Immunology

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.

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.

Extracellular microvesicles (MVs) are released into the circulation in large numbers during acute systemic inflammation, yet little is known of their intravascular cell/tissue-specific interactions under these conditions. We recently described a dramatic increase in the uptake of intravenously injected MVs by monocytes marginated within the pulmonary vasculature, in a mouse model of low-dose lipopolysaccharide-induced systemic inflammation. To investigate the mechanisms of enhanced MV uptake by monocytes, we developed an in vitro model using in vivo derived monocytes. Although mouse blood is a convenient source, monocyte numbers are too low for in vitro experimentation. In contrast, differentiated bone marrow monocytes are abundant, but they are rapidly mobilized during systemic inflammation, and thus no longer available. Instead, we developed a protocol using marginated monocytes from the pulmonary vasculature as an anatomically relevant and abundant source. Mice are sacrificed by terminal anesthesia, the lungs inflated and perfused via the pulmonary artery. Perfusate cell populations are evaluated by flow cytometry, combined with in vitro generated fluorescently labelled MVs, and incubated in suspension for up to one hour. Washed cells are analyzed by flow cytometry to quantify MV uptake and confocal microscopy to localize MVs within cells (O'Dea et al., 2020). Using this perfusion-based method, substantial numbers of marginated pulmonary vascular monocytes are recovered, allowing multiple in vitro tests to be performed from a single mouse donor. As MV uptake profiles were comparable to those observed in vivo, this method is suitable for physiologically relevant high throughput mechanistic studies on mouse monocytes under in vitro conditions.

Graphic abstract:

Figure 1. Schematic of lung perfusate cell harvest and co-incubation with in vitro generated MVs. Created with BioRender.com.

Recent advances in single-cell RNA-sequencing (scRNA-seq) technologies provide unprecedented opportunities to identify new cell types and characterize cell states. One of the most important requirements for performing scRNA-seq is to obtain high-quality single cells in suspension. Recently, we used this approach to characterize Drosophila blood cells (hemocytes). Here, we provide a detailed protocol for obtaining single hemocytes in suspension, which can be used for microfluidics-based scRNA-seq platforms. This protocol involves the simple bleeding of third instar larvae and the subsequent purification of the hemolymph using either Optiprep-based gradient centrifugation or traditional centrifugation methods to obtain single hemocytes of high quality for scRNA-seq. Importantly, this method for single-hemocyte preparation is straightforward and reproducible, with negligible issues associated with cell viability as the entire procedure involves no enzymatic dissociation.

Graphic abstract:

Workflow for the preparation of Drosophila larval blood cells in suspension. Hemocytes (blood cells) of the sessile and circulatory compartments of larvae are derived by simple bleeding and purification using gradient centrifugation. Blood cells are counted and subsequently encapsulated by microfluidics-based scRNA-seq platforms. Blood cells represented in the schematic are derived from third instar larvae of the genotype Hemolectin-GAL4.Delta, UAS-2xEGFP (BDSC stock #30140).

Microglia are a unique type of tissue-resident innate immune cell found within the brain, spinal cord, and retina. In the healthy nervous system, their main functions are to defend the tissue against infectious microbes, support neuronal networks through synapse remodeling, and clear extracellular debris and dying cells through phagocytosis. Many existing microglia isolation protocols require the use of enzymatic tissue digestion or magnetic bead-based isolation steps, which increase both the time and cost of these procedures and introduce variability to the experiment. Here, we report a protocol to generate single-cell suspensions from freshly harvested murine brains or spinal cords, which efficiently dissociates tissue and removes myelin debris through simple mechanical dissociation and density centrifugation and can be applied to rat and non-human primate tissues. We further describe the importance of including empty channels in downstream flow cytometry analyses of microglia single-cell suspensions to accurately assess the expression of protein targets in this highly autofluorescent cell type. This methodology ensures that observed fluorescence signals are not incorrectly attributed to the protein target of interest by appropriately taking into account the unique autofluorescence of this cell type, a phenomenon already present in young animals and that increases with aging to levels that are comparable to those observed with antibodies against highly abundant antigens.

Mycobacterium tuberculosis (Mtb) is transmitted by aerosol and can cause serious bacterial infection in the lung that can be fatal if left untreated. Mtb is now the leading cause of death worldwide by an infectious agent. Characterizing the early events of in vivo infection following aerosol challenge is critical for understanding how innate immune cells respond to infection but is technically challenging due to the small number of bacteria that initially infect the lung. Previous studies either evaluated Mtb-infected cells at later stages of infection when the number of bacteria in the lung is much higher or used in vitro model systems to assess the response of myeloid cells to Mtb. Here, we describe a method that uses fluorescent bacteria, a high-dose aerosol infection model, and flow cytometry to track Mtb-infected cells in the lung immediately following aerosol infection and fluorescence-activated cell sorting (FACS) to isolate naïve, bystander, and Mtb-infected cells for downstream applications, including RNA-sequencing. This protocol provides the ability to monitor Mtb-infection and cell-specific responses within the context of the lung environment, which is known to modulate the function of both resident and recruited populations. Using this protocol, we discovered that alveolar macrophages respond to Mtb infection in vivo by up-regulating a cell protective transcriptional response that is regulated by the transcription factor Nrf2 and is detrimental to early control of the bacteria.

In homeostasis, the liver is critical for the metabolism of nutrients including sugars, lipids, proteins and iron, for the clearance of toxins, and to induce immune tolerance to gut-derived antigens. These functions predispose the liver to infection by blood-borne pathogens, and to a variety of diseases ranging from toxin and medication-induced disorders (CCl₄, acetaminophen) to metabolic disorders (steatohepatitis, alcoholic liver disease, biliary obstruction, cholestasis) or autoimmunity. Chronic liver injury often progresses to life threatening fibrosis and can end in liver cirrhosis and hepatocellular carcinoma (Pellicoro et al., 2014).

The liver contains parenchymal cells or hepatocytes that make up the majority of hepatic cells. It also contains non-parenchymal structural cells such as sinusoidal endothelial cells and a large number of non-parenchymal innate immune cells, mainly monocytes, neutrophils, macrophages, DCs, NK and NKT cells that can trigger an adaptive immune response in the case of infections or other pathogenic insults (Jenne and Kubes, 2013). How this immune response is regulated determines the extent of acute and chronic liver injury (Stijlemans et al., 2014). In this context, liver macrophages have been demonstrated to play central but divergent (from initiating to resolving) functions in liver injury (Sica et al., 2014). It has become clear in the last years that hepatic macrophages consist of two classes, tissue-resident macrophages, the Kupffer cells (KCs) originating from yolk sac/fetal liver progenitors and tissue-infiltrating macrophages originating from bone marrow-derived Ly6C^Hi monocytes (Jinhoux and Jung, 2014; Tacke and Zimmerman, 2014). Distinguishing the activities of KCs from those of monocyte-derived macrophages during liver injury or repair is currently a frontline research topic in the macrophage field. Indeed, considering that clinical management of liver failure remains problematic, a better understanding of the immune mechanisms regulating liver injury is expected to allow the development of new therapeutic modalities. Here, we describe an isolation technique for liver non-parenchymal polymorphonuclear (PMN) and mononuclear myeloid cells permitting their molecular and functional characterization.