微生物学

Dolichol diphosphate-linked oligosaccharides (LLO) are the sugar donors in N-glycosylation, a fundamental protein post-translational modification of the eukaryotic secretory pathway. Defects in LLO biosynthesis produce human Congenital Disorders of Glycosylation Type I. The synthesis of LLOs and the transfer reactions to their protein acceptors is highly conserved among animal, plant, and fungi kingdoms, making the fission yeast Schizosaccharomyces pombe a suitable model to study these processes. Here, we present a protocol to determine the LLO patterns produced in vivo by S. pombe cells that may be easily adapted to other cell types. First, exponentially growing cultures are labeled with a pulse of [¹⁴C]-glucose. LLOs are then purified by successive extractions with organic solvents, and glycans are separated from the lipid moieties in mild acid hydrolysis and a new solvent extraction. The purified glycans are then run on paper chromatography. We use a deconvolution process to adjust the profile obtained to the minimal number of Gaussian functions needed to fit the data and determine the proportion of each species with respect to total glycan species present in the cell. The method we provide here might be used without any expensive or specialized equipment. The deconvolution process described here might also be useful to analyze species in non-completely resolved chromatograms.

Graphical abstract:

Workflow for the labeling, extraction, separation, and identification of LLO species in S. pombe.

(A) Radioactive pulse of S. pombe cells with [¹⁴C]-glucose for 15 min at 28 °C. (B) Organic extraction of LLOs from labeled yeasts sequentially using methanol, chloroform, H₂O, chloroform:methanol:H₂O (1:1:0.3), 0.02 M HCl (to separate glycans from dolichol), and chloroform:methanol:H₂O (1:16:16). (C) Preparation of the sample for chromatography on paper: drying by airflow and radioactivity check. (D) Loading of samples in chromatographic paper and descendent chromatography in a glass chamber. The obtained plots (CPM versus running distance) need to be analyzed to identify single glycan species.

The diversity of lipid structures, properties, and combinations in biological tissues makes their extraction and analysis an experimental challenge. Accordingly, even for one of the simplest single-cellular fungi, the budding yeast (Saccharomyces cerevisiae), numerous extraction and analysis protocols have been developed to separate and quantitate the different molecular lipid species. Among them, most are quite sophisticated and tricky to follow. Herein, we describe a yeast total lipids extraction procedure with a relatively good yield, which is appropriate for subsequent thin-layer chromatography (TLC) or liquid chromatography-mass (LC-MS) analysis. We then discuss the most widely used solvent systems to separate yeast phospholipids and neutral lipids by TLC. Finally, we describe an easy and rapid method for silica gel staining by a Coomassie Brilliant Blue-methanol mixture. The stained lipid species can then be quantitated using imaging software such as ImageJ. Overall, the methods described in this protocol are time-saving and novice-friendly.

Palmitoylation refers to the modification of the cysteine thiols in proteins by fatty acids, most commonly palmitic acid, through ‘thioester bond’ formation. In vivo, palmitoylation of proteins is catalyzed by palmitoyl acyltransferases (PATs or DHHC-PATs). Palmitoylation has recently emerged as a crucial post-translational modification in malarial parasites. The expression and activity of palmitoyl transferases vary across different developmental stages of the malarial parasite’s life cycle. The abundance of palmitoylated proteins at a given stage is a measure of overall PAT activity. The PAT activity can also change in response to external signals or inhibitors. Here, we describe a protocol to ‘image’ palmitoyl-transferase activity during the asexual stages using Click Chemistry and fluorescence microscopy. This method is based on metabolic labeling of a clickable analog of palmitic acid by parasitic cells, followed by CuAAC (Copper-catalyzed Alkyne-Azide Cycloaddition reaction) Click Chemistry to render palmitoylated proteins fluorescent. Fluorescence allows the quantitation of intracellular palmitoylation in parasite cells across various development stages. Using this method, we observed that intracellular palmitoylation increases as the parasite transitions from ring to schizont stages and appears to be most abundant during the schizont stages in Plasmodium falciparum.

Peptidoglycan encases the bacterial cytoplasmic membrane to protect the cell from lysis due to the turgor. The final steps of peptidoglycan synthesis require a membrane-anchored substrate called lipid II, in which the peptidoglycan subunit is linked to the carrier lipid undecaprenol via a pyrophosphate moiety. Lipid II is the target of glycopeptide antibiotics and several antimicrobial peptides, and is degraded by ‘attacking’ enzymes involved in bacterial competition to induce lysis. Here we describe two protocols using thin-layer chromatography (TLC) and high pressure liquid chromatography (HPLC), respectively, to assay the digestion of lipid II by phosphatases such as Colicin M or the LXG toxin protein TelC from Streptococcus intermedius. The TLC method can also monitor the digestion of undecaprenyl (pyro)phosphate, whereas the HPLC method allows to separate the di-, mono- or unphosphorylated disaccharide pentapeptide products of lipid II.

Triacylglycerols (TAGs) are esters formed from one glycerol and three fatty acids. TAGs are induced to accumulate in algal cells under environmental stress conditions including nutrient-limitation, hyperosmosis, and low temperature, for the storage of metabolic energy and carbon, and also for the consumption of excess energy (e.g., Hirai et al., 2016; Hayashi et al., 2017). Beside their physiological significance, the commercial utilization of algal TAG has been expected for the production of biodiesel, the methyl esters of fatty acids, from the aspect of carbon-neutral conception. The amounts of TAGs can be determined through quantitative measurement of their constituent fatty acids. This protocol consists of the following three parts: the first is the extraction of total lipids from algal cells with the use of organic solvents, chloroform and methanol, according to the method of Bligh and Dyer (1959), the second is the separation of TAG from the other lipid classes by thin-layer chromatography (TLC), and the third is the production of methyl-esterified derivatives of their constitutive fatty acids and subsequent quantitation of them by capillary gas-liquid chromatography (GLC). This protocol adapted from Sato and Tsuzuki (2011) is used for TAG analysis in a green alga, Chlorella kessleri.

Exosomes have emerged as an important mediator of intercellular communication. They are present in extracellular milieu and therefore, easily accessible by neighboring or distant cells. They carry mRNA, microRNAs and proteins within their vesicles and once internalized by recipient cells; they can modulate multiple signaling pathways with pleiotropic effects from inducing antiviral state to disease progression. We have previously shown that hepatitis C virus (HCV) infected hepatocytes or hepatoma cells harboring genome-length replicon secrete exosomes in culture supernatants. These exosomes are taken up by hepatic stellate cells (HSC) and activate them to induce fibrosis during HCV infection. Here, we describe detailed protocols for exosomes isolation and uptake of BODIPY labeled exosomes by hepatic stellate cells.

This protocol provides an easy and rapid method to prepare lipopolysaccharide from the gastric pathogen Helicobacter pylori for visualization on acrylamide gels by silver staining and for detecting the presence of Lewis antigens by Western blot. The silver staining is a four-step procedure, involving a 20 min-oxidation step, a 10 min-silver staining step, a 2-10 min color development step and finally a 1-min color termination step. Lipopolysaccharide from H. pylori wild-type and corresponding mutants analyzed by this method are described in a recent publication (Li et al., 2017). This crude preparation of LPS for silver staining is also applicable in other Gram-negative bacteria.

Proteins may have three dimensional structural or amino acid features that suggest a role in targeting and disrupting lipids within cell membranes. It is often necessary to experimentally investigate if these proteins and biomolecules are able to disrupt membranes in order to conclusively characterize the function of these biomolecules. Here, we describe an in vitro assay to evaluate the membrane lytic properties of proteins and biomolecules. Large unilamellar vesicles (liposomes) containing carboxyfluorescein at fluorescence-quenching concentrations are treated with the biomolecule of interest. A resulting increase in fluorescence due to leakage of the dye from liposomes and subsequent dilution in the buffer demonstrates that the biomolecule is sufficient for disrupting liposomes and membranes. Additionally, since liposome disruption may occur via pore-formation or via general solubilization of lipids similar to detergents, we provide a method to distinguish between these two mechanisms. Pore-formation can be identified and evaluated by examining the blockade of carboxyfluorescein release with dextran molecules that fit the pore. The methods described here were used to determine that the malaria vaccine candidate CelTOS and proapoptotic Bax disrupt liposomes by pore formation (Saito et al., 2000; Jimah et al., 2016). Since membrane lipid binding by a biomolecule precedes membrane disruption, we recommend the companion protocol: Jimah et al., 2017.

Efficient delivery of oligonucleotide therapeutics, i.e., siRNAs, to the central nervous system represents a significant barrier to their clinical advancement for the treatment of neurological disorders. Small, endogenous extracellular vesicles were shown to be able to transport lipids, proteins and RNA between cells, including neurons. This natural trafficking ability gives extracellular vesicles the potential to be used as delivery vehicles for oligonucleotides, i.e., siRNAs. However, robust and scalable methods for loading of extracellular vesicles with oligonucleotide cargo are lacking. We describe a detailed protocol for the loading of hydrophobically modified siRNAs into extracellular vesicles upon simple co-incubation. We detail methods of the workflow from purification of extracellular vesicles to data analysis. This method may advance extracellular vesicles-based therapies for the treatment of a broad range of neurological disorders.

The protocol for obtaining electrically sealed membrane vesicles from E. coli cells is presented. Proton pumps such as Complex I, quinol oxidase, and ATPase are active in the obtained vesicles. Quality of the preparation was tested by monitoring the electric potential generated by these pumps.