Cell Biology

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.

Adult muscle stem cells (MuSCs) are the key cellular source for regenerating skeletal muscle in vertebrates. MuSCs are typically identified in skeletal muscle by the expression of the paired box protein 7 (PAX7) protein. Here, we developed a combined RNA fluorescent in situ hybridization (FISH) using RNAscope technology and an immunofluorescence (IF) protocol for the simultaneous detection of Pax7 mRNA and PAX7 protein in individual MuSCs in vivo. Interestingly, we show that while most PAX7⁺ (protein) MuSCs express Pax7 mRNA, there is a subset of Pax7⁺ (mRNA) cells that do not express PAX7 protein. Altogether, we developed a combined FISH/IF protocol that allows for the co-detection of mRNA and protein in MuSCs in vivo, a strategy that can be applied to any target gene. The functional significance of the Pax7-expressing subset of cells lacking PAX7 protein prior to injury remains unknown.

High-throughput sequencing has created a tremendous amount of information about the genes expressed in various cell types and tissues throughout the body. As such, there is a need for a quick and effective method to knock down genes of interest in order to investigate their roles. While there are many approaches for this in mammalian models, there are limited ways to knock down genes of interest in adult zebrafish. Unlike mammals, zebrafish have the natural ability to regenerate their neurons after injury or disease is detected, making them a staple in regenerative studies. Unfortunately, current approaches for gene knockdown in the retina of adult zebrafish are costly and provide a barrier for many scientists. We provide two cost-effective approaches for targeted gene knockdowns in adult zebrafish retinas. We describe this approach through the use of Vivo-morpholinos and lipid-encapsulated siRNAs that target the expression of the proliferating cell nuclear antigen (PCNA) gene in adult zebrafish. We also describe how to collect and process retina samples for downstream immunohistochemistry, imaging, and quantification. Overall, this protocol will provide researchers with a straightforward, cheap, and effective method to perform targeted gene knockdowns in adult zebrafish retinas.

The female reproductive tract is comprised of different regions, each with distinctive physiological characteristics. One of them is the fallopian tubes, which are vital for human reproductive health and success. The ability to model their function and physiology is of utmost importance. So far, in vitro models have been based on a few immortalized or cancer cell lines derived from fallopian tube cells that lacked differentiated, specialized cell types and did not allow for the study of cancer initiation due to their implicit biases. Organoids, in contrast, overcome these limitations and provide an advanced, three-dimensional system for the study of healthy fallopian tube physiology and pathology. Fallopian tube organoids are comprised of epithelial progenitors that can be enriched using chemical or hormonal treatment into the different cell types that are found in the in vivo tissue, namely detyrosinated-tubulin-positive ciliated cells or paired-box protein 8 (PAX8)-positive secretory cells. This protocol provides a step-by-step guide for the establishment and maintenance of a long-term culture of organoids from healthy human fallopian tube tissue. The organoid model described here closely mimics the in vivo physiology and anatomy of human fallopian tube epithelium and provides a comprehensive basis for future studies on its underlying molecular characteristics and possible pathology.

Cell transplantation is a promising strategy for treating age-related muscle atrophy, but its critical application remains limited. Cultured myoblasts, unlike freshly isolated muscle stem cells, show poor engraftment efficiency and fail to contribute effectively to muscle regeneration. Moreover, successful engraftment generally requires prior muscle injury, as skeletal muscle regeneration is typically triggered by a damaged microenvironment. These limitations present major obstacles for applying cell therapy to sarcopenia, where muscle degeneration occurs without injury. In this protocol, we describe a novel approach that enables the transplantation of cultured myoblasts into intact skeletal muscle without the need for preexisting injuries or genetic modification. By combining myoblasts with extracellular matrices (ECM), such as Matrigel, which mimic the native muscle niche and support cell survival, adhesion, proliferation, and differentiation, we achieve efficient engraftment and increased muscle mass without the need for preexisting injury. The ECM also provides a scaffold and retains bioactive factors that enhance the regenerative capacity of transplanted cells. This is the first protocol that enables robust myoblast engraftment in non-injury muscle conditions, offering a practical tool for studying and potentially treating sarcopenia.

Chemically induced murine colitis models are widely used to understand intestinal homeostasis and inflammatory responses during acute and chronic gut inflammation, such as inflammatory bowel disease (IBD). Resident populations of immune cells, together with those recruited during an inflammatory response, maintain intestinal immunity by mounting an effective immune response to enteropathogenic microbes while at the same time maintaining tolerance against commensals. To better understand the disease mechanism, studying different immune cell populations and their dynamic changes during infection and inflammation is essential. However, isolating healthy and viable immune populations, particularly hyperactivated neutrophils and macrophages from the inflamed gut (i.e., active disease site), is challenging as tissues are usually subjected to rigorous enzymatic digestion for an extended period. Here, we describe a method that uses a cell dissociator (Medimachine II from Syntec International) to separate intestinal tissue after short enzymatic digestion to obtain a single-cell suspension. This technique facilitates the isolation of immune cells from mouse intestinal tissues in high quantity and with superior viability in a very short time frame. This protocol delivers 80%–90% cell viability, which is 1.5 to 2-fold higher than conventional methods of isolating cells from inflamed mouse colons. The composition, phenotype, activation state, and gene expression profile of cells isolated using this protocol can be assessed by using multiple methods, including, but not limited to, flow cytometry, quantitative PCR, immunoblotting, mass spectrometry, single-cell RNA sequencing, and functional readouts such as reactive oxygen species (ROS) production.

Protein–protein interactions facilitate cellular functions through the creation of networks and multi-protein complexes. Mapping the interactions within and between protein networks and elucidating the composition of protein complexes provides critical insight into biological processes. Interactions among soluble cytoplasmic proteins have been extensively investigated through the application of immunoaffinity capture as well as conventional nuclear two-hybrid testing. The integrated membrane yeast two-hybrid provides a method to investigate protein–protein interactions between integral membrane proteins in their native membrane environment. This procedure makes use of the ability of the amino-terminal fragment of ubiquitin (Nub) and the carboxyl-terminal fragment of ubiquitin (Cub) to refold reconstituting functional ubiquitin, which can be recognized by a ubiquitin peptidase. Appending a fusion protein composed of Cub fused to LexA and VP16 (CLV) to a candidate "bait" protein and Nub to candidate "prey" proteins allows a test of their interaction. If the two proteins interact closely, the CLV fragment is cleaved and enters the nucleus to activate the expression of reporter genes, signaling the interaction. When the bait and prey proteins are tagged with CLV and NubG, respectively, at their genomic loci, they are only copies of the bait and prey in the cell and are expressed under the regulation of their native promoters. This avoids overexpression artifacts that can occur if the tagged proteins are expressed from plasmids while the untagged chromosomally encoded copies of the bait and prey continue to be expressed.

This protocol describes an ex vivo co-culture method to assess CD8⁺ T-cell activation, proliferation, and cytotoxic potential using bulk splenocytes isolated from immunocompetent mice. Mouse splenocytes are stimulated with anti-CD3 and anti-CD28 antibodies to activate CD8⁺ T cells, which are then co-incubated with either cancer cells or cancer cell–derived conditioned media (CM) to evaluate tumor-driven modulation of immune cell functions. The use of unfractionated splenocytes preserves physiological cell–cell interactions, eliminating the need for exogenous interleukin (IL-2) and bypassing flow sorting, which simplifies the workflow and reduces experimental variability. CD8⁺ T-cell responses are measured via flow cytometry, using markers of proliferation (CFSE dilution), activation (CD69), and effector function (Granzyme B and IFNγ). Additionally, immune-mediated tumor cell death is evaluated by Annexin-V/7-AAD staining. Together, this experimental platform supports the investigation of both cell contact-dependent and contact-independent mechanisms of immune cell modulation in a cost-effective and reproducible setting.

Regulatory T cells (Tregs) are essential for maintaining immune balance by controlling the activation and expansion of other immune cells. Conventional suppression assays often rely on co-culturing purified cell populations, which limits multiplexed phenotyping and physiological relevance. This protocol describes a high-dimensional, single-cell assay for profiling Treg-mediated suppression within a peripheral blood mononuclear cell (PBMC) system. Tregs are first isolated by cell sorting and then reintroduced into autologous PBMCs at defined ratios. A 52-marker mass cytometry (CyTOF) panel is used to quantify cell division and phenotypic responses across multiple immune subsets. This approach allows for integrated analysis of Treg function with broad compatibility for patient profiling and drug evaluation.