The work was funded by the German research council (DFG) with grants No. SFB-938-M and SA 393/3-4.

1. Preparation of Pan-leukocytes
<ol>
	<li>Draw 1 mL of peripheral blood from a healthy donor (or patient) in a heparinized syringe. Make sure to have approval by the responsible ethics committee for the blood donation.</li>
	<li>Mix 1 mL of human peripheral blood with 30 mL of ACK lysis buffer (150 mM NH4Cl, 1 mM KHCO3, and 0.1 mM EDTA, pH 7.0) in a 50 mL tube and incubate for 8 min at room temperature.</li>
	<li>Fill the tubes with PBS and centrifuge at 300 x g for 6 min. Aspirate the supernatant and resuspend the pellet in 30 mL of ACK lysis buffer.</li>
	<li>Repeat steps 1.2 and 1.3 until the supernatant is clear. Finally, wash the cells in PBS, centrifuge at 300 x g for 6 min at room temperature, and resuspend the cell pellet in 2 mL of culture medium (RPMI1640 + 10% FCS). Incubate the cells at 37 &#176;C for 60 min.</li>
</ol>
2. Loading of Raji Cells with SEB
<ol>
	<li>Prepare two 15 mL Falcon tubes with 1.5 x 106 Raji cells per tube. Spin down the cells (300 x g for 6 min at room temperature) and discard the supernatant.</li>
	<li>Resuspend the cells in residual medium (about 50&#8211;100 &#181;L), add 1.9 &#181;L (1.9 &#181;G) SEB, and incubate at room temperature for 15 min. Add 5 mL of culture medium, spin down the cells (300 x g for 6 min at room temperature), and resuspend the pellet in culture medium (RPMI1640 + 10% FCS) at a density of 1 x 106 cells/mL.</li>
</ol>
3. Induction of Immune Synapses and Staining Protocol
<ol>
	<li>Pipette 500 &#181;L of the preparation of SEB-loaded Raji cells into a FACS tube and 500 &#181;L of the preparation of unloaded Raji cells into another FACS tube. Add 650 &#181;L of pan-leukocytes to each tube and spin down the cells (300 x g for 10 min at room temperature). Discard the supernatant and resuspend the pellet in 150 &#181;L of culture medium (RPMI1640 + 10% FCS). Incubate at 37 &#176;C (typically for 45 min).</li>
	<li>Gently vortex the cells (10 s at 1,000 rpm) and add 1.5 mL of paraformaldehyde (1.5%) during the vortex to fix the cells. Stop the fixation by adding 1 mL of PBS + 1% BSA. Pellet the cells (300 x g for 10 min at room temperature and resuspension the cell pellet in 1 mL of PBS + 1% BSA after incubating at room temperature for 10 min.</li>
	<li>Pellet (300 x g for 10 min at room temperature) and resuspend the cells in 100 &#181;L of PBS + 1% BSA + 0.1% saponin for 15 min at room temperature to permeabilize the cells (96-well plate, U-shaped).</li>
	<li>Wash the cells in PBS + 1% BSA + 0.1% saponin with centrifugation (300 x g for 10min at room temperature) and resuspend the cell pellet in 50 &#181;L of PBS + 1% BSA + 0.1% saponin containing fluorophore-labelled antibodies or compounds (CD3-PE-TxRed (1:30), Phalloidin-AF647 (1:150), and DAPI (1:3,000)).</li>
	<li>Incubate the cells at room temperature in the dark for 30 min. Wash the cells 3 times by adding 1 mL of PBS + 1% BSA + 0.1% saponin. Centrifuge at 300 x g for 10 min at room temperature. Re-resuspend the cells in 60 &#181;L of PBS for imaging flow cytometry.</li>
</ol>
4. Image Acquisition Using a Flow Cytometer
NOTE: The following image acquisition procedure and data analysis are based on imaging flow cytometry using software such as imagestream (IS100), INSPIRE, and IDEAS. However, other flow cytometers and analysis software can also be used.
<ol>
	<li>Open the analysis software on the computer connected to the imaging flow cytometer and click on Initialize Fluidics of the Instrument menu. Apply the beads on the right port when prompted to do so.</li>
	<li>Load the default template from the File menu and click on Run/Setup. Choose Beads from the View dropdown menu.</li>
	<li>Adjust the bright-field illuminator by clicking on Set Intensity if the indicated value is below 200.</li>
	<li>Run the calibration and test routine in the Assist tab by clicking on Start All.</li>
	<li>Click on Flush/Lock/Load and apply the samples in the left port when prompted to do so. After loading the cells in the flow cytometer, open the Cell Classifier and adjust the values as follows: peak intensity upper limit at 1,022 for each channel, peak intensity lower limit at 50 for channel 2 (DAPI) and channel 5 (CD3-Pe-TxR), area lower limit at 50 for channel 1 (side scatter), and upper limit at 1,500.</li>
	<li>Change the excitation laser power to 405 nm (15 mW), 488 nm (200 mW), and 647 nm (90 mW) in the Setup tab.</li>
	<li>Switch the View dropdown menu between Cells and Beads to evaluate the cell classifier and laser power adjustments. 
	NOTE: Make sure that all cells and cell couples are found the Cell View and that cell clumps, debris, and images with saturated pixels are found in the Debris View by changing the cell classifiers and/or the excitation laser powers.</li>
	<li>Define the sample name and the amount of images to acquire (15,000&#8211;25,000 for samples and 500 for compensation controls) in the Setup tab. Click on Run/Setup to start the acquisition.</li>
</ol>
5. Data analysis
<ol>
	<li>Transfer the raw image files (.rif) to the data analysis computer and open the analysis software.</li>
	<li>Produce a compensation matrix following the instructions of the Compensation dropdown. Save the compensation matrix as comp_Date.ctm.</li>
	<li>Open a sample raw image file (.rif) and apply the comp_Date.ctm in the window that appears to produce the compensated image files (.cif) and the default data analysis file (.daf).</li>
	<li>Open the compensated image file. Convert the images to color mode and adjust the lookup tables to obtain optimal visible colors in the Image Gallery Properties toolbar. Obtain an RGB-merged image using the Composite tab of the Image Gallery Properties toolbar.</li>
	<li>Open the Mask Manager from the Analysis dropdown. Create masks to define the T cells and the immune synapse, as follows:
	<ol>
		<li>Select the T-cell mask: &#34;(Fill(Threshold_Ch05, 60).&#34; Select the valley mask: &#34;Valley(Ch02,3).&#34; Select the T-cell synapse mask: &#34;T-cell mask AND Valley(M02,Ch02, 3).&#34;</li>
	</ol>
	</li>
	<li>Open the Feature Manager from the Analysis dropdown. Calculate the following features:
	<ol>
		<li>For the total CD3 expression in T cells, select &#34;Intensity_T-cells_Ch5.&#34; For the total amount of F-actin in the T cells, select &#34;Intensity_T-cells_Ch6.&#34; For CD3 expression in the immune synapse, select &#34;Intensity_T-cell synapse_Ch5.&#34; For the amount of F-actin in the immune synapse, select &#34;Intensity_T-cell synapse_Ch6.&#34;</li>
		<li>To calculate the T-cell area, select &#34;Area_T-cells.&#34; To calculate the T-cell immune synapse area, select &#34;Area_T-cell synapse.&#34;</li>
	</ol>
	</li>
	<li>Determine the F-actin and CD3 enrichment in the immune synapse by using the equation in the Feature Manager: 
	<img alt="Equation" src="/files/ftp_upload/55345/55345eq1.jpg" /></li>
	<li>Apply the following gating strategy by using histograms and dot plots from the Analysis area (for further details, see References 17 and 19):
	<ol>
		<li>Discard out-of-focus cells by plotting the &#34;Gradient RMS_M2_Ch2&#34; in a histogram; set the threshold at 15.</li>
		<li>Plot the SSC versus CD3 intensity in a dot plot. Set a gate on CD3-positive events.</li>
		<li>Plot the &#34;Aspect ratio&#34; of M02 (DAPI stain) versus the area of M02 (Dapi stain). Gate on T-cell singlets and cell couples accordingly. Correct for true cell couples using the area of the synapse mask, as described previously17,19.</li>
		<li>Determine the amount of F-actin in T-cell singlets and T cells of T-cell/APC couples and the percent of F-actin in the immune synapse.</li>
	</ol>
	</li>
</ol>

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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cofilin:+a+redox+sensitive+mediator+of+actin+dynamics+during+T-cell+activation+and+migration.">Cofilin: a redox sensitive mediator of actin dynamics during T-cell activation and migration.</a> Immunol Rev. 256 (1), 30-47 (2013).</li><li>Comrie, W. A., Burkhardt, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Action+and+Traction:+Cytoskeletal+Control+of+Receptor+Triggering+at+the+Immunological+Synapse.">Action and Traction: Cytoskeletal Control of Receptor Triggering at the Immunological Synapse.</a> Front Immunol. 7, 68 (2016).</li><li>Piragyte, I., Jun, C. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Actin+engine+in+immunological+synapse.">Actin engine in immunological synapse.</a> Immune Netw. 12 (3), 71-83 (2012).</li><li>Rossy, J., Pageon, S. V., Davis, D. M., Gaus, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Super-resolution+microscopy+of+the+immunological+synapse.">Super-resolution microscopy of the immunological synapse.</a> Curr Opin Immunol. 25 (3), 307-312 (2013).</li><li>Mace, E. M., Orange, J. 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The authors have nothing to disclose.

The workflow presented here enables the quantification of immune synapses between human T cells (ex vivo) and APCs. Notably, erythrocyte-lysed pan-leukocytes were used as T-cell sources, making T-cell purification steps dispensable. The B-cell lymphoma cell line Raji served as surrogate APCs. This bears significant advantages, since it allows comparisons between blood donors of the T-cell side of the immune synapse. Furthermore, autologous DCs are hardly available directly from peripheral human blood. The production of m...

T cells are major regulators of the adaptive immune system and are activated through antigenic peptides that are presented in the context of major histocompatibility complexes (MHC). Full T-cell activation requires two signals, the competence signal via the antigen-specific T-cell receptor (TCR)/CD3 complex and the costimulatory signal via accessory receptors. Both signals are generated through the direct interaction of T cells with antigen-presenting cells (APCs). Mature APCs provide the competence signal for T-cell activation through MHC-peptide complexes, and they express costimulatory ligands (e.g., CD80 or CD86) to assure the progression of T-cell activation1. One important function of costimulation is the rearrangement of the actin cytoskeleton2,3,4. The cortical F-actin is relatively static in resting T cells. T-cell stimulation through antigen-bearing APCs leads to a profound rearrangement of the actin cytoskeleton. Actin dynamics (i.e., fast actin polymerization/depolymerization circles) enable the T cells to create forces that are used to transport proteins or organelles, for example. Moreover, the actin cytoskeleton is important for developing a special contact zone between T cells and APCs, called the immune synapse. Due to the importance of the actin cytoskeleton to the immune synapse, it has become essential to develop methods to quantify changes in the actin cytoskeleton of T cells5,6,7,8,9.
By means of actin cytoskeletal aid, surface receptors and signaling proteins are segregated in supramolecular activation clusters (SMACs) within the immune synapse. The stability of the immune synapse is assured by the binding of receptors to F-actin bundles that increase the elasticity of the actin cytoskeleton. Immune synapse formation has been shown to be critical for the generation of the adaptive immune responses. The detrimental effects of a defective immune synapse formation in vivo were first realized in patients suffering from Wiskott Aldrich Syndrome (WAS), a disease in which actin polymerization and, concomitantly, immune synapse formation are disturbed10. WAS patients can suffer from eczema, severe recurrent infections, autoimmune diseases, and melanomas. Despite this finding, it is currently not known whether immune synapse formation differs in the T cells of healthy individuals and patients suffering from immune defects or autoimmune diseases.
Fluorescence microscopy, including confocal, TIRF, and super-resolution microscopy, were used to uncover the architecture of the immune synapse11,12,13,14. The high resolution of these systems and the possibility of performing live-cell imaging enables the collection of exact, spatio-temporal information about the actin cytoskeleton and surface or intracellular proteins in the immune synapse. Many results, however, are based on the analysis of only a few tens of T cells. Moreover, T cells must be purified for these types of fluorescence microscopy. However, for many research questions, the use of unpurified cells rather than the highest-possible resolution is of the utmost importance. This is relevant if T cells from patients are analyzed, since the amount of donated blood is limited and there might be the need to process many samples in parallel.
We established microscopic methods that allow the analysis of the actin cytoskeleton in the immune synapse in the human system15,16,17. These methods are based on imaging flow cytometry, also called In-Flow microscopy18. As a hybrid between multispectral flow cytometry and fluorescence microscopy, imaging flow cytometry has its strengths in analyzing morphological parameters and protein localization in heterogeneous cell populations, such as pan-leukocytes from the peripheral blood. We introduced a methodology that enables us to quantify F-actin in T-cell/APC conjugates of human T cells from whole-blood samples, without the need of time-consuming and costly purification steps17. The technique presented here comprises the whole workflow, from getting the blood sample to the quantification of F-actin in the immune synapse.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>&#62;Multifuge 3 SR</td>
			<td>Heraeus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI 1640</td>
			<td>LifeTechnologies</td>
			<td>#11875085</td>
			<td>500 mL</td>
		</tr>
		<tr>
			<td>FCS</td>
			<td>Pan Biotech#3302-P101102</td>
		</tr>
		<tr>
			<td>Polystyrene Round Bottom Tube</td>
			<td>Falcon</td>
			<td>#352054</td>
			<td>5 mL</td>
		</tr>
		<tr>
			<td>Kulturflasche</td>
			<td>Thermo Scientific</td>
			<td>#178883</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Phosphate Buffered Saline</td>
			<td>Sigma</td>
			<td>D8662</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Roth</td>
			<td>#8076.3</td>
			<td></td>
		</tr>
		<tr>
			<td>Saponin</td>
			<td>Sigma</td>
			<td>S7900</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma Aldrich</td>
			<td>#16005</td>
			<td></td>
		</tr>
		<tr>
			<td>FACS Wash Saponin</td>
			<td></td>
			<td>PBS 1% BSA 0.15 Saponin</td>
		</tr>
		<tr>
			<td>ReaktiosgefaB</td>
			<td>Sarstedt</td>
			<td>72.699.002</td>
			<td>0.5 mL</td>
		</tr>
		<tr>
			<td>Speed Bead</td>
			<td>Amnis</td>
			<td>#400041</td>
			<td></td>
		</tr>
		<tr>
			<td>Minishaker MS1</td>
			<td>IKA Works</td>
			<td>&#160;MS1</td>
			<td></td>
		</tr>
		<tr>
			<td>Mikrotiterplatte</td>
			<td>Greiner Bio One</td>
			<td>#650101</td>
			<td>96U</td>
		</tr>
		<tr>
			<td>Enterotoxin SEB</td>
			<td>Sigma Aldrich</td>
			<td>S4881</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma Aldrich</td>
			<td>D9542</td>
			<td>1:3000</td>
		</tr>
		<tr>
			<td>CD3-PeTxR</td>
			<td>Invitrogen</td>
			<td>MHCD0317</td>
			<td>1:30</td>
		</tr>
		<tr>
			<td>Phalloidin-AF647</td>
			<td>Molecular Probes</td>
			<td>A22287</td>
			<td>1:150</td>
		</tr>
		<tr>
			<td>IS100</td>
			<td>Amnis</td>
			<td></td>
			<td>Imaging flow cytometer</td>
		</tr>
		<tr>
			<td>IDEAS</td>
			<td>Amnis</td>
			<td></td>
			<td>Software</td>
		</tr>
		<tr>
			<td>INSPIRE</td>
			<td>Amnis</td>
			<td></td>
			<td>Software</td>
		</tr>
	</tbody>
</table>

qualitative and quantitative analysis of the immune synapse in the human system using imaging flow cytometry

The immune synapse is the area of communication between T cells and antigen-presenting cells (APCs). T cells polarize surface receptors and proteins towards the immune synapse to assure a stable binding and signal exchange. Classical confocal, TIRF, or super-resolution microscopy have been used to study the immune synapse. Since these methods require manual image acquisition and time-consuming quantification, the imaging of rare events is challenging. Here, we describe a workflow that enables the morphological analysis of tens of thousands of cells. Immune synapses are induced between primary human T cells in pan-leukocyte preparations and Staphylococcus aureus enterotoxin B (SEB)-loaded Raji cells as APCs. Image acquisition is performed with imaging flow cytometry, also called In-Flow microscopy, which combines features of a flow cytometer and a fluorescence microscope. A complete gating strategy for identifying T cell/APC couples and analyzing the immune synapses is provided. As this workflow allows the analysis of immune synapses in unpurified pan-leukocyte preparations and hence requires only a small volume of blood (i.e., 1 mL), it can be applied to samples from patients. Importantly, several samples can be prepared, measured, and analyzed in parallel.

A major goal of the method described here is the quantification of protein enrichment (e.g., F-actin) in the immune synapse between surrogate APCs (Raji cells) and T cells in unpurified pan-leukocytes taken from low-volume (1 mL) human blood samples. The screenshot in Figure 1 gives an overview of the critical gating strategy of this method. It shows the image gallery on the left and the analysis area on the right (Figure 1). The image gallery sh...

Watch this Scientific Journal Video about Qualitative and Quantitative Analysis of the Immune Synapse in the Human System Using Imaging Flow Cytometry at JoVE.com

Qualitative and Quantitative Analysis of the Immune Synapse in the Human System Using Imaging Flow Cytometry

Here, we describe a complete workflow for the qualitative and quantitative analysis of immune synapses between primary human T cells and antigen-presenting cells. The method is based on imaging flow cytometry, which allows the acquisition and evaluation of several thousand cell images within a relatively short period of time.

The immune synapse is the area of communication between T cells and antigen-presenting cells (APCs). T cells polarize surface receptors and proteins ...

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Immunology and Infection

11.3K Views.  University of Heidelberg. Here, we describe a complete workflow for the qualitative and quantitative analysis of immune synapses between primary human T cells and antigen-presenting cells. The method is based on imaging flow cytometry, which allows the acquisition and evaluation of several thousand cell images within a relatively short period of time.

Video: Qualitative and Quantitative Analysis of the Immune Synapse in the Human System Using Imaging 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, ...

Flow Cytometry Analysis of Immune Cells Within Murine Aortas

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam cells, immune cells, vascular endothelial and smooth muscle cells, platelets, extracellular matrix, and a lipid-rich core with extensive necrosis and fibrosis of surrounding tissues.1 The innate and adaptive arms of the immune response are involved in the initiation, development and persistence of atherosclerosis.2, 3 There is a significant body of evidence that different subsets of the immune cells, such as macrophages, dendritic cells, T and B lymphocytes, are present within the aortas of healthy and atherosclerosis-prone mice4. Additionally, immune cells are found in the surrounding aortic adventitia which suggests an important role of this tissue in atherogenesis.2
For some time, the quantitative detection of different types of immune cells, their activation status, and the cellular composition within the aortic wall was limited by RT-PCR and immunohistochemical methods for the study of atherosclerosis. Few attempts were made to perform flow cytometry using human aortas, and a number of problems, such as a high autofluorescence, have been reported5,6. Human atherosclerotic plaques were digested with collagenase 1, and free cells were collected and stained for CD14+/CD11c+ to highlight macrophage-derived foam cells. In this study, a "mock" channel was used to avoid false-positive staining.6 Necrotic materials accumulating during the digestion process give rise in a large amount of debris that generates a high autofluorescence in aortic samples. To resolve this problem, a panel of negative and positive controls has been proposed, but only double staining could be applied in these samples. We have developed a new flow cytometry-based method7 to analyze the immune cell composition and characterize the activation, proliferation, differentiation of immune cells in healthy and atherosclerosis-prone aorta. This method allows the investigation of the immune cell composition of the aortic wall and opens possibilities to use a broad spectrum of immunological methods for investigations of immune aspects of this disease.

This paper presents a flow cytometry-based method to investigate the immune composition of aortas. The paper also illustrates an additional technique that allows examining surrounding adventitia and vessel wall separately. This method opens possibilities to perform phenotypical analyses of aortic leukocytes and apply several immunological assays for atherosclerosis studies.

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam ...

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 ...

Intravital Imaging of the Mouse Thymus using 2-Photon Microscopy

Two-photon Microscopy (TPM) provides image acquisition in deep areas inside tissues and organs. In combination with the development of new stereotactic tools and surgical procedures, TPM becomes a powerful technique to identify "niches" inside organs and to document cellular "behaviors" in live animals. While intravital imaging provides information that best resembles the real cellular behavior inside the organ, it is both more laborious and technically demanding in terms of required equipment/procedures than alternative ex vivo imaging acquisition. Thus, we describe a surgical procedure and novel "stereotactic" organ holder that allows us to follow the movements of Foxp3+ cells within the thymus.
Foxp3 is the master regulator for the generation of regulatory T cells (Tregs). Moreover, these cells can be classified according to their origin: ie. thymus-differentiated Tregs are called "naturally-occurring Tregs" (nTregs), as opposed to peripherally-converted Tregs (pTregs). Although significant amount of research has been reported in the literature concerning the phenotype and physiology of these T cells, very little is known about their in vivo interactions with other cells. This deficiency may be due to the absence of techniques that would permit such observations. The protocol described in this paper provides a remedy for this situation.
Our protocol consists of using nude mice that lack an endogenous thymus since they have a punctual mutation in the DNA sequence that compromises the differentiation of some epithelial cells, including thymic epithelial cells. Nude mice were gamma-irradiated and reconstituted with bone marrows (BM) from Foxp3-KIgfp/gfp mice. After BM recovery (6 weeks), each animal received embryonic thymus transplantation inside the kidney capsule. After thymus acceptance (6 weeks), the animals were anesthetized; the kidney containing the transplanted thymus was exposed, fixed in our organ holder, and kept under physiological conditions for in vivo imaging by TPM. We have been using this approach to study the influence of drugs in the generation of regulatory T cells.

We have developed novel laboratory tools and protocols for intravital imaging acquisition of the thymus. Our technique should help in the identification of &#x201C;niches&#x201D; within the thymus where T cell development occurs.

Two-photon Microscopy (TPM) provides image acquisition in deep areas inside tissues and organs. In combination with the development of new stereotactic ...

A Simple and Rapid Protocol to Non-enzymatically Dissociate Fresh Human Tissues for the Analysis of Infiltrating Lymphocytes

The ability of malignant cells to evade the immune system, characterized by tumor escape from both innate and adaptive immune responses, is now accepted as an important hallmark of cancer. Our research on breast cancer focuses on the active role that tumor infiltrating lymphocytes play in tumor progression and patient outcome. Toward this goal, we developed a methodology for the rapid isolation of intact lymphoid cells from normal and abnormal tissues in an effort to evaluate them proximate to their native state. Homogenates prepared using a mechanical dissociator show both increased viability and cell recovery while preserving surface receptor expression compared to enzyme-digested tissues. Furthermore, enzymatic digestion of the remaining insoluble material did not recover additional CD45+ cells indicating that quantitative and qualitative measurements in the primary homogenate likely genuinely reflect infiltrating subpopulations in the tissue fragment. The lymphoid cells in these homogenates can be easily characterized using immunological (phenotype, proliferation, etc.) or molecular (DNA, RNA and/or protein) approaches. CD45+ cells can also be used for subpopulation purification, in vitro expansion or cryopreservation. An additional benefit of this approach is that the primary tissue supernatant from the homogenates can be used to characterize and compare cytokines, chemokines, immunoglobulins and antigens present in normal and malignant tissues. This protocol functions extremely well for human breast tissues and should be applicable to a wide variety of normal and abnormal tissues.

This protocol describes the rapid non-enzymatic dissociation of fresh human tissue fragments for qualitative and quantitative assessment of CD45+ cells (lymphocytes/leukocytes) present in various normal and malignant human tissues. Additionally, the supernatant obtained from the primary tissue homogenate can be collected and stored for further analysis or experimentation.

The ability of malignant cells to evade the immune system, characterized by tumor escape from both innate and adaptive immune responses, is now accepted ...

Qualitative and Quantitative Assays for Detection and Characterization of Protein Antimicrobials

Initial evaluations of large microbial libraries for potential producers of novel antimicrobial proteins require both qualitative and quantitative methods to screen for target enzymes prior to investing greater research effort and resources. The goal of this protocol is to demonstrate two complementary assays for conducting these initial evaluations. The microslide diffusion assay provides an initial or simple detection screen to enable the qualitative and rapid assessment of proteolytic activity against an array of both viable and heat-killed bacterial target substrates. As a counterpart, the increased sensitivity and reproducibility of the dye-release assay provides a quantitative platform for evaluating and comparing environmental influences affecting the hydrolytic activity of protein antimicrobials. The ability to label specific heat-killed cell culture substrates with Remazol brilliant blue R dye expands this capability to tailor the dye-release assay to characterize enzymatic activity of interest.

Here, we present protocols to rapidly screen and characterize enzymes for antimicrobial activity. The microslide diffusion assay and the dye-release assay utilize target bacterial substrates for qualitative and quantitative enzymatic activity evaluation.

Initial evaluations of large microbial libraries for potential producers of novel antimicrobial proteins require both qualitative and quantitative methods ...

Assessing Retinal Microglial Phagocytic Function In Vivo Using a Flow Cytometry-based Assay

Microglia are the tissue resident macrophages of the central nervous system (CNS) and they perform a variety of functions that support CNS homeostasis, including phagocytosis of damaged synapses or cells, debris, and/or invading pathogens. Impaired phagocytic function has been implicated in the pathogenesis of diseases such as Alzheimer's and age-related macular degeneration, where amyloid-&#946; plaque and drusen accumulate, respectively. Despite its importance, microglial phagocytosis has been challenging to assess in vivo. Here, we describe a simple, yet robust, technique for precisely monitoring and quantifying the in vivo phagocytic potential of retinal microglia. Previous methods have relied on immunohistochemical staining and imaging techniques. Our method uses flow cytometry to measure microglial uptake of fluorescently labeled particles after intravitreal delivery to the eye in live rodents. This method replaces conventional practices that involve laborious tissue sectioning, immunostaining, and imaging, allowing for more precise quantification of microglia phagocytic function in just under six hours. This procedure can also be adapted to test how various compounds alter microglial phagocytosis in physiological settings. While this technique was developed in the eye, its use is not limited to vision research.

Assessing Retinal Microglial Phagocytic Function In Vivo Using a Flow Cytometry-based Assay

Microglial phagocytosis is critical for the maintenance of tissue homeostasis and inadequate phagocytic function has been implicated in pathology. However, assessing microglia function in vivo is technically challenging. We have developed a simple but robust technique for precisely monitoring and quantifying the phagocytic potential of microglia in a physiological setting.

Microglia are the tissue resident macrophages of the central nervous system (CNS) and they perform a variety of functions that support CNS homeostasis, ...

Analysis of Microglia and Monocyte-derived Macrophages from the Central Nervous System by Flow Cytometry

Numerous studies have demonstrated the role of immune cells, in particular macrophages, in central nervous system (CNS) pathologies. There are two main macrophage populations in the CNS: (i) the microglia, which are the resident macrophages of the CNS and are derived from yolk sac progenitors during embryogenesis, and (ii) the monocyte-derived macrophages (MDM), which can infiltrate the CNS during disease and are derived from bone marrow progenitors. The roles of each macrophage subpopulation differ depending on the pathology being studied. Furthermore, there is no consensus on the histological markers or the distinguishing criteria used for these macrophage subpopulations. However, the analysis of the expression profiles of the CD11b and CD45 markers by flow cytometry allows us to distinguish the microglia (CD11b+CD45med) from the MDM (CD11b+CD45high). In this protocol, we show that the density gradient centrifugation and the flow cytometry analysis can be used to characterize these CNS macrophage subpopulations, and to study several markers of interest expressed by these cells as we recently published. Thus, this technique can further our understanding of the role of macrophages in mouse models of neurological diseases and can also be used to evaluate drug effects on these cells.

This protocol provides an analysis of the macrophage subpopulations in the adult mouse central nervous system by flow cytometry and is helpful for the study of multiple markers expressed by these cells.

Numerous studies have demonstrated the role of immune cells, in particular macrophages, in central nervous system (CNS) pathologies. There are two main ...

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 ...