细胞生物学

Exosomes are a subpopulation of the heterogenous pool of extracellular vesicles that are secreted to the extracellular space. Exosomes have been purported to play a role in intercellular communication and have demonstrated utility as biomarkers for a variety of diseases. Despite broad interest in exosome biology, the conditions that regulate their secretion are incompletely understood. The goal of this procedure is to biochemically reconstitute exosome secretion in Streptolysin O (SLO)-permeabilized mammalian cells. This protocol describes the reconstitution of lyophilized SLO, preparation of cytosol and SLO-permeabilized cells, assembly of the biochemical reconstitution reaction, and quantification of exosome secretion using a sensitive luminescence-based assay. This biochemical reconstitution reaction can be utilized to characterize the molecular mechanisms by which different gene products regulate exosome secretion.

Key features

• This protocol establishes a functional in vitro system to reconstitute exosome secretion in permeabilized mammalian cells upon addition of cytosol, ATP, GTP, and calcium (Ca²⁺).

Graphical overview

Schematic overview of the exosome secretion biochemical reconstitution protocol. Streptolysin O (SLO) is prepared as described in Procedure A. Cytosol is isolated from HCT116 WT cells as described in Procedure B. HCT116 CD63-Nluc cells are permeabilized by SLO as detailed in Procedure C. The assembly of the exosome secretion reactions are described in Procedure D. Quantification of CD63-Nluc secretion is detailed in Procedure E (Modified from Williams et al., 2023).

Extracellular vesicles (EVs), such as exosomes, are produced by all known eukaryotic cells, and constitute essential means of intercellular communication. Recent studies have unraveled the important roles of EVs in migrating to specific sites and cells. Functional studies of EVs using in vivo and in vitro systems require tracking these organelles using fluorescent dyes or, alternatively, transfected and fluorescent-tagged proteins, located either intravesicularly or anchored to the EV bilayer membrane. Due to design simplicity, the fluorescent dye might be a preferred method if the cells are difficult to modify by transfection or when the genetic alteration of the mother cells is not desired. This protocol describes techniques to label cultured cell-derived EVs, using lipophilic DiR [DiIC18(7) (1,1'-Dioctadecyl-3,3,3',3'-Tetramethylindotricarbocyanine Iodide)] fluorophore. This technique can be used to study the cellular uptake and intracellular localization of EVs, and their biodistribution in vivo, which are crucial evaluations of any isolated EVs.

Throughout their life cycle, bacteria shed portions of their outermost membrane comprised of proteins, lipids, and a diversity of other biomolecules. These biological nanoparticles have been shown to have a range of highly diverse biological activities, including pathogenesis, community regulation, and cellular defense (among others). In recent publications, we have isolated and characterized membrane vesicles (MVs) from several species of Lactobacilli, microbes classified as commensals within the human gut microbiome (Dean et al., 2019 and 2020). With increasing scientific understanding of host-microbe interactions, the gut-brain axis, and tailored probiotics for therapeutic or performance increasing applications, the protocols described herein will be useful to researchers developing new strategies for gut community engineering or the targeted delivery of bio-active molecules.

Graphic abstract:

Figure 1. Atomic force microscopic image of Lactobacillus casei ATCC 393 bacteria margins (white arrows) and membrane vesicles (black arrows)

Giant unilamellar vesicles (GUVs) are a widely used model system for a range of applications including membrane biophysics, drug delivery, and the study of actin dynamics. While several protocols have been developed for their generation in recent years, the use of these techniques involving charged lipid types and buffers of physiological ionic strength has not been widely adopted. This protocol describes the generation of large numbers of free-floating GUVs, even for charged lipid types and buffers of higher ionic strength, using a simple approach involving soft polyacrylamide (PAA) gels. This method entails glass cover slip functionalization with (3-Aminopropyl)trimethoxysilane (APTES) and glutaraldehyde to allow for covalent bonding of PAA onto the glass surface. After polymerization of the PAA, the gels are dried in vacuo. Subsequently, a lipid of choice is evenly dispersed on the dried gel surface, and buffers of varying ionic strength can be used to rehydrate the gels and form GUVs. This protocol is robust for the production of large numbers of free-floating GUVs composed of different lipid compositions under physiological conditions. It can conveniently be performed with commonly utilized laboratory reagents.

Small extracellular vesicles (sEVs) encompass a variety of distinct vesicles that are secreted to the extracellular space. Many methodologies currently used for EV isolation (e.g., differential ultracentrifugation concluding in a high-speed pellet, precipitation by macromolecular crowding agents or size excusion chromatography–SEC) do not fractionate distinct sEV sub-populations. Samples obtained by the aforementioned methods are usually used for characterization and physiological studies. However the fraction that contains the molecule of interest or is the carrier of a specific activity is unknown. Therefore isolating distinct sEV sub-populations is critical to understand EV function. The goal of this procedure is to purify distinct sEV sub-populations based on slight differences in their buoyant density. Moreover, this technique also allows sEVs purification from vesicle-free RNA-protein complexes co-isolating in the high-speed pellet or by the use of crowding agents. This protocol describes cultivation of mammalian cells for sEV collection, sEV sedimentation, buoyant density fractionation of sEV sub-populations and immunoblots for sEV markers. This protocol can be used to fractionate distinct sEV sub-populations produced by a variety of mammalian cells.

Cell signalling, cell secretion, and plasma membrane repair are processes that critically rely on intracellular vesicles, important components of the endocytic and secretory pathways. More specifically, the strategic distribution of intracellular vesicles is important for diverse cellular processes. The method presented here is a simple, affordable, and efficient tool to analyze the distribution of intracellular vesicles such as lysosomes, endosomes, Golgi vesicles or secretory granules under different experimental conditions. The method is an accessible way to analyze the density and dispersion of intracellular vesicles by combining immunofluorescence with pixel-based quantification software (e.g., ImageJ/FIJI). This protocol can be used widely within the scientific community because it utilizes ImageJ/FIJI, an open source software that is free. By tracking fluorescent vesicles based on their position relative to cell nuclei we are able to quantify and analyze their distribution throughout the cell.

Milk is a complex fluid that contains various types of proteins and extracellular vesicles (EVs). Some proteins can mingle with EVs, and interfere with their isolation. Among these proteins, caseins form micelles of a size comparable to milk EVs, and can thus be co-isolated with EVs. Preliminary steps that affect milk are crucial for EV isolation and impact the purity and abundance of isolated EVs. In the course of our previous works on cow’s milk EVs, we found that sodium citrate (1% final), which is a biocompatible reagent capable of breaking down casein micelles into 40-nm monomers, allowed the isolation of high quantities of EVs with low coprecipitation of caseins or other contaminating proteins. Using this protocol, we successfully separated different EV subsets, characterized in depth their morphology, protein content and small RNA enrichment patterns. We were also able to describe their biological function in a mouse model of intestinal inflammation. We, hereby, detail the differential ultracentrifugation procedure that leads to high quantify, medium specificity, isolation of different milk EV subsets from the same sample. More specifically, we highlight the use of sodium citrate as a standardized approach to isolate and study milk EVs and its potential for isolation techniques other than differential ultracentrifugation.

Extracellular vesicles (EVs) are membranous structures that cells massively release in extracellular fluids. EVs are cargo of cellular components such as lipids, proteins, and nucleic acids that can work as a formidable source in liquid biopsy studies searching for disease biomarkers. We recently demonstrated that nickel-based isolation (NBI) is a valuable method for fast, efficient, and easy recovery of heterogeneous EVs from biological fluids. NBI exploits nickel cations to capture negatively charged vesicles. Then, a mix of balanced chelating agents elutes EVs while preserving their integrity and stability in solution. Here, we describe steps and quality controls to functionalize a matrix of agarose beads, obtain an efficient elution of EVs, and extract nucleic acids carried by them. We demonstrate the versatility of NBI method in isolating EVs from media of primary mouse astrocytes, from human blood, urine, and saliva processed in parallel, as well as outer membrane vesicles (OMVs) from cultured Gram-negative bacteria.

Extracellular vesicles (EVs) are produced by all domains of life including Bacteria, Archaea and Eukarya. EVs are critical for cellular physiology and contain varied cargo: virulence factors, cell wall remodeling enzymes, extracellular matrix components and even nucleic acids and metabolites. While various protocols for isolating EVs have been established for mammalian cells, the field is actively developing tools to study EVs in other organisms. In this protocol we describe our methods to perform density gradient purification of EVs in bacterial cells, allowing for separation of EV subpopulations, followed by protection assays for EV cargo characterization. Furthermore, we devised a protocol which incorporates a fluorescent conjugate of fatty acids into EVs, the first to allow live-cell EV tracking to observe release of EVs, including during infection of mammalian cells by pathogenic bacteria. These protocols are powerful tools for EV researchers as they enable the observation of EV release and the study of the mechanisms of their formation and release.

Function of extracellular vesicles such as exosomes and microvesicles is determined by their wide ranges of cargoes inside them. Even in the pure exosomes or microvesicles the cargo contents are very heterogeneous. To understand this heterogeneous nature of extracellular vesicles, we need information of the vesicles, which will give us some parameters including vesicle size, number and cargo content of each vesicle. Here, we describe a new method to quantify cargo density in single-extracellular vesicles. Staining of extracellular vesicles in a membrane lipid content-proportionate manner and immobilization of extracellular vesicles onto glass substrate allow us to obtain cargo density information of single-extracellular vesicles. This protocol will be useful to analyze the effects of various drugs or genetic manipulation on vesicle generation and maturation including cargo sorting into heterogeneous extracellular vesicles.