Cell Biology

Extracellular vesicles (EVs) are lipid bilayer–enclosed vesicles released by diverse cell types and found in various body fluids. Because their composition and cargo dynamically respond to physiological and environmental cues, EVs hold promise both as biomarkers and as carriers for therapeutic delivery. Skeletal muscle functions as an endocrine organ, secreting myokines and EVs that modulate a wide range of cellular processes. The murine C2C12 cell line is a widely used in vitro model for investigating muscle biology. Here, we describe a protocol for isolating EVs from differentiated C2C12 myocytes. The isolated EVs are characterized and validated using western blotting, transmission electron microscopy (TEM), and dynamic light scattering (DLS) analysis. This workflow provides a robust platform for studying the molecular composition and functional roles of muscle-derived EVs.

Plant-derived extracellular vesicles (PDEVs) have emerged as important mediators of intercellular communication and hold growing potential in therapeutic applications. However, standardized methods for their isolation, particularly from Piper betle leaves (PBL), remain unexplored. Existing apoplastic fluid washing (AFW) extraction techniques typically rely on manual syringe infiltration, which often leads to inconsistent pressure control, variable yields, and increased risk of tissue damage. This protocol describes a vacuum-assisted AFW extraction method optimized for the recovery of intact extracellular vesicles (EVs) from PBL. The workflow features controlled negative pressure using a vacuum pump and chamber to achieve more efficient leaf infiltration compared to infiltration using the syringe method and reproducible apoplastic fluid (AF) collection with subsequent low-speed centrifugation steps, to ensure minimal contamination and preservation of vesicle integrity. Piper betle–derived extracellular vesicle (PBdEV) isolation and purification steps are performed using size exclusion chromatography (SEC). The size and concentration of PBdEVs were confirmed using nanoparticle tracking analysis (NTA), whereas the cup-shaped and lipid bilayer morphology of the EVs were confirmed using transmission electron microscopy (TEM). The method is scalable and adaptable to various leaf morphologies and physiological states, making it suitable for both exploratory and high-throughput studies. Overall, this protocol provides a more consistent, efficient, and tissue-preserving alternative to traditional syringe-based AF extraction methods, offering higher-quality EV preparations for plant EV research.

Aloe vera has long been used for its diverse pharmacological properties, motivating continued interest in isolating and preserving the bioactive molecules responsible for its therapeutic potential. More recently, Aloe vera–derived extracellular vesicles (Av-EVs) have emerged as nanoscale, cell-free carriers capable of retaining and delivering these properties, making them attractive for various biomaterials, nanomedicine, and regenerative medicine applications. Multiple techniques are available for extracellular vesicle isolation. These include ultracentrifugation, polymer-based precipitation, size-exclusion chromatography, immunoaffinity capture, ultrafiltration, density gradient separation, and emerging microfluidic platforms. Each method presents distinct trade-offs in purity, yield, scalability, and downstream compatibility. Despite this diversity, standardized workflows tailored to Av-EV isolation remain limited, and the influence of homogenization-induced shear forces and plant maturity on vesicle recovery and characterization has not been systematically addressed. Here, we present a reproducible protocol for isolating Av-EVs from Aloe vera gel employing two distinct homogenization strategies: manual, no-shear force (NB EVs), and blender-based shear-force homogenization (B EVs). The workflow covers gel preparation, serial centrifugation for debris removal, ultracentrifugation as the gold standard for vesicle enrichment, and final sterile filtration. This protocol enables consistent recovery of Av-EVs suitable for physicochemical characterization and functional analyses. It is simple and relies on commonly available laboratory equipment, facilitating broad adoption by ultracentrifugation users and offering adaptability to diverse research projects involving purified Aloe vera gel and Av-EVs, including studies focused on wound healing, fibrotic scarring, and regenerative processes, where coordinated antioxidant, anti-inflammatory, antimicrobial, immunomodulatory, and moisturizing responses are of interest.

Extracellular vesicles (EVs) are critical mediators of cell–cell communication and play a key role in male reproductive biology by modulating sperm function. This protocol describes a robust and reproducible workflow for isolating EVs from ram seminal plasma using size-exclusion chromatography (SEC) and assessing their uptake by ram spermatozoa. In contrast to ultracentrifugation-based methods, SEC provides a gentle and more efficient isolation approach that preserves EV integrity and functionality. A central innovation of this protocol is the use of carboxyfluorescein succinimidyl ester (CFSE)-labeled seminal plasma EVs (SP-EVs) to evaluate their incorporation into sperm cells through two complementary detection platforms: (i) flow cytometry with standard resolution and (ii) confocal microscopy, for spatial confirmation of EV–sperm interactions. By bridging the gap between EV isolation and functional analysis, this protocol provides a valuable tool for investigating the role of EV–cell interactions. Specifically, it offers potential applications in male fertility preservation, biomarker discovery, and the development of EV-based therapeutic strategies in reproductive medicine.

Extracellular vesicles (EVs) circulating in blood serve as non-invasive “liquid biopsies,” carrying molecular cargo that reflects the physiological and pathological state of distant cells. Their analysis is crucial for understanding disease mechanisms and discovering novel biomarkers. Clinically, blood EVs hold significant promise for early disease diagnosis, prognostic assessment, and monitoring treatment response in diverse areas such as organ transplantation, cancer, and neurological disorders. Current EV isolation techniques, beyond ultracentrifugation, include size exclusion chromatography (separation by size for high purity) and immunoaffinity capture (using antibodies for high specificity). Here, we present a simplified, rapid, and reproducible method for isolating EVs from small-volume blood samples. This protocol consistently yields a concentrated EV pellet covering 50–300 nm EVs, amenable to direct downstream analysis. Developed and validated in our laboratory using human, porcine, and murine blood samples, this method has proven instrumental in identifying EV-based biomarkers for predicting outcomes related to organ transplantation. The protocol’s adaptability and reliance on readily prepared, cost-effective reagents further enhance its utility. This scalable approach can be further integrated with subsequent purification or enrichment steps to optimize sample preparation for protein and nucleic acid assays.

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.