细胞生物学

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.

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.

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.

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.

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.

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.

Proper brain function depends on the integrity of the blood–brain barrier (BBB), which is formed by a specialized network of microvessels in the brain. Reliable isolation of these microvessels is crucial for studying BBB composition and function in both health and disease. Here, we describe a protocol for the mechanical dissociation and density-based separation of microvessels from fresh or frozen human and murine brain tissue. The isolated microvessels retain their molecular integrity and are compatible with downstream applications, including fluorescence imaging and biochemical analyses. This method enables direct comparisons across species and disease states using the same workflow, facilitating translational research on BBB biology.

Protein synthesis and degradation (i.e., turnover) forms an important part of protein homeostasis and has been implicated in many age-associated diseases. Different cellular locations, such as organelles and membraneless compartments, often contain individual protein quality control and degradation machineries. Conventional methods to assess protein turnover across subcellular compartments require targeted genetic manipulation or isolation of specific organelles. Here we describe a protocol for simultaneous proteome localization and turnover (SPLAT) analysis, which combines protein turnover measurements with unbiased subcellular spatial proteomics to measure compartment-specific protein turnover rates on a proteome-wide scale. This protocol utilizes dynamic stable isotope labeling of amino acids in cell culture (dynamic SILAC) to resolve the temporal information of protein turnover and multi-step differential ultracentrifugation to assign proteins to multiple subcellular localizations. We further incorporate 2D liquid chromatography fractionation to greatly increase analytical depth while multiplexing with tandem mass tags (TMT) to reduce acquisition time 10-fold. This protocol resolves the spatial and temporal distributions of proteins and can also reveal temporally distinct spatial localizations within a protein pool.

Accurate identification of cell cycle stages is essential for investigating fundamental biological processes such as proliferation, differentiation, and tumorigenesis. While flow cytometry remains a widely used technique for such analyses, it is limited by its lack of single-cell resolution and its requirement for large sample sizes due to its population-based approach. These limitations underscore the need for alternative or complementary methods that offer single-cell precision with compatibility for small-scale applications. We present ImmunoCellCycle-ID, an immunofluorescence-based method that leverages the spatial distribution of endogenous markers, such as DNA, proliferating cell nuclear antigen (PCNA), centromere protein F (CENP-F), and centromere protein C (CENP-C), to reliably distinguish G1, early S, late S, early G2, late G2, and all mitotic sub-stages. This technique does not rely on precise signal quantification and utilizes standard immunofluorescence protocols alongside conventional laboratory microscopes, ensuring broad accessibility. Importantly, ImmunoCellCycle-ID detects endogenous proteins without the need for genetic modification, making it readily applicable to a wide range of human cell lines. Beyond its utility for single-cell resolution, the method can be scaled for population-level analyses, similar to flow cytometry. With its precision, versatility, and ease of implementation, ImmunoCellCycle-ID offers a powerful tool for high-resolution cell cycle profiling across diverse experimental platforms.

Studying G protein-coupled receptor (GPCR) activation of heterotrimeric G proteins is crucial for understanding diverse physiological processes and developing novel therapeutics. Traditional methods to assay GPCR activation of G proteins, including assays of second messengers and biosensors, involve complex or indirect procedures. However, second messengers like cAMP and calcium are not direct readouts of GPCR activity due to signaling crosstalk, while biosensors can have undesired consequences due to structural alteration caused by fluorescent protein insertion. Here, we present a streamlined protocol employing GST-tagged bait proteins and epitope-embedded Gα subunits to achieve direct monitoring of Gα activity within cells. This method involves purification of GST-tagged bait constructs from bacteria and subsequent direct interaction studies with GluGlu-tagged Gα proteins expressed in any human cells of interest by including GST-tagged bait proteins in the cell lysis buffer. The approach enables sensitive detection of activated Gα within cells following extracellular stimulation. Advantages of this protocol include high sensitivity, enhanced monitoring of GPCR signaling dynamics under physiologically relevant conditions with minimum alteration in Gα, and the ability to distinguish between highly homologous isoforms within the same Gα family.