细胞生物学

Zebrafish are a powerful model for investigating vascular and lymphatic biology due to their genetic tractability and optical transparency. While translating ribosome affinity purification (TRAP) has been widely applied in other systems, its application in zebrafish has remained limited. Here, we present an optimized TRAP protocol for isolating ribosome-associated mRNAs from endothelial cells in vivo, without the need for cell dissociation or sorting. Using a novel transgenic zebrafish line, which expresses HA-tagged Rpl10a under the mrc1a promoter, we enriched actively translating endothelial transcripts. Differential expression analysis revealed robust upregulation of vascular and lymphatic genes including flt4, kdrl, and lyve1b. This approach captures the endothelial cell translatome with high specificity and offers a robust platform for investigating the molecular mechanisms of endothelial biology under genetic, environmental, or toxicological perturbations.

Lipid droplets have emerged as dynamic organelles involved in diverse cellular processes beyond simple lipid storage. In plants and cyanobacteria, growing evidence highlights their importance in stress adaptation and signaling, yet methods to study their structure and purity remain limited. Traditionally, in situ transmission electron microscopy (TEM) has been used to visualize lipid droplets within intact cells. While powerful, this approach cannot easily evaluate isolated lipid droplets or confirm their purity. In this protocol, we describe a rapid method for preparing and visualizing cyanoglobule lipid droplets isolated from cyanobacteria. The isolated droplets are directly processed for TEM using negative staining with uranyl acetate, providing a straightforward and efficient workflow. The procedure can be applied broadly to lipid droplets from diverse organisms, independent of species or cellular origin. This protocol offers a simple, fast, and widely applicable approach to assessing lipid droplets, expanding the toolkit for researchers studying their structure and function.

This protocol describes the isolation and flow cytometric analysis of extracellular vesicles (EVs) derived from red blood cells, endothelial cells, and platelets in human peripheral blood. The protocol includes steps for preparing platelet-free plasma, fluorescent antibody staining, gating strategies, and technical controls. This protocol was developed within a study on EV release in snakebite-associated thrombotic microangiopathy; the protocol addresses challenges such as variable autofluorescence and heterogeneity in EV origin. It is flexible and can be adapted for alternative antibody panels targeting different cell populations or EV subtypes, including leukocyte-derived EVs.

Extracellular vesicles (EVs) have emerged as promising carriers for the targeted delivery of therapeutic proteins to specific cells. Previously, we demonstrated that genetically engineered EVs can be used for targeted protein delivery. This protocol details the generation of mannose receptor (CD206)-targeted EVs using a modular plasmid system optimized for production in HEK293T cells. Three plasmids enable customizable EV budding, cargo loading, and surface modification for targeting to antigen-presenting cells (APCs). EVs are isolated via differential centrifugation and chromatography, characterized using transmission electron microscopy (TEM) and nanoparticle tracking analysis (NTA), and validated through functional uptake assays in primary human activated dendritic cells. Our approach combines flexibility in engineering required EVs with robust, reproducible isolation and characterization workflows. Its modularity allows easy adaptation to alternative targets or cargoes, which can be validated immediately through in vitro testing.

Mitochondria are dynamic organelles with essential roles in energetics and metabolism. Several metabolites are common to both the cytosolic and mitochondrial fractions of the cell. The compartmentalization of metabolites within the mitochondria allows specialized uses for mitochondrial metabolism. Inorganic phosphate (Pi) is one such critical metabolite required for ATP synthesis, via glycolysis and mitochondrial oxidative phosphorylation. Estimating total cellular Pi levels cannot distinguish the distribution of Pi pools across different cellular compartments, such as the cytosol and mitochondria, and therefore separate the contributions made toward glycolysis or other cytosolic metabolic processes vs. mitochondrial outputs. Quantifying Pi pools in mitochondria can therefore be very useful toward understanding mitochondrial metabolism and phosphate homeostasis. Here, we describe a protocol for the fairly rapid, efficient isolation of mitochondria from Saccharomyces cerevisiae by immunoprecipitation for quantitative estimation of mitochondrial and cytosolic Pi pools. This method utilizes magnetic beads to capture FLAG-tagged mitochondria (Tom20-FLAG) from homogenized cell lysates. This method provides a valuable tool to investigate changes in mitochondrial phosphate dynamics. Additionally, this protocol can be coupled with LC–MS approaches to quantitatively estimate mitochondrial metabolites and proteins and can be similarly used to assess other metabolite pools that are partitioned between the cytosol and mitochondria.

Analysis of mitochondrial function has broad applicability in many research specialties. Neurodegenerative disorders such as chemotherapy-induced peripheral neuropathy (CIPN) often exhibit damaged mitochondria or reduced mitochondrial respiratory capacity. Isolation of intact mitochondria for protein analysis or respiration measurements has been previously reported in numerous model organisms. Here, we describe an adaptation of previous protocols to isolate intact functional mitochondria from Drosophila melanogaster for use in a model of CIPN. Whole Drosophila are ground in isolation buffer, and mitochondria are purified using differential centrifugation through a sucrose and mannitol solution. The intact mitochondria are plated as a monolayer for measurements of mitochondrial oxygen consumption rates and response to inhibitor compounds on an Agilent Seahorse analyzer. This experimental protocol is quick and yields a purified population of intact mitochondria that may be used for functional assays for several hours after isolation. The isolated mitochondria may be used for respiration measurements, which reflect their health, and stored for protein or genetic analysis. Mitochondrial populations from multiple strains or treatment groups can be easily compared simultaneously. The rapid biochemical assessment of mitochondria, in combination with the utility of Drosophila as an in vivo genetic model system, offers great potential for researchers to probe the impact of genetics and pharmacologic interventions on mitochondrial respiratory capacity.

Lysosome-related organelles (LROs) are a class of heterogeneous subcellular organelles conserved in eukaryotes, performing various functions. An important function of LROs is to mediate phosphorus and metal homeostasis. Chlamydomonas reinhardtii serves as a model organism for investigating metal ion metabolism. Considering that LROs contain polyphosphate and various metal elements, the purification strategy is based on their higher density by fractionating cell lysate through OptiPrep density gradient ultracentrifugation. Here, we optimized a method for purifying LROs from C. reinhardtii cells that have reached stationary phase (sta-LROs) or are overloaded with iron (Fe-LROs). Our protocol provides technical support for further investigations on the biogenesis and function of LROs in C. reinhardtii.

Mitochondria are vital organelles essential for cellular functions, but their lipid composition and response to stressors are not fully understood. Recent advancements in lipidomics reveal insights into lipid functions, especially their roles in metabolic perturbations and diseases. Previous methods have focused on the protein composition of mitochondria and mitochondrial-associated membranes. The advantage of our technique is that it combines organelle isolation with targeted lipidomics, offering new insights into the composition and dynamics of these organelles in pathological conditions. We developed a mitochondria isolation protocol for L6 myotubes, enabling lipidomics analysis of specific organelles without interference from other cellular compartments. This approach offers a unique opportunity to dissect lipid dynamics within mitochondria and their associated ER compartments under cellular stress.

Key features

• Analysis and quantification of lipids in mitochondria–ER fraction through liquid chromatography–tandem mass spectrometry-based lipidomics (LC-MS/MS lipidomics).

• LC-MS/MS lipidomics provide precise and unbiased information on the lipid composition in in vitro systems.

• LC-MS/MS lipidomics facilitates the identification of lipid signatures in mammalian cells.

Extracellular vesicles (EVs) are a heterogeneous group of nanoparticles possessing a lipid bilayer membrane that plays a significant role in intercellular communication by transferring their cargoes, consisting of peptides, proteins, fatty acids, DNA, and RNA, to receiver cells. Isolation of EVs is cumbersome and time-consuming due to their nano size and the co-isolation of small molecules along with EVs. This is why current protocols for the isolation of EVs are unable to provide high purity. So far, studies have focused on EVs derived from cell supernatants or body fluids but are associated with a number of limitations. Cell lines with a high passage number cannot be considered as representative of the original cell type, and EVs isolated from those can present distinct properties and characteristics. Additionally, cultured cells only have a single cell type and do not possess any cellular interactions with other types of cells, which normally exist in the tissue microenvironment. Therefore, studies involving the direct EVs isolation from whole tissues can provide a better understanding of intercellular communication in vivo. This underscores the critical need to standardize and optimize protocols for isolating and characterizing EVs from tissues. We have developed a differential centrifugation-based technique to isolate and characterize EVs from whole adipose tissue, which can be potentially applied to other types of tissues. This may help us to better understand the role of EVs in the tissue microenvironment in both diseased and normal conditions.

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