Cell Biology

Prostate carcinoma (PCa) progression is strongly influenced by the surrounding tumor microenvironment, where cancer-associated fibroblasts (CAFs) represent the most abundant and functionally relevant stromal population. Despite their importance, the lack of stable cell lines representing CAF phenotypes limits the study of stromal–tumor interactions. To address this limitation, we provide an optimized protocol for isolating CAFs from fresh human PCa biopsies based on a mechanical procedure exploiting the specific CAF ability to migrate out from the tumor explants. This approach preserves tissue architecture and maintains CAF viability and phenotype. The resulting ex vivo CAF cultures provide a suitable model to investigate CAF biology within the tumor microenvironment.

Endocytosis is an essential membrane transport mechanism that is indispensable for the maintenance of life. It is responsible for the selective internalization and subsequent degradation or recycling of specific extracellular proteins and nutrients, thereby facilitating cellular nutrient supply, modulation of receptor signaling, and clearance of foreign substances. However, methods for the quantitative analysis of lysosomal degradation of extracellular proteins via endocytosis remain limited. This protocol describes a method for purifying the protein-of-interest (POI)–red fluorescent protein (RFP)–green fluorescent protein (GFP) fusion protein, which is modified with specific mammalian cell glycans or other modifications, from the conditioned medium of mammalian cell cultures. Subsequently, the protocol details a quantitative approach for evaluating its internalization and lysosomal degradation within cells using the RFP–GFP tandem fluorescent reporter. Following the addition of POI-RFP-GFP to the medium, cells can be subjected to cell biological assays, such as flow cytometry, as well as biochemical analyses, such as immunoblotting. This protocol is broadly applicable to studies of the internalization of extracellular proteins.

Obtaining articular cartilage-derived cells (chondroprogenitors) by explant methodology is a reliable approach for isolating migratory progenitor cells that retain strong chondrogenic potential. This method allows cells to emerge naturally from small cartilage fragments without enzymatic digestion. The procedure consists of plating cartilage explants on a plastic surface with culture medium, from which cells subsequently migrate and adhere to the substrate. Compared with enzymatic isolation, the explant approach minimizes cellular stress and better reproduces the physiological microenvironment of cartilage tissue. This protocol can be applied to both osteoarthritic and non-osteoarthritic samples, enabling comparative studies on disease-related phenotypic differences. Overall, this technique offers a reproducible, straightforward, and minimally invasive strategy for obtaining functional chondroprogenitor cells suitable for cartilage regeneration research.

Organelle abundance is a key microscopic readout of organelle formation and, in many cases, function. Quantification of organelle abundance using confocal microscopy requires estimating their area based on the fluorescence intensity of compartment-specific markers. This analysis usually depends on a user-defined intensity threshold to distinguish organelle regions from the surrounding cytoplasm, which introduces potential bias and variability. To address this issue, we present a machine learning–assisted algorithm that allows for the quantification of organelle density using the open-source Fiji platform and WEKA segmentation. Our method enables the automated quantification of organelle number, area, and density by learning from training data. This standardizes threshold selection and minimizes user intervention. We demonstrate the utility of this approach for both membrane and non-membrane organelles, such as peroxisomes, lipid droplets, and stress granules, in human cells and whole fish samples.

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.

Breast cancer remains one of the most prevalent and deadly malignancies affecting women worldwide. Its progression and metastatic behavior are driven by complex mechanisms. To develop more effective therapeutic strategies, it is crucial to understand tumor growth, angiogenesis, and microenvironmental interactions. Although traditional in vivo models such as murine xenografts have long been used to study tumor biology, these approaches are often time-consuming, costly, and ethically constrained. In contrast, the chick embryo chorioallantoic membrane (CAM) assay offers a rapid, cost-effective, and ethically flexible alternative for evaluating tumor development and angiogenesis. This protocol describes an in ovo CAM-based xenograft model in which human breast cancer cells are implanted onto the vascularized CAM of chick embryos. This method enables real-time evaluation of tumor growth. Furthermore, the model allows for manipulation of experimental conditions, including pharmacological treatments or genetic modifications, to study specific molecular mechanisms involved in breast cancer progression. The major advantages of this protocol lie in its simplicity, reduced cost, and capacity for high-throughput screening, making it a valuable tool for translational cancer research.

In the Japanese rhinoceros beetle Trypoxylus dichotomus, gene function studies have relied mainly on systemic larval RNA interference (RNAi), as gain-of-function techniques remain underdeveloped and germline transgenesis is impractical given the species’ approximately one-year generation time. In addition, because larval RNAi is systemic, it has been difficult to analyze the function of lethal genes. Here, we present a simple and efficient protocol for the direct introduction of exogenous DNA into T. dichotomus larvae via in vivo electroporation. This protocol includes optimized procedures for adult breeding and egg collection, as well as a rigorously parameterized electroporation technique that delivers a piggyBac transposon vector into region-specific larval tissues. Within one day after electroporation, treated larvae exhibit mosaic expression of a reporter gene, enabling rapid tissue-specific functional analysis without the need to establish stable germline transgenic lines. Moreover, the key promoter used in this system (T. dichotomus actinA3 promoter) is effective across diverse insect species, indicating that the method can be readily adapted to other non-model insects. Overall, this electroporation-based approach provides a valuable gain-of-function tool for T. dichotomus and potentially many other insect species.

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.

Time-lapse into immunofluorescence (TL into IF) imaging combines the wealth of information acquired during live-cell imaging with ease of access for static immunofluorescence markers. In the field of mechanobiology, connecting live and static imaging to visualize cell biology dynamics is often troublesome. For instance, nuclear blebs are deformations of the nucleus that often rupture spontaneously, leading to changes in the molecular composition of the nucleus and the nuclear bleb. Current techniques to connect cellular dynamics and their downstream effects via live-cell imaging, followed by immunofluorescence, often require third-party analysis programs or stage position measurements to accurately track cells. This protocol simplifies the connection between live and static imaging by utilizing a gridded imaging dish. In our protocol, cells are plated on a dish with an engraved coordinate plane. Individual cells are then matched from when the time-lapse ends to the immunofluorescence images simply by their known coordinate location. Overall, TL into IF offers a straightforward method for connecting dynamic live-cell with static immunofluorescence imaging, in an easy and accessible tool for cell biologists.

This protocol describes an easy, quick, cheap, and effective method for the purification and concentration of bacteriophages (phages) produced in rich culture media, meeting the quality criteria required for structural analyses. It is based on a tube dialysis system that replaces the classical but expensive and tedious density gradient ultracentrifugation step. We developed this protocol for the Oenococcus oeni bacteriophage OE33PA from its amplification to imaging by negative stain electron microscopy (NS-EM). The host bacterium, O. oeni, is a lactic acid bacterium that lives in harsh oenological ecosystems and grows only in rich and complex media such as Man–Rogosa–Sharpe (MRS) or fruit juice-based media in laboratory conditions. This raises experimental challenges in pure and concentrated phage preparations for further uses such as structure-function studies.