Among the latter, our flavivirus-activatable GFP and mNeptune reporters contain a quenching peptide (QP) joined to the fluorescent protein by a linker consisting of a cleavage site for the flavivirus NS2B-NS3 proteases (AAQRRGRIG). When the viral protease cleaves the linker, the quenching peptide is removed, and the fluorescent protein adopts a conformation promoting fluorescence. Here we provide a detailed protocol for the generation, selection and implementation of stable BHK-21 cells expressing our flavivirus genetically-encoded molecular reporters, suitable to monitor the viral infection by live-cell imaging. We also describe the image analysis procedures and provide the required software pipelines. Our reporter cells allow the implementation of single-cell infection kinetics as well as plaque assays for both reference and native strains of flaviviruses by live-cell imaging.

Graphic abstract:

Workflow for the generation and implementation of reporter BHK-21 cells for live imaging of flavivirus infection.

Bacteria are surrounded by a protective peptidoglycan cell wall. Provided that this structure and the enzymes involved are the preferred target for our most successful antibiotics, determining its structural and chemical complexity is of the highest interest. Traditionally, high-performance liquid chromatography (HPLC) analyses have been performed, but these methods are very time consuming in terms of sample preparation and chromatographic separation. Here we describe an optimized method for preparation of Gram-negative bacteria peptidoglycan and its subsequent analysis by ultra-performance liquid chromatography (UPLC). The use of UPLC in peptidoglycan analyses provides a dramatic reduction of the sample volume and hands-on time required and, furthermore, permits in-line mass spectrometry (MS) of the UPLC resolved muropeptides, thus facilitating their identification. This method improves our capability to perform high throughput analysis to better understand the cell-wall biology.

Human liver is the primary and obligatory site for malaria infection where sporozoites invade host hepatocytes. Malaria hepatic stages are asymptomatic and represent an attractive target for development of anti-malarial interventions and vaccines. However, owing to lack of robust and reproducible in vitro culture system, it is difficult to target and study this imperative malaria liver stage. Here, we describe a procedure that allow cultivation and visualization of malaria hepatic stages including dormant hypnozoites using primary simian hepatocytes. This method enables sensitive and quantitative assessment of different hepatic stages in vitro.

The intracellular pH of yeast is a tightly regulated physiological cue that changes in response to growth state and environmental conditions. Fluorescent reporters, which have altered fluorescence in response to local pH changes, can be used to measure intracellular pH. While microscopy is often used to make such measurements, it is relatively low-throughput such that collecting enough data to fully characterize populations of cells is challenging. Flow cytometry avoids this drawback, and is a powerful tool that allows for rapid, high-throughput measurement of fluorescent readouts in individual cells. When combined with pH-sensitive fluorescent reporters, it can be used to characterize the intracellular pH of large populations of cells at the single-cell level. We adapted microscopy and flow-cytometry based methods to measure the intracellular pH of yeast. Cells can be grown under near-native conditions up until the point of measurement, and the protocol can be adapted to single-point or dynamic (time-resolved) measurements during changing environmental conditions.

The latent HIV-1 viral reservoir in resting CD4⁺ (rCD4⁺) T cells represents a major barrier to an HIV-1 cure. There is an ongoing effort to identify therapeutic approaches that will eliminate or reduce the size of this reservoir. However, clinical investigators lack an assay to determine whether or not a decrease in the latent reservoir has been achieved. Therefore, it is critical to develop assays that can reproducibly quantify the reservoir size and changes therein, in participant’s blood during a therapeutic trial. Quantification of the latent HIV viral reservoir requires a highly sensitive, cost-effective assay capable of measuring the low frequency of rCD4⁺ T cells carrying functional provirus. Preferably, such an assay should be such that it can be adopted for high throughput and could be adopted under conditions for use in large-scale clinical trials. While PCR-based assays are commonly used to quantify pro-viral DNA or intracellular RNA transcript, they cannot distinguish between replication-competent and defective proviruses. We have recently published a study where a reporter cell-based assay (termed TZA or TZM-bl based quantitative assay) was used to quantify inducible replication-competent latent HIV-1 in blood. This assay is more sensitive, cost-efficient, and faster than available technology, including the quantitative viral outgrowth assay or the Q-VOA. Using this assay, we show that the size of the inducible latent HIV-1 reservoir in virally suppressed participants on ART is approximately 70-fold larger than previous estimates. We describe here in detail an optimized method to quantitate latently infected cells using the TZA.