This work was supported by the Canadian Institutes of Health Research (148808 and 148792), SE was supported by a Heart and Stroke Foundation, personnel award from the Ontario Provincial Office, March of Dimes, Ted Rogers Centre for Heart Research and the Peter Munk Cardiac Centre. XCC holds a CIHR Banting Fellowship. LA holds a Heart &amp; Stroke/Richard Lewar Studentship Award.

Ethics statement: This protocol has been reviewed and approved by the Animal Care Committee at the University Health Network (Toronto, Canada) and is in compliance with the Canadian Council on Animal Care.
1. Buffer Preparation
<ol>
	<li>Prepare HBB buffer (Hank&#39;s Balanced Salt Solution (HBSS), 2% heat inactivated bovine serum, 0.2% bovine serum albumin). Add 10 mL of heat inactivated bovine serum and 1 g of bovine serum albumin to 500 mL of HBSS. Filter sterilize through a 0.2 &#181;m filter and store at 4 &#176;C.</li>
	<li>Prepare FACS buffer (Phosphate buffered saline (PBS), 2% heat inactivated bovine serum, 1 mM Ethylenediaminetetraacetic acid (EDTA, pH 8.0)). Add 10 mL heat inactivated bovine serum and 1 mL of 0.5 M EDTA to 500 mL of PBS. Filter sterilize through a 0.2 &#181;m filter and store at 4 &#176;C. 
	NOTE: Solutions can be stored at 4 &#176;C for up to 1 month.</li>
</ol>
2. Labelling of Circulating Leukocytes
<ol>
	<li>Aseptically prepare anti-mouse CD45 injection with 1 &#181;g of antibody diluted in sterile phosphate buffered saline (PBS) for a final injection volume of 200 &#181;L/mouse. Load volume into a 28.5G insulin syringe and keep away from light. 
	NOTE: Use two different fluorochrome-labelled anti-CD45 antibodies in order to differentiate the intravascular (stained through i.v. administration) from the intracardiac staining (stained after digestion; see step 4).</li>
	<li>Pre-warm mice (8&#8211;12 weeks old) by exposing the tail to a red heat lamp for 5 min. 
	CAUTION: Do not overheat the animal. Keep the lamp at a safe distance (&#62;30 cm).
	<ol>
		<li>Restrain the animal in the sternal position using an appropriate device. Expose and stabilize the tail with the non-dominant hand by immobilizing the base of the tail with the index and middle finger and the mid-distal region of the tail with the thumb and ring finger. 
		CAUTION: Do not apply too much force at the tip of the tail as the skin can be pulled off. Insert the needle with the bevel facing up into the vein. Keep the needle parallel to the vein at all times as it is easy to puncture the vein. 
		NOTE: Blood entering the syringe may occur and is a sign of a good injection. Bending the needle to a 90&#176; angle can facilitate i.v. administration. Confirm with your safety office as bending needles may represent a hazard.</li>
		<li>Inject 200 &#181;L intravenously into the tail vein. The fluid should flow easily and temporarily clear the vein. 
		CAUTION: Do not force fluid in if resistance is met. This is sign of misplacement of the needle.</li>
		<li>Allow antibody to circulate for 5 min before humanely euthanizing mouse.</li>
	</ol>
	</li>
</ol>
3. Heart Isolating and Digestion
<ol>
	<li>Euthanize mice using CO2 or by other institutionally accepted euthanasia procedure.
	<ol>
		<li>For CO2 euthanasia, place the mouse in the CO2 chamber and set the fill rate to 20% of the chamber volume per minute (i.e., if the chamber is 10 L, fill at 2 L/min). Run for 2 min and monitor mouse until breathing has ceased.</li>
		<li>Shut off the CO2 and leave the mouse for an additional 1-2 min. Ensure breathing has ceased before opening the chamber and perform a toe pinch to confirm the mouse is dead before proceeding to process the mouse.</li>
	</ol>
	</li>
	<li>Spray the mouse with 70% ethanol. Lift the skin at the abdomen and dissect the mouse by cutting along the ventral midline beginning at the level of the hips up to the sternum. Open the abdomen and move the liver to expose the diaphragm.</li>
	<li>Cut the diaphragm, then the ribs on both sides of the midline, being careful to avoid puncturing the lungs and heart.
	<ol>
		<li>Expose the heart by peeling back the ribs, then cut first the left atria followed by the right atria. 
		&#8203;CAUTION: Use caution when cutting the atria, ensuring no arteries or veins are injured, as this can decrease the efficiency of flushing the blood contained in the organs.</li>
		<li>Perfuse the heart with 15&#8211;20 mL of cold PBS using a 60 mL syringe fitted with a 21 G needle and cold PBS. Gently grasp the base of the heart with forceps and insert the needle into the left ventricle near the apex. Repeat the procedure in the right ventricle. A proper injection results in white lungs and pale liver. 
		&#8203;NOTE: Perfusion should be performed within 5 minutes after euthanasia in order to avoid blood coagulation, which can contaminate the organs of interest with circulating cells.</li>
	</ol>
	</li>
	<li>Hold the base of the heart with forceps and gently pull the heart up, then detach the heart by cutting the major pulmonary arteries and veins, aorta and the pericardial sack.
	<ol>
		<li>Clean the heart by gently holding the heart between the thumb and index fingers in a clean lint-free paper towel and pull off the atria and remnants of major arteries and veins and any other contaminants with forceps. 
		&#8203;NOTE: Ensure the entirety of the atria is removed and no other tissues, such as the lung or blood vessels, are present as these tissues have their own resident immune cells. The lung in particular has a large immune cell population that can mask the smaller populations of those within the myocardium.</li>
	</ol>
	</li>
	<li>Place the heart in a 5 cm plastic dish with 50 &#181;L of cold PBS.
	<ol>
		<li>Mince the heart using curved dissecting scissors until there are no visible pieces and it has a smooth consistency. 
		&#8203;NOTE: Try to keep the heart in a contained area of the dish during chopping to minimize tissue loss.</li>
		<li>Transfer thetrituratedheart tissue using curved forceps into 2 mL microcentrifuge tube with 800 &#181;L cold Dulbecco&#39;s Modified Eagle Medium (DMEM).</li>
	</ol>
	</li>
	<li>Add 450 U/mL ofcollagenase I, 60 U/mL of hyaluronidase type I-S, and 60 U/mL of DNase-I to each sample.
	<ol>
		<li>Incubate samples at 37 &#176;C on a rocking shaker at 50 rpm for 45&#8211;60 min.</li>
	</ol>
	</li>
	<li>After digestion, briefly vortex samples (20 s) and immediately place on ice to keep samples at 4 &#176;C.
	<ol>
		<li>Using a 1 mL pipettor, pipette samples up and down until tissue samples are homogeneous and do not clog the pipette tip. 
		&#8203;NOTE: Defining a specific number of times to pipette may increase reproducibility. If the hearts are not optimally digested, pieces of myocardium may obstruct the pipette. Gently grinding the pipette tip against the microcentrifuge tube will help homogenize bigger pieces of tissue.</li>
	</ol>
	</li>
	<li>Pre-wet a 40 &#181;m cell strainer placed on a 50 mL conical tube with 1 mL of cold HBB.
	<ol>
		<li>Add homogenized sample to the filter and wash through by slowly pouring 12&#8211;14 mL of cold HBB.</li>
		<li>Transfer filtered samples to a labelled 15 mL conical tube and keep at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Centrifuge samples at 400 x g for 5 min at 4 &#176;C. Aspirate off supernatant.
	<ol>
		<li>Lyse red blood cells by adding 1 mL of Ammonium-Chloride-Potassium (ACK) lysis buffer to each sample. Resuspend samples by vortexing and incubate at room temperature for 5 min.</li>
		<li>Once lysis has completed, add 5&#8211;8 mL of cold Hank&#39;s Balanced Salt Solution (HBSS) to stop hemolysis.</li>
	</ol>
	</li>
	<li>Centrifuge samples at 400 x gfor 5 min at 4 &#176;C. Aspirate off supernatant.</li>
	<li>Add 1 mL of FACS buffer to each sample and transfer to a labelled 1.5 mL microcentrifuge tube. 
	NOTE: The protocol can be paused here for up to 2 h with samples stored at 4 &#176;C.</li>
</ol>
4. Flow Cytometry Staining and Analysis
<ol>
	<li>Centrifuge samples at 400 x g for 5 min at 4 &#176;C.
	<ol>
		<li>To reduce non-specific antibody binding, dilute 1:100 anti-CD16/CD32 and Monocyte Blocker (1:20) (see Table of Materials) accounting for 50 &#181;L FACS per sample. Incubate at room temperature for 15 min.</li>
		<li>Prepare the antibody dilution (1:250 for each antibody) accounting for 50 &#181;L of FACS per sample. 
		NOTE: A viability dye can be included in the antibody cocktail in order to exclude dead cells. Alternatively, small size exclusion by flow analysis can be used (Figure 1C and D).</li>
		<li>Without washing, add 50 &#181;L of antibody master mix to each sample and mix.</li>
		<li>Incubate samples at 4 &#176;C for 30 min in the dark.</li>
	</ol>
	</li>
	<li>Add 600 &#181;L FACS to each sample to wash and centrifuge samples at 400 x g for 5 min at 4 &#176;C.</li>
	<li>Aspirate off supernatant and resuspend pellet in 300 &#181;L FACS. Keep samples at 4 &#176;C in the dark. 
	NOTE: Some viability dyes, such as DAPI, require staining within 10 min prior to running flow cytometry.</li>
	<li>Analyze by flow cytometry (see Figures 1 and 2) within the next 3 h. If a longer time is required, short term fixation using a low concentration paraformaldehyde (0.5&#8211;1% in PBS) is recommended to preserve staining. 
	NOTE: It is highly recommended to filter samples through a 40 &#181;m cell strainer prior to initiate flow cytometry analysis in order to prevent obstruction of the fluidics of the flow cytometer.</li>
</ol>

<ol>
	<li>Marboe, C. C., Fenoglio, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pathology+and+natural+history+of+human+myocarditis.">Pathology and natural history of human myocarditis.</a> Pathology and Immunopathology Research. 7 (4), 226-239 (1988).</li><li>Epelman, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Embryonic+and+adult-derived+resident+cardiac+macrophages+are+maintained+through+distinct+mechanisms+at+steady+state+and+during+inflammation.">Embryonic and adult-derived resident cardiac macrophages are maintained through distinct mechanisms at steady state and during inflammation.</a> Immunity. 40 (1), 91-104 (2014).</li><li>Pinto, A. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+abundant+tissue+macrophage+population+in+the+adult+murine+heart+with+a+distinct+alternatively-activated+macrophage+profile.">An abundant tissue macrophage population in the adult murine heart with a distinct alternatively-activated macrophage profile.</a> PLoS One. 7 (5), 36814 (2012).</li><li>Lavine, K. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+macrophage+lineages+contribute+to+disparate+patterns+of+cardiac+recovery+and+remodeling+in+the+neonatal+and+adult+heart.">Distinct macrophage lineages contribute to disparate patterns of cardiac recovery and remodeling in the neonatal and adult heart.</a> Proceedings of the National Academy of Sciences of the United States of America. 111 (45), 16029-16034 (2014).</li><li>Fujiu, K., Wang, J., Nagai, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardioprotective+function+of+cardiac+macrophages.">Cardioprotective function of cardiac macrophages.</a> Cardiovascular Research. 102 (2), 232-239 (2014).</li><li>Hulsmans, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Macrophages+Facilitate+Electrical+Conduction+in+the+Heart.">Macrophages Facilitate Electrical Conduction in the Heart.</a> Cell. 169 (3), 510-522 (2017).</li><li>Nahrendorf, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+healing+myocardium+sequentially+mobilizes+two+monocyte+subsets+with+divergent+and+complementary+functions.">The healing myocardium sequentially mobilizes two monocyte subsets with divergent and complementary functions.</a> Journal of Experimental Medicine. 204 (12), 3037-3047 (2007).</li><li>Clemente-Casares, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+CD103(+)+Conventional+Dendritic+Cell+Surveillance+System+Prevents+Development+of+Overt+Heart+Failure+during+Subclinical+Viral+Myocarditis.">A CD103(+) Conventional Dendritic Cell Surveillance System Prevents Development of Overt Heart Failure during Subclinical Viral Myocarditis.</a> Immunity. 47 (5), 974-989 (2017).</li><li>Van der Borght, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myocardial+Infarction+Primes+Autoreactive+T+Cells+through+Activation+of+Dendritic+Cells.">Myocardial Infarction Primes Autoreactive T Cells through Activation of Dendritic Cells.</a> Cell Reports. 18 (12), 3005-3017 (2017).</li><li>Galkina, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preferential+migration+of+effector+CD8++T+cells+into+the+interstitium+of+the+normal+lung.">Preferential migration of effector CD8+ T cells into the interstitium of the normal lung.</a> Journal of Clinical Investigation. 115 (12), 3473-3483 (2005).</li><li>Edelson, B. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peripheral+CD103++dendritic+cells+form+a+unified+subset+developmentally+related+to+CD8alpha++conventional+dendritic+cells.">Peripheral CD103+ dendritic cells form a unified subset developmentally related to CD8alpha+ conventional dendritic cells.</a> Journal of Experimental Medicine. 207 (4), 823-836 (2010).</li></ol>

The authors have nothing to disclose.

Myocardial inflammation, or myocarditis, is a feature of most cardiovascular diseases. However, the myocardium is not devoid of its own immune components in non-disease states. During steady state, many immune cells reside in the myocardium and play the essential roles of maintenance and protection. The characterization of these diverse populations of cells would not have been possible without methods such as the one presented in this protocol.
A combination of mechanical and enzymatic digestion of the myocardium permits the isolation of a single cell suspension that can be used in combination with other methods of analysis, such as flow cytometry or immunomagnetic bead isolation. Several considerations regarding the present method must be taken into account. First, all of our analyses refer to the populations within the myocardium; we exclude the atria from our analysis as these are specialized tissues with its own idiosyncrasies. Second, the amounts of enzymes used are expressed as activity units to help in the set-up of digestions using enzymes from other sources. However, empiric optimization of the final concentration of enzymes and the time of digestion is highly recommended, as limiting enzyme can significantly reduce the cell yield and too much digestion can drastically affect viability. Third, the optional step involving intravascular administration of labelled antibodies is especially recommended when studying uncommon populations in the normal myocardium (such as lymphocytes, monocytes or neutrophils)2 or under circumstances when numbers of populations in the blood may change (monocytosis, granulocytosis or lymphocytosis) which may give a false impression of changes within the myocardium that need to be confirmed using this method or, alternatively, microscopy. However, this step is of limited utility when analyzing macrophages or mature DCs, as these cells are rarely found in significant quantities in the blood (Figure 2D). It is important to optimize the quantity of antibody administered and the incubation time before sacrificing the mouse, as long term incubation can allow for some antibody diffusion into the heart, especially in an inflamed myocardium. Finally, it is very important to be meticulous in the isolation of the heart, eliminating remnants of major arteries, veins and or any other tissue, such as the lung. These contaminant tissues have highly elevated numbers of immune cells that can easily mask the myocardial indigenous populations.
It is very important to perform exclusion of dead cells, whether using viability dyes or through small size exclusion during flow cytometry analysis, as digestion (or tissue damage in disease models) can cause a relatively large amount of dead cells. We have compared the efficiency of dead cell exclusion by size and using a viability label (Figure 1B and D), demonstrating their near equivalence. Importantly, size exclusion cannot be used after fixation and permeabilization as this procedure significantly reduces cell size.
Although autofluorescence is rarely a problem to identify cardiac macrophages using flow cytometry, this represents a more important problem when trying to characterize the expression of some markers, especially with fluorochromes excited by high energy (shorter) light wavelengths (350&#8211;500 nm). In these experiments, it is highly recommended to use isotype controls, avoid the blue and violet lasers (fluorescein isothiocyanate, phycoerythrin, Pacific Blue/Brilliant Violet 421) and use fluorochromes with emissions above 600 nm, such as allophycocyanin.
The use of protocols as the one presented herein are becoming more important as an increasing amount of literature is identifying the role of intensity and type of inflammation in evaluating and characterizing the outcomes of cardiovascular disease. This method permits higher levels of cell characterization than histology. For instance, the use of lineage tracing allowed for the identification of ontogenically different subsets of macrophages with their programmed different functions2. Similarly, this method can be used to study T cell responses during myocardial inflammation and combined with ex vivo stimulation to study T cell polarization8.

Different forms of myocardial stress or injury, including ischemic (ischemia reperfusion or myocardial infarction) and non-ischemic (hypertension or myocarditis), promote the recruitment of inflammatory cells with reparative and protective, but also pathogenic properties. As early as 1891, Romberg first described the presence of cellular infiltrates in the myocardium of patients infected with typhus and scarlet fever1. However, the detailed study of cardiac immune cells required the development of more advanced immunophenotyping techniques. As a consequence, only recently have we started to understand that a diverse population of immune cells with essential maintenance roles during steady state resides within the myocardium.
Histology has been, and still is, the most common method to characterize cardiac immune cells during inflammation. However, while histology represents a valuable diagnostic tool, its use in the study of cardiac immune cells has important limitations. The immune cells residing in the myocardium represent a very small proportion of the total cells and are more or less evenly distributed in a proportionally immense space. Similarly, several forms of cardiac inflammation, such as viral myocarditis, show focal patterns of inflammation. This means that accurate analysis of the immune system of the heart often require higher degrees of histological sampling, increasing the cost to reduce bias. Moreover, histology provides a very limited amount of usable information in cell identification (e.g. number of parameters). With an ever increasing number of cell subsets that require the use of multiple expression markers (including surface markers, transcription factors or secreted molecules), flow cytometry has established itself as the most powerful tool for immunophenotyping, due to low-cost and high-throughput.
The use of immunophenotyping techniques is closely dependent on the development of efficient digestion protocols that allowed for single-cells analysis of large numbers of cells. The development of exhaustive protocols of cell extraction opened a new window of opportunities in the study of cardiac immunology. Through the combination of digestion, flow cytometry and transcriptomics, the major populations of macrophages and dendritic cells residing in the heart have been characterized2,3. The most abundant population of cardiac immune cells during steady state are CD64+MerTK+ macrophages, which can be further divided based on their expression of CCR2 or CD11c2. CCR2+ macrophages originate from adult bone marrow hematopoiesis, whereas the majority of CCR2- macrophages, which can express high or low levels of MHC-II, are mainly of prenatal origin. Macrophages perform key functions such as repair4 and removal of debris5 during injury or supporting electrical connectivity6 in steady state. During different forms of inflammation an important influx of Ly6Chi MHC-IIlo CD64int monocytes occur, which later differentiate into Ly6Chi CCR2+ macrophages2,7. A significantly smaller, but important, population of cells residing in the myocardium of healthy individuals is composed of dendritic cells8,9. The two major subsets of cardiac conventional DCs (cDCs) have been recently characterized: cDC1 (CD103+ DCs) and cDC2 (CD11b+ DCs). DCs play important roles in defense against infection8 but can also promote self-injury during inflammation, particularly during myocardial infarction9.
Here, we describe a simple method for isolation of viable cardiac immune cells from mouse myocardium. The method combines enzymatic and mechanical digestion with a cell-strainer filtration in order to obtain a single cell suspension that can be analyzed, or further purified, by flow cytometry sorting or magnetic bead enrichment. Accurate measurements of extravascular myocardial cells require cardiac perfusion in order to remove possible contaminants from the bloodstream of the cardiac microvasculature. In addition, we present an optional step of intravascular labeling of immune cells that can be used to further differentiate myocardial cells from intravascular contaminants, based on the Galkina et al. protocol10. Finally, we present a basic flow cytometry analysis to identify the major macrophage and cDC subpopulations.

<table border="0" cellpadding="0" cellspacing="0" style="width:815px;" width="814">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:340px;">Phosphate buffered saline</td>
			<td style="width:123px;">Wisent</td>
			<td style="width:185px;">311-010-CL</td>
			<td style="width:167px;">1x PBS</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">21Gx 1 1/2 (0.8mmx40mm) PrecisionGlide Needle</td>
			<td>BD</td>
			<td>305167</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Hank&#8217;s Balanced Salt Solution</td>
			<td>Wisent</td>
			<td>311-511-CL</td>
			<td>1x HBSS</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Bovine serum albumin</td>
			<td>Sigma-Aldrich</td>
			<td>A4503-50G</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Bovine serum</td>
			<td>Sigma</td>
			<td>B9433</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">0.5M Ethylenediaminetetraacetic acid</td>
			<td>BioShop</td>
			<td>EDT111</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Vacuum filter , 0.2 &#181;m Filtropur V50</td>
			<td>Sarstedt</td>
			<td>83.1823.001</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">28G 1/2 1 cc insulin syringe</td>
			<td>BD</td>
			<td>329424</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">60 mL syringe</td>
			<td>BD</td>
			<td>309653</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Dulbecco&#39;s Modified Eagle Medium</td>
			<td>Wisent</td>
			<td>319-005-CL</td>
			<td>1x DMEM with 4.5 g/L glucose and L-Glutamine and Sodium Pyruvate</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Collagenase I</td>
			<td>Sigma</td>
			<td>C0130</td>
			<td>from Clostridium histolyticum</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Hyaluronidase type I-S</td>
			<td>Sigma</td>
			<td>H3506</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">DNase-I</td>
			<td>Sigma</td>
			<td>D4513</td>
			<td>from bovine pancreas</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Cell strainer, 40 &#181;m Nylon</td>
			<td>Falcon</td>
			<td>352340</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Ammonium-Chloride- Potassium (ACK) lysis buffer</td>
			<td>Lonza</td>
			<td>10-546E</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Alexa Fluor 700 anti-mouse/human CD11b</td>
			<td>Biolegend</td>
			<td>101222</td>
			<td>1:250 dilution</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">APC/Cy7 anti-mouse Ly-6c</td>
			<td>Biolegend</td>
			<td>128025</td>
			<td>1:250 dilution</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">APC anti-mouse CD103</td>
			<td>Biolegend</td>
			<td>121414</td>
			<td>1:250 dilution</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Brilliant Violet 605 anti-mouse CD11c</td>
			<td>Biolegend</td>
			<td>117334</td>
			<td>1:250 dilution</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">PE anti-mouse Ly-6G</td>
			<td>Biolegend</td>
			<td>127607</td>
			<td>1:250 dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pacific Blue anti-mouse I-Ab</td>
			<td>Biolegend</td>
			<td>116422</td>
			<td>1:250 dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">FITC anti-mouse CD64 (FcgRI)</td>
			<td>Biolegend</td>
			<td>139315</td>
			<td>1:250 dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PE/Cy7 anti-mouse CD45</td>
			<td>Biolegend</td>
			<td>103113</td>
			<td>1:250 dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PerCP/Cy5.5 anti-mouse CD45</td>
			<td>Biolegend</td>
			<td>103132</td>
			<td>1:40 dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">TruStain fcX (anti-mouse CD16/32)</td>
			<td>Biolegend</td>
			<td>101320</td>
			<td>1:100 dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">True-Stain Monocyte Blocker</td>
			<td>Biolegend</td>
			<td>426101</td>
			<td>1:20 dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">FlowJo V10</td>
			<td>TreeStar Inc</td>
			<td></td>
			<td>https://www.flowjo.com/solutions/flowjo</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Mouse: Batf3-/-</td>
			<td>The Jackson Laboratory</td>
			<td>JAX: 013755</td>
			<td></td>
		</tr>
	</tbody>
</table>

isolation and identification of extravascular immune cells of the heart

The immune system is an essential component of a healthy heart. The myocardium is home to a rich population of different immune cell subsets with functional compartmentalization both during steady state and during different forms of inflammation. Until recently, the study of immune cells in the heart required the use of microscopy or poorly developed digestion protocols, which provided enough sensitivity during severe inflammation but were unable to confidently identify small &#8212; but key &#8212; populations of cells during steady state. Here, we discuss a simple method combining enzymatic (collagenase, hyaluronidase and DNAse) and mechanical digestion of murine hearts preceded by intravascular administration of fluorescently-labelled antibodies to differentiate small but unavoidable intravascular cell contaminants. This method generates a suspension of isolated viable cells that can be analyzed by flow cytometry for identification, phenotyping and quantification, or further purified with fluorescence-activated cell sorting or magnetic bead separation for transcriptional analysis or in vitro studies. We include an example of a step-by-step flow cytometric analysis to differentiate the key macrophage and dendritic cell populations of the heart. For a medium sized experiment (10 hearts) the completion of the procedure requires 2&#8211;3 h.

To date, no good method has been designed to isolate immune cells from other cardiac cell components, such as cardiomyocytes. Hence, analysis of the cardiac single cell suspension by flow cytometry requires a pre-gating with CD45 to identify the immune cell populations, followed by single cell and small size exclusion gating (Figure 1A). Alternatively, a viability staining can be performed to exclude dead cells. Small size exclusion and viability dye staining represent nearly equivalent methods to exclude dead cells (Figure 1B and 1D). It is highly recommended to proceed with the antibody staining soon after the digestion procedure and always keep the cell suspension at 4 &#176;C, as the viability of the cells may decay. This procedure typically yields about 80&#8211;90% of viable immune cells.
The myocardium houses a diverse colony of immune cells, many of which are still pending identification and characterization. Macrophages, identified as CD64+ cells, are the major immune population of the heart, representing 60&#8211;70% of all immune cells (Figure 1C, top). The digestion of an adult murine heart yields about 3&#8211;5 x 104 macrophages, depending of the heart&#39;s size. Cardiac macrophages can be further subdivided based on their expression of CD11c or CCR2 and their levels of MHC-II2.
The other cardiac immune subpopulation that has been well characterized are conventional dendritic cells. Conventional DCs are CD64- cells that express intermediate to high levels of MHC-II and high levels of CD11c (Figure 1C, bottom). These cells can be further divided based on their expression of either CD103 or CD11b8,9. DCs represent a much smaller cardiac subpopulation compared to macrophages. The digestion of an adult murine heart yields between 150 and 500 DCs of each subset.
Perfusion of the heart with cold PBS is essential when identifying immune cell populations within the myocardium. However, imperfect perfusion may increase the amounts of intravascular contaminants in the samples, leading to important bias that can be based on peripheral changes (blood) rather than changes in the cardiac infiltrates. In these cases, intravascular administration of fluorescent-labelled anti-CD45 antibodies, which labels all immune cells circulating through macro- and microvasculature (Figure 2A), can help differentiate myocardial cells from intravascular contaminants. Figure 2B shows an example of an imperfectly perfused heart in which 20% of immune cells are intravascular contaminants. Importantly, this level of contamination has little effect on macrophage and DC analysis, since these cells are rarely found in blood (Figure 2D).
The use of flow cytometry of a single cell suspension of myocardium allows for the study of the small subpopulations of the heart. For example, the deficiency of the transcription factor BATF3 has been shown to prevent the development of mature CD103+ DCs in several tissues11. In order to study if this is also true in the case of the myocardium, single cell suspensions of cardiac cells from 8&#8211;15-week-old BATF3-deficient or control littermates (in C57BL/6 background) were analyzed using flow cytometry. Due to the small population size and number of markers needed to differentiate the DC subsets, histology could not have provided enough power for analysis to reach a definitive answer. However, using flow cytometry analysis to compare the immune cells of the myocardium of wild-type versus a Batf3-/- mice, a lack of CD103+ DCs can be confirmed in the latter (Figure 3A and B). In addition, we observed that BATF3-deficiency did not significantly alter the composition of other cardiac populations (Figure 3C and D). These results exemplify the degree of sensitivity that can be achieved by using this combined method of enzymatic and mechanical digestion to achieve a single cell suspension.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58114/58114fig1.jpg" /> 
Figure 1: Gating strategy for cardiac macrophages and DCs. Isolated hearts from C57BL/6 mice were digested as described, and single cell suspension was analyzed by flow cytometry. (A) Gating of CD45+ live singlet cardiac cells. Use CD45 labeling to identify immune cells from the cardiac single cell homogenate, which includes very high levels of non-immune cells (i.e., cardiomyocytes, endothelial cells and fibroblasts). Exclude doublets by comparing SSC-H vs SSC-A followed by FSC-H vs FSC-A. Note: Any other comparison between SSC-A, SSC-H and SSC-W or FSC-A, FSC-H and FSC-W are similarly useful. Selection of live cells can be performed by excluding very small cells (SSC-H vs FSC-H gate). (B) Comparison of dead cell exclusion using size vs viability staining (DAPI). (C) Gating strategy for identification of cardiac macrophages (CD11b+ CD64+ cells) and DCs (CD64- MHC-IIhi CD11c+). Cardiac macrophages can be further classified based on their expression of MHC-II and CD11c. Cardiac DCs can be subdivided based on their expression of CD103 or CD11b. (D) Representative plots of DAPI+ dead cells within the macrophage and DC compartments after small size exclusion. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58114/58114fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58114/58114fig2.jpg" /> 
Figure 2: Labelling of intravascular cells contaminants. (A) Comparison of intravascular CD45 labelling of blood neutrophils and Ly6Chi monocytes in anti-CD45 antibody treated (red) or non-treated (blue) mice. (B) Identification of intravascular contaminants in a cardiac preparation by intravascular CD45 labelling. (C) Representative flow plots of myocardial and intravascular cells. (D) Representative plots of labelled cells within the macrophage and DC compartments after anti-CD45 i.v. administration. Note that macrophages and DCs are exclusively extravascular. Intravascular contaminants are typically lymphocytes, monocytes and neutrophils2. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58114/58114fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58114/58114fig3.jpg" /> 
Figure 3: Quantification of cardiac macrophages and DCs in BATF3-deficient mice. (A) Representative flow plots of cardiac DCs in Batf3+/+ and Batf3-/- mice. (B) Quantification of cardiac DCs in Batf3+/+ and Batf3-/- mice. Note the absence of CD103+ DCs in Batf3-/- mice. (C and D) Quantification of cardiac macrophages (MF) in Batf3+/+ and Batf3-/- mice. Error bars represent SEM. Red symbols represent individual animals. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58114/58114fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Isolation and Identification of Extravascular Immune Cells of the Heart at JoVE.com

Isolation and Identification of Extravascular Immune Cells of the Heart

This protocol presents a simple and efficient method to isolate, identify and quantify immune cells residing in the myocardium of mice during steady state or inflammation. The protocol combines enzymatic and mechanical digestion for the generation of a single cell suspension that can be further analyzed by flow cytometry.

The immune system is an essential component of a healthy heart. The myocardium is home to a rich population of different immune cell subsets with ...

isolation-identification-extravascular-immune-cells

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

12.4K Views.  University Health Network (UHN). This protocol presents a simple and efficient method to isolate, identify and quantify immune cells residing in the myocardium of mice during steady state or inflammation. The protocol combines enzymatic and mechanical digestion for the generation of a single cell suspension that can be further analyzed by flow cytometry.

Video: Isolation and Identification of Extravascular Immune Cells of the Heart

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, ...

Seven Steps to Stellate Cells

Hepatic stellate cells are liver-resident cells of star-like morphology and are located in the space of Disse between liver sinusoidal endothelial cells and hepatocytes1,2. Stellate cells are derived from bone marrow precursors and store up to 80% of the total body vitamin A1, 2. Upon activation, stellate cells differentiate into myofibroblasts to produce extracellular matrix, thus contributing to liver fibrosis3. Based on their ability to contract, myofibroblastic stellate cells can regulate the vascular tone associated with portal hypertension4. Recently, we demonstrated that hepatic stellate cells are potent antigen presenting cells and can activate NKT cells as well as conventional T lymphocytes5. 

 Here we present a method for the efficient preparation of hepatic stellate cells from mouse liver. Due to their perisinusoidal localization, the isolation of hepatic stellate cells is a multi-step process. In order to render stellate cells accessible to isolation from the space of Disse, mouse livers are perfused in situ with the digestive enzymes Pronase E and Collagenase P. Following perfusion, the liver tissue is subjected to additional enzymatic treatment with Pronase E and Collagenase P in vitro. Subsequently, the method takes advantage of the massive amount of vitamin A-storing lipid droplets in hepatic stellate cells. This feature allows the separation of stellate cells from other hepatic cell types by centrifugation on an 8% Nycodenz gradient. The protocol described here yields a highly pure and homogenous population of stellate cells. Purity of preparations can be assessed by staining for the marker molecule glial fibrillary acidic protein (GFAP), prior to analysis by fluorescence microscopy or flow cytometry. Further, light microscopy reveals the unique appearance of star-shaped hepatic stellate cells that harbor high amounts of lipid droplets. 


 Taken together, we present a detailed protocol for the efficient isolation of hepatic stellate cells, including representative images of their morphological appearance and GFAP expression that help to define the stellate cell entity.

Here we describe a method for the isolation of hepatic stellate cells from mouse liver. For stellate cell purification, mouse livers are digested in situ and in vitro by pronase-collagenase treatment prior to density gradient centrifugation. This technique yields highly pure hepatic stellate cells.

Hepatic stellate cells are liver-resident  cells of star-like morphology and are located in the space of Disse between  liver sinusoidal endothelial cells ...

Isolation of Functional Cardiac Immune Cells

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5 These immune cells, which include mast cells, T-cells and macrophages; store and release a variety of biologically active mediators including cytokines and proteases such as tryptase.6-8 These mediators have been shown to be key players in extracellular matrix metabolism by activating matrix metalloproteinases or causing collagen accumulation by modulating the cardiac fibroblasts' function.9-11 However, available techniques for isolating cardiac immune cells have been problematic because they use bacterial collagenase to digest the myocardial tissue. This technique causes activation of the immune cells and thus a loss of function. For example, cardiac mast cells become significantly less responsive to compounds that cause degranulation.12 Therefore, we developed a technique that allows for the isolation of functional cardiac immune cells which would lead to a better understanding of the role of these cells in cardiac disease.13, 14
This method requires a familiarity with the anatomical location of the rat's xiphoid process, axilla and falciform ligament, and pericardium of the heart. These landmarks are important to increase success of the procedure and to ensure a higher yield of cardiac immune cells. These isolated cardiac immune cells can then be used for characterization of functionality, phenotype, maturity, and co-culture experiments with other cardiac cells to gain a better understanding of their interactions.

This method for isolating functional immune cells from the heart provides an alternative to the conventional methods of collagenase digestion, which causes unwanted immune cell activation, resulting in a decreased responsiveness of these cells. Our method of isolation yields functional cardiac immune cells by avoiding problems associated with enzymatic digestion.

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5  These immune cells, which ...

An Introduction to Parasitic Wasps of Drosophila and the Antiparasite Immune Response

Most known parasitoid wasp species attack the larval or pupal stages of Drosophila. While Trichopria drosophilae infect the pupal stages of the host (Fig. 1A-C), females of the genus Leptopilina (Fig. 1D, 1F, 1G) and Ganaspis (Fig. 1E) attack the larval stages. We use these parasites to study the molecular basis of a biological arms race. Parasitic wasps have tremendous value as biocontrol agents. Most of them carry virulence and other factors that modify host physiology and immunity. Analysis of Drosophila wasps is providing insights into how species-specific interactions shape the genetic structures of natural communities. These studies also serve as a model for understanding the hosts' immune physiology and how coordinated immune reactions are thwarted by this class of parasites.
The larval/pupal cuticle serves as the first line of defense. The wasp ovipositor is a sharp needle-like structure that efficiently delivers eggs into the host hemocoel. Oviposition is followed by a wound healing reaction at the cuticle (Fig. 1C, arrowheads). Some wasps can insert two or more eggs into the same host, although the development of only one egg succeeds. Supernumerary eggs or developing larvae are eliminated by a process that is not yet understood. These wasps are therefore referred to as solitary parasitoids.
Depending on the fly strain and the wasp species, the wasp egg has one of two fates. It is either encapsulated, so that its development is blocked (host emerges; Fig. 2 left); or the wasp egg hatches, develops, molts, and grows into an adult (wasp emerges; Fig. 2 right). L. heterotoma is one of the best-studied species of Drosophila parasitic wasps. It is a "generalist," which means that it can utilize most Drosophila species as hosts1. L. heterotoma and L. victoriae are sister species and they produce virus-like particles that actively interfere with the encapsulation response2. Unlike L. heterotoma, L. boulardi is a specialist parasite and the range of Drosophila species it utilizes is relatively limited1. Strains of L. boulardi also produce virus-like particles3 although they differ significantly in their ability to succeed on D. melanogaster1. Some of these L. boulardi strains are difficult to grow on D. melanogaster1 as the fly host frequently succeeds in encapsulating their eggs. Thus, it is important to have the knowledge of both partners in specific experimental protocols.
In addition to barrier tissues (cuticle, gut and trachea), Drosophila larvae have systemic cellular and humoral immune responses that arise from functions of blood cells and the fat body, respectively. Oviposition by L. boulardi activates both immune arms1,4. Blood cells are found in circulation, in sessile populations under the segmented cuticle, and in the lymph gland. The lymph gland is a small hematopoietic organ on the dorsal side of the larva. Clusters of hematopoietic cells, called lobes, are arranged segmentally in pairs along the dorsal vessel that runs along the anterior-posterior axis of the animal (Fig. 3A). The fat body is a large multifunctional organ (Fig. 3B). It secretes antimicrobial peptides in response to microbial and metazoan infections.
Wasp infection activates immune signaling (Fig. 4)4. At the cellular level, it triggers division and differentiation of blood cells. In self defense, aggregates and capsules develop in the hemocoel of infected animals (Fig. 5)5,6. Activated blood cells migrate toward the wasp egg (or wasp larva) and begin to form a capsule around it (Fig. 5A-F). Some blood cells aggregate to form nodules (Fig. 5G-H). Careful analysis reveals that wasp infection induces the anterior-most lymph gland lobes to disperse at their peripheries (Fig. 6C, D).
We present representative data with Toll signal transduction pathway components Dorsal and Sp&auml;tzle (Figs. 4,5,7), and its target Drosomycin (Fig. 6), to illustrate how specific changes in the lymph gland and hemocoel can be studied after wasp infection. The dissection protocols described here also yield the wasp eggs (or developing stages of wasps) from the host hemolymph (Fig. 8).

An Introduction to Parasitic Wasps of Drosophila and the Antiparasite Immune Response

Parasitoid (parasitic) wasps constitute a major class of natural enemies of many insects including Drosophila melanogaster. We will introduce the techniques to propagate these parasites in Drosophila spp. and demonstrate how to analyze their effects on immune tissues of Drosophila larvae.

Most known parasitoid wasp species attack the larval or pupal stages of Drosophila. While Trichopria drosophilae infect the pupal stages of the host (Fig. ...

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 ...

Differentiating Functional Roles of Gene Expression from Immune and Non-immune Cells in Mouse Colitis by Bone Marrow Transplantation

To understand the role of a gene in the development of colitis, we compared the responses of wild-type mice and gene-of-interest deficient knockout mice to colitis. If the gene-of-interest is expressed in both bone marrow derived cells and non-bone marrow derived cells of the host; however, it is possible to differentiate the role of a gene of interest in bone marrow derived cells and non- bone marrow derived cells by bone marrow transplantation technique. To change the bone marrow derived cell genotype of mice, the original bone marrow of recipient mice were destroyed by irradiation and then replaced by new donor bone marrow of different genotype. When wild-type mice donor bone marrow was transplanted to knockout mice, we could generate knockout mice with wild-type gene expression in bone marrow derived cells. Alternatively, when knockout mice donor bone marrow was transplanted to wild-type recipient mice, wild-type mice without gene-of-interest expressing from bone marrow derived cells were produced. However, bone marrow transplantation may not be 100% complete. Therefore, we utilized cluster of differentiation (CD) molecules (CD45.1 and CD45.2) as markers of donor and recipient cells to track the proportion of donor bone marrow derived cells in recipient mice and success of bone marrow transplantation. Wild-type mice with CD45.1 genotype and knockout mice with CD45.2 genotype were used. After irradiation of recipient mice, the donor bone marrow cells of different genotypes were infused into the recipient mice. When the new bone marrow regenerated to take over its immunity, the mice were challenged by chemical agent (dextran sodium sulfate, DSS 5%) to induce colitis. Here we also showed the method to induce colitis in mice and evaluate the role of the gene of interest expressed from bone-marrow derived cells. If the gene-of-interest from the bone derived cells plays an important role in the development of the disease (such as colitis), the phenotype of the recipient mice with bone marrow transplantation can be significantly altered. At the end of colitis experiments, the bone marrow derived cells in blood and bone marrow were labeled with antibodies against CD45.1 and CD45.2 and their quantitative ratio of existence could be used to evaluate the success of bone marrow transplantation by flow cytometry. Successful bone marrow transplantation should show a vast majority of donor genotype (in term of CD molecule marker) over recipient genotype in both the bone marrow and blood of recipient mice.

Bone marrow transplantation provides a way to change the genotype of the bone marrow derived cells. If the gene of interest is expressed in both bone marrow derived cells and non-bone marrow derived cells, bone marrow transplantation can change the bone marrow derived cells to a different genotype without changing the non-bone marrow derived cell genotype.

To understand the role of a gene in the development of colitis, we compared the responses of wild-type mice and gene-of-interest deficient knockout mice ...

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 ...

Isolation of Adipose Tissue Immune Cells

The discovery of increased macrophage infiltration in the adipose tissue (AT) of obese rodents and humans has led to an intensification of interest in immune cell contribution to local and systemic insulin resistance. Isolation and quantification of different immune cell populations in lean and obese AT is now a commonly utilized technique in immunometabolism laboratories; yet extreme care must be taken both in stromal vascular cell isolation and in the flow cytometry analysis so that the data obtained is reliable and interpretable. In this video we demonstrate how to mince, digest, and isolate the immune cell-enriched stromal vascular fraction. Subsequently, we show how to antibody label macrophages and T lymphocytes and how to properly gate on them in flow cytometry experiments. Representative flow cytometry plots from low fat-fed lean and high fat-fed obese mice are provided. A critical element of this analysis is the use of antibodies that do not fluoresce in channels where AT macrophages are naturally autofluorescent, as well as the use of proper compensation controls.

Adipose tissue (AT) is a site of intense immune cell activation and interaction. Almost all cells of the immune system are present in AT and their ratios are altered by obesity. Proper isolation, quantification, and characterization of AT immune cell populations are critical for understanding their role in immunometabolic disease.

The discovery of increased macrophage infiltration in  the adipose tissue (AT) of obese rodents and humans has led to an  intensification of interest in ...

Isolation, Identification, and Purification of Murine Thymic Epithelial Cells

The thymus is a vital organ for T lymphocyte development. Of thymic stromal cells, thymic epithelial cells (TECs) are particularly crucial at multiple stages of T cell development: T cell commitment, positive selection and negative selection. However, the function of TECs in the thymus remains incompletely understood. In the article, we provide a method to isolate TEC subsets from fresh mouse thymus using a combination of mechanical disruption and enzymatic digestion. The method allows thymic stromal cells and thymocytes to be efficiently released from cell-cell and cell-extracellular matrix connections and to form a single-cell suspension. Using the isolated cells, multiparameter flow cytometry can be applied to identification and characterization of TECs and dendritic cells. Because TECs are a rare cell population in the thymus, we also describe an effective way to enrich and purify TECs by depleting thymocytes, the most abundant cell type in the thymus. Following the enrichment, cell sorting time can be decreased so that loss of cell viability can be minimized during purification of TECs. Purified cells are suitable for various downstream analyses like Real Time-PCR, Western blot and gene expression profiling. The protocol will promote research of TEC function and as well as the development of in vitro T cell reconstitution.

Here we describe an efficient method for isolation, identification, and purification of mouse thymic epithelial cells (TECs). The protocol can be utilized for studies of thymus function for normal T cell development, thymus dysfunction, and T cell reconstitution.

The thymus is a vital organ for T lymphocyte development. Of thymic stromal cells, thymic epithelial cells (TECs) are particularly crucial at multiple ...

Isolation and Flow Cytometric Analysis of Immune Cells from the Ischemic Mouse Brain

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident cells. A key mechanism of this inflammatory response is the migration of circulating immune cells to the ischemic brain facilitated by chemokine release and increased endothelial adhesion molecule expression. Brain-invading leukocytes are well-known contributing to early-stage secondary ischemic injury, but their significance for the termination of inflammation and later brain repair has only recently been noticed.
Here, a simple protocol for the efficient isolation of immune cells from the ischemic mouse brain is provided. After transcardial perfusion, brain hemispheres are dissected and mechanically dissociated. Enzymatic digestion with Liberase&#160;is followed by density gradient (such as Percoll) centrifugation to remove myelin and cell debris. One major advantage of this protocol is the single-layer density gradient procedure which does not require time-consuming preparation of gradients and can be reliably performed. The approach yields highly reproducible cell counts per brain hemisphere and allows for measuring several flow cytometry panels in one biological replicate. Phenotypic characterization and quantification of brain-invading leukocytes after experimental stroke may contribute to a better understanding of their multifaceted roles in ischemic injury and repair.

Inflammation plays a central role in the pathogenesis of ischemic stroke. Increasing evidence suggests that it acts as a double-edged sword which exacerbates early brain injury, but also contributes to later repair. This protocol describes the isolation of immune cells from the ischemic brain and their subsequent flow cytometric phenotyping.

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident ...