The authors would like to thank Bing Zhang for his technical assistance. This work was funded by the Canadian Institutes for Health Research (MOP-86595). T.A.B is a CIHR New Investigator and AHFMR Scholar. Q.H. is supported by a CIHR Canada Graduate Scholarship - Doctoral and an AIHS Full-time Studentship. S.A.N. is supported by a Queen Elizabeth II Graduate Scholarship. A.Y.W.S. is supported by an NSERC Postgraduate Scholarship - Doctoral.

Refer to Figure 1 for an overall scheme of the experimental protocol.
1. Dissection
<ol>
 <li>Place sterile steel mesh screen into 60 x 15 mm Petri dish. One unit is needed per tissue sample.</li>
 <li>Add 5 ml of Hank's balanced salt solution (HBSS) to each dish. Keep dishes on ice.</li>
 <li>Euthanize mice with CO2.</li>
 <li>Secure mouse to dissection surface, ventral side facing up. Spray mouse with 70% ethanol for sterilization and to ensure that the fur is matted down.</li>
 <li>Using surgical scissors, begin dissection by making a ventral incision in the abdominal skin just above the genitalia. Extend the incision upwards to the chin. </li>
 <li>From the midline, extend the incision down along all limbs. Pull back the loose skin and pin it down to the dissection surface.</li>
 <li>Harvest the thymus:
 <ol>
 <li>Lift the bottom tip of the sternum to make an incision.</li>
 <li>Avoiding the liver, cut the diaphragm to detach the rib cage and then cut the rib cage to each side. Cut the ribcage upwards on each side, taking care to avoid the lungs and heart.</li>
 <li>Gently pull the ribcage back with forceps. The thymus is a white, bilobed organ located above the heart. Using the flat edge of the forceps to grasp the bottom of the lobes, gently pull the thymus and place it on the mesh screen.</li>
 </ol>
 </li>
 <li>Harvest the spleen:
 <ol>
 <li>The spleen is a red, surfboard-shaped organ located on the left side of the mouse's abdominal cavity, below the liver.</li>
 <li>Make an incision into the abdominal cavity. Gently pull out the spleen with one pair of forceps, using the second pair to tease away connective tissue. Place the spleen on a separate mesh screen.</li>
 </ol>
 </li>
</ol>
2. Cell Preparation
<ol>
 <li>Using a plunger from a 3 ml syringe, grind organs into the mesh screen until only connective tissue and fat remains. Rinse mesh with HBSS from the dish three times. Use a new plunger for each tissue sample.
 <ol>
 <li>Other options for homogenizing tissue can be used in place of this method.</li>
 </ol>
 </li>
 <li>Count cells using a hemocytometer.</li>
 <li>Pellet cells by centrifugation at 335 x g for 5 min at 4 &deg;C.</li>
 <li>Resuspend splenocytes in 500 &mu;l of ACK lysis buffer (0.15 M NH4Cl, 10 mM KHC03, 0.1 mM Na2EDTA, pH 7.2) for 10 min at room temperature. Resuspend thymocytes at 20 x 106 cells/ml in FACS buffer (PBS, 1% FCS, 0.02% sodium azide) and set aside on ice.</li>
 <li>Return splenocytes to isotonicity by adding 5 ml of HBSS. Pellet splenocytes by centrifugation and resuspend at 20 x 106 cells/ml in FACS buffer. If cells need to remain sterile, you may use sterile RPMI + 10% FCS.</li>
</ol>
3. Staining Cells for Flow Cytometry
The purpose of this protocol is to outline strategies for the analysis of positive and negative selection using wildtype (WT) and HYcd4 mice. For general considerations regarding flow cytometry experimental design, acquisition, and analysis, we refer readers to an excellent review by Tung et al.11
<ol>
 <li>Aliquot 4 x 106 thymocytes per flow cytometry sample to each well of a 96-well plate, as well as 1 x 106 WT splenocytes per compensation control. If using transgenic mice, you should include a WT mouse as a control in your experiment.</li>
 <li>Block Fc receptors by incubating cells with anti-CD16/32 (clone 2.4G2) for 10 min on ice.</li>
 <li>Spin plate at 335 x g at 4 &deg;C for 5 min. Remove lid and dispel liquid from wells by flicking the plate once, face down into a sink. Resuspend each well in 200 &mu;l of FACS buffer. Repeat the wash.</li>
 <li>Prepare antibody cocktails as listed below, using the optimal concentration of each antibody based on dilution in 200 &mu;l of FACS buffer per well. The optimal concentration of each antibody is defined as the lowest concentration needed to give the largest separation of positive and negative populations. If there is no negative population for a given antibody, an isotype antibody should be included. The total volume per well can be scaled down (e.g. to 100 &mu;l per well) if desired.
 <ol>
 <li>WT thymus: anti-TCR&beta;, anti-CD4, anti-CD8, anti-CD69 or anti-CD5, anti-CD24</li>
 <li>HYcd4 thymus: anti-HY TCR (T3.70), anti-CD4, anti-CD8, anti-CD69 or anti- CD5, anti-CD24 
To obtain better separation of CD69lo and CD69hi populations in thymocytes, it is recommended that a biotinylated anti-CD69 primary antibody be used, followed by a secondary stain containing fluorochrome-conjugated streptavidin. We use the same strategy for CD5 staining.
 </li>
 </ol>
 </li>
 <li>Incubate cells with 200 &mu;l of antibody cocktail in FACS buffer for 30 min on ice in the dark.</li>
 <li>Concurrent with antibody cocktail staining, stain each compensation control with a single fluorochrome-conjugated antibody. Ideally, compensation staining involves the same antibodies as used in the antibody cocktail. However, staining for each individual antigen may not be feasible for larger experiments when many antigens are being assayed. Therefore, staining for either anti-CD4 or anti-CD8 is recommended due to high expression of these antigens, or the use of compensation beads. Stain in FACS buffer for 30 min on ice in the dark. Leave one compensation control unstained. </li>
 <li>Wash cells twice with FACS buffer.</li>
 <li>Resuspend cells in FACS buffer and transfer to FACS tubes.</li>
 <li>Acquire samples on flow cytometer. Include acquisition of FSC-A and FSC-W to allow for doublet discrimination.</li>
</ol>
4. Analysis of Flow Cytometry Data - Non-TCR Transgenic Mice
We use FlowJo for flow cytometry data analysis. Refer to Figure 2A for the gating strategy for non-TCR transgenic mice.
<ol>
 <li>Using FSC-A by SSC, electronically gate on the "lymphocyte" population.</li>
 <li>Within the "lymphocyte" population, use FSC-A by FSC-W to electronically gate the FSC-Wlo population to exclude cell doublets and aggregrates. This is the "singlet" gate.</li>
 <li>Using the events in the "singlet" gate, plot CD8 by CD4. Draw gates for DN, DPdull, DPbright, CD4+CD8lo, CD4SP and CD8SP populations as depicted in Figure 2B.</li>
 <li>Under the CD4SP and CD8SP gates, create TCR&beta; by CD24 plots. Use the quadrant gating tool to draw gates as depicted in Figure 2C.</li>
 <li>Create a new plot from the "singlet" gate: TCR&beta; by CD69. Draw gates A, B, C, D as depicted in Figure 2D.</li>
 <li>Create another plot from the "singlet" gate: TCR&beta; by CD5. Draw gates i, ii, iii, iv as depicted in Figure 2E. This is another strategy for examining positive selection.</li>
 <li>Apply DN, DPdull, DPbright, CD4int, CD4SP and CD8SP gates from under "singlet" to gates i, ii, iii, iv, A, B, C, D (Figure 2D, E).</li>
 <li>Calculate the number of cells in a particular subset by multiplying organ cellularity by the frequency of each subsequent gate until you reach your target population. For example, the number of TCR&beta;hiCD69- CD8SP thymocytes would be determined by thymus cellularity x %TCRbhi CD69- (fraction D) x %CD8SP.
 <ol>
 <li>Counting already takes into account the live and dead cells so do not include the "lymphocyte" gate in your calculations.</li>
 </ol>
 </li>
</ol>
5. Analysis of Flow Cytometry Data - TCR Transgenic Mice
Refer to Figure 3A for the gating strategy for TCR transgenic mice. 
<ol>
 <li>Follow steps 4.1 and 4.2 as in the previous section.</li>
 <li>Under the "singlet" gate, create a T3.70 by SSC plot and gate on the T3.70+ cell population to analyze antigen-specific T cells (Figure 3B). In some circumstances, it may be of interest to compare this population to the non-antigen specific (T3.70-) T cell population.</li>
 <li>Within the T3.70+ population, draw DN, DP, CD4SP and CD8SP gates (Figure 3C).
 <ol>
 <li>For WT control samples, draw these gates under the &#x201C;singlet&#x201D; gate or create a T3.70- gate as there will be very few T3.70+ cells.</li>
 </ol>
 </li>
 <li> Under the CD8SP gate, create a T3.70 by CD24 plot. Use the quadrant gating tool to divide cells into four populations (Figure 3F).</li>
 <li>Create an overlaid histogram depicting CD69 expression in the DP compartment of WT mice and T3.70+ DP compartment of HYcd4 mice (Figure 4A). Repeat to examine CD5 expression (Figure 4B).</li>
 <li>Calculate the absolute number of cells in each subset of interest as in 4.8.</li>
</ol>

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No conflicts of interest declared.

The protocol presented here can be used to examine positive and negative selection in non-TCR transgenic and TCR transgenic mice. This protocol describes the staining of surface antigens. For further analysis of molecular mechanisms, it is often necessary to perform intracellular staining. We use the BD Biosciences Cytofix/Cytoperm Kit for most intracellular proteins and the BD Biosciences Foxp3 Staining Kit for transcription factors. We usually acquire our samples immediately after staining. However, samples can ...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>HyClone Hank's balanced salt solution</td>
 <td>Thermo Scientific</td>
 <td>SH30030.02</td>
 <td></td>
 </tr>
 <tr>
 <td>Metal mesh screens</td>
 <td>Cedarlane</td>
 <td>CX-0080-E-01</td>
 <td></td>
 </tr>
 <tr>
 <td>Petri dishes (60 x 15 mm)</td>
 <td>Fisher Scientific</td>
 <td>877221</td>
 <td></td>
 </tr>
 <tr>
 <td>Syringes (3 ml)</td>
 <td>BD Biosciences</td>
 <td>309657</td>
 <td></td>
 </tr>
 <tr>
 <td>Conical tubes (15 ml)</td>
 <td>Sarstedt</td>
 <td>62.554.205</td>
 <td></td>
 </tr>
 <tr>
 <td>Microscope</td>
 <td>Zeiss - Primo Star</td>
 <td>415500-00XX-000</td>
 <td></td>
 </tr>
 <tr>
 <td>Hemocytometer </td>
 <td>Hausser Scientific</td>
 <td>3110</td>
 <td></td>
 </tr>
 <tr>
 <td>96-well plate</td>
 <td>Sarstedt</td>
 <td>82.1582.001</td>
 <td></td>
 </tr>
 <tr>
 <td>Multichannel pipette</td>
 <td>Fisherbrand</td>
 <td>21-377-829</td>
 <td></td>
 </tr>
 <tr>
 <td>Fetal calf serum</td>
 <td>PAA</td>
 <td>A15-701</td>
 <td></td>
 </tr>
 <tr>
 <td>Phosphate buffered saline</td>
 <td>Fisher Scientific</td>
 <td>SH3025802</td>
 <td></td>
 </tr>
 <tr>
 <td>Sodium azide</td>
 <td>IT Baker Chemical Co.</td>
 <td>V015-05</td>
 <td></td>
 </tr>
 <tr>
 <td>FcR blocking reagent</td>
 <td></td>
 <td></td>
 <td>Clone 2.4G2</td>
 </tr>
 <tr>
 <td>Anti-mouse HY TCR </td>
 <td>eBioscience</td>
 <td>XX-9930-YY*</td>
 <td>Clone T3.70</td>
 </tr>
 <tr>
 <td>Anti-mouse CD4</td>
 <td>eBioscience</td>
 <td>XX-0042-YY*</td>
 <td>Clone RM4-5</td>
 </tr>
 <tr>
 <td>Anti-mouse CD8&alpha; </td>
 <td>eBioscience</td>
 <td>XX-0081-YY*</td>
 <td>Clone 53-6.7</td>
 </tr>
 <tr>
 <td>Anti-mouse CD24</td>
 <td>eBioscience</td>
 <td>XX-0242-YY*</td>
 <td>Clone M1/69</td>
 </tr>
 <tr>
 <td>Anti-mouse TCR&beta; </td>
 <td>eBioscience</td>
 <td>XX-5961-YY*</td>
 <td>Clone H57-597</td>
 </tr>
 <tr>
 <td>Anti-mouse CD69 Biotinylated</td>
 <td>eBioscience</td>
 <td>13-0691-YY*</td>
 <td>Clone H1.2F3</td>
 </tr>
 <tr>
 <td>Anti-mouse CD5 Biotinylated</td>
 <td>eBioscience</td>
 <td>13-0051-YY*</td>
 <td>Clone 53-7.3</td>
 </tr>
 <tr>
 <td>Streptavidin</td>
 <td>eBioscience</td>
 <td>XX-4217-YY*</td>
 <td></td>
 </tr>
 <tr>
 <td>Flow cytometer</td>
 <td>BD Biosciences - FACS Canto </td>
 <td>338962</td>
 <td></td>
 </tr>
 <tr>
 <td>FACS tubes</td>
 <td>BD Biosciences</td>
 <td>352052</td>
 <td></td>
 </tr>
 <tr>
 <td>Flow cytometry analysis software</td>
 <td>TreeStar - Flowjo</td>
 <td>FlowJo v7/9</td>
 <td></td>
 </tr>
 <tr>
 <td>HyClone RPMI - 1640 medium</td>
 <td>Thermo Scientific</td>
 <td>SH30027.01</td>
 <td></td>
 </tr>
 <tr>
 	<td></td>
 <td></td>
 <td></td>
 <td>*XX varies by fluorochrome and YY varies by vial size.</td>
 </tr>
 </tbody>
</table>

examination of thymic positive and negative selection by flow cytometry

A healthy immune system requires that T cells respond to foreign antigens while remaining tolerant to self-antigens. Random rearrangement of the T cell receptor (TCR) &alpha; and &beta; loci generates a T cell repertoire with vast diversity in antigen specificity, both to self and foreign. Selection of the repertoire during development in the thymus is critical for generating safe and useful T cells. Defects in thymic selection contribute to the development of autoimmune and immunodeficiency disorders1-4. 
T cell progenitors enter the thymus as double negative (DN) thymocytes that do not express CD4 or CD8 co-receptors. Expression of the &alpha;&beta;TCR and both co-receptors occurs at the double positive (DP) stage. Interaction of the &alpha;&beta;TCR with self-peptide-MHC (pMHC) presented by thymic cells determines the fate of the DP thymocyte. High affinity interactions lead to negative selection and elimination of self-reactive thymocytes. Low affinity interactions result in positive selection and development of CD4 or CD8 single positive (SP) T cells capable of recognizing foreign antigens presented by self-MHC5.
Positive selection can be studied in mice with a polyclonal (wildtype) TCR repertoire by observing the generation of mature T cells. However, they are not ideal for the study of negative selection, which involves deletion of small antigen-specific populations. Many model systems have been used to study negative selection but vary in their ability to recapitulate physiological events6. For example, in vitro stimulation of thymocytes lacks the thymic environment that is intimately involved in selection, while administration of exogenous antigen can lead to non-specific deletion of thymocytes7-9. Currently, the best tools for studying in vivo negative selection are mice that express a transgenic TCR specific for endogenous self-antigen. However, many classical TCR transgenic models are characterized by premature expression of the transgenic TCR&alpha; chain at the DN stage, resulting in premature negative selection. Our lab has developed the HYcd4 model, in which the transgenic HY TCR&alpha; is conditionally expressed at the DP stage, allowing negative selection to occur during the DP to SP transition as occurs in wildtype mice10.
Here, we describe a flow cytometry-based protocol to examine thymic positive and negative selection in the HYcd4 mouse model. While negative selection in HYcd4 mice is highly physiological, these methods can also be applied to other TCR transgenic models. We will also present general strategies for analyzing positive selection in a polyclonal repertoire applicable to any genetically manipulated mice.

In physiological TCR transgenic models and WT mice, positive selection begins at the DPbright stage before moving into the DPdull stage after antigen encounter. DPdull thymocytes then enter a transitional CD4+CD8lo stage before becoming CD4SP or CD8SP thymocytes (Figure 2B). Mature SP thymocytes are characterized by high TCR expression and loss of CD24 (Figure 2C). While the CD8 by CD4 profile can reveal defects in positive se...

Watch this Scientific Journal Video about Examination of Thymic Positive and Negative Selection by Flow Cytometry at JoVE.com

Examination of Thymic Positive and Negative Selection by Flow Cytometry

We present a flow cytometry-based method to examine T cell development in vivo using genetically manipulated mice on a wildtype or T cell receptor transgenic background.

A healthy immune system requires that T cells respond to foreign  antigens while remaining tolerant to self-antigens. Random rearrangement of the  T cell ...

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Immunology and Infection

21.6K Views.  University of Alberta. We present a flow cytometry-based method to examine T cell development in vivo using genetically manipulated mice on a wildtype or T cell receptor transgenic background.

Video: Examination of Thymic Positive and Negative Selection by Flow Cytometry

Measuring Calpain Activity in Fixed and Living Cells by Flow Cytometry

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, including signaling, cytoskeletal remodeling, regulation of gene expression, apoptosis and cell cycle progression 1. Calpains have been implicated in many pathologies including muscular dystrophies, cancer, diabetes, Alzheimer's disease and multiple sclerosis 1. Calpain regulation is complex and incompletely understood. mRNA and protein levels correlate poorly with activity, limiting the use of gene or protein expression techniques to measure calpain activity. This video protocol details a flow cytometric assay developed in our laboratory for measuring calpain activity in fixed and living cells. This method uses the fluorescent substrate BOC-LM-CMAC, which is cleaved specifically by calpain, to measure calpain activity. 2 In this video, calpain activity in fixed and living murine 32Dkit leukemia cells, alone or as part of a splenocyte population is measured using an LSRII (BD Bioscience). 32Dkit cells are shown to have elevated activity compared to normal splenocytes.

This article will detail the protocol for measuring calpain activity in fixed and living cells using flow cytometry.

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, ...

Human In Vitro Suppression as Screening Tool for the Recognition of an Early State of Immune Imbalance

Regulatory T cells (Tregs) are critical mediators of immune tolerance to self-antigens. In addition, they are crucial regulators of the immune response following an infection. Despite efforts to identify unique surface marker on Tregs, the only unique feature is their ability to suppress the proliferation and function of effector T cells. While it is clear that only in vitro assays can be used in assessing human Treg function, this becomes problematic when assessing the results from cross-sectional studies where healthy cells and cells isolated from subjects with autoimmune diseases (like Type 1 Diabetes-T1D) need to be compared. There is a great variability among laboratories in the number and type of responder T cells, nature and strength of stimulation, Treg:responder ratios and the number and type of antigen-presenting cells (APC) used in human in vitro suppression assays. This variability makes comparison between studies measuring Treg function difficult. The Treg field needs a standardized suppression assay that will work well with both healthy subjects and those with autoimmune diseases. We have developed an in vitro suppression assay that shows very little intra-assay variability in the stimulation of T cells isolated from healthy volunteers compared to subjects with underlying autoimmune destruction of pancreatic &beta;-cells. The main goal of this piece is to describe an in vitro human suppression assay that allows comparison between different subject groups. Additionally, this assay has the potential to delineate a small loss in nTreg function and anticipate further loss in the future, thus identifying subjects who could benefit from preventive immunomodulatory therapy1. Below, we provide thorough description of the steps involved in this procedure. We hope to contribute to the standardization of the in vitro suppression assay used to measure Treg function. In addition, we offer this assay as a tool to recognize an early state of immune imbalance and a potential functional biomarker for T1D.

Human In Vitro Suppression as Screening Tool for the Recognition of an Early State of Immune Imbalance

Tregs are potent suppressors of the immune system. There is a lack of unique surface markers to define them, hence, definitions of Tregs are primarily functional. Here we describe an optimized in vitro assay capable of identifying immune imbalance in subjects at risk to develop T1D.

Regulatory T cells (Tregs) are critical mediators of immune tolerance to self-antigens. In addition, they are crucial regulators of the immune response ...

Detection of Fluorescent Nanoparticle Interactions with Primary Immune Cell Subpopulations by Flow Cytometry

Engineered nanoparticles are endowed with very promising properties for therapeutic and diagnostic purposes. This work describes a fast and reliable method of analysis by flow cytometry to study nanoparticle interaction with immune cells. Primary immune cells can be easily purified from human or mouse tissues by antibody-mediated magnetic isolation. In the first instance, the different cell populations running in a flow cytometer can be distinguished by the forward-scattered light (FSC), which is proportional to cell size, and the side-scattered light (SSC), related to cell internal complexity. Furthermore, fluorescently labeled antibodies against specific cell surface receptors permit the identification of several subpopulations within the same sample. Often, all these features vary when cells are boosted by external stimuli that change their physiological and morphological state. Here, 50 nm FITC-SiO2 nanoparticles are used as a model to identify the internalization of nanostructured materials in human blood immune cells. The cell fluorescence and side-scattered light increase after incubation with nanoparticles allowed us to define time and concentration dependence of nanoparticle-cell interaction. Moreover, such protocol can be extended to investigate Rhodamine-SiO2 nanoparticle interaction with primary microglia, the central nervous system resident immune cells, isolated from mutant mice that specifically express the Green Fluorescent Protein (GFP) in the monocyte/macrophage lineage. Finally, flow cytometry data related to nanoparticle internalization into the cells have been confirmed by confocal microscopy.

Analysis of nanoparticle interaction with defined subpopulations of immune cells by flow cytometry.

Engineered nanoparticles are endowed with very promising properties for therapeutic and diagnostic purposes. This work describes a fast and reliable ...

In Vivo Assessment of Rodent Plasmodium Parasitemia and Merozoite Invasion by Flow Cytometry

During blood stage infection, malaria parasites invade, mature, and replicate within red blood cells (RBCs). This results in a regular growth cycle and an exponential increase in the proportion of malaria infected RBCs, known as parasitemia. We describe a flow cytometry based protocol which utilizes a combination of the DNA dye Hoechst, and the mitochondrial membrane potential dye, JC-1, to identify RBCs which contain parasites and therefore the parasitemia, of in vivo blood samples from Plasmodium chabaudi adami DS infected mice. Using this approach, in combination with fluorescently conjugated antibodies, parasitized RBCs can be distinguished from leukocytes, RBC progenitors, and RBCs containing Howell-Jolly bodies (HJ-RBCs), with a limit of detection of 0.007% parasitemia. Additionally, we outline a method for the comparative assessment of merozoite invasion into two different RBC populations. In this assay RBCs, labeled with two distinct compounds identifiable by flow cytometry, are transfused into infected mice. The relative rate of invasion into the two populations can then be assessed by flow cytometry based on the proportion of parasitized RBCs in each population over time. This combined approach allows the accurate measurement of both parasitemia and merozoite invasion in an in vivo model of malaria infection.

In Vivo Assessment of Rodent Plasmodium Parasitemia and Merozoite Invasion by Flow Cytometry

The malaria parasite invades and replicates within red blood cells. The accurate assessment of merozoite invasion and parasitemia is therefore crucial in assessing the course of malaria infection. Here we describe a flow cytometry based protocol for the measurement of these parameters in a mouse model of malaria.

During blood stage infection, malaria parasites invade, mature, and replicate within red blood cells (RBCs). This results in a regular growth cycle and an ...

Phenotypic Characterization of Macrophages from Rat Kidney by Flow Cytometry

There is increasing evidence suggesting the important role of inflammation and, subsequently, macrophages in the development and progression of renal disease. Macrophages are heterogeneous cells that have been implicated in kidney injury. Macrophages may be classified into two different phenotypes: classically activated macrophages (M1 macrophages), that release pro-inflammatory cytokines and promote fibrosis; and alternatively activated macrophages (M2 macrophages) that are associated with immunoregulatory and tissue-remodeling functions. These macrophage phenotypes need to be discriminated and analyzed to determine their contribution to renal injury. However, there are scarce studies reporting consistent phenotypic and functional information about macrophage subtypes in inflammatory renal disease models, especially in rats. This fact may be related to the limited macrophage markers used in rats, contrary to mice. Therefore, novel strategies are necessary to quantify and characterize the renal content of these infiltrating cells in a reliable way. This manuscript details a protocol for kidney digestion and further phenotypic and quantitative analysis of macrophages from rat kidneys by flow cytometry. Briefly, kidneys were incubated with collagenase and total macrophages were identified according to the dual presence of CD45 (leukocytes common antigen) and CD68 (PAN macrophage marker) in live cells.This was followed by surface staining of CD86 (M1 marker) and CD163 (M2 marker). Rat peritoneal macrophages were used as positive control for macrophage marker detection by flow cytometry. Our protocol resulted in low cellular mortality and allowed characterization of different intracellular and surface protein markers, thus limiting the loss of cellular integrity observed in other protocols. Moreover, this procedure allows the use of macrophages for further techniques, including cell sorting and mRNA or protein expression studies, among others.

This manuscript describes a detailed protocol for phenotypic and quantitative analysis of resident macrophages from rat kidneys by flow cytometry. The resulting stained cells can be also used for other applications, including cell sorting, gene expression analysis or functional studies, thus increasing the information obtained in the experimental model.

There is increasing evidence suggesting the important role of inflammation and, subsequently, macrophages in the development and progression of renal ...

Fluorescence-activated Cell Sorting for Purification of Plasmacytoid Dendritic Cells from the Mouse Bone Marrow

Fluorescence-activated cell sorting (FACS) is a technique to purify specific cell populations based on phenotypes detected by flow cytometry. This method enables researchers to better understand the characteristics of a single cell population without the influence of other cells. Compared to other methods of cell enrichment, such as magnetic-activated cell sorting (MCS), FACS is more flexible and accurate for cell separation due to the ability of phenotype detection by flow cytometry. In addition, FACS is usually capable of separating multiple cell populations simultaneously, which improves the efficiency and diversity of experiments. Although FACS has some limitations, it has been broadly used to purify cells for functional studies in both in vitro and in vivo settings. Here we report a protocol using fluorescence-activated cell sorting to isolate a very rare population of immune cells, plasmacytoid dendritic cells (pDC), with high purity from the bone marrow of lupus-prone mice for in vitro functional studies of pDC.

We report a protocol using fluorescence-activated cell sorting to isolate plasmacytoid dendritic cells (pDC) with high purity from the bone marrow of lupus-prone mice for functional studies of pDC.

Fluorescence-activated cell sorting (FACS) is a technique to purify specific cell populations based on phenotypes detected by flow cytometry. This method ...

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and their important role in lung development and function, as well as their lung-localized responses to infection and inflammation. To date, no unified method for identification, isolation, and handling of alveolar macrophages from humans and mice exists. Such a method is needed for studies on these important innate immune cells in various experimental settings. The method described here, which can be easily adopted by any laboratory, is a simplified approach to harvesting alveolar macrophages from bronchoalveolar lavage fluid or from lung tissue and maintaining them in vitro. Because alveolar macrophages primarily occur as adherent cells in the alveoli, the focus of this method is on dislodging them prior to harvest and identification. The lung is a highly vascularized organ, and various cell types of myeloid and lymphoid origin inhabit, interact, and are influenced by the lung microenvironment. By using the set of surface markers described here, researchers can easily and unambiguously distinguish alveolar macrophages from other leukocytes, and purify them for downstream applications. The culture method developed herein supports both human and mouse alveolar macrophages for in vitro growth, and is compatible with cellular and molecular studies.

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

This communication describes methodologies for isolation and culture of alveolar macrophages from humans and murine models for experimental purposes.

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and ...

Characterization of Human Monocyte Subsets by Whole Blood Flow Cytometry Analysis

Monocytes are key contributors in various inflammatory disorders and alterations to these cells, including their subset proportions and functions, can have pathological significance. An ideal method for examining alterations to monocytes is whole blood flow cytometry as the minimal handling of samples by this method limits artifactual cell activation. However, many different approaches are taken to gate the monocyte subsets leading to inconsistent identification of the subsets between studies. Here we demonstrate a method using whole blood flow cytometry to identify and characterize human monocyte subsets (classical, intermediate, and non-classical). We outline how to prepare the blood samples for flow cytometry, gate the subsets (ensure contaminating cells have been removed), and determine monocyte subset expression of surface markers &#8212; in this example M1 and M2 markers. This protocol can be extended to other studies that require a standard gating method for assessing monocyte subset proportions and monocyte subset expression of other functional markers.

Here we present a protocol for characterizing monocyte subsets by whole blood flow cytometry. This includes outlining how to gate the subsets and assess their expression of surface markers and giving an example of the assessment of the expression of M1 (inflammatory) and M2 markers (anti-inflammatory).

Monocytes are key contributors in various inflammatory disorders and alterations to these cells, including their subset proportions and functions, can ...

Discrimination of Seven Immune Cell Subsets by Two-fluorochrome Flow Cytometry

Immune cell characterization heavily relies on multicolor flow cytometry to identify subpopulations based on differential expression of surface markers. Setup of a classic multicolor panel requires high-end instruments, custom labeled antibodies, and careful study design to minimize spectral overlap. We developed a multiparametric analysis to identify major human immune populations (CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, NK cells and monocytes) in peripheral blood by combining seven lineage markers using only two fluorochromes. Our strategy is based on the observation that lineage markers are constantly expressed in a unique combination by each cell population. Combining this information with a careful titration of the antibodies allows investigators to record five additional markers, expanding the optical limit of most flow cytometers. Head-to-head comparison demonstrated that the vast majority of immune cell populations in peripheral blood can be characterized with comparable accuracy between our method and the classic &#34;one fluorochrome-one marker approach&#34;, although the latter is still more precise for identifying populations such as NKT cells and &#947;&#948; T cells. Combining seven markers using two fluorochromes allows for the analysis of complex immune cell populations and clinical samples on affordable 6-10 fluorochrome flow cytometers, and even on 2-3 fluorochrome field instruments in areas with limited resources. High-end instruments can also benefit from this approach by using extra fluorochromes to accomplish deeper flow cytometry analysis. This approach is also very well suited for screening several cell populations in the case of clinical samples with limited number of cells.

Here, we present a flow cytometric protocol to identify CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, NK cells and monocytes in human peripheral blood by using only two fluorochromes instead of seven. With this approach, five additional markers can be recorded on most flow cytometers.

Immune cell characterization heavily relies on multicolor flow cytometry to identify subpopulations based on differential expression of surface markers. ...

Preparation of Whole Bone Marrow for Mass Cytometry Analysis of Neutrophil-lineage Cells

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a myeloid-biased 39-antibody CyTOF panel to evaluate the hematopoietic system with a focus on the neutrophil-lineage cells by using this protocol. The CyTOF result was analyzed with an open-resource dimensional reduction algorithm, viSNE, and the data was presented to demonstrate the outcome of this protocol. We have discovered new neutrophil-lineage cell populations based on this protocol. This protocol of fresh whole BM preparation may be used for 1), CyTOF analysis to discover unidentified cell populations from whole BM, 2), investigating whole BM defects for patients with blood disorders such as leukemia, 3), assisting optimization of fluorescence-activated flow cytometry protocols that utilize fresh whole BM.

Here, we present a protocol to process fresh bone marrow (BM) isolated from mouse or human for high-dimensional mass cytometry (Cytometry by Time-Of-Flight, CyTOF) analysis of neutrophil-lineage cells.

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a ...