Immunology

Macrophages are at the center of innate immunity and are the main target cells of the intracellular pathogen Salmonella enterica serovar Typhi. The production of reactive oxygen and nitrogen species (ROS/RNS) is the host’s early response to invading microbes, as oxidative stress is highly toxic for bacteria. Adequate ROS/RNS production in infected macrophages is critical for the clearance of intracellular pathogens; this is achieved by several enzymes, including inducible NADPH phagocyte oxidase (NOX) and nitric oxide synthase (iNOS), respectively. The pro-inflammatory cytokine interferon gamma (IFNγ), primarily produced by activated natural killer cells and T-helper cells type 1, is a potent inducer of iNOS. Therefore, it is crucial for infection control through oxidative microbicidal activity.

To characterize the early oxidative stress response via ROS formation, which is critical for the reduction of Salmonella proliferation within macrophages, we established an in vitro model of murine macrophages infected with Salmonella enterica serovar Typhimurium (S.tm). This serovar induces a systemic infection in mice that is frequently used as a model for typhoid fever, which, in human subjects, is caused by Salmonella Typhi.

We generated bone marrow–derived macrophages (BMDM) from C57BL/6N wildtype mice using macrophage colony-stimulating factor (M-CSF) stimulation for six days. Thereafter, we infected BMDM with S.tm for one hour. Shortly before infection, cells were stained with CellROX^TM Deep Red reagent. In its reduced form, CellROX^TM is non-fluorescent. As a result of oxidation by ROS, this reagent exhibits strong fluorescence and persists within the cells. Subsequently, changes as a result of the oxidative stress response can be measured with a TECAN Spark microplate reader over time.

We designed this protocol to measure oxidative stress in macrophages through the course of an infection with an intracellular bacterium. The protocol has several advantages over established techniques. First, it allows to continuously monitor and quantify ROS production in living cells from the very start of the infection to the final clearance of the intracellular pathogen. Second, this protocol enables efficient ROS detection without stressing the cells by detaching or staining procedures.

Graphical abstract

Accessible chromatin regions modulate gene expression by acting as cis-regulatory elements. Understanding the epigenetic landscape by mapping accessible regions of DNA is therefore imperative to decipher mechanisms of gene regulation under specific biological contexts of interest. The assay for transposase-accessible chromatin sequencing (ATAC-seq) has been widely used to detect accessible chromatin and the recent introduction of single-cell technology has increased resolution to the single-cell level. In a recent study, we used droplet-based, single-cell ATAC-seq technology (scATAC-seq) to reveal the epigenetic profile of the transit-amplifying subset of thymic epithelial cells (TECs), which was identified previously using single-cell RNA-sequencing technology (scRNA-seq). This protocol allows the preparation of nuclei from TECs in order to perform droplet-based scATAC-seq and its integrative analysis with scRNA-seq data obtained from the same cell population. Integrative analysis has the advantage of identifying cell types in scATAC-seq data based on cell cluster annotations in scRNA-seq analysis.

Type 1 regulatory T (Tr1) cells are an immunoregulatory CD4⁺ Foxp3^- IL-10^high T cell subset with therapeutic potential for various inflammatory diseases. Retroviral (RV) transduction has been a valuable tool in defining the signaling pathways and transcription factors that regulate Tr1 differentiation and suppressive function. This protocol describes a method for RV transduction of naïve CD4⁺ T cells differentiating under Tr1 conditions, without the use of reagents such as polybrene or RetroNectin. A major advantage of this protocol over others is that it allows for the role of genes of interest on both differentiation and function of Tr1 cells to be interrogated. This is due to the high efficiency of RV transduction combined with the use of an IL10^GFP/Foxp3^RFP dual reporter mouse model, which enables successfully transduced Tr1 cells to be identified and sorted for functional assays. In addition, this protocol may be utilized for dual/multiple transduction approaches and transduction of other lymphocyte populations, such as CD8⁺ T cells.

Genome-editing technologies, especially CRISPR (clustered regularly interspaced short palindrome repeats)/Cas9 (CRISPR-associated protein 9), endows researchers the ability to make efficient, simple, and precise genomic DNA changes in many eukaryotic cell types. CRISPR/Cas9-mediated efficient gene knockout holds huge potential to improve the efficacy and safety of chimeric antigen receptor (CAR) T cell-based immunotherapies. Here, we describe an optimized approach for a complete loss of endogenous T cell receptor (TCR) protein expression, by CRISPR/Cas9-mediated TCR α constant (TRAC) and TCR β constant (TRBC) gene knockout, followed by subsequent CD3 negative selection in engineered human orthoCAR19 T cells. We believe this method can be expanded beyond CAR T cell application, and target other cell surface receptors.

Graphical abstract:

Schematic overview of the two-step process of endogenous TCR depletion in engineered human orthoCAR19 T cells using (1) CRISPR/Cas9-mediated gene knockout followed by (2) CD3 negative selection.

CD4⁺ T cells are essential players in orchestrating the specific immune response against intracellular pathogens, and in inhibiting tumor development in an early stage. The activation of T cells is triggered by engagement of T cell receptors (TCRs). Here, CD3 and CD28 molecules are key factors, (co)stimulating signaling pathways essential for activation and proliferation of CD4⁺ T cells. T cell activation induces the formation of a tight mechanical bond between T cell and target cell, the so-called immunological synapse (IS). Due to this, mechanical cell properties, including stiffness, play a significant role in modulating cell functions. In the past, many approaches were made to investigate mechanical properties of immune cells, including micropipette aspiration, microplate-based rheometry, techniques based on deformation during cytometry, or the use of optical tweezers. However, the stiffness of T lymphocytes at a subcellular level at the IS still remains largely elusive.

With this protocol, we introduce a method based on atomic force microscopy (AFM), to investigate the local cellular stiffness of T cells on functionalized glass/Polydimethylsiloxan (PDMS) surfaces, which mimicks focal stimulation of target cells inducing IS formation by T cells. By applying the peak force nanomechanical mapping (QNM) technique, cellular surface structures and the local stiffness are determined simultaneously, with a resolution of approximately 60 nm. This protocol can be easily adapted to investigate the mechanical impact of numerous factors influencing IS formation and T cell activation.

Graphical abstract:

Overview of the experimental workflow.

Individual experimental steps are shown on the left, hands on and incubation times for each step are shown right.

As a model to interrogate human macrophage biology, macrophages differentiated from human induced pluripotent stem cells (hiPSCs) transcend other existing models by circumventing the variability seen in human monocyte-derived macrophages, whilst epitomizing macrophage phenotypic and functional characteristics over those offered by macrophage-like cell lines (Mukherjee et al., 2018). Furthermore, hiPSCs are amenable to genetic manipulation, unlike human monocyte-derived macrophages (MDMs) (van Wilgenburg et al., 2013; Lopez-Yrigoyen et al., 2020), proposing boundless opportunities for specific disease modelling.

We outline an effective and efficient protocol that delivers a continual production of hiPSC-derived-macrophages (iMACs), exhibiting human macrophage surface and intracellular markers, together with functional activity.

The protocol describes the resuscitation, culture, and differentiation of hiPSC into mature terminal macrophages, via the initial and intermediate steps of expansion of hiPSCs, formation into embryoid bodies (EBs), and generation of hematopoietic myeloid precursors.

We offer a simplified, scalable, and adaptable technique that advances upon other protocols, utilizing feeder-free conditions and reduced growth factors, to produce high yields of consistent iMACs over a period of several months, economically.

The human immunodeficiency virus (HIV)-1 viral inhibition assay (VIA) measures CD8⁺ T cell-mediated inhibition of HIV replication in CD4⁺ T cells and is increasingly used for clinical testing of HIV vaccines and immunotherapies. Different VIAs that differ in length of CD8:CD4 T cell culture periods (6–13 days), purity of CD4 cultures [isolated CD4⁺ T cells or CD8⁺ depleted peripheral blood mononuclear cells (PBMCs)], HIV strains (laboratory strains, isolates, reporter viruses) and read-outs of virus inhibition (p24 ELISA, intracellular measurement of p24, luciferase reporter expression, and viral gag RNA) have been reported.

Here, we describe multiple modifications to a 7-day VIA protocol, the most impactful being the introduction of independent replicate cultures for both HIV infected-CD4 (HIV-CD4) and HIV-CD4:CD8 T cell cultures. Virus inhibition was quantified using a ratio of weighted averages of p24⁺ cells in replicate cultures and the corresponding 95% confidence intervals. We identify methodological and analysis changes that could be incorporated into other protocols to improve assay reproducibility. We found that in people living with HIV (PLWH) on antiretroviral therapy (ART), CD8 T cell virus inhibition was largely stable over time, supporting the use of this assay and/or analysis methods to examine therapeutic interventions.

Graphic abstract:

When the body mounts an immune response against a foreign pathogen, the adaptive arm of the immune system relies upon clonal expansion of antigen-specific T cell populations to exercise acquired effector and cytotoxic functions to clear it. However, T cell expansion must be modulated to effectively combat the perceived threat without inducing excessive collateral damage to host tissues. Restimulation-induced cell death (RICD) is an apoptotic program triggered in activated T cells when an abundance of antigen and IL-2 are present, imposing a negative feedback mechanism that constrains the growing T cell population. This autoregulatory process can be detected via increases in caspase activation, Annexin V binding, and loss of mitochondrial membrane potential. However, simple changes in T cell viability through flow cytometric analysis can reliably measure RICD sensitivity in response to T-cell receptor (TCR) restimulation. This protocol describes the in vitro polyclonal activation, expansion, and restimulation of human primary T cells isolated from donor peripheral blood mononuclear cells (PBMC). This simple procedure allows for accurate quantification of RICD via flow cytometry. We also describe strategies for interrogating the role of specific proteins and pathways that may alter RICD sensitivity. This straightforward protocol provides a quick and dependable tool to track RICD sensitivity in culture over time while probing critical factors that control the magnitude and longevity of an adaptive immune response.

Graphic abstract:

In-vitro simulation of restimulation-induced cell death in activated human T cells.

Blood endothelial cells (ECs) constitute the primary physical barrier to be crossed by circulating leukocytes, once attracted to a site of ongoing inflammation/infection. Upon a pro-inflammatory stimulus, such as tumor necrosis factor (TNF), ECs upregulate adhesion molecule expression to favor the adhesion and, subsequently, the transendothelial migration of the attracted lymphocytes. To address the ability of a cell to transmigrate through a monolayer of ECs, the classical transmigration assay is usually performed (Muller and Luscinskas, 2008). In the present protocol, adapted from Safuan et al. (2012), we describe an in vitro assay for assessing the functionality of the second step of the transendothelial migration, i.e., the firm adhesion of peripheral blood mononuclear cells (PBMCs) to ECs, under static conditions. By pre-incubating primary human umbilical cord ECs (HUVECs) with either innate lymphoid cell progenitors (ILCPs) or TNF, we were able to upregulate adhesion molecules on the EC surface. Then, by adding total PBMCs, we were able to both quantitatively and qualitatively analyze the cellular subtype and number of PBMCs that adhered to the pre-treated ECs. The important advantage of this technique is the possibility to perform functional studies on ECs biology since, differently from transwell-based strategies, it allows a good recovery of ECs at the end of the assay. Overall, this assay enables to interrogate how/if different stimulations/cell types can influence EC ability to retain PBMCs in vitro, under static conditions.

Graphic abstract:

The workflow of the Static Adhesion Assay.

Natural killer (NK) cells are large granular lymphocytes that keep in check the health of neighboring cells through a large array of intrinsically expressed germline-coded receptors. Most importantly, CD16 is a low affinity Fc receptor for IgG that mediates the antibody-dependent cellular cytotoxicity (ADCC) of NK cells, bridging the innate and adaptive immunities. There has been a significant interest in genetically engineering NK cells to enhance its ADCC, with the ultimate goal to produce off-the-shelf NK cell therapy products that can be combined with target-specific monoclonal antibodies to improve clinical outcomes. Previous protocols of ADCC assays use complex cell-based antigen-antibody models, which are both costly and time-consuming. This current protocol is devoid of target cells and uses plate-bound immobilized anti-CD16 antibodies as the trigger. It greatly shortens the experimental time, while faithfully evaluating NK cells ADCC.

Graphic abstract:

Workflow of stimulating NK cells via CD16 by plate-bound anti-CD16 mAb.