Cell Biology

Here, we present a protocol for implementing the fluorogen-activating protein FAST (fluorescence-activating and absorption-shifting tag) in fluorescence lifetime imaging microscopy (FLIM), which allows separating fluorescent species in the same spectral channel based on fluorescence lifetime properties. Previous studies have demonstrated FLIM multiplexing using various combinations of synthetic probes, fluorescent proteins, or self-labeling tags. In this protocol, we utilize engineered FAST point mutation variants that bind fluorogen HBR-2,5-DM. The designed probes possess nearly identical, compact protein sizes (14 kDa), and the resulting protein–fluorogen complexes demonstrate comparable steady-state optical properties and exhibit distinct fluorescence lifetimes, displaying monoexponential fluorescence decay kinetics. When FAST variants are expressed with localization signals, these properties facilitate robust signal separation in regions with co-localized or spatially overlapping labels (nucleus and cytoskeleton in this protocol) in live mammalian cells. This method can be applied to separate other overlapping cellular compartments, such as the nucleus and Golgi apparatus, or mitochondria and cytoskeleton.

High-content analysis (HCA) is a powerful image-based approach for phenotypic profiling and drug discovery, enabling the extraction of multiparametric data from individual cells. Traditional HCA protocols often rely on fixed-cell imaging, with assays like cell painting widely adopted as standard. While these methods provide rich morphological information, the integration of live-cell imaging expands analytical capabilities by enabling the study of dynamic biological processes and real-time cellular responses. This protocol presents a simple, cost-effective, and scalable method for live-cell HCA using acridine orange (AO), a metachromatic fluorescent dye that highlights cellular organization by staining nucleic acids and acidic compartments. The assay provides visualization of distinct subcellular structures, including nuclei and cytoplasmic organelles, using a two-channel fluorescence readout. Compatible with high-throughput microscopy and computational analysis, the method supports diverse applications such as phenotypic screening, cytotoxicity assessment, and morphological profiling. By preserving cell viability and enabling dynamic, real-time measurements, this live-cell imaging approach complements existing fixed-cell assays and offers a versatile platform for uncovering complex cellular phenotypes.

Formalin-fixed paraffin-embedded (FFPE) slides are essential for histological and immunohistochemical analyses of organoids. Conventional preparation of FFPE slides from organoids embedded in basement membrane extract (BME) presents several challenges. During the fixation step, dehydration often causes collapse of the BME, which normally supports the three-dimensional architecture of organoids. As a result, organoids may lose their original morphology, particularly in the case of cystic or structurally delicate types, leading to distortion and reduced reliability in downstream histological evaluation. Here, we introduce a straightforward protocol that improves the reliability of FFPE slide preparation for BME-based organoids by enhancing sample integrity and sectioning quality. By using 2% agarose as a mold during the embedding process, organoids grown in BME were effectively stabilized, enabling reliable preservation of their morphology throughout FFPE slide preparation. This method effectively addresses the difficulties in processing structurally delicate organoids and allows robust preparation of diverse cancer organoid morphologies—such as cystic, dense, and grape-like structures—while maintaining their native three-dimensional architecture. Our approach simplified the technical process while ensuring reliable histopathological analysis, making it a valuable tool for cancer research and personalized medicine.

Inherited germline variants are now recognized as important contributors to hematologic myeloid malignancies, but their reliable detection depends on obtaining uncontaminated germline DNA. In solid tumors, peripheral blood remains free of tumor cells and therefore serves as a standard source for germline testing. In contrast, peripheral blood often contains neoplastic or clonally mutated cells in hematologic malignancies, making it impossible to distinguish somatic from germline variants. This unique challenge necessitates using an alternative, non-hematopoietic tissue source for accurate germline assessment in patients with hematologic myeloid malignancies. Cultured skin fibroblasts derived from punch biopsies have long been considered the gold standard for this purpose. Nevertheless, most existing protocols are optimized for research settings and lack detailed, patient-centric workflows for routine clinical use. Addressing this translational gap, we present a robust, enzyme-free protocol for culturing dermal fibroblasts from skin punch biopsies collected at the bedside during routine bone marrow procedures. The method details practical bedside collection, sterile transport, mechanical dissection without enzymatic digestion, plating strategy, culture expansion, and high-yield DNA isolation with validated purity. By integrating this standardized approach into routine hematopathology workflows, the protocol ensures reliable germline material with minimal patient discomfort and a turnaround time suitable for clinical diagnostics.

Intestinal organoids are generated from intestinal epithelial stem cells, forming 3D mini-guts that are often used as an in vitro model to evaluate and manipulate the regenerative capacities of intestinal epithelial stem cells. Plating 3D organoids on different substrates transforms organoids into 2D monolayers, which self-organize to form crypt-like regions (which contain stem cells and transit amplifying cells) and villus-like regions (which contain differentiated cells). This “open lumen” organization facilitates multiple biochemical and biomechanical studies that are otherwise complex in 3D organoids, such as drug applications to the cell’s apical side or precise control over substrate protein composition or substrate stiffness. Here, we describe a protocol to generate homogenous intestinal monolayers from single-cell intestinal organoid suspension, resulting in de novo crypt formation. Our protocol results in higher viability of intestinal cells, allowing successful monolayer formation.

Rapid and uniform labeling of plasma membrane proteins is essential for high-resolution imaging of dynamic membrane topologies and intercellular communication in live mammalian cells. Existing strategies for labeling live cell membranes, such as fluorescent fusion proteins, enzyme-mediated tags, metabolic bioorthogonal labeling, and lipophilic dyes, face trade-offs in the requirement of genetic manipulation, the presence of non-uniform labeling, the need for extensive preparation times, and limited choices of fluorophores. Here, we present a streamlined protocol that leverages N-hydroxysuccinimide (NHS)-ester chemistry to achieve rapid (≤5 min), covalent conjugation of synthetic small-molecule dyes to surface-exposed primary amines, enabling pan-membrane-protein labeling. This workflow covers dye stock preparation, labeling for suspension and adherent cells, multiplex live-cell imaging, fusion protein co-staining (including insulin-triggered receptor endocytosis), 3D membrane visualization, and in vivo assays for visualizing membrane-derived material transfers between donor and recipient cells using a lymphoma T-cell mouse model. This high-density labeling approach is compatible with various cell types across diverse imaging platforms. Its speed, versatility, and stability make it a broadly applicable tool for studying plasma membrane dynamics and intercellular membrane trafficking.

Autophagy plays a crucial role in cellular homeostasis and is responsible for removing and degrading damaged cytoplasmic cargo. This lysosome-mediated catabolic process removes defective organelles and misfolded proteins, and impaired autophagy has been directly linked to ageing and numerous diseases. This emphasises the importance of developing intervention methods to counteract this dysregulation. One promising intervention is thermal therapy, specifically hyperthermia, which is described in this protocol. In order to investigate this form of treatment, a rapid and reliable detection method is required to allow comparison of autophagy status under different conditions. While methods such as transmission electron microscopy (TEM) or western blotting can provide valuable structural analysis, they are often time-consuming and expensive, and are not suitable for small, round cells such as peripheral blood mononuclear cells (PBMCs). The method described in this protocol enables absolute quantification of PBMCs using the Guava^® Autophagy Detection kit after heat treatment with water-filtered infrared-A radiation (wIRA), compared with an untreated control. This method is based on antibody labelling, and subsequent flow cytometric analysis enables the number of autophagosomes to be determined by measuring the FITC intensity. This protocol provides rapid, reliable results and can be adapted to investigate not only heat therapy, but also other interventions, such as caloric restriction.

Crypts at the base of intestinal villi contain intestinal stem cells (ISCs) and Paneth cells, the latter of which work as niche cells for ISCs. When isolated and cultured in the presence of specific growth factors, crypts give rise to self-renewing 3D structures called organoids that are highly similar to the crypt-villus structure of the small intestine. However, the organoid culture from whole crypts does not allow investigators to determine the contribution of their individual components, namely ISCs and Paneth cells, to organoid formation efficiency. Here, we describe the method to isolate Paneth cells and ISCs by flow cytometry and co-culture them to form organoids. This approach allows the determination of the contribution of Paneth cells or ISCs to organoid formation and provides a novel tool to analyze the function of Paneth cells, the main component of the intestinal stem cell niche.

Cell–surface and cell–cell interaction assays are fundamental for studying receptor–ligand interactions and characterizing cellular responses and functions. They play a critical role in diagnostics and in modulating immune system activity for therapeutic applications, notably in cancer immunotherapy. By providing time-lapsed and cell-level direct observation of the sample, optical microscopy offers strong advantages compared to current go-to techniques, which are typically either ensemble methods (e.g., measuring cell populations) or indirect readouts (e.g., impedance for adherent cells). This protocol describes two complementary microscopy-based assays: (1) a cell–surface ligand binding assay to quantify dynamic interactions between human primary Natural Killer (NK) cells and a cancer-mimicking surface, and (2) a cell–cell interaction assay to evaluate antibody-dependent cell cytotoxicity (ADCC) mediated by NK cells targeting tumor cells. Additionally, the protocol uses Celldetective, a new open graphical user interface for quantitative analysis of cell interaction dynamics from 2D time-lapse microscopy datasets. Although applied here to primary immune cells, these methods are adaptable to various cell types, including other immune cells, fibroblasts, and cancer cells. This approach enables direct observation and quantification of cellular morphology, motility, cell–cell interactions, and dynamic behaviors at single-cell resolution over time, facilitating detailed analysis of mechanisms such as cell death, migration, and immune synapse formation.

Proper genome organization is essential for genome function and stability. Disruptions to this organization can lead to detrimental effects and the transformation of cells into diseased states. Individual chromosomes and their subregions can move or rearrange during transcriptional activation, in response to DNA damage, and during terminal differentiation. Techniques such as fluorescence in situ hybridization (FISH) and chromosome conformation capture (e.g., 3C and Hi-C) have provided valuable insights into genome architecture. However, these techniques require cell fixation, limiting studies of the temporal evolution of chromatin organization in detail. Our understanding of the heterogeneity and dynamics of chromatin organization at the single-cell level is still emerging. To address this, clustered regularly interspaced short palindromic repeats (CRISPR)/dead Cas9 (dCas9) systems have been repurposed for precise live-cell imaging of genome dynamics. This protocol uses a system called CRISPRainbow, a powerful tool that allows simultaneous targeting of up to seven genomic loci and tracks their locations over time using spectrally distinct fluorescent markers to study real-time chromatin organization. Multiple single-guide RNA (sgRNA), carrying specific RNA aptamers for labeling, can be cloned into a single vector to improve transfection efficiency in human cells. The precise targeting of CRISPRainbow offers distinct advantages over previous techniques while also complementing them by validating findings in live cells.