This work was supported by National Institutes of Health grant: PO1 HL55798 (to K.L.) and American Heart Association Scientist Development Grant 0525532U (to E.G.).

1. Isolation of murine aortas 
Institutional IACUC committee approval of the procedure is required to work with mice. 
<ol>
<li>Prepare heparinzed PBS by adding 1000 units of heparin sodium to 50 ml PBS and inverting the tube to mix. Prepare an empty collection tube for the blood and a collection tube containing PBS for each aorta to be collected. Keep all tubes on ice.</li>
<li>Heparinize a syringe for drawing blood (0.1ml of 1000 U/ml Heparin sodium), prepare surgical tools (two pairs of curved forceps, a pair of dissecting scissors, and a pair of microshears), and a dissecting stage for the dissection.</li>
<li>Euthanize a mouse using carbon dioxide in an approved chamber following NIH and Institutional IACUC committee policies regarding rodent euthanasia. Check for effectiveness before transferring the mouse to the dissecting stage. </li>
<li>Briefly soak the mouse with 70% ethanol and fasten the mouse to the dissection stage. Draw blood from the mouse via cardiac puncture.</li>
<li>Open the abdominal and chest cavities. Using a 10 ml syringe with a 25 g needle, completely perfuse the vasculature with PBS containing 2% of heparin to completely remove blood from the vessels by cardiac puncture. Make sure that there is no blood within the aortic tissues. The perfusion should be performed slowly with little pressure to ensure that all plaques in the vessel wall remain intact.</li>
<li>Dissect and remove the visceral organs, genitourinary organs, diaphragm and spleen, leaving the kidneys, heart, and aorta intact.</li>
<li>Carefully dissect adipose tissues and para-aortic lymph nodes away from the aorta, leaving the aorta and adventitia intact. Collect the whole aorta including aortic arch, ascending, descending, thoracic, and abdominal portions. Place the isolated aorta in a collection tube with PBS. During the isolation procedure, try to keep the vessel moist. </li>
</ol>
2. Preparation of single cell suspensions
<ol>
	<li>Create a 1X Aorta Dissociation Enzyme stock Solution (ADES) (125 U/ml Collagenase type XI, 60 U/ml Hyaluronidase type 1-s, 60 U/ml DNase I, and 450 U/ml Collagenase type I, in 2.5 mls of PBS, modified from Galkina et al.7). All enzymes are from Sigma-Aldrich. Place the stock enzyme solution on ice.</li>
	<li> Remove the PBS from the collection tube containing the aorta. Add 2.5 ml of 1X ADES to each aorta. Cut the aortas into small pieces to facilitate enzymatic digestion or leave the whole aorta intact. Incubate the aortas with the 1X enzyme solution for 1 hour at 37&deg;C (slower shaking is optional). </li>
	<li>Following the 1 hour incubation, prepare single cell suspensions from the digested aorta by shearing the aortas apart and passing them through a 70 &mu;m cell strainer into 5ml polypropylene FACS tubes (BD Falcon). Pellet the cells by centrifugation (400xg, 5 minutes, 4&deg;C).</li>
	<li> Resuspend the cells in 1ml of FACS buffer (PBS supplemented with 1% BSA and 0.05% NaN3) and determine how many cells are present in the aortic cell suspension using trypan blue, a hemocytometer, and a light microscope. The total number of cells obtained after the digestion will depend on mouse age and diet or severity of atherosclerosis.</li>	
</ol>
3. Flow cytometry staining
<ol>
<li>Label an appropriate number of new FACS tubes. In general, flow cytometry experiments should have a non-stained control tube, set of single color tubes, appropriate fluorescence-minus-one controls (FMO)8, appropriate isotype controls, and a set of experimental tubes. Since there are limited numbers of aortic leukocytes that can be isolated, splenic leukocytes can be used to perform single control staining.</li>
<li>We use a standard flow cytometry protocol to stain aortic cell suspensions. Briefly, transfer an aliquot of 0.5-1x106 cells from an aortic cell suspension into a FACS tube(s). Add 1ml of FACS buffer and pellet the cells by centrifugation. Remove the supernatant from the pelleted cells by decanting.</li>
<li>Prepare Fc block solution and antibody cocktails for the experimental tubes, fluorescence-minus-one tubes, and isotype tubes. Add 100&mu;l of Fc block in FACS buffer (14.2&mu;g/ml, clone 2.4G2) to all of the tubes and finger flick or gently vortex to resuspend the cells. Incubate the samples for 15-20 minutes at room temperature.</li>
<li>Prepare antibody staining, isotype staining or FMO control staining cocktails. Determine the optimal concentration of antibodies in your preliminary titration experiments. In the presence of Fc block, add antibody cocktails (100ul/0.5-1.0x106 cells) to the sample tubes. Incubate for 20-30 min at 4&deg;C in the dark. </li>
<li>Add 1ml of FACS buffer to each tube, vortex to mix, and pellet the cells by centrifugation. Repeat the procedure one more time. </li>
<li>Decant the supernatant and resuspend the pelleted cells in 300&mu;l of 2% PFA. Run the samples on a flow cytometer.
<ul>
<li>Note: Typically anti-CD45 antibody (common leukocyte antigen marker) is added to all of the samples in order to gate CD45+ leukocytes later.7 In addition, CD45 FMO and isotype controls should be used initially to place the CD45+ leukocyte gate as CD45 expression can vary amongst leukocytes.</li>
<li>Note: To detect low density expressing surface antigens or low event antigens, use "bright fluorochromes" such as R-Phycoerythrin (PE) or Allophycocyanin (APC). In addition, rare events can be detected by pooling several matched aortas together; however if more than one million cells are used, the amount of antibodies used per test should be increased proportionally. </li>
<li>Note: To ensure that there are minimal blood contaminations in the isolated aortas and consequently in the aortic cell suspension, in some experiments additional staining was performed for TER-119,7 an antigen expressed by red blood cells (RBC). The number of RBC to white blood cells (WBC) in blood is roughly 10x106 cell/&mu;l to 8x103cell/&mu;l. Based on the expression of TER-119 in aortic samples, the percentage of blood-derived white blood cells in the sample can be calculated. Typically we have less than 0.02% of blood-derived WBC in aortic samples. (Fig.1) </li>
<li>Note: Since the enzyme treatment may affect the expression of surface antigens, the resistance to enzyme digestion should be determined for antigens of interest. Briefly, peripheral lymph node (pLN) or small pieces of spleen are incubated with or without enzyme cocktail for 1 hour at 37&deg;C. After 1 hour, the expression of antigens is determined by flow cytometry. The enzyme cocktail has no effect on multiple surface antigens (Fig.2)7. In some other cases, the enzyme treatment resulted in a significant loss of antigen expression. This problem can be circumvented by using alternative cell markers. </li>
<li>Note: Live/dead cell viability staining can be performed after step 4 if desired. Typically use Live/Dead Fixable Dead Cell Stain kit (Invitrogen, Molecular Probes). To stain the cells with Live/Dead dye, create a 1x Live/Dead solution by adding 1&mu;l of re-dissolved live/dead dye in 1ml of PBS and vortex to mix. Add 100-200 &mu;l of 1x Live/Dead dye to the live/dead single, cocktail, FMO and isotype control samples. Incubate the samples with the dye at room temperature for 10 minutes in the dark. The single control tube for Live/Dead should be incubated at 56&deg;C in the dark for 10 minutes to kill the cells by heat shock. Wash the cells with 1 ml of PBS and pellet the cells by centrifugation. Resume the protocol at step 5. </li>
<li>Note: Intracellular staining for intracellular antigens and cytokines is compatible with this protocol. Briefly, intracellular staining can be performed using cell fixation/permeabilization reagents from BD Pharmingen, eBioscience, or Caltag laboratories &#x2013; depending on the antigens of interest. To fix and permeabilize the cells for intracellular staining, perform the fixation/permeabilization step, following the manufacturer&#x2019;s instructions, after the extracellular staining step (section 3 step 5). Following the intracellular wash step, resume our protocol at step 6 (section 3 step 6).</li>
</ul>
</li>
</ol>
4. Flow cytometry analysis of isolated surrounding adventitia and vessel wall
As leukocytes can migrate to the aortic adventitia as well as atherosclerotic plaques within the aortic wall, to examine adventitial and aortic leukocytes by flow cytometry a protocol for isolating and performing flow cytometry on these two anatomical sites was developed.9 Briefly, before the whole aortas are digested with ADES (Section 2, step 2) the aortic adventitia is partially digested and removed from the rest of the vessel. Once the adventitia is removed and set aside, the rest of the aorta is digested with ADES to liberate leukocytes from the vessel wall.
<ul>
<li>Aortic Adventitia Isolation Protocol</li>
</ul>
<ol>
<li>Following the preparation of 1X ADES (section 2 step 1), prepare 2.5 ml/aorta of 1X Aortic adventitia digestion enzyme solution (AADES) (781.25 U Collagenase II and 14.0625 U Elastase in 2.5mls PBS (Worthington biochemical Corp., Lakewood, NJ)).9 Place the stock solution on ice until use.</li>
<li>Remove the PBS from the collection tube containing the aorta. Add 2.5 ml of 1X AADES to each aorta. Do not cut the aortas into small pieces. Incubate the aortas with the 1X adventitia enzyme solution for 10-20 minutes at 37&deg;C.</li>
<li>Following the 10-20 minute digestion, transfer the partially digested aorta out of the 1X AADES to a petri dish with fresh PBS. Very carefully, using two pairs of curved forceps, peel the adventitial layer away from the aorta as a single unit.
	<ul>
 <li>Note: Longer digestion times make it easier to remove the adventitia; however, if the adventitia is over-digested it will tear.</li>
 </ul>
</li>
<li>When the adventitia has been completely removed, transfer the adventitia and the aorta to separate FACS tubes. Add 1ml of PBS to the adventitia tube, and place the tube on ice. Add 2.5 mls of 1X aorta dissociation enzyme solution to the aorta tube and incubate the aorta for 40 minutes at 37&deg;C.
<ul><li>Note: To facilitate the enzymatic digestion, the aorta can be divided up into smaller pieces at this point. </li></ul>
</li>
<li>Following the 40 minute digestion, place the aorta tube on ice and prepare single cell suspensions from the aorta and adventitia by shearing the aortas and adventitia apart and passing them through a 70 &mu;m cell strainer into 5ml polypropylene FACS tubes (BD Falcon). Pellet the cells by centrifugation (400xg, 5 minutes, 4&deg;C). Return to section 2 step 4 and resume the protocol. </li>
</ol>
5. Representative Results
Here we present a number of figures that demonstrates flow cytometry staining to analyze the immune composition of whole aortas, the aortic vessel wall and the surrounding aortic adventitia. First, we demonstrate a representative FACS plot that shows a TER-119 staining on the whole blood and isolated aortic cell suspension (Fig. 1). TER-119 positive red blood cells accounted for 18% of the cells in the aortic cell suspension, which indicates that only 0.014% of the cells isolated from aortic cell suspensions are blood-derived. This is an important control experiment that clearly demonstrates that digested vessels, but not circulating peripheral blood is the source for most leukocytes analyzed in the aortic cell suspension. In addition, to validate the method, we assessed the effects of the aorta dissociation enzyme cocktail on splenocyte surface antigens (Fig. 2). The enzyme cocktail has no effects on the expression of several antigens, including, CD45, CD19, CD3, TCR&alpha;&beta;, TCR&gamma;&delta; and several other surface antigens7. 
CD45 staining in conjunction with Live/Dead viability dye and appropriate isotype controls are used to detect and sub-gate major populations of interest and to exclude cellular debris during analysis. In the Fig.3, live aortic CD45+ leukocytes were gated to determine the percentage of IFN&Upsilon;+ T cells within the pooled aortas of two young Apoe-/- mice. To emphasize the versatility of this method, using the gating scheme presented in Fig.3, we present in Fig.4 representative intracellular staining for CD68+CD11b+ macrophages and IFN&gamma;+TCR&alpha;&beta;+ cells from two Apoe-/- mice fed western diet for 12 weeks. As expected, in western diet fed Apoe-/- aortas, the majority of the leukocytes within the aorta are macrophages (40% CD68+CD11b+) or other myeloid cells (CD11b+CD68low and CD11b+CD68-). In addition, Th1 cells comprise a major portion of the aortic infiltrating TCR&alpha;&beta; T cells (Fig.4). As the overall percentage of aortic leukocytes and leukocyte subsets varies depending on the age of the mouse and severity of atherosclerosis, the optimal gating strategy for major populations of interest should be empirically determined 
To demonstrate the feasibility of isolating aortic adventitia for flow cytometry staining, we present representative CD45+ leukocyte staining for aortic and adventitial cell suspensions (Fig 5). Briefly, three aged Apoe-/- mouse aortas were digested and pooled together as described above (Sections 2 and 4). Infiltrating CD45+ leukocytes were detected in both the adventitia (18% of the cell suspension) and the remaining aorta (11% of the cell suspension). 
<img src="/files/ftp_upload/2848/2848fig1.jpg" alt="Figure 1"/> 
Figure 1.TER-119 staining in digested aortic cell suspension. Aorta was perfused by cardiac puncture with PBS containing 2% heparin. Then the aorta was digested with the enzyme cocktail for 1 hr at 37&deg;C. Aortic cells suspension and blood sample (as a positive control) were stained with anti-TER-119-PE Abs and analyzed by flow cytometry. TER-119-positive red blood cells account for 18% of all cells in the aortic cell suspension indicating that only 0.014% of all leukocytes isolated from aortas are likely to be blood-derived leukocytes. 
<img src="/files/ftp_upload/2848/2848fig2.jpg" alt="Figure 2"/> 
Figure 2.Enzyme-cocktail treatment has no effects on CD45 (top) or CD19 (bottom) expression on lymphocytes. Cell suspensions from untreated (A, B) and treated with enzyme-cocktail (C, D) LN were obtained, and stained with APC-Cy7-conjugated anti-CD45 and APC-conjugated anti-CD19 mAbs. (A, C) The numbers represent the percentages of CD45+ leukocytes in R1 gate. (B, D) The numbers represent the percentages of CD45+/CD19+ lymphocytes. Profiles are gated on CD45+ leukocytes.
<img src="/files/ftp_upload/2848/2848fig3.jpg" alt="Figure 3"/> 
Figure 3. Gating strategy for the analysis of aortic leukocytes. Two aortas were isolated and pooled from atherosclerotic prone Apoe-/- mice and aortic cell suspensions were prepared as described above. The cell suspensions were stained for CD45 (PerCP), TCR&alpha;&beta; (FITC), IFN&gamma; (eFluor 450), and Live/Dead Aqua, and analyzed using a Cytek DXP 8 Color upgraded BD FACS Calibur. Briefly, CD45+ Leukocytes were gated (B) and further analyzed (C-F). Dead cells were removed from the analysis based on Live/Dead Aqua staining (D) and FSC plots (E). Live aortic leukocytes were then examined for TCR&alpha;&beta; and IFN&gamma; expression (F). 
<img src="/files/ftp_upload/2848/2848fig4.jpg" alt="Figure 4"/> 
Figure 4. Intracellular staining for intracellular antigens and cytokines.
Cell suspensions were prepared from a whole Apoe-/- aorta and spleen as described. Aortic (A) and splenic (B, C) cell suspensions were stained for CD45, CD11b, and CD68 or an isotype control using BD Cytofix/Cytoperm&trade; Kit (BD Biosciences). Cells were gated on CD45+ leukocytes and debris was excluded based on the forward and side scatter profiles. For intracellular IFN&gamma; staining, aortic and splenic cell suspensions were cultured for five hours in RPMI 1640 supplemented with Golgi stop, PMA, and Ionomycin C, as earlier described.9 Stimulated single aortic (D) and splenic (E, F) cell suspensions were subsequently stained with CD45(PerCP), TCR&alpha;&beta; (FITC), CD3 (APC-Cy7), Live Dead Aqua, and IFN&gamma; (eFluor 450; E) or an isotype (IgG1-eF450, F). T cells (D-F) were gated from live CD45+CD3+ leukocytes (CD45+Live/Dead Aqua-) and examined for TCR&alpha;&beta; and IFN&gamma;.
<img src="/files/ftp_upload/2848/2848fig5.jpg" alt="Figure 5"/> 
Figure 5. Representative image of isolated murine aortic adventitia and aortic vessel wall. Representative flow cytometry counter plot demonstrates the presence of CD45+ T cells in the adventitia and aortic wall of aged Apoe-/- mice. SSC- side scatter. To eliminate autofluorescence from debris and necrotic tissues, plots were gated for FSC&gt;750. To avoid additional autofluorescence from doublets the gates were also set up as FSC&lt;3500, SSC&lt;3500. The percentages indicate CD45+ cells in the gates.

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	<li>Lusis, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atherosclerosis.">Atherosclerosis.</a> Nature. 407, 233-2341 (2000).</li><li>Galkina, E., Ley, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immune+and+inflammatory+mechanisms+of+atherosclerosis.">Immune and inflammatory mechanisms of atherosclerosis.</a> Annu Rev Immunol. 27, 165-197 (2009).</li><li>Hansson, G. K., Libby, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+immune+response+in+atherosclerosis:+a+double-edged+sword.">The immune response in atherosclerosis: a double-edged sword.</a> Nat Rev Immunol. 6, 508-519 (2006).</li><li>Galkina, E., Ley, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Leukocyte+influx+in+atherosclerosis.">Leukocyte influx in atherosclerosis.</a> Curr Drug Targets. 812, 1239-1248 (2007).</li><li>Bonanno, E., Mauriello, A., Partenzi, A., Anemona, L., Spagnoli, L. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flow+cytometry+analysis+of+atherosclerotic+plaque+cells+from+human+carotids:+a+validation+study.">Flow cytometry analysis of atherosclerotic plaque cells from human carotids: a validation study.</a> Cytometry. 39, 158-165 (2000).</li><li>Liu-Wu, Y., Svenningsson, A., Stemme, S., Holm, J., Wiklund, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+and+analysis+of+macrophage-derived+foam+cells+from+human+atherosclerotic+lesions+by+using+a+"mock"+FL3+channel+in+flow+cytometry.">Identification and analysis of macrophage-derived foam cells from human atherosclerotic lesions by using a "mock" FL3 channel in flow cytometry.</a> Cytometry. 29, 155-164 (1997).</li><li>Galkina, E., Kadl, A., Sanders, J., Varughese, D., Sarembock, I. J., Ley, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lymphocyte+recruitment+into+the+aortic+wall+before+and+during+development+of+atherosclerosis+is+partially+L-selectin+dependent.">Lymphocyte recruitment into the aortic wall before and during development of atherosclerosis is partially L-selectin dependent.</a> J Exp Med. 203, 1273-1282 (2006).</li><li>Tung, J. W., Parks, D. R., Moore, W. A., Herzenberg, L. A., Herzenberg, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+approaches+to+fluorescence+compensation+and+visualization+of+FACS+data.">New approaches to fluorescence compensation and visualization of FACS data.</a> Clin Immunol. 110, 277-283 (2004).</li><li>Smith, E., Prasad, K. M., Butcher, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blockade+of+interleukin-17A+results+in+reduced+atherosclerosis+in+apolipoprotein+E-deficient+mice.">Blockade of interleukin-17A results in reduced atherosclerosis in apolipoprotein E-deficient mice.</a> Circulation. 121, 1746-1755 (2010).</li><li>Eid, R. E., Rao, D. A., Zhou, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interleukin-17+and+interferon-gamma+are+produced+concomitantly+by+human+coronary+artery-infiltrating+T+cells+and+act+synergistically+on+vascular+smooth+muscle+cells.">Interleukin-17 and interferon-gamma are produced concomitantly by human coronary artery-infiltrating T cells and act synergistically on vascular smooth muscle cells.</a> Circulation. 119, 1424-1432 (2009).</li></ol>

No conflicts of interest declared.

Here, we present a flow-cytometry-based method for the investigation of the immune cell composition of murine aortas. The major advantage of this method is the ability to analyze aortic immune cells at a single cell levels and to characterize the activation status of aortic leukocytes. This method is not restricted to murine aortas and we (unpublished data) and others10 used this approach to analyze human specimens such as internal mammary artery, aortic valves and coronary arteries. One limitation of this met...

<table border="1">
	<tbody>
		<tr>
			<td>Material</td>
			<td>Catalog Number</td>
			<td>Company</td>
		</tr>
		<tr>
			<td>Collagenase XI</td>
			<td>C7657</td>
			<td>Sigma-Aldrich</td>
		</tr>
		<tr>
			<td>Hyaluronidase</td>
			<td>H3506</td>
			<td>Sigma-Aldrich</td>
		</tr>
		<tr>
			<td>DNase I, type 2</td>
			<td>D4527</td>
			<td>Sigma-Aldrich</td>
		</tr>
		<tr>
			<td>Collagenase I</td>
			<td>C0130</td>
			<td>Sigma-Aldrich</td>
		</tr>
		<tr>
			<td>Collagenase II</td>
			<td>LS004174</td>
			<td>Worthington Biochemical Corporation</td>
		</tr>
		<tr>
			<td>Elastase</td>
			<td>LS002292</td>
			<td>Worthington Biochemical Corporation</td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Flow Cytometry Analysis of Immune Cells Within Murine Aortas at JoVE.com

Flow Cytometry Analysis of Immune Cells Within Murine Aortas

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 ...

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Immunology and Infection

34.3K Views.  Eastern Virginia Medical School. 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.

Video: Flow Cytometry Analysis of Immune Cells Within Murine Aortas

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 ...

Flow Cytometric Isolation of Primary Murine Type II Alveolar Epithelial Cells for Functional and Molecular Studies

Throughout the last years, the contribution of alveolar type II epithelial cells (AECII) to various aspects of immune regulation in the lung has been increasingly recognized. AECII have been shown to participate in cytokine production in inflamed airways and to even act as antigen-presenting cells in both infection and T-cell mediated autoimmunity 1-8. Therefore, they are especially interesting also in clinical contexts such as airway hyper-reactivity to foreign and self-antigens as well as infections that directly or indirectly target AECII. However, our understanding of the detailed immunologic functions served by alveolar type II epithelial cells in the healthy lung as well as in inflammation remains fragmentary. Many studies regarding AECII function are performed using mouse or human alveolar epithelial cell lines 9-12. Working with cell lines certainly offers a range of benefits, such as the availability of large numbers of cells for extensive analyses. However, we believe the use of primary murine AECII allows a better understanding of the role of this cell type in complex processes like infection or autoimmune inflammation. Primary murine AECII can be isolated directly from animals suffering from such respiratory conditions, meaning they have been subject to all additional extrinsic factors playing a role in the analyzed setting. As an example, viable AECII can be isolated from mice intranasally infected with influenza A virus, which primarily targets these cells for replication 13. Importantly, through ex vivo infection of AECII isolated from healthy mice, studies of the cellular responses mounted upon infection can be further extended.
Our protocol for the isolation of primary murine AECII is based on enzymatic digestion of the mouse lung followed by labeling of the resulting cell suspension with antibodies specific for CD11c, CD11b, F4/80, CD19, CD45 and CD16/CD32. Granular AECII are then identified as the unlabeled and sideward scatter high (SSChigh) cell population and are separated by fluorescence activated cell sorting 3.
In comparison to alternative methods of isolating primary epithelial cells from mouse lungs, our protocol for flow cytometric isolation of AECII by negative selection yields untouched, highly viable and pure AECII in relatively short time. Additionally, and in contrast to conventional methods of isolation by panning and depletion of lymphocytes via binding of antibody-coupled magnetic beads 14, 15, flow cytometric cell-sorting allows discrimination by means of cell size and granularity. Given that instrumentation for flow cytometric cell sorting is available, the described procedure can be applied at relatively low costs. Next to standard antibodies and enzymes for lung disintegration, no additional reagents such as magnetic beads are required. The isolated cells are suitable for a wide range of functional and molecular studies, which include in vitro culture and T-cell stimulation assays as well as transcriptome, proteome or secretome analyses 3, 4.

We describe the rapid isolation of primary murine type II alveolar epithelial cells (AECII) by flow cytometric negative selection. These AECII show high viability and purity and are suitable for a wide range of functional and molecular studies regarding their role in respiratory conditions such as autoimmune or infectious diseases.

Throughout  the last years, the contribution of alveolar type II epithelial cells (AECII)  to various aspects of immune regulation in the lung has been ...

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 ...

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.

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 ...

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 ...

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 ...

Isolation of Mouse Kidney-Resident CD8+ T cells for Flow Cytometry Analysis

Tissue-resident memory T cell (TRM) is a rapidly expanding field of immunology research. Isolating T cells from various non-lymphoid tissues is one of the key steps to investigate TRMs. There are slight variations in lymphocyte isolation protocols for different organs. Kidney is an essential non-lymphoid organ with numerous immune cell infiltration especially after pathogen exposure or autoimmune activation. In recent years, multiple labs including our own have started characterizing kidney resident CD8+ T cells in various physiological and pathological settings in both mouse and human. Due to the abundance of T lymphocytes, kidney represents an attractive model organ to study TRMs in non-mucosal or non-barrier tissues. Here, we will describe a protocol commonly used in TRM-focused labs to isolate CD8+ T cells from mouse kidneys following systemic viral infection. Briefly, using an acute lymphocytic choriomeningitis virus (LCMV) infection model in C57BL/6 mice, we demonstrate intravascular CD8+ T cell labeling, enzymatic digestion, and density gradient centrifugation to isolate and enrich lymphocytes from mouse kidneys to make samples ready for the subsequent flow cytometry analysis.

Isolation of Mouse Kidney-Resident CD8+ T cells for Flow Cytometry Analysis

Following viral infection, kidney harbors a relatively large number of CD8+ T cells and offers an opportunity to study non-mucosal TRM cells. Here, we describe a protocol to isolate mouse kidney lymphocytes for flow cytometry analysis.

Tissue-resident memory T cell (TRM) is a rapidly expanding field of immunology research. Isolating T cells from various non-lymphoid tissues is one of the ...

Analysis of SAMHD1 Restriction by Flow Cytometry in Human Myeloid U937 Cells

Sterile &#945;-motif/histidine-aspartate domain-containing protein 1 (SAMHD1) inhibits replication of HIV-1 in quiescent myeloid cells. U937 cells are widely used as a convenient cell system for analyzing SAMHD1 activity due to a low level of SAMHD1 RNA expression, leading to undetectable endogenous protein expression. Based on similar assays developed in the Stoye laboratory to characterize other retroviral restriction factors, the Bishop lab developed a two-color restriction assay to analyze SAMHD1 in U937 cells. Murine Leukaemia Virus-like particles expressing SAMHD1, alongside YFP expressed from an IRES, are used to transduce U937 cells. Cells are then treated with phorbol myristate acetate to induce differentiation to a quiescent phenotype. Following differentiation, cells are infected with HIV-1 virus-like particles expressing a fluorescent reporter. After 48 h, cells are harvested and analyzed by flow cytometry. The proportion of HIV-infected cells in the SAMHD1-expressing population is compared to that in internal control cells lacking SAMHD1. This comparison reveals a restriction ratio. SAMHD1 expression leads to a five-fold reduction in HIV infection, corresponding to a restriction ratio of 0.2. Our recent substitution of RFP for the original GFP as the reporter gene for HIV infection has facilitated flow cytometry analysis.
This assay has been successfully used to characterize the effect of amino acid substitutions on SAMHD1 restriction by transducing with viruses encoding altered SAMHD1 proteins, derived from site-directed mutagenesis of the expression vector. For example, the catalytic site substitutions HD206-7AA show a restriction phenotype of 1, indicating a loss of restriction activity. Equally, the susceptibility of different tester viruses can be determined. The assay can be further adapted to incorporate the effect of differentiation status, metabolic status, and SAMHD1 modifiers to better understand the relationship between SAMHD1, cell metabolic state, and viral restriction.

Described here is an established method to determine the extent of HIV-1 restriction by the cellular inhibitory protein SAMHD1. Human myeloid lineage U937 cells are transduced with a SAMHD1 expression vector co-expressing YFP, differentiated and then challenged with HIV-RFP. The level of restriction is determined by flow cytometry analysis.

Sterile &#945;-motif/histidine-aspartate domain-containing protein 1 (SAMHD1) inhibits replication of HIV-1 in quiescent myeloid cells. U937 cells are ...