免疫学

Macrophages are at the center of innate immunity and iron metabolism. In the case of an infection, macrophages adapt their cellular iron metabolism to deprive iron from invading bacteria to combat intracellular bacterial proliferation. A concise evaluation of the cellular iron content upon an infection with bacterial pathogens and diverse cellular stimuli is necessary to identify underlying mechanisms concerning iron homeostasis in macrophages. For the characterization of cellular iron levels during infection, we established an in vitro infection model where the murine macrophage cell line J774A.1 is infected with Salmonella enterica serovar Typhimurium (S.tm), the mouse counterpart to S. enterica serovar Typhi, under normal and iron-overload conditions using ferric chloride (FeCl₃) treatment. To evaluate the effect of infection and iron stimulation on cellular iron levels, the macrophages are stained with FerroOrange. This fluorescent probe specifically detects Fe²⁺ ions and its fluorescence can be quantified photometrically in a plate reader. Importantly, FerroOrange fluorescence does not increase with chelated iron or other bivalent metal ions. In this protocol, we present a simple and reliable method to quantify cellular Fe²⁺ levels in cultured macrophages by applying a highly specific fluorescence probe (FerroOrange) in a TECAN Spark microplate reader. Compared to already established techniques, our protocol allows assessing cellular iron levels in innate immune cells without the use of radioactive iron isotopes or extensive sample preparation, exposing the cells to stress.

Key features

• Easy quantification of Fe²⁺ in cultured macrophages with a fluorescent probe.

• Analysis of iron in living cells without the need for fixation.

• Performed on a plate reader capable of 540 nm excitation and 585 nm emission by trained employees for handling biosafety level 2 bacteria.

Graphical overview

Clearance of dying cells, named efferocytosis, is a pivotal function of professional phagocytes that impedes the accumulation of cell debris. Efferocytosis can be experimentally assessed by differentially tagging the target cells and professional phagocytes and analyzing by cell imaging or flow cytometry. Here, we describe an assay to evaluate the uptake of apoptotic cells (ACs) by human macrophages in vitro by labeling the different cells with commercially available dyes and analysis by flow cytometry. We detail the methods to prepare and label human macrophages and apoptotic lymphocytes and the in vitro approach to determine AC uptake. This protocol is based on previously published literature and allows for in vitro modeling of the efficiency of AC engulfment during continual efferocytosis process. Also, it can be modified to evaluate the clearance of different cell types by diverse professional phagocytes.

Graphical overview

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.

An inflammasome is an intracellular multiprotein complex that plays important roles in host defense and inflammatory responses. Inflammasomes are typically composed of the adaptor protein apoptosis-associated speck-like protein containing a CARD (ASC), cytoplasmic sensor protein, and the effector protein pro-caspase-1. ASC assembly into a protein complex termed ASC speck is a readout for inflammasome activation. Here, we provide a step-by-step protocol for the detection of ASC speck by confocal microscopy in Bone marrow derived macrophages (BMBDs) triggered by chemical stimuli and bacterial pathogens. We also describe the detailed procedure for the generation of BMDMs, stimulating conditions for inflammasome activation, immunofluorescence cell staining of ASC protein, and microscopic examination. Thus far, this method is a simple and reliable manner to visualize and quantify the intracellular localization of ASC speck.

Graphic abstract:

Figure 1. Confocal microscopy detection of ASC speck formation in untreated WT BMDMs and WT BMDMs stimulated with LPS and ATP, transfected with dsDNA, and infected with F. novicida or Salmonella as indicated. Arrow indicates the ASC speck. Scale bars: 10 μm.

Over the last decade, lipids have emerged as possessing an ever-increasing number of key functions, especially in membrane trafficking. For instance, phosphatidic acid (PA) has been proposed to play a critical role in different steps along the secretory pathway or during phagocytosis. To further investigate in detail the precise nature of PA activities, we need to identify the organelles in which PA is synthesized and the PA subspecies involved in these biological functions. Indeed, PA, like all phospholipids, has a large variety based on its fatty acid composition. The recent development of PA sensors has helped us to follow intracellular PA dynamics but has failed to provide information on individual PA species. Here, we describe a method for the subcellular fractionation of RAW264.7 macrophages that allows us to obtain membrane fractions enriched in specific organelles based on their density. Lipids from these membrane fractions are precipitated and subsequently processed by advanced mass spectrometry-based lipidomics analysis to measure the levels of different PA species based on their fatty acyl chain composition. This approach revealed the presence of up to 50 different species of PA in cellular membranes, opening up the possibility that a single class of phospholipid could play multiple functions in any given organelle. This protocol can be adapted or modified and used for the evaluation of other intracellular membrane compartments or cell types of interest.

The ability to conduct in vivo macrophage-specific depletion remains an effective means to uncover functions of macrophages in a wide range of physiological contexts. Compared to the murine model, zebrafish offer superior imaging capabilities due to their optical transparency starting from a single-cell stage to throughout larval development. These qualities become important for in vivo cell specific depletions so that the elimination of the targeted cells can be tracked and validated in real time through microscopy. Multiple methods to deplete macrophages in zebrafish are available, including genetic (such as an irf8 knockout), chemogenetic (such as the nitroreductase/metronidazole system), and toxin-based depletions (such as using clodronate liposomes). The use of clodronate-containing liposomes to induce macrophage apoptosis after phagocytosing the liposomes is effective in depleting macrophages as well as testing their ability to phagocytose. Here we describe a detailed protocol for the systemic depletion of macrophages in zebrafish larvae by intravenous injection of liposomal clodronate supplemented with fluorescent dextran conjugates. Co-injection with the fluorescent dextran allows tracking of macrophage depletion in real time starting with verifying the successful intravenous injection to macrophage uptake of molecules and their eventual death. To verify a high degree of macrophage depletion, the level of brain macrophage (microglia) elimination can be determined by a rapid neutral red vital dye staining when clodronate injection is performed at early larval stages.

Graphical abstract:

Experimental workflow for in vivo macrophage-specific depletion by liposomal clodronate in larval zebrafish

Glomerulonephritis (GN) is a common pathological condition in chronic kidney diseases that often leads to end stage renal failure. Mac-1 (CD11b/CD18)-mediated neutrophil, macrophage, and dendritic cell glomerular infiltration leading to cellular dysfunction and destruction is an important disease mechanism. The cellular distribution and dynamics of the expression of Mac-1 ligands ICAM-1 and ICAM-2 in GN have not been well studied because of the difficulties in tissue staining and colocalizing glomerular cells with surface antigens. To improve the visualization of cell surface marker and antigen expression in kidney compartments, we have devised an even but mild fixation procedure employing p-formaldehyde-lysine-periodate (PLP) perfusion. A large panel of antibodies (Ab) against cell surface markers was used to identify kidney cell types and adhesion molecules. When confocal microscopy was used in visualizing glomerular adhesion molecule staining, the endothelial cells were found to specifically express CD31, and these cells express ICAM-2 constitutively. Though ICAM-1 was not expressed by glomerular endothelial cells in homeostasis, it was highly upregulated in mice with chronic GN and severe proteinuria. VCAM-1, a ligand for VLA-4 important in leukocyte migration, was not expressed in the glomerulus. The results highlight the importance of ICAM-1 in the infiltration of macrophages and dendritic cells in cGN. This report will provide a widely applicable procedure for yielding high quality confocal images and for the identification and quantitation of receptors and other cellular antigens expressed in different kidney compartments and cell types.

Methods to test both the functionality and mechanism of action for human recombinant proteins and antibodies in vitro have been limited by multiple factors. To test the functionality of a recombinant protein or antibody, the receptor, the receptor-associated ligand, or both must be expressed by the cells present within the in vitro culture. While the use of transfected cell lines can circumvent this gap, the use of transfected cell lines does not allow for studying the native signaling pathway(s) modulated by the specific recombinant protein or antibody in primary cells. The present protocol utilizes sort purified CD14⁺ monocytes and T cells, both CD4⁺ T cells and CD8⁺ T cells, from healthy donors in a co-culture system. This methodology is particularly relevant for testing recombinant proteins or antibodies that are putative therapeutics for the treatment of autoimmune disease and cancer. While the current protocol focuses on co-cultures containing B7-H4 expressing monocytes plus either autologous CD4⁺ T cells or CD8⁺ T cells, the protocol can be modified for the user’s specific needs.

Tissue-resident macrophages are pivotal for a tightly-regulated iron metabolism at a cellular and systemic level, since subtle iron alterations increase the susceptibility for microbial infections or drive multiple diseases. However, research on cellular iron homeostasis in macrophages remains challenging due to the limited amount of available methods using radioactive ⁵⁹Fe isotopes or strong iron chelators, which might be inapplicable in certain experimental settings. This protocol describes the analysis of the quenchable iron pool (QIP) in macrophages by loading these cells with exogenous iron-complexes. Thereby, the cytoplasmic iron pool can be determined, since the iron uptake ability of macrophages inversely correlates with intracellular iron levels. Thus, this assay enables the accurate analysis of even minor alterations in cytoplasmic iron fluxes and is applicable in almost every laboratory environment. In addition, the protocol can also be adopted for other immune cell types in vitro and in vivo.

Chlamydia trachomatis (C.t.) is an obligate intracellular pathogen that cannot be cultured axenically and must be propagated within eukaryotic host cells. There are at least 15 distinct chlamydial serovariants that belong to 2 major biovars commonly referred to as trachoma and lymphogranuloma venereum (LGV). The invasive chlamydia LGV serovar L2 is the most widely used experimental model for studying C.t. biology and infection and is the only strain with reliable genetic tools available. New techniques to genetically manipulate C.t. L2 have provided opportunities to make mutants using TargeTron and allelic exchange as well as strains overexpressing epitope-tagged proteins, in turn necessitating the regular purification of transformant and mutant clones. Purification of C.t. is a labor-intensive exercise and one of the most common reagents classically used in the purification process, Renografin, is no longer commercially available. A similar formulation of diatrizoate meglumine called Gastrografin is readily available and we as well as others have had great success using this in place of Renografin for chlamydial purifications. Here, we provide a detailed general protocol for infection, propagation, purification, and titering of Chlamydia trachomatis serovar L2 with additional notes specifically pertaining to mutants or recombinant DNA carrying clones.