免疫学

Accurate, sensitive quantification of B-lineage cells is critical for pharmacodynamic evaluation of B cell–targeted therapies in lupus nephritis (LN) clinical trials. While high-dimensional discovery platforms offer broad profiling, they often lack the sensitivity, quantitative rigor, and throughput needed for precise cell enumeration in renal trial needle biopsies. Traditional immunostaining is hampered by CD20-directed therapeutic masking or downregulation, inadequate sensitivity of CD19 in FFPE tissue, and confounding renal tubular CD138 expression. This protocol details an automated, fit-for-purpose, 5-plex sequential tyramide signal amplification (TSA)-based immunofluorescence assay (CD38, CD79a, CD19, Ki-67, CD138) developed on the Ventana Discovery Ultra platform for deployment on single tissue sections. The workflow anchors B-cell detection on CD79a to ensure sensitivity and utilizes CD38 as an obligate co-marker for CD138+ antibody-secreting cells (ASCs) to definitively exclude the CD138+ epithelial background. Following acquisition via fluorescence whole-slide imaging, a digital analysis pipeline utilizing InstanSeg-based automated segmentation rigorously classifies cell phenotypes to generate precise spatial densities (cells/mm²). This validated protocol maximizes data yield from scarce clinical biopsies while providing high-precision quantitative monitoring of longitudinal therapeutic depletion in the renal microenvironment.

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 PrimeFlow^TM assay is a flow cytometry–based method for the co-detection of RNAs and proteins in cells. When combined with cell characterization by immunophenotyping, PrimeFlow^TM can be used to simultaneously detect RNA and proteins in a cell type–specific manner in complex heterogeneous samples, offering an advantage over bulk tissue analysis methods. Here, we describe the implementation of the PrimeFlow^TM assay protocol for the detection of transgene mRNA and protein expression in spleen samples from mice treated in vivo with luciferase mRNA-lipid nanoparticles (LNPs). This protocol involves spleen tissue dissociation for cell isolation, followed by cell fixation and permeabilization to allow immunolabeling of intracellular luciferase protein. The immunophenotyping strategy is based on immunolabeling with mouse CD marker antibodies for the identification of T cells, B cells, monocytes, granulocytes/macrophages, NK cells, and non-hematopoietic cells. The RNAs of luciferase and a housekeeping gene, β-actin, are detected with sequence-specific probe sets by employing sequential oligonucleotide annealing steps and fluorescent labeling using a branched DNA (bDNA) technology. Samples are analyzed by flow cytometry. Based on our analysis, we conclude it is feasible to apply the PrimeFlow^TM approach for evaluating successful drug targeting to the cell types of interest and any potential differences in the kinetics of RNA delivery and protein expression in various tissue cells, supporting the discovery and development of RNA therapeutics.

Natural killer (NK) cells are crucial innate immune effectors, mediating cytotoxicity against cancer and infected cells through receptors such as NKG2D. Reliable quantification of NK cell subsets is essential for evaluating NK cell-based immune responses in cancer research. Unlike other assays, including traditional flow cytometry used in assessing NK cells, imaging flow cytometry (IFC) is a simple and direct method for quantitative analysis of NK cells. This protocol describes the necessary procedures, including harvesting splenocytes, acquiring these cells labeled with NKG2D antibodies, and analyzing IFC data with IDEAS^® software. We applied this protocol to quantitatively assess the number of splenic NKG2D⁺ NK cells in mice injected with SVTneg2 cancer cells (which carry the p53 G242A missense mutation) and compared them to mice injected with EMT6 cancer cells (which have wild-type p53) or normal fibroblasts. We found that the SVTneg2 cancer cells significantly decreased the number of NKG2D⁺ NK cells in mice by approximately 2-fold (933 cells vs. 2360 cells, p < 0.001) compared with mice injected with EMT6 cancer cells. This IFC protocol can be applied to directly quantify NK cells in vivo. This quantitative protocol allows novices to quickly handle the analysis of cytotoxic NK cells with a single NKG2D marker. Further multicolor flow cytometry and cytokine assay may be required to precisely define the subtypes and effects of NK cells in anticancer immunity.

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.

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.

The human T-lymphotropic virus type-1 (HTLV-1) is an oncogenic retrovirus that predominantly spreads through cell-to-cell contact due to the limited infectivity of cell-free viruses. Among various modes of intercellular transmission, HTLV-1 biofilms emerge as adhesive structures, polarized at the cell surface, which encapsulate virions within a protective matrix. This biofilm is supposed to facilitate simultaneous virion delivery during infection. Yet, the molecular and functional intricacies of viral biofilms remain largely unexplored, despite their pivotal role in understanding retroviral pathogenesis. In this study, we optimized a protocol to isolate HTLV-1 biofilms from chronically infected T cells, facilitating their structural and molecular characterization using proteomic and super-resolution microscopy analyses. This protocol involves cultivating HTLV-1 chronically infected T cells at high density to facilitate the natural detachment of viral biofilms into the supernatant. Then, employing successive centrifugations, the cells are separated from the detached biofilms, and these structures are pelleted at medium speed (10,000× g). This method circumvents the need for mechanical, chemical, or enzymatic biofilm detachment, bypasses the use of ultracentrifugation, and enables us to resuspend the biofilms in the appropriate buffer for subsequent analyses such as western blotting or super-resolution microscopy imaging as presented.

Neutrophils, constituting 50%–70% of circulating leukocytes, play crucial roles in host defense and exhibit anti-tumorigenic properties. An elevated peripheral blood neutrophil-to-lymphocyte ratio is associated with decreased survival rates in cancer patients. In response to exposure to various antigens, neutrophils release neutrophil granular proteins, which combine to form web-like structures known as neutrophil extracellular traps (NETs). Previously, the relative percentage of NETs was found to be increased in resected tumor tissue samples from patients with gastrointestinal malignancies. The presence of NETs in peripheral blood is indicative of underlying pathological conditions. Hence, employing a non-invasive method to detect NETs in peripheral blood, along with other diagnostic tests, shows potential as a valuable tool not just for identifying different inflammatory disorders but also for assessing disease severity and determining patient suitability for surgical resection. While reliable methods exist for identifying NETs in tissue, accurately quantifying them in whole blood remains challenging. Many previous methods are time-consuming and rely on a limited set of markers that are inadequate for fully characterizing NETs. Therefore, we established a unique sensitive smear immunofluorescence assay based on blood smears to identify NETs in only as little as 2 μL of whole blood. To identify the NET complexes that have enhanced specificities, this combines the use of various antibodies against neutrophil-specific CD15, NET-specific myeloperoxidase (MPO), citrullinated histone H3 (Cit H3), and nuclear DNA. This protocol offers an easy, affordable, rapid, and non-invasive method for identifying NETs; thus, it can be utilized as a diagnostic marker and targeted through various therapeutic approaches for treating human malignancies.

Inflammatory bowel disease (IBD) is characterized by an aberrant immune response against microbiota. It is well established that T cells play a critical role in mediating the pathology. Assessing the contribution of each subset of T cells in mediating the pathology is crucial in order to design better therapeutic strategies. This protocol presents a method to identify the specific effector T-cell population responsible for intestinal immunopathologies in bone marrow–engrafted mouse models. Here, we used anti-CD4 and anti-CD8β depleting antibodies in bone marrow–engrafted mouse models to identify the effector T-cell population responsible for intestinal damage in a genetic mouse model of chronic intestinal inflammation.

Key features

• This protocol allows addressing the role of CD4+ or CD8αβ+ in an engrafted model of inflammatory bowel disease (IBD).

• This protocol can easily be adapted to address the role of other immune cells or molecules that may play a role in IBD.

B cells play a critical role in host defense, producing antibodies in response to microbial infection. An inability to produce an effective antibody response leaves affected individuals prone to serious infection; therefore, proper B-cell development is essential to human health. B-cell development begins in the bone marrow and progresses through various stages until maturation occurs in the spleen. This process involves several sequential, complex events, starting with pre- and pro-B cells, which rearrange the heavy and light chain genes responsible for producing clonally diverse immunoglobulin (Ig) molecules. These cells then differentiate into immature B cells, followed by mature B cells. The bone marrow is a complex ecological niche of supporting stromal cells, extracellular matrix components, macrophages, and hematopoietic precursor cells influencing B-cell development, maturation, and differentiation. Once fully mature, B cells circulate in peripheral lymphoid organs and can respond to antigenic stimuli. As specific cell surface markers are expressed during each stage of B-cell development, researchers use flow cytometry as a powerful tool to evaluate developmental progression. In this protocol, we provide a step-by-step method for bone marrow isolation, cell staining, and data analysis. This tool will help researchers gain a deeper understanding of the progression of B-cell development and provide a pertinent flow gating strategy.