细胞生物学

The cellular compartments of eukaryotic cells are defined by their specific protein compositions. Different strategies are used for the identification of the subcellular proteomes, such as fractionation by differential centrifugation of cellular extracts. The localization of mitochondrial proteins is particularly challenging, as mitochondria consist of two membranes of different protein composition and two aqueous subcompartments, the intermembrane space (IMS) and the matrix. Previous studies identified subcompartment-specific proteomes by using combinations of hypotonic swelling and protease digestion followed by mass spectrometry. Here, we present an alternative, more unbiased method to identify the proteomes of mitochondrial subcompartments by use of an improved ascorbate peroxidase (APEX2) that is targeted to the IMS and the matrix. This method allows the subcompartment-specific labeling of proteins in mitochondria isolated from cells of the baker’s yeast Saccharomyces cerevisiae, followed by their purification on streptavidin beads. With this method, the proteins located in the different mitochondrial subcompartments of yeast cells can be efficiently and comprehensively identified.

Mitophagy is a highly conserved process among eukaryotic cells, playing a primordial role in mitochondrial quality control and overall cellular homeostasis. In Saccharomyces cerevisiae, Atg32 is the only identified mitophagy receptor localized to the mitochondrial outer membrane, making this yeast a particularly powerful model for molecular studies of mitophagy that require the isolation of intact mitochondria. However, traditional methods for isolating mitochondria from yeast often rely on enzymatic cell wall digestion and homogenization, which can compromise the stability of mitochondrial surface proteins such as Atg32. In this protocol, we describe an optimized mechanical approach for yeast cell disruption using glass beads in a cold, protease-inhibited buffer to preserve mitochondrial integrity and facilitate the detection of Atg32. Subsequent differential centrifugation and washing steps yield mitochondrial fractions suitable for downstream biochemical analyses. This workflow eliminates enzymatic digestion steps, reduces sample variability, and allows parallel processing of multiple strains or experimental conditions. Overall, this method offers a rapid, low-cost, and reproducible alternative for crude mitochondrial isolation, ensuring excellent preservation of Atg32 and broad compatibility with quantitative and comparative studies.

It is common practice for laboratories to discard clotted blood or freeze it for future DNA extraction after extracting serum from a serum-separating tube. If freezing for DNA extraction, the blood clot is not usually cryopreserved, which leads to cell membrane fragility. In this protocol, we describe steps to isolate high-quality nuclei from leukocytes derived from whole blood samples frozen without a cryoprotective medium. Nuclei isolated from this protocol were able to undergo ATAC (assay for transposase-accessible chromatin) sequencing to obtain chromatin accessibility data. We successfully characterized and isolated B cells and T cells from leukocytes isolated from previously frozen blood clot using Miltenyi’s gentleMACS Octo Dissociator coupled with flow sorting. Nuclei showed round, intact nuclear envelopes suitable for downstream applications, including bulk sequencing of nuclei or single-cell nuclei sequencing. We validated this protocol by performing bulk ATAC-seq.

Single-cell and single-nucleus RNA sequencing are revolutionizing our understanding of cellular biology. The identification of molecular markers, single-cell transcriptomic profiling, and differential gene expression at the cellular level has revealed key functional differences between cells within the same tissue. However, tissue dissociation remains challenging for non-model organisms and for tissues with unique biochemical properties. For example, the mosquito fat body, which serves functions analogous to mammalian adipose and liver tissues, consists of trophocytes—large, adipocyte-like cells whose cytoplasm is filled with lipid droplets. Conventional enzymatic dissociation methods are often too harsh for these fragile cells, and their high lipid content can interfere with reagents required for single-cell transcriptomic analysis. Single-nucleus RNA sequencing (snRNA-seq) offers an alternative strategy when intact cells with high-quality RNA cannot be obtained by enzymatic or mechanical dissociation. Here, we present an optimized reproducible methodology for nuclei isolation from the fat body of Anopheles gambiae mosquitoes, enabling high-quality snRNA-seq. Our approach involves tissue fixation and lipid removal, followed by cell lysis and nuclei purification using a sucrose cushion. We validated this protocol on both sugar-fed and blood-fed samples, established quality metrics to remove potential ambient RNA contamination, and demonstrated that snRNA-seq using this method yields high-quality sequencing results.

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.