Nowadays, the use of 3D cultures (organoids) is considered a valuable experimental tool to model physiological and pathological conditions of organs and tissues. Organoids, retaining cellular heterogeneity with the presence of stem, progenitor, and differentiated cells, allow the faithful in vitro reproduction of structures resembling the original tissue. In this context, the growth of endometrial organoids allows the generation of 3D cultures characterized by a hollow lumen, secretory activity, and apicobasal polarity and displaying phenotypical modification in response to hormone stimulation. However, a limitation in currently used models is the absence of stromal cells in their structure; as a result, they miss epithelial–stromal interactions, which are crucial in endometrial physiology. We developed a novel 3D model to generate endometrial organoids grown in floating MatrigelTM droplets in the presence of standard culture medium. From a structural point of view, these novel floating 3D cultures develop as gland-like structures constituted by epithelial cells organized around a central lumen and retain the expression of endometrial and decidual genes, like previously published organoids, although with a phenotype resembling hormonally differentiated structures. Importantly, floating organoids retain stromal cells which grow in close contact with the epithelial cells, localized within the internal or external portion of the organoid structure. In summary, we present a simple and rapid model for generating 3D endometrial organoids that preserve epithelial–stromal cell interactions, promoting the formation of differentiated organoids and enabling the study of reciprocal modulation between epithelium and stroma.Key features• Development of 3D endometrial organoids using 10% FBS-DMEM/F12 complete medium in less than a month.• Preservation of stromal and epithelial cell populations, favoring the study of their interaction.• Development of differentiated endometrial organoids without the need for in vitro hormonal stimulation.

Biomolecular condensates organize cellular processes through liquid–liquid phase separation, creating membrane-less compartments enriched in specific proteins and RNAs. Understanding their RNA composition is essential for elucidating plant stress responses, yet capturing these transiently associated RNAs remains technically challenging. We present Turbo-RIP (TurboID-based proximity labeling with RNA immunopurification), a comprehensive protocol for identifying condensate-associated RNAs in plants. Turbo-RIP employs the biotin ligase TurboID to label proximal proteins at 22 °C, followed by formaldehyde crosslinking and streptavidin-based capture of protein–RNA complexes. We provide detailed procedures for three cloning strategies, transformation of Nicotiana benthamiana and Arabidopsis thaliana, validation of TurboID activity, and RNA recovery. The protocol successfully captured processing body–associated RNAs with minimal background. Turbo-RIP enables systematic mapping of RNA populations within plant condensates under diverse conditions. The protocol requires 3–5 days from sample preparation to RNA isolation, with construct validation taking 2–4 weeks. All procedures use standard laboratory equipment, making Turbo-RIP accessible for plant molecular biology laboratories.Key features• Apply TurboID-based proximity labeling specifically for RNA capture in plant condensates.• Optimize experimental conditions for different plant species and condensate types.• Implement quality control measures and data analysis pipelines.

Our genome is duplicated during every round of cell division through the process of DNA replication, but this fundamental process is subjected to various stresses arising from endogenous or exogenous sources. Thus, studying replication dynamics is crucial for understanding the mechanisms underlying genome duplication in physiological and replication stress conditions. Earlier, radioisotope-based autoradiography and density-labeling methods were used to study replication dynamics, which were limited in spatial resolution, representing only average estimates from many DNA samples. Here, we describe a DNA fiber assay that utilizes different thymidine analog incorporation, like 5-chloro-2’-deoxyuridine (CldU) and 5-iodo-2’-deoxyuridine (IdU), into replicating DNA. Such labeled DNA can be stretched and fixed on silanized glass slides, which are denatured with mild acidic treatment to expose the labeled nascent DNA. This DNA can then be visualized by using primary antibodies against CldU and IdU, followed by fluorophore-conjugated secondary antibodies, and observing them using a fluorescence microscope. The DNA fiber assay allows the visualization of individually replicating DNA at a single-molecular resolution and is highly quantitative, high-throughput, and easily reproducible. This technique offers insights into different replication parameters, like rate of DNA synthesis, extent of reversed fork protection, restart of stalled forks, and fork asymmetry under untreated or replication stress conditions at a single-molecule level.Key features• Single-molecule resolution of DNA replication dynamics.• Diverse replication parameters can be quantified with variations of the labeling protocol.• Easily reproducible across different cell lines in any lab with a basic molecular biology setup.• Effective in studying the effects of different genotoxic treatments on replication.

The extracellular matrix (ECM) critically shapes melanoma progression and therapeutic response, yet commonly used matrices such as Matrigel fail to capture tissue- and disease-specific ECM properties. This protocol provides a streamlined and scalable method for generating murine, tissue-specific ECM hydrogels from skin, lung, and melanoma tumors, therefore overcoming the restricted materials of mouse-derived ECM. The workflow integrates tissue-tailored decellularization, lyophilization, mechanical fragmentation, pepsin digestion, and physiological polymerization to produce hydrogels that reliably preserve fibrillar collagen architecture and organ-specific ECM cues. Decellularization efficiency and ECM integrity are validated by DNA quantification, H&E staining, and Picrosirius Red staining analysis. These hydrogels provide a species- and tissue-matched platform for studying melanoma–ECM–immune interactions, pre-metastatic niche features, and therapy-induced ECM remodeling. Overall, this protocol offers a reproducible and physiologically relevant ECM model that expands experimental capabilities for melanoma biology and treatment-resistance research and that can be easily extended to other tumors and tissues.Key features• A miniaturized, tissue-specific workflow for generating ECM hydrogels from small murine skin, lung, and melanoma tissues, overcoming size limitations of existing protocols.• Preservation of native ECM architecture using tailored decellularization steps validated by DNA quantification, H&E, and Picrosirius Red staining.• A standardized digestion–gelation process optimized for heterogeneous and lipid-rich murine tissues, enabling reproducible hydrogel formation at defined ECM concentrations.• A physiologically relevant platform capturing melanoma- and organ-specific ECM cues for studying ECM–tumor–immune interactions and therapy-induced remodeling.

Toxoplasma gondii is an apicomplexan parasite that infects a wide variety of eukaryotic hosts and causes toxoplasmosis. The cell cycle of T. gondii exhibits a distinct architecture and regulation that differ significantly from those observed in well-studied eukaryotic models. To better understand the tachyzoite cell cycle, we developed a fluorescent ubiquitination-based cell cycle indicator (FUCCI) system that enables real-time visualization and quantitative assessment of the different cell cycle phases via immunofluorescence microscopy. Quantitative immunofluorescence and live-cell imaging of the ToxoFUCCIS probe with specific cell cycle markers revealed substantial overlap between cell cycle phases S, G2, mitosis, and cytokinesis, further confirming the intricacy of the apicomplexan cell cycle. Key features • This protocol describes the development of the transgenic lines capable of detecting individual cell cycle phases and processes of the Toxoplasma tachyzoite cell cycle. • Quantitative immunofluorescence analysis and real-time microscopy enable the measurement of each cell cycle phase. • The ToxoFUCCIS probe helps to gain new insights into the highly flexible, overlapping nature of cell cycle organization in apicomplexan parasites.

To study gene function in regulating rodent retinal waves during development, an efficient method for gene delivery into whole-mount retinas is required while preserving circuit functionality for physiological studies. We present an optimized electroporation protocol for developing rodent retinal explants. The procedure includes the fabrication of horizontally aligned platinum electrodes and the placement of retinal explants between them to generate a uniform electric field for high transfection efficiency. The entire process-dissection and electroporation-can be completed within 1-2 h. Successful transfection is verified by fluorescence microscopy, and physiological assays such as patch-clamp recordings and live imaging can be performed within 1-4 days following electroporation. This rapid and reliable protocol enables functional analysis for a specific gene in regulating retinal waves and can be adapted to other organotypic slice cultures. Key features • Incorporates horizontally aligned platinum electrodes and enables cell type-specific promoters to drive gene expression for physiological studies. • Preserves retinal wave activity while markedly improving transfection efficiency in whole-mount postnatal rodent retinas. • Requires only 1-2 h from retinal dissection to electroporation. • Allows completion of functional experiments within four days after electroporation.

Underwater noise is a growing source of anthropogenic pollution in aquatic environments. However, few studies have evaluated the impact of underwater noise on aquatic invertebrates. More importantly, studies involving early developmental stages have been poorly addressed. Significant limitations are due to the lack of standardized protocols for working in the laboratory. Particularly, the design of uniform procedures in the laboratory is important when working with species that inhabit short-term changing habitats, such as estuaries, which makes it difficult to carry out repeated experiments in the natural habitat. Besides, controlling for environmental variables is also important when assessing the effect of a stressor on the physiological parameters of individuals. This experimental protocol addresses that gap by offering an adaptable laboratory-based method to evaluate sublethal physiological responses to sound exposure under highly controlled conditions. Here, we present a reproducible and accessible laboratory protocol to expose crabs to recorded boat noise and evaluate physiological responses using oxidative stress biomarkers. The method is designed for ovigerous females, as we evaluated the effects on embryos and early life stages (i.e., larvae), but it can be readily adapted to different life stages of aquatic invertebrates. A key strength of this protocol is its simplicity and flexibility: animals are exposed to noise using submerged transducers under well-controlled laboratory conditions, ensuring consistency and repeatability. Following exposure, tissues or whole-body samples can be processed for a suite of oxidative stress biomarkers-glutathione-S-transferase (GST), catalase (CAT), lipid peroxidation (LPO), and protein oxidation. These biomarkers are highly responsive, cost-effective indicators that provide a sensitive and early readout of sublethal stress. Together, the exposure and analysis steps described in this protocol offer a powerful and scalable approach for investigating the physiological impacts of underwater noise in crustaceans and other aquatic invertebrates. Key features • Enables measurement of oxidative stress markers across different life stages-from embryos to larvae and adult tissues-offering a complete view of physiological impact. • Ensures consistent, reproducible conditions through standardized exposure and sampling, supporting reliable comparisons across experiments. • Flexible protocol adaptable to Neohelice granulata and other estuarine decapods or marine benthic invertebrates, broadening its applicability.

Adipogenic differentiation efficiency remains highly variable across laboratories and cellular models, underscoring a critical need for a robust and standardized protocol. Here, we describe an optimized and highly effective protocol for inducing adipogenesis in multiple models, including murine 3T3-L1 preadipocytes, stromal vascular fraction (SVF) from neonatal and adult mice, and human adipose-derived stem cells (hADSCs). Systematic optimization was performed on key parameters such as initial cell confluence, induction timing, inducer composition, and culture surface coating. We show that high cell density, rosiglitazone supplementation, and an extended primary induction phase combine to promote lipid accumulation. Notably, we introduce a crucial modification-prolonged low-dose insulin stimulation during the maintenance phase-that is essential for the efficient differentiation of adult SVF. Furthermore, when applied to hADSCs, the protocol consistently induced robust adipogenesis, confirming its cross-species applicability. Taken together, this comprehensive and reproducible protocol serves as a valuable tool for advancing in vitro adipogenesis research. Key features • Extend a robust, standardized adipogenic differentiation protocol from 3T3-L1 preadipocytes to clinically relevant models, including hADSCs and the heterogeneous SVF. • Identify key optimized parameters-cell density, induction timing, and inducer composition-enabling highly reproducible differentiation across species.

Flagellate stages of green microalgae such as Trebouxia are only partially characterised, with recent evidence suggesting that they are involved in both sexual and asexual reproduction. Conventional methods based on fixed samples in light, confocal, or electron microscopy provide only static observations and prevent real-time monitoring of living cells. To overcome this limitation, we have developed a simple and cost-effective protocol for observing Trebouxia flagellate cells over several days by coating microscopy slides with Bold’s basal medium. The method preserves cell viability and allows repeated imaging of motile cells in the same areas so that their behaviour and development can be continuously observed. In this way, qualitative observations, such as flagellate cell release, motility, and gamete fusion, can be combined with quantitative analyses of cell morphology. The protocol has proven to be robust and reproducible and was applied to several Trebouxia species. Compared to existing techniques, it allows the monitoring of dynamic processes and provides a powerful tool to study specific life stages not only in Trebouxia but also in other unicellular and colonial green algae.Key features• This protocol allows real-time monitoring over several days of Trebouxia flagellate cells with standard light microscopy.• This protocol preserves cell viability and motility for repeated daily observations of the same cell groups.• This protocol is simple, low-cost, and adaptable to other motile algal cells.• This protocol is based on the methodology described in [1], where it was originally applied and validated.

Although protein–protein interactions (PPIs) are central to nearly all biological processes, identifying and engineering high-affinity intracellular binders remains a significant challenge due to the complexity of the cellular environment and the folding constraints of proteins. Here, we present a two-stage complementary platform that combines magnetic-activated cell sorting (MACS)-based yeast surface display with functional ligand-binding identification by twin-arginine translocation (Tat)-based recognition of associating proteins (FLI-TRAP), a bacterial genetic selection system for efficient screening, validation, and optimization of PPIs. In the first stage, MACS-based yeast display enables the rapid high-throughput identification of candidate binders for a target antigen from a large synthetic-yeast display library through extracellular interaction screening. In the second stage, an antigen-focused library is subcloned into the FLI-TRAP system, which exploits the hitchhiker export process of the Escherichia coli Tat pathway to evaluate binder–antigen binding in the cytoplasm. This stage is achieved by co-expressing a Tat signal peptide–tagged protein of interest with a β-lactamase-tagged antigen target, such that only binder–antigen pairs with sufficient affinity are co-translocated into the periplasm, thus rendering the bacterium β-lactam antibiotic resistant. Because Tat-dependent export requires fully folded and soluble proteins, FLI-TRAP further serves as a stringent in vivo filter for intracellular compatibility, folding, and stability. Therefore, this approach provides a powerful and cost-effective pipeline for discovering and engineering intracellular protein binders with high affinity, specificity, and functional expression in bacterial systems. This workflow holds promise for several applications, including synthetic biology and screening of theragnostic proteins and PPI inhibitors.Key features• Combines a single round of MACS enrichment with FLI-TRAP for high-throughput Nb discovery.• Reduces time and resource demands compared to traditional workflows involving multiple rounds of MACS/FACS.• Enables in vivo selection of high-affinity, functional binders via Tat-dependent export linked to β-lactam resistance, correlating binding affinity and solubility with antibiotic resistance.