免疫学

Neuroinflammation disrupts blood–brain barrier (BBB) integrity, promoting leukocyte recruitment into the central nervous system and contributing to the progression of neurological disorders. This protocol describes a reproducible macrophage adhesion assay to evaluate interactions between immune cells and brain endothelial cells and to screen compounds with potential anti-inflammatory activity. Murine brain endothelial cells (bEnd.3) were cultured to confluency and exposed to inflammatory stimuli, such as lipopolysaccharide (LPS) or pilocarpine, a cholinergic muscarinic receptor agonist reported to induce inflammatory responses through seizure-associated neuroinflammatory mechanisms, in the presence or absence of candidate therapeutic compounds. In these studies, the natural flavonoid quercetin and the synthetic alkyl-lysophospholipid edelfosine were tested for their effects on macrophage adhesion. After 48 h of treatment, fluorescently labeled murine macrophages (RAW 264.7) were added to the endothelial monolayer, and adherent cells were quantified by fluorescence microscopy. The assay was validated using dexamethasone as an anti-inflammatory control and inflammatory stimulation with LPS or pilocarpine. As expected, dexamethasone reduced macrophage adhesion, whereas both LPS and pilocarpine significantly increased adhesion, demonstrating the assay's sensitivity to changes in endothelial inflammatory status. Overall, this protocol provides a reliable and accessible platform for investigating endothelial–immune cell interactions under neuroinflammatory conditions and for evaluating therapeutic compounds that may preserve BBB function and reduce inflammatory cell recruitment in neurological disease models.

Surface receptor engagement governs T-cell activation. Since these surface receptors undergo extensive glycosylation, lectin-mediated crosslinking of these glycosylated surface receptors has the potential to modulate signaling. Here, we systematically evaluate the abilities of recombinant human galectins in triggering immune responses. We describe how to apply the human galectins to modulate Jurkat E6-1 cell activation by measuring the expression level of cellular surface CD69 and the mRNA of IL-2. To validate the protocol, we confirmed that galectin-3 and galectin-8 variants 1 and 2 reproducibly induce CD69 and IL-2 expression on Jurkat E6-1 cells. Our approach offers a galectin-based toolset to study how glycosylation modulates human adaptive immunity.

This protocol describes an optimized strategy for the efficient generation of peptide-loaded major histocompatibility complex (MHC) class II (pMHC) tetramers, which are essential tools for detecting and characterizing antigen-specific T cells in immunological research. Traditional methods require separate expression of MHC proteins followed by in vitro biotinylation—a multi-step process that is time-consuming and prone to protein loss. Here, we present an integrated approach based on co-expression of MHC monomers and BirA biotin ligase in Expi293F T cells, enabling site-specific biotinylation in vivo during protein synthesis. At the same time, the incorporation of a thrombin-cleavable class II–associated invariant chain peptide (CLIP) peptide into the MHC construct allows flexible loading of any antigenic peptide of interest without the need for re-cloning or re-expression of the MHC molecule. Pre-biotinylated MHC molecules are subsequently purified, loaded with antigenic peptides, and assembled into fluorescent tetramers via streptavidin conjugation. This streamlined workflow significantly reduces handling steps, improves protein yield, and enhances reproducibility. The resulting tetramers are suitable for sensitive detection and isolation of antigen-specific T cells by flow cytometry, supporting applications in T-cell immunogenicity studies, vaccine development, and autoimmune disease research.

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.

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.

Human mononuclear cells derived from peripheral blood and bone marrow are valuable resources for the study of hematological malignancies, including acute myeloid leukemia (AML) and chronic myelomonocytic leukemia (CMML). Cryopreservation enables long-term storage of patient samples for downstream assays; while thawing protocols have been described, subsequent recovery of viable cells after thawing can be challenging, particularly for fragile blast and monocyte populations. Here, we describe a reliable protocol for thawing cryopreserved AML and CMML mononuclear cells designed to preserve post-thaw viability, recovery, and functional integrity. The method incorporates controlled dilution of cells out of cryoprotectant with anticoagulant-supplemented thaw buffer, DNase I treatment, and gentle resuspension steps. Using this approach, post-thaw viability consistently exceeded 80% with a mean recovery of 55.6% across samples. Recovered cells retained functional capacity, as demonstrated by colony-forming assays, and maintained immunophenotypic characteristics by flow cytometry. This protocol provides a robust and reproducible method for the recovery of cryopreserved AML and CMML mononuclear cells and may be broadly applicable to other fragile or monocyte-rich patient-derived hematopoietic samples.

Macrophage efferocytosis is a previously unrecognized key pathogenic event, engulfing apoptotic targets, preventing inflammation and necrosis, and maintaining immune homeostasis. The phagocytic function can be disrupted by harmful factors and toxic substances. This protocol describes a versatile visualized in vitro method that can be used for the detection of general efferocytosis. This method is applicable to a wide range of research scenarios. As a representative application, it can be used to evaluate macrophage efferocytosis dysfunction in diseases linked to harmful exposures, including atherosclerosis, chronic inflammation, and malignant tumors. Among them, the detection of the effects of oxidized low-density lipoprotein (ox-LDL) and arsenite on macrophage efferocytosis capacity is an exemplary application of this protocol. Primary macrophages collected from mice were labeled with a cell-tracking dye and exposed to ox-LDL or arsenite, then co-cultured with apoptotic thymocytes or hepatocytes (labeled with another cell-tracking dye) for 2 h at a ratio of 5:1. Macrophage efferocytosis was visualized using a laser confocal microscope. The results indicate that arsenite impaired macrophage efferocytosis, leading to insufficient clearance of apoptotic thymocytes or hepatocytes. This method can be extended to subsequent studies, including those involving different types of phagocytes, apoptotic cell models, and research related to exposure to various factors.

Detecting the proliferation of cells with copper(I)-catalyzed azide-alkyne cycloaddition (click chemistry) and the thymidine analogue, 5-ethynyl-2’-deoxyuridine (EdU), is a simpler and more versatile method than traditional antibody-based approaches. Instead of the harsh series of steps typically used for 5-bromo-2’-deoxyuridine (BrdU) detection, detecting EdU does not require DNA denaturation and is suitable for use with other applications. This approach was implemented in an animal model of ischemic stroke. The following protocol details how to use EdU to label, track, and visualize leukocyte recruitment for flow cytometry and fluorescence microscopy, including the processes for EdU injection and blood and tissue sample preparation. Considerations for timing, dosing, and cell viability are also outlined to tailor the protocol to experimental needs. This method could be applied to various models that require extended tracking periods, as the signal from EdU can last several cell divisions, depending on cell type and condition.

Protein phosphorylation is a dynamic post-translational modification that regulates fundamental processes, including signal transduction, cell proliferation, differentiation, and effector function of immune cells. The Janus Kinase/Signal Transducer and Activator of Transcription (JAK/STAT) pathway is a key mediator of cytokine responses, essential for maintaining immune cell homeostasis and determining cell fate across diverse immune subsets. Dysregulation of JAK/STAT signaling has been linked to a broad spectrum of pathologies, including monogenic immune disorders, autoimmunity, and cancer. Platforms facilitating single-cell analysis of protein phosphorylation offer the ability to reveal subtle signaling defects and dissect the pleiotropy in cellular composition and phosphorylation status, providing insights into immune phenotype and function, while identifying potential therapeutic targets. While an application of cytometry-by-time-of-flight, termed phospho-CyTOF, has proven invaluable for studying protein phosphorylation in cryopreserved peripheral blood mononuclear cells (cPBMCs), its application is limited by cell loss and signaling artifacts stemming from isolation and cryopreservation. Conversely, whole blood (WB) approaches, preserving the native immune cell composition and signaling context, offer a more physiological representation but necessitate robust and consistent protocols for broad application. Herein, we present optimized dual phospho-CyTOF workflows tailored for both cPBMCs and whole blood, building upon established protocols for cytokine stimulation of both samples. These workflows facilitate comprehensive, high-dimensional profiling of JAK/STAT signaling in response to pleiotropic cytokines such as Type I interferons (IFN-α), Type II interferons (IFN-γ), and Interleukin-21 (IL-21). By leveraging CyTOF's capacity for high-dimensional profiling using pure heavy metal–labeled antibodies, these protocols aim to identify pathway-specific alterations in STAT phosphorylation across major immune subsets that may be overlooked by traditional flow cytometry. Together, these optimized dual workflows provide scalable, translationally relevant tools for dissecting the subtle and differential JAK/STAT-driven immune responses in both clinical and research settings, while also being compatible with the simultaneous assessment of crosstalk with alternative immune cell signaling pathways.

Recent advances in single-cell technologies have provided limited insight into the role of protein phosphorylation in T-cell fate and function. Dysregulated protein phosphorylation is associated with adverse clinical outcomes, emphasizing the need for reliable methods to unravel the complexities of T-cell signal transduction and disease-related alterations. While flow cytometry is widely used, it is constrained by spectral overlap, limiting the number of protein targets for simultaneous analysis. To overcome this, we present a robust protocol for whole blood T-cell stimulation and subsequent analysis using mass cytometry by time-of-flight (CyTOF). CyTOF minimizes spillover into adjacent channels by employing highly pure, stable, heavy metal–conjugated antibodies for protein detection. This protocol offers a high-dimensional approach for phenotypic and phospho-protein characterization of key signaling pathways, including JAK/STAT, MAPK, PI3K/mTOR, PKC, and NF-κB. A key feature is the T-cell stimulation reagent, which mimics endogenous activation by engaging the T-cell receptor (TCR)/CD3 complex and providing co-stimulation via an anti-CD28 antibody. Further, we enhance reproducibility and enable batch processing through the implementation of the Prot1/Thaw-Lyse system for immediate cryopreservation of stimulated blood samples. By employing CyTOF, this method permits the simultaneous analysis of 31 protein targets with single-cell resolution, minimizing spillover and providing superior specificity, sensitivity, and resolution over flow cytometric methods. This approach facilitates the robust assessment of TCR activation and its effect on bystander populations, which has been challenging with spectral flow cytometry due to the limited availability of methanol-resistant fluorophores. This protocol is a precise and reproducible method for elucidating the downstream effects of T-cell stimulation and immune status, with significant potential for clinical applications, including the assessment of T-cell-targeted therapies.