Immunology

γδ T cells play a critical role in homeostasis and diseases such as infectious diseases and tumors in both mice and humans. They can be categorized into two main functional subsets: IFN-γ-producing γδT1 cells and IL-17-producing γδT17 cells. While CD27 expression segregates these two subsets in mice, little is known about human γδT17 cell differentiation and expansion. Previous studies have identified γδT17 cells in human skin and mucosal tissues, including the oral cavity and colon. However, human γδ T cells from peripheral blood mononuclear cells (PBMCs) primarily produce IFN-γ. In this protocol, we describe a method for in vitro expansion and polarization of human γδT17 cells from PBMCs.

Key Features

• Expansion of γδ T cells from peripheral blood mononuclear cells.

• Human IL-17A-producing γδ T-cell differentiation and expansion using IL-7 and anti-γδTCR.

• Analysis of IL-17A production post γδ T-cell expansion.

Medullary thymic epithelial cells (mTEC) are bona fide antigen-presenting cells that play a crucial role in the induction of T-cell tolerance. By their unique ability to express a broad range of tissue-restricted self-antigens, mTEC control the clonal deletion (also known as negative selection) of potentially hazardous autoreactive T cells and the generation of Foxp3⁺ regulatory T cells. Here, we describe a protocol to assess major histocompatibility complex (MHC) class II antigen-presentation capacity of mTEC to CD4⁺ T cells. We detail the different steps of thymus enzymatic digestion, immunostaining, cell sorting of mTEC and CD4⁺ T cells, peptide-loading of mTEC, and the co-culture between these two cell types. Finally, we describe the flow cytometry protocol and the subsequent analysis to assess the activation of CD4⁺ T cells. This rapid co-culture assay enables the evaluation of the ability of mTEC to present antigens to CD4⁺ T cells in an antigen-specific context.

Key features

• This protocol builds upon the method used by Lopes et al. (2018 and 2022) and Charaix et al. (2022).

• This protocol requires transgenic mice, such as OTIIxRag2^-/- mice and the cognate peptide OVA_323–339, to assess mTEC antigen presentation to CD4⁺ T cells.

• This requires specific equipment such as a Miltenyi Biotec AutoMACS^® Pro Separator, a BD FACSAria^TM III cell sorter, and a BD^® LSR II flow cytometer.

Graphical overview

Dendritic cells have been investigated for cell-based immunotherapy for various applications. The low abundance of dendritic cells in blood hampers their clinical application, resulting in the use of monocyte-derived dendritic cells as an alternative cell type. Limited knowledge is available regarding blood-circulating human dendritic cells, which can be divided into three subsets: type 2 conventional dendritic cells, type 1 conventional dendritic cells, and plasmacytoid dendritic cells. These subsets exhibit unique and desirable features for dendritic cell-based therapies. To enable efficient and reliable human research on dendritic cell subsets, we developed an efficient isolation protocol for the three human dendritic cell subsets, resulting in pure populations. The sequential steps include peripheral blood mononuclear cell isolation, magnetic-microbead lineage depletion (CD14, CD56, CD3, and CD19), and individual magnetic-microbead isolation of the three human dendritic cell subsets.

Graphical overview

Scheme of the dendritic cell (DC) isolation protocol. Starting material for this process is human blood (buffy coat or aphaeresis). From that, peripheral blood mononuclear cells (PBMCs) are isolated by using ficoll gradient centrifugation. Then, an enrichment for DCs is performed using semi-automated equipment. From the enriched fraction, DC subsets are obtained by magnetic cell sorting.

Immune cell trafficking in steady-state conditions and inflammatory cell recruitment into injured tissues is crucial for the surveillance of the immune system and the maintenance of body homeostasis. Tracking the cell journey from the infection site in the skin to lymphoid tissues has been challenging, and is typically determined using fluorescent cell tracers, antibodies, or photoconvertible models. Here, we describe the detailed method to track Leishmania-infected myeloid cells migrating from the skin to lymphatic tissues by multiparametric flow cytometry. These methods involve labeling of infective Leishmania donovani parasites with fluorescent cell tracers and phenotyping of myeloid cells with fluorescent antibodies, to determine the infection status of migratory myeloid cells. We also describe the detailed protocol to trace donor monocytes transferred intradermally into recipient mice in Leishmania donovani infection. These protocols can be adapted to study skin-lymphoid tissue migration of dendritic cells, inflammatory monocytes, neutrophils, and other phagocytic myeloid cells in response to vaccine antigens and infection.

Key features

• Cell-tracking of cell-trace-labeled parasites and monocytes from the skin to lymphatic tissues after transference into donor mice.

• Identification of migratory cells labeled with fluorescent cell tracers and antibodies by flow cytometry.

• Isolation, labeling, and transference of bone marrow monocytes from donor mice into the skin of recipient mice.

• Description of a double-staining technique with fluorescent cell tracers to determine cell and parasite dissemination from the skin to lymphoid tissues.

Graphical overview

Overview of the methods to trace the migration of Leishmania and monocytes from the skin to lymphatic tissues by flow cytometry. Infective metacyclic promastigotes (from axenic culture) and monocytes (isolated from the bone marrow of donor mice) are labeled with fluorescent cell tracers. After intradermal injection into the test mouse (1, 2), migratory cells and infected cells are isolated from the skin and lymphoid tissues of the test mouse. These cells are then labeled with fluorescent antibodies against myeloid cells and recognized according to the differential excitation/emission wavelengths of the fluorochromes by flow cytometry.

During life, the embryonic alveolar macrophage (AM) population undergoes successive waves of depletion and replenishment in response to infectious and inflammatory episodes. While resident AMs are traditionally described as from embryonic origin, their ontogeny following inflammation or infection is much more complex. Indeed, it appears that the contribution of monocytes (MOs) to the AM pool is variable and depends on the type of inflammation, its severity, and the signals released in the microenvironment of the pulmonary niche (peripheral imprinting) and/or in the bone marrow (central imprinting). Deciphering the cellular and molecular mechanisms regulating the differentiation of MOs into AMs remains an area of intense investigation, as this could potentially explain part of the inter-individual susceptibility to respiratory immunopathologies. Here, we detail a relevant ex vivo co-culture model to investigate how lung epithelial cells (ECs) and group 2 lung innate lymphoid cells (ILC2s) contribute to the differentiation of recruited MOs into AMs. Interestingly, the presence of lung ILC2s and ECs provides the necessary niche signals to ensure the differentiation of bone marrow MOs into AMs, thus establishing an accessible model to study the underlying mechanisms following different infection or inflammation processes.

Key features

• Ex vivo co-culture model of the alveolar niche.

• Deciphering the particular niche signals underlying the differentiation of MO into AMs and their functional polarization.

Graphical overview
This protocol described the isolation of bone marrow monocytes (MOs), lung epithelial cells (ECs), and lung group 2 lung innate lymphoid cells (ILC2s) and the ex vivo co-culture of these cells to drive the differentiation of bone marrow MOs into alveolar macrophages (AMs).

This co-culture experiment is composed of three steps (Graphical overview):
1. Identification and FACS-sorting of ECs and MOs isolated from the lung and the bone marrow of naive mice, respectively.
2. Culture of these ECs and bone marrow MOs for three days.
3. Addition of ILC2s isolated from the lung of naïve mice or mice subjected to a treatment/infection of interest.

Invariant natural killer T (iNKT) cells are a non-conventional T-cell population expressing a conserved semi-invariant T-cell receptor (TCR) that reacts to lipid antigens, such as α-galactosyl ceramide (α-GalCer), presented by the monomorphic molecule CD1d. iNKT cells play a central role in tumor immunosurveillance and represent a powerful tool for anti-cancer treatment, notably because they can be efficiently redirected against hematological or solid malignancies by engineering with tumor-specific chimeric antigen receptors (CARs) or TCRs. However, iNKT cells are rare and require specific ex vivo pre-selection and substantial in vitro expansion to be exploited for adoptive cell therapy (ACT). This protocol describes a robust method to obtain a large number of mouse iNKT cells that can be effectually engineered by retroviral (RV) transduction. A major advantage of this protocol is that it requires neither particular instrumentation nor a high number of mice. iNKT cells are enriched from the spleens of iVα14-Jα18 transgenic mice; the rapid purification protocol yields a highly enriched iNKT cell population that is activated by anti-CD3/CD28 beads, which is more reproducible and less time consuming than using bone marrow–derived dendritic cells loaded with α-GalCer, without risks of expanding contaminant T cells. Forty-eight hours after activation, iNKT cells are transduced with the selected RV by spin inoculation. This protocol allows to obtain, in 15 days, millions of ready-to-use, highly pure, and stably transduced iNKT cells that might be exploited for in vitro assays and ACT experiments in preclinical studies.

The nematode Haemonchus placei is a pathogenic parasite, the most seriously affecting ruminant’s health and being responsible for enormous economic losses all over the world. The present protocol describes different in vitro techniques to select potential candidate antigens with immune-protective activity from excretory and secretory products (ESP) from H. placei transitory infective larvae (xL₃). ESP from xL₃ were obtained from the in vitro infective larvae (L₃) maintained in Hank’s medium at 37 °C with 5% CO₂ for 48 h. Then, the presence of ESP proteins was confirmed by SDS-PAGE to be used in an in vitro proliferation assay with bovine peripheral blood mononuclear cells (PBMCs). The ESP were exposed to the PBMCs during two different periods (24 and 48 h). Genes associated with immune response against the nematode were analyzed using relative gene expression and bioinformatic tools. These are simple, economic, and helpful tools to identify potential immune-protective molecules under in vitro conditions for confirming the efficacy of future in vivo assays.

Graphical overview

T cells localized to the kidneys and vasculature/perivascular adipose tissue (PVAT) play an important role in hypertension and vascular injury. CD4⁺, CD8⁺, and γδ T-cell subtypes are programmed to produce interleukin (IL)-17 or interferon-γ (IFNγ), and naïve T cells can be induced to produce IL-17 via the IL-23 receptor. Importantly, both IL-17 and IFNγ have been demonstrated to contribute to hypertension. Therefore, profiling cytokine-producing T-cell subtypes in tissues relevant to hypertension provides useful information regarding immune activation. Here, we describe a protocol to obtain single-cell suspensions from the spleen, mesenteric lymph nodes, mesenteric vessels and PVAT, lungs, and kidneys, and profile IL-17A- and IFNγ-producing T cells using flow cytometry. This protocol is different from cytokine assays such as ELISA or ELISpot in that no prior cell sorting is required, and various T-cell subsets can be identified and individually assessed for cytokine production simultaneously within an individual sample. This is advantageous as sample processing is kept to a minimum, yet many tissues and T-cell subsets can be screened for cytokine production in a single experiment. In brief, single-cell suspensions are activated in vitro with phorbol 12-myristate 13-acetate (PMA) and ionomycin, and Golgi cytokine export is inhibited with monensin. Cells are then stained for viability and extracellular marker expression. They are then fixed and permeabilized with paraformaldehyde and saponin. Finally, antibodies against IL-17 and IFNγ are incubated with the cell suspensions to report cytokine production. T-cell cytokine production and marker expression is then determined by running samples on a flow cytometer. While other groups have published methods to perform T-cell intracellular cytokine staining for flow cytometry, this protocol is the first to describe a highly reproducible method to activate, phenotype, and determine cytokine production by CD4, CD8, and γδ T cells isolated from PVAT. Additionally, this protocol can be easily modified to investigate other intracellular and extracellular markers of interest, allowing for efficient T-cell phenotyping.

Macrophages are a heterogeneous class of innate immune cells that offer a primary line of defense to the body by phagocytizing pathogens, digesting them, and presenting the antigens to T and B cells to initiate adaptive immunity. Through specialized pro-inflammatory or anti-inflammatory activities, macrophages also directly contribute to the clearance of infections and the repair of tissue injury. Macrophages are distributed throughout the body and largely carry out tissue-specific functions. In skeletal muscle, macrophages regulate tissue repair and regeneration; however, the characteristics of these macrophages are not yet fully understood, and their involvement in skeletal muscle aging remains to be elucidated. To investigate these functions, it is critical to efficiently isolate macrophages from skeletal muscle with sufficient purity and yield for various downstream analyses. However, methods to prepare enriched skeletal muscle macrophages are scarce. Here, we describe in detail an optimized method to isolate skeletal muscle macrophages from mice. This method has allowed the isolation of CD45⁺/CD11b⁺ macrophage-enriched cells from young and old mice, which can be further used for flow cytometric analysis, fluorescence-activated cell sorting (FACS), and single-cell RNA sequencing.

During adaptive immune responses, germinal centers (GC) appear as transient microstructures, in which antigen-specific B and T cells interact with each other. Because only the antigen-activated B and T cells, such as Plasmablasts or follicular T helper (Tfh) cells, are present in GC, the in depth-analysis of GC is of great interest. To identify the cells that reside within GC, the majority of studies use the expression of specific surface molecules for analysis by flow cytometry. To do so, the tissue has to be disrupted for the preparation of single-cell suspensions. Thereby, the local information regarding neighborhoods of B cells and T cells and their potential interaction is lost. To study GC in vivo within their original microenvironment, we established a protocol for the isolation of GC by laser microdissection. To enable the identification of GC for subsequent transcriptomic analysis, the degradation of mRNA was diminished by using frozen tissues and by establishing a rapid staining protocol. This procedure enables histological and transcriptomic analysis of individual GC even within one lymphoid organ.