细胞生物学

Cyclic peptides are emerging as a promising class of recognition modules for chimeric antigen receptor (CAR) engineering. Compared with single-chain variable fragment (scFv)-based CARs, disulfide-directed multicyclic peptides (DDMPs) represent a novel alternative, offering a markedly smaller molecular size (<5 kDa), enhanced structural stability through disulfide-directed cyclization, and broad tolerance to sequence diversification that supports systematic affinity and specificity optimization. DDMP-based CAR T cells leverage these properties to mediate antigen-dependent cytotoxicity while exhibiting an attenuated cytokine secretion profile, supporting the development of potentially safer immunotherapies for solid tumors. Here, we present a comprehensive workflow spanning CAR construct design and generation through in vitro and in vivo functional evaluation. While DDMPs are used as the exemplar recognition module, sections A and C–L of the protocol are directly applicable to any CAR format, including scFv- and nanobody-based designs with minimal modifications, making the workflow accessible to the broader CAR T-cell research community. The protocol includes the generation of Jurkat NFAT reporter cell lines and luciferase-expressing tumor target lines, which are widely used in different assays. Together, these standardized readouts enable rigorous, objective comparison of CAR T-cell efficacy and safety across tumor models.

When the function of cardiac capillaries is impaired, cardiac function declines, and the risk of disease increases. No reliable assay has been developed to detect or evaluate the level of material exchange of capillaries deep within healthy heart tissue. In this study, we develop a new method to detect and evaluate molecules leaking from capillaries in cardiac tissue. By administering fluorescent dextran to mice via the tail vein, followed by rapid processing of the heart tissue, we have detected leaking fluorescent material from intracardiac microvessels. By comparing the detected images with those taken during the negative-control administration, using the image processing software LAS X and ImageJ, we detected trace amounts of fluorescent material that had leaked from the capillaries. We calculated the area of tissue where fluorescence was detected to perform a quantitative assessment, which we used as an indicator of capillary permeability. This new method of indexing will provide a different perspective on the factors contributing to the decline in cardiac function and the increased risk of disease with aging.

Protein kinase B, more commonly known as Akt, is a family of three serine/threonine kinases (Akt1, Akt2, and Akt3) that play a central role in regulating processes such as proliferation, survival, metabolism, and migration through phosphorylation of downstream targets. Given its involvement in numerous cellular processes, aberrant Akt signaling is prevalent across multiple cancer types, underscoring the need for Akt kinase assays to assess activity, regulatory mechanisms, and the efficacy of targeted interventions. Most existing Akt kinase assays rely on expensive commercial kits, some of which employ pre-purified, constitutively active Akt expressed in insect cells, bypassing physiologic autoinhibition of Akt; therefore, they are not suitable for evaluating allosteric inhibitors or context-dependent regulation. Here, we describe a detailed, step-by-step protocol for a nonradioactive Akt kinase assay using epitope-tagged, recombinant Akt1 expressed in a mammalian cell line and isolated by immunoprecipitation. This method eliminates the need to co-express Akt with upstream regulatory kinases or to purify active enzyme from insect cells, a time-consuming and technically demanding process, particularly when analyzing multiple Akt mutants. Because Akt is assayed in a regulated, autoinhibited state, this protocol enables direct evaluation of allosteric inhibitors that cannot be assessed using active Akt purified from insect cells. We note, however, that Akt1 kinase activity in this assay is measured from epitope-tagged, transiently overexpressed protein, which could influence cellular signaling dynamics. Despite this limitation, the cellular context preserves key regulatory features of Akt1 autoinhibition and membrane-dependent activation that are absent in assays using purified, pre-activated kinase. Together, this protocol supports analysis of Akt kinase activity under diverse experimental conditions, including receptor stimulation, pharmacologic treatment, allosteric inhibitor exposure, and mutations, using an accessible, economical, and physiologically relevant approach.

Whole-mount techniques are widely used in medical and biological research to analyze protein expression and tissue organization in intact specimens. Traditional approaches for protein localization include section-based immunohistochemistry and in situ hybridization; however, these methods can be limited by tissue disruption and loss of spatial context. Whole-mount protocols generally involve tissue fixation, permeabilization, and staining with specific probes, but their effectiveness varies depending on the antigen–antibody combination and the specimen type. Consequently, no universal protocol is suitable for all experimental conditions. This protocol presents a detailed whole-mount immunostaining protocol for evaluating tyrosine hydroxylase (TH) expression, a key marker of dopaminergic neurons, in zebrafish (Danio rerio) larvae. The procedure outlines critical steps from sample preparation to staining optimization to ensure reproducible and specific signal detection. This approach enables accurate visualization and analysis of dopaminergic neuron distribution in intact larvae. The protocol offers a reliable and adaptable approach that preserves tissue integrity, enables three-dimensional visualization, and is particularly well-suited for developmental and neurobiological studies using zebrafish larvae.

The binding of transmembrane (TM) ligands to their cognate TM receptors on neighboring cells governs intercellular adhesion and direct cell–cell communication. However, these interactions are difficult to study in vitro because they depend on membrane presentation, ligand orientation, receptor clustering, and avidity, features often not captured by soluble recombinant ligands or cell-free assays. Here, we describe a flow cytometry–based assay using fluorescent, lentiviral virus-like particles (VLPs) displaying TM ligands to quantify binding to their receptors on target cells. Fluorescent VLPs are generated in-house by plasmid transfection in HEK293T cells and enable direct fluorescent detection without fluorochrome-conjugated secondary antibodies. The system is modular and readily accommodates engineered ligand constructs, including patient-derived variants. We applied this platform to generate ICAM-1-displaying fluorescent VLPs and to study human LFA-1 function in patient-derived leukocytes. This protocol provides a detailed workflow for VLP production and in vitro binding assays, offering a simple, quantitative, and cost-effective approach for studying TM ligand–receptor interactions in a membrane context. The system is well-suited for mechanistic studies, functional assessment of patient-derived variants, and direct binding assays using patient-derived cells. Integrating the assay into multicolor flow cytometry panels enables simultaneous immunophenotyping and quantification of up to four ligand–receptor interactions at single-cell resolution.

Thin membrane protrusions in cells help them communicate, create traction forces during their movement, and coordinate complex development in multicellular organisms. These structures include cytonemes, tunneling nanotubes, and microtubule-based nanotubes (MT-nanotubes), each with a different cytoskeletal constitution and function. Actin-based cytonemes help deliver signaling molecules, while microtubule-based nanotubes assist with transporting vesicles and organelles. Despite their physiological role, we still do not fully understand how these thin membrane protrusions form and function. In this study, we introduce an improved live-cell imaging method to observe polar cell protrusions during micropyle morphogenesis in developing Drosophila eggs. This technique combines precise developmental staging, careful dissection, and optimized ex vivo culture conditions to maintain tissue health during extended imaging. We also fine-tuned the imaging settings to reduce phototoxicity and thermal stress. This allows for continuous, high-resolution tracking of protrusion dynamics in real time. Our protocol addresses major drawbacks of fixed-tissue methods by capturing the entire process of protrusion formation, extension, and remodeling in intact living tissue. Additionally, it works well with drugs, making it a useful tool for functional studies. Overall, this approach builds a strong foundation for exploring membrane protrusion biology. It can also be applied to investigate similar developmental processes in other systems, aiding our understanding of normal development and diseases.

In vivo imaging of brown algal cells in 3D is extremely challenging because of the presence of pigments, such as fucoxanthin and chlorophyll, that diffract light. Moreover, brown algae live in seawater, a high ionic environment that can change the fluorochrome behavior or cause aggregates. Despite the importance of in vivo monitoring the developmental process of brown algal tissues, 4D imaging (x, y, z, t) on a conventional fluorescence microscope is limited. Here, we propose a detailed protocol using a new orange-emitting fluorochrome, styryl benzoindoleninium sulfonate (SBIS), suitable for labeling the plasma membrane of brown algal cells and multicolor in vivo imaging in 3D using confocal and light sheet microscopy. Unlike calcofluor white (CFW), SBIS enables the observation of brown algal cells at thicknesses up to 25 μm and over periods up to 7 days on brown algae such as Ectocarpus sp., Sphacelaria rigidula, and Saccharina latissima. This step-by-step protocol includes labeling of brown algal tissues, mounting for 3D confocal time-lapse microscopy, and mounting for 3D time-lapse light sheet microscopy. The imaging setup and parameters have been optimized for minimizing toxicity for brown algal tissues, improving signal-to-noise ratio, and enabling detailed visualization of cell shape. Therefore, this protocol provides robust and multiplexed imaging with 4D visualization of brown algal cell shape throughout the brown algae growth, offering broad applications to brown algae study at the cellular level.

Anoikis resistance, or the ability of cancer cells to evade cell death triggered by immediate detachment from the extracellular matrix, is a critical established hallmark of metastatic cancer. While suspension culture models have been used to study anoikis, most focus on defined single time points or prolonged suspension that may not recapitulate the effects of repeated stress that tumor cells experience during metastatic dissemination. Here, we describe a detailed protocol for generating anoikis-resistant (AnR) cancer cells that have adapted to such stress through exposure to repeated cycles of suspension stress on poly-HEMA-coated plates, followed by recovery under standard attached conditions. The protocol includes methods for determining baseline anoikis sensitivity, generating AnR cells over 7–9 attachment-detachment cycles, assessing the stability and reversion of the anoikis-resistant phenotype, and characterizing AnR cells using Live/Dead staining of spheroids, flow cytometry–based apoptosis assays, and immunofluorescence for proliferation markers. This approach produces a non-genetic, reversible anoikis-resistant state that models the adaptive transcriptional reprogramming underlying metastatic progression, providing a reproducible and physiologically relevant in vitro system for studying anoikis resistance mechanisms and evaluating therapeutic strategies for prevention and reversal of such adaptations.

Autoreactive CD4⁺ T cells are shaped by MHC class II–dependent selection, and HLA-DQ8 is a major susceptibility allele for type 1 diabetes and celiac disease. To define how HLA-DQ8 influences the autoreactive CD4⁺ T-cell repertoire, we generated T-cell hybridomas from HLA-DQ8 humanized mice using a BW5147 Nur77-GFP (BW-GFP) platform that enables sensitive quantification of antigen-induced T-cell receptor (TCR) signaling. The frequency of autoreactive conventional CD4⁺ hybridomas observed in HLA-DQ8 mice was higher than previously reported in C57BL/6 mice in our earlier study, suggesting that HLA-DQ8 shapes an autoreactive repertoire. However, because antigen presentation in this system is restricted by human HLA-DQ8 while hybridomas express murine CD4, we considered that CD4-MHC interspecies mismatch might affect signal strength and influence the apparent magnitude of autoreactivity. To address this limitation, we engineered a BW-GFP fusion partner expressing an optimized version of human CD4 (hCD4), restoring optimal CD4-HLA-DQ8 interactions. Hybridomas generated with this modified platform from both regulatory (Treg) and conventional (non-Treg) CD4⁺ T cells exhibited enhanced responses to HLA-DQ8/peptide complexes compared with hybridomas that do not express hCD4. This approach improves the reactivity and physiological accuracy of screening mouse-derived CD4 hybridomas specific to self and foreign antigens presented by human class II MHC complexes.

Determining the recruitment relationships of nuclear proteins is essential for understanding the mechanisms underlying nuclear complex assembly and gene regulation. A widely used method for studying recruitment is chromatin immunoprecipitation (ChIP), but it requires fixation, chromatin shearing, and specific antibodies and cannot easily resolve recruitment directionality. Other systems like lacO/LacI are restricted to a limited number of specialized cell lines containing this lacO array’s integration. To overcome these limitations, we developed a novel microscopy-based assay, CRISPR-PITA (protein interaction and telomere recruitment assay), to assess whether a nuclear protein can recruit other nuclear factors in living cells. The protein of interest is targeted to repetitive genomic loci (e.g., telomeres) using catalytically inactive Cas9 (dCas9) fused to a SunTag array, resulting in visible nuclear foci. Recruitment of endogenous proteins is evaluated by immunofluorescence. For proof-of-concept, we tested the Kaposi’s sarcoma herpesvirus (KSHV) latency-associated nuclear antigen (LANA). CRISPR-PITA revealed that LANA recruits known interactors, such as ORC2 and SIN3A, but not MeCP2. Conversely, MeCP2 recruits LANA, indicating a unidirectional recruitment relationship. Similarly, MeCP2 could recruit HDAC1, while HDAC1 could not recruit MeCP2, further supporting directional nuclear interactions. Here, we present an easy, straightforward protocol applicable to any transfectable cell line, enabling researchers to dissect recruitment dynamics at high spatial resolution. CRISPR-PITA provides a powerful, flexible, and accessible platform to interrogate recruitment directionality between nuclear proteins in their native cellular context.