Immunology

CRISPR/Cas9 genome editing is a widely used tool for creating genetic knock-ins, which allow for endogenous tagging of genes. This is in contrast with random insertion using viral vectors, where expression of the inserted transgene changes the total copy number of a gene in a cell and does not reflect the endogenous chromatin environment or any trans-acting regulation experienced at a locus. There are very few protocols for endogenous fluorescent tagging in macrophages. Here, we describe a protocol to design and test CRISPR guide RNAs and donor plasmids, to transfect them into RAW 264.7 mouse macrophage-like cells using the Neon transfection system and to grow up clonal populations of cells containing the endogenous knock-in at various loci. We have used this protocol to create endogenous fluorescent knock-ins in at least six loci, including both endogenously tagging genes and inserting transgenes in the Rosa26 and Tigre safe harbor loci. This protocol uses circular plasmid DNA as the donor template and delivers the sgRNA and Cas9 as an all-in-one expression plasmid. We designed this protocol for fluorescent protein knock-ins; it is best used when positive clones can be identified by fluorescence. However, it may be possible to adapt the protocol for non-fluorescent knock-ins. This protocol allows for the fairly straightforward creation of clonal populations of macrophages with tags at the endogenous loci of genes. We also describe how to set up imaging experiments in 24-well plates to track fluorescence in the edited cells over time.

Key features

• CRISPR knock-in of fluorescent proteins in RAW 264.7 mouse macrophages at diverse genomic loci.

• This protocol is optimized for the use of the Neon transfection system.

• Includes instructions for growing up edited clonal populations from single cells with one single-cell sorting step and efficient growth in conditioned media after cell sorting.

• Designed for knocking in fluorescent proteins and screening transfected cells byFACS, but modification for non-fluorescent knock-ins may be possible.

Graphical overview

During the onset of autoimmune diabetes, nerve–immune cell interactions seem to play an important role; however, there are currently no models to follow and interfere with these interactions over time in vivo or in vitro. Two-dimensional in vitro models provide insufficient information and microfluidics or organs on a chip are usually challenging to work with. We present here what we believe to be the first simple model that provides the opportunity to co-culture pancreatic islets with sympathetic nerves and immune cells. This model is based on our stamping device that can be 3D printed (STL file provided). Due to the imprint in the agarose gel, sympathetic neurons, pancreatic islets, and macrophages can be seeded in specific locations at a level that allows for confocal live-cell imaging. In this protocol, we provide the instructions to construct and perform live cell imaging experiments in our co-culture model, including: 1) design for the stamping device to make the imprint in the gel, 2) isolation of sympathetic neurons, pancreatic islets, and macrophages, 3) co-culture conditions, 4) how this can be used for live cell imaging, and 5) possibilities for wider use of the model. In summary, we developed an easy-to-use co-culture model that allows manipulation and imaging of interactions between sympathetic nerves, pancreatic islets, and macrophages. This new co-culture model is useful to study nerve– immune cell– islet interactions and will help to identify the functional relevance of neuro-immune interactions in the pancreas.

Key features

• A novel device that allows for 3D co-culture of sympathetic neurons, pancreatic islets, and immune cells

• The device allows the capture of live interactions between mouse sympatheticneurons, pancreatic islets, and immune cells in a controlled environment after six days of co-culturing.

• This protocol uses cultured sympathetic neurons isolated from the superior cervical ganglia using a previously established method (Jackson and Tourtellotte, 2014) in a 3D co-culture.

• This method requires 3D printing of our own designed gel-stamping device (STL print file provided on SciLifeLab FigShare DOI: 10.17044/scilifelab.24073062).

Graphical overview

Graphical overview of co-culture model. 1) Print the stamp with a 3D printer. 2) Isolate neurons, islets, and macrophages. 3) Use the stamp to make the imprint in the agarose gel. 4) Seed the macrophages and islets in the agarose gel on their seeding points. 5) Place the coverslip with neurons on top. 6) Incubate the culture for six days. 7) Image the co-culture. Images adapted from BioRender.

Current means to quantify cells, gene expression, and fibrosis of liver histological slides are not standardized in the research community and typically rely upon data acquired from a selection of random regions identified in each slide. As such, analyses are subject to selection bias as well as limited subsets of available data elements throughout the slide. A whole-slide analysis of cells and fibrosis would provide for a more accurate and complete quantitative analysis, along with minimization of intra- and inter-experimental variables. Herein, we present LiverQuant, a method for quantifying whole-slide scans of digitized histologic images to render a more comprehensive analysis of presented data elements. After loading images and preparing the project in the QuPath program, researchers are provided with one to two scripts per analysis that generate an average intensity threshold for their staining, automated tissue annotation, and downstream detection of their anticipated cellular matrices. When compared with two standard methodologies for histological quantification, LiverQuant had two significant advantages: increased speed and a 50-fold greater tissue area coverage. Using publicly available open-source code (GitHub), LiverQuant improves the reliability and reproducibility of experimental results while reducing the time scientists require to perform bulk analysis of liver histology. This analytical process is readily adaptable by most laboratories, requires minimal optimization, and its principles and code can be optimized for use in other organs.

Graphical overview

CD45 is a pan-leukocyte marker, and CD45 stain is widely used to determine the extent of inflammatory cell infiltration and its association with tissue injury. In this manuscript, we share a reliable immunohistochemistry (IHC) protocol for CD45 staining in sections of paraffin-embedded mouse kidney. A rat anti-CD45 antibody was used as primary antibody, and a mouse adsorbed biotin-conjugated goat anti-rat IgG was selected as secondary antibody. A horseradish peroxidase (HRP)-linked avidin/biotin detection system was used to amplify the signal, which was detected with 3,3′-Diaminobenzidine (DAB). With this protocol, we show that the CD45 antibody recognizes cells of hematolymphoid lineage in bone marrow, as well as monocyte/macrophages in liver and lung tissue. The utility of this protocol in pathology research was indicated by dramatically increased CD45-positive (CD45⁺) cells in the kidneys of a mouse model of diabetes. Double staining for CD45 and injury marker KIM-1 showed accumulated CD45⁺ cells around injured tubular cells. CD45 and F4/80 macrophage staining on adjacent tissue sections revealed overlap of CD45⁺ cells with other inflammatory cells.

Elevations in cytosolic calcium (Ca²⁺) drive a wide array of immune cell functions, including cytokine production, gene expression, and cell motility. Live-cell imaging of cells loaded with ratiometric chemical Ca²⁺ indicators remains the gold standard for visualization and quantification of intracellular Ca²⁺ signals; ratiometric imaging can be accomplished with dyes such as Fura-2, the combination of Fluo-4 and Fura-Red, or, alternatively, by expressing genetically-encoded Ca²⁺ indicators (GECI) such as GCaMPs. Here, we describe a detailed protocol for Ca²⁺ imaging of T cells in vitro using genetically encoded or chemical indicators that can also be applied to a wide variety of cell types. The protocol addresses the challenge of facilitating T cell attachment on various substrates prepared on glass-bottom dishes to enable T cell imaging on an inverted microscope. The protocol also emphasizes cell preparation steps that ensure optimal cell viability – an essential requirement for recording dynamic changes in cytosolic Ca²⁺ levels – and that ensure reproducibility between multiple samples. Finally, we describe a simple algorithm to analyze single-cell Ca²⁺ signals over time using Fiji (ImageJ) software.

Neutrophils are one of the first innate immune cells recruited to tissues during inflammation. An important function of neutrophils relies on their ability to release extracellular structures, known as Neutrophil Extracellular Traps or NETs, into their environment. Detecting such NETs in humans has often proven challenging for both biological fluids and tissues; however, this can be achieved by quantitating NET components (e.g., DNA or granule/histone proteins) or by directly visualizing them by microscopy, respectively. Direct visualization by confocal microscopy is preferably performed on formalin-fixed paraffin-embedded (FFPE) tissue sections stained with a fluorescent DNA dye and antibodies directed against myeloperoxidase (MPO) and citrullinated histone 3 (Cit-H3), two components of NETs, following paraffin removal, antigen retrieval, and permeabilization. NETs are defined as extracellular structures that stain double-positive for MPO and Cit-H3. Here, we propose a novel software-based objective method for NET volume quantitation in tissue sections based on the measurement of the volume of structures exhibiting co-localization of Cit-H3 and MPO outside the cell. Such a technique not only allows the unambiguous identification of NETs in tissue sections but also their quantitation and relationship with surrounding tissues.

Graphic abstract:

Graphical representation of the methodology used to stain and quantitate NETs in human lung tissue.

During immune responses, B cells home to lymph nodes (LN), where they encounter antigens. Homing starts with capture and L-selectin-dependent rolling on the activated endothelium of high endothelial venules (HEV). After recognition of chemokines presented on HEV, activation of B cell integrins occurs mediating firm arrest. Subsequently, B cells crawl to the spot of extravasation to enter the LN. Extravasation can be visualized and quantified in vivo by intravital microscopy (IVM) of the inguinal LN. Here, we describe an established protocol that permits detailed in vivo analysis of B cell recruitment to LN under sterile inflammatory conditions. We describe data acquisition, exportation, quantification, and statistical analysis using specialized software. IVM of LN is a powerful technique that can provide a better understanding of B cell migratory behavior during inflammation in vivo.

Supramolecular signaling assemblies are of interest for their unique signaling properties. A µm scale signaling assembly, the central supramolecular signaling cluster (cSMAC), forms at the center interface of T cells activated by antigen presenting cells (APC). The adaptor protein linker for activation of T cells (LAT) is a key cSMAC component. The cSMAC has widely been studied using total internal reflection fluorescence microscopy of CD4⁺ T cells activated by planar APC substitutes. Here we provide a protocol to image the cSMAC in its cellular context at the interface between a T cell and an APC. Super resolution stimulated emission depletion microscopy (STED) was utilized to determine the localization of LAT, that of its active, phosphorylated form and its entire pool. Agonist peptide-loaded APCs were incubated with TCR transgenic CD4⁺ T cells for 4.5 min before fixation and antibody staining. Fixed cell couples were imaged using a 100x 1.4 NA objective on a Leica SP8 AOBS confocal laser scanning microscope. LAT clustered in multiple supramolecular complexes and their number and size distributions were determined. Using this protocol, cSMAC properties in its cellular context at the interface between a T cell and an APC could be quantified.

B lymphocyte activation is regulated by its membrane-bound B cell receptors (BCRs) upon recognizing diverse antigens. It is hypothesized that antigen binding would trigger conformational changes within BCRs, followed by a series of downstream signaling activation. To measure the BCR conformational changes in live cells, a fluorescent site-specific labeling technique is preferred. Genetically encoded fluorescent tags visualize the location of the target proteins. However, these fluorescent proteins are large (~30 kDa) and would potentially perturb the conformation of BCRs. Here, we describe the general procedures of utilizing short tag-based site-specific labeling methodologies combining with fluorescence resonance energy transfer (FRET) assay to monitor the conformational changes within BCR extracellular domains upon antigen engagement.

T follicular helper (Tfh) cells regulate B cell selection for entry into the germinal center (GC) reaction or for differentiation into antibody forming cells. This process takes place at the border between the T and B zones in lymphoid organs and involves physical contacts between T and B cells. During these interactions, T cells endow the B cells with selection signals that promote GC seeding or plasmablast differentiation based on their B cell receptor affinity. In Peyer’s patches (PPs), T cells promote B cell colonization of the subepithelial dome (SED) without effective affinity-based clonal selection. To specifically characterize the T cell population that resides within the SED niche, we performed ex vivo photoactivation of the SED compartment followed by flow cytometry analysis of the labeled cells, as described in this protocol. This technique integrates both spatial and cellular information in studies of immunological niches and can be adapted to various experimental systems.