This study was funded by NHLBI grants 5K08HLO145108-03 and 1R01HL160716-01 awarded to Luigi Adamo.
The Aurora Flow Cytometer used to develop this study was funded by NIH Grant S10OD026859. We acknowledge the support of the JHU Ross Flow Cytometry Core.

All experiments described in this manuscript were performed with the approval of the IACUC at Johns Hopkins University School of Medicine.
1. Preparations
<ol>
	<li>Prepare the FACS Buffer, as described in Table 1.</li>
	<li>Ensure there is enough CO2 to euthanize the animals.</li>
	<li>Prepare the dissection space (place the bench pad and place the tape and dissection tools nearby).</li>
	<li>Label the 15 mL tubes, put 3 mL of HBSS with calcium and magnesium in each tube (see Table of Materials) and place them on ice (one tube per heart and an extra one for the Reference group/flow cytometry controls). Also, fill the small (35 mm) Petri dishes with HBSS.</li>
	<li>Turn on the temperature-controlled shaker, set it to 37 &#176;C, and pre-chill the centrifuge at a temperature of 4 &#176;C.</li>
	<li>Prepare the syringe pump and set the flow rate to 3 mL/min. Fill a 20 mL syringe with HBSS, prime the tube line with buffer, and remove any bubbles.</li>
</ol>
2. Intravascular B-cell staining (in vivo)
<ol>
	<li>Weigh a 12-week-old male mouse of the C57BL/6J strain. Load a 3/10 cc insulin syringe with 100 &#181;L of the B220 antibody dilution (1:10, in PBS). Cover the syringe with aluminum foil to protect from light.</li>
	<li>Anesthetize one mouse.
	<ol>
		<li>Apply an intraperitoneal injection of 2% 2,2,2-tribromoethanol with a dose of 400 mg/kg and wait 10-20 min. 
		NOTE: Monitor the mouse&#39;s breathing and movement. Once the mouse stops moving, stimulate pain by pinching the toe; the absence of a withdrawal reflex indicates adequate anesthesia.</li>
		<li>If adequate anesthesia is not achieved within the course of 20 min, an additional dose of anesthetic at 100 mg/kg can be administered, and an assessment of mouse kinetics, withdrawal reflex, and respiration should be made every 5 min.</li>
	</ol>
	</li>
	<li>Place the mouse on the lab bench on a bench pad inclined toward one side, gently pull down the skin of the back, and slightly press the body against the bench to make the eye protrude.</li>
	<li>Inject the diluted B220 antibody from step 2.1 by retroorbital injection.
	<ol>
		<li>Introduce the syringe into the eye at 45&#176; from the sagittal plane of the mouse&#39;s head. Slowly inject the solution. If resistance is felt, remove the syringe and enter again. Wait for 2 min. 
		NOTE: Prolonged incubation time would jeopardize the ability to discriminate intra-vascular vs extra-vascular B cells as it would give time to the injected antibody to diffuse outside of the vasculature.&#160;</li>
	</ol>
	</li>
	<li>Euthanize the mouse as per the IACUC regulations.
	<ol>
		<li>Open the CO2 flow to the euthanasia chamber at a rate of 0.5 L/min, or the recommended flow rate based on the size of your chamber.</li>
		<li>Place the mouse in the chamber for the recommended time, ensuring it is no longer breathing. Remove it and perform cervical dislocation as a secondary method of euthanasia.</li>
		<li>Once the mouse is euthanized, promptly proceed to organ harvesting and perfusion.</li>
	</ol>
	</li>
</ol>
3. Heart collection
<ol>
	<li>Chest opening.
	<ol>
		<li>Hold the skin of the mouse with forceps at the epigastric area. Make a 2 mm opening in the skin using scissors and use the forceps to peel the skin away to reveal the fascia layer, a shiny translucent layer, of the chest and abdomen. Cut at the epigastrium through the fascia and peritoneum to open the abdominal wall and reveal the xiphoid process. Clamp the xiphoid process using hemostatic forceps.</li>
		<li>With the scissors, make a vertical cut through the chest wall at the level of the mid-clavicular line on each side and lift the anterior chest wall to reveal the mediastinum content, using hemostatic forceps as a weight to maintain the opening.</li>
	</ol>
	</li>
	<li>Heart perfusion and extraction.
	<ol>
		<li>Using forceps, remove any fat or thymic tissue surrounding the heart.</li>
		<li>Hold the aorta securely from behind using forceps. Introduce the syringe needle (25 G) into the apex of the heart. Perfuse with 3 mL of HBSS at a rate of 3 mL/min. 
		NOTE: Monitor the perfusion. The liver should fill, and blood in the coronary vessels should be entirely removed.&#160;Use of a syringe pump calibrated to 1 mL/ minute is recommended to maintain a steady flow.</li>
		<li>Cut the aorta using scissors and take out the heart. Wash the heart in the Petri dish with HBSS to remove any remaining blood. Using scissors and forceps, remove the fat, connective tissue and thymus, and dry the heart. Weigh the isolated heart and note the weight in milligrams. Mince the heart at room temperature into small pieces (1-2 mm) with a blade. 
		NOTE: Adult mouse hearts may weigh between 0.090-0.210 mg.</li>
		<li>Measure the mass of heart tissue and place 60 mg of the minced heart to be digested in a 15 mL tube with 3 mL of HBSS. Retain the remaining heart tissue and place in a tube labelled &#34;Reference control tissue&#34;; this will be used for compensation control for live-dead signal and autofluorescence.</li>
	</ol>
	</li>
</ol>
4. Heart digestion and cell staining
<ol>
	<li>Add enzymes to each tube containing minced tissue (see Table 1). Add 0.5 &#181;L/mg of DNase 1 (300 Units for 60 mg of heart tissue), 0.5 &#181;L/mg of collagenase II (625 Units for 60 mg of heart tissue), and 0.2 &#181;L/mg of hyaluronidase (50 Units for 60 mg of heart tissue) to the tube.</li>
	<li>Place the digestion reaction in the shaker for 30 min at 300 rpm. Adjust the shaker rack to about 25&#176; of incline.</li>
	<li>Add 7 mL of HBSS to each of the 15 mL reaction tubes and centrifuge at 250 x g for 5 min at 4 &#176;C with acceleration and breaking set to moderate or slow. Decant the supernatant and resuspend each pellet in 5 mL of ACK lysis buffer to remove red blood cells. Incubate in the ACK lysis buffer for 5 min at room temperature.</li>
	<li>Add 10 mL of PBS to each tube, mix, and then filter into a 50 mL tube with a 40 &#181;m strainer. Ensure that all the debris is inside the filter chamber. Smash any debris on the filter with a syringe plunger until it is completely suspended within the filter chamber.</li>
	<li>Add PBS through the filter until the volume reaches 25 mL per heart; this will allow the suspended cells to pass through the filter. Add an additional 25 mL of PBS to the Reference control tissue tube and divide the volume into different tubes (25 mL each). Label one of the tubes &#34;Unstained&#34; and the other one as &#34;Live-dead&#34;. Centrifuge at 250 x g for 5 min at 4 &#176;C. 
	NOTE: At this point, prepare the dilution for the live-dead stain (dilution: 1:500 in 1x PBS).</li>
	<li>Decant the supernatant, resuspend the pellet in the Unstained tube in 100 &#181;L of PBS and each of the other pellets in 100 &#181;L of the live-dead stain dilution. Incubate on ice for 30 min in the dark, covering the ice bucket with aluminum foil.</li>
	<li>Add 500 &#181;L of FACS buffer to each sample and transfer it into a labelled FACS tube through a 40 &#181;m filter. Centrifuge the FACS tubes at 250 x g for 5 min at 4 &#176;C. Discard the supernatant. Resuspend the pellet in 500 &#181;L of FACS buffer.</li>
	<li>Add 50 &#181;L of Fc blocking solution (see Table 1) to each tube, except for the Unstained and Live-dead tubes. Mix and incubate for 5 min.</li>
	<li>Add 50 &#181;L of antibody mix solution (see Table 1) to each tube, except for the Unstained and Live-dead tubes, and incubate for 30 min in the dark, covering the ice bucket with aluminum foil.</li>
	<li>Add 2 mL of FACS buffer to each tube and centrifuge at 250 x g for 5 min at 4 &#176;C. Remove the supernatant and resuspend in 300-500 &#181;L of FACS buffer.</li>
</ol>
5. Flow cytometry
<ol>
	<li>Setup instrument control settings. 
	NOTE:&#160;In this protocol the sample is analyzed with a 4 lasers Cytek Aurora flow cytometer. Another appropriate flow cytometer could be used to detect the markers of interest.&#160;
	<ol>
		<li>Open the instrument control software (see Table of Materials). At the top of the window, select the Acquisition tab and click on New +. A window called Create New Experiment will display.</li>
		<li>In the Library section, in the field Type to filter, type PE, PerCP-Cy5.5, BV421, Alexa Fluor 700, and Zombie Aqua, clicking Add after typing each. The fluorophore names should be seen to the right. Then click Next to proceed to the Group tab.</li>
		<li>In the Group tab, click the symbol with a tube to add the heart for the analysis.</li>
		<li>Click on the + Reference group and a window will display. In the fluorescent tag column select Zombie Aqua, and in the control type column select Cells, then click Save.</li>
		<li>Click Next &#62; Markers tab; a table will display with the names of the fluorophores. Type the corresponding cell marker in the first row of each fluorophore column and type Enter: CD45 for PerCP-Cy5.5; CD19 for BV421; CD11b for Alexa Fluor 700; B220 for PE; and live-dead for Zombie Aqua.</li>
		<li>Click Next twice to go to the Acquisition tab. In Events to Record of the experiment group type 10,000,000, and in the same field for the reference group line type 200,000. Click Save and Open. The other settings should remain as default, Stopping Time to 10,000, and Stopping Volume to 3,000.</li>
		<li>On the bottom left click on Instrument control. Set Voltage to FSC-50, SSC-175, SSC-B-175. Then, select the Acquisition Control. For the Flow rate option, select High from the drop-down menu.</li>
	</ol>
	</li>
	<li>Acquire reference samples with a cytometer and unmix in software.
	<ol>
		<li>Vortex the Unstained heart tube and place it securely on the sample injection port (SIP). Ensure the green indicator arrow is pointing to the correct sample, then click Start in the Acquisition Control panel of the cytometer software and wait until the events are recorded. Do the same for the Live-dead-stained heart tissue.</li>
		<li>Select the Unmix menu and set the following controls:
		<ol>
			<li>Select the checkboxes for PE, PerCP-Cy5.5, BV421, and Alexa Fluor 700 in the column from library. Ensure Zombie Aqua is unchecked in the same column. At the bottom, check the option of Autofluorescence as a Fluorescent Tag.</li>
			<li>Click Next to proceed to the Identify Positive/Negative Populations tab. In this tab, a plot of FSC-A vs. SSC-B-A will be displayed. Click and drag the points of the polygon to select an area between 1.0 M and 4.0 M on the FSC-A axis, and between 0.2 M and 3.0 M on the SSC-B-A axis.</li>
			<li>In the next plot to the right, V5-A will be displayed on the X-axis. Move the gates to select half of the peak on the far right as positive and a region on the left side of the plot as negative. In the plot titled Reference Group - Unstained, select a population in a similar range. For this the unstained no positive-negative has to be set.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Acquire experimental samples.
	<ol>
		<li>For each tube containing a sample for an individual mouse, vortex the tube and place it securely on the SIP. Ensure the green indicator arrow is pointing to the correct sample, then click Start in the Acquisition Control panel of the cytometer software, and wait until the events are recorded.</li>
	</ol>
	</li>
	<li>Gating strategy.
	<ol>
		<li>Select the Default Unmixed Worksheet tab. Create an FSC-A vs. SSC-A plot using the Dot plot tool on top. Select events higher than 0.4 M on the FSC-A axis and higher than 0.8 M on the SSC-A axis using the Polygon selection tool. Double click in this selection and a new plot will display.</li>
		<li>From the newly created plot, right-click on the X axis label and select AF-A; right-click on the Y axis label and select Live-dead stain. Click on the Rectangle selection tool on top and select all the events below 105 on the live-dead axis&#160;that correspond to the lower live-dead signal. Double click in this selection and a new plot will display.</li>
		<li>In the new plot, right-click to select PerCP-Cy5.5-A for the X axis and SSC-A for the Y axis. Using the Rectangle selection tool, select the events higher than 104 on the PerCP-Cy5.5-A axis that correspond to the CD45 positive cells. Double click on the selection and a new plot will display.</li>
		<li>On the new plot, right-click to select FSC-A for the X axis and FSC-H for the Y axis. Using the Polygon selection tool&#160;select the events that follow a trend on which FSC-A value and SSC-A value are the same; with this the cell doublets will be removed and the population in the selection will be referred to as the CD45+ population. Double click on the selection to display a new plot with the CD45+ population.</li>
		<li>From the CD45+ population plot, right-click to select BV421 on the X axis vs. SSC-A on the Y axis. Using the Polygon selection tool, select the population to the right on the BV421 axis. This is the B-cell population. Double click twice in this population to create two new plots with the B-cell population.
		<ol>
			<li>Right-click to setup the first B-cell plot with PE-A at the X axis and BV421-A on the Y axis. The population to the right corresponds to the intravascular B-cells, and the population to the left represents the interstitial B-cells. Use the Polygon selection tool to make a selection of both.</li>
			<li>Right-click to setup the second B-cell plot with Alexa Fluor 700-A on the X axis and SSC-A on the Y axis. The population at the left corresponds to the CD11b- cells, and the events at the right (if any) correspond to the CD11b+ cells.</li>
			<li>Right click on each selection and then click Gate Properties. Click on Count and % Parent to display the sizes and percentages. Record the number of events and percentages for further data analysis.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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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=A+protocol+for+the+comprehensive+flow+cytometric+analysis+of+immune+cells+in+normal+and+inflamed+murine+non-lymphoid+tissues.">A protocol for the comprehensive flow cytometric analysis of immune cells in normal and inflamed murine non-lymphoid tissues.</a> PLoS One. 11 (3), 0150606 (2016).</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>Horckmans, 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=Pericardial+adipose+tissue+regulates+granulopoiesis,+fibrosis+and+cardiac+function+after+myocardial+infarction.">Pericardial adipose tissue regulates granulopoiesis, fibrosis and cardiac function after myocardial infarction.</a> Circulation. 137 (9), 948-960 (2017).</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. 111 (45), 16029-16034 (2014).</li><li>Bajpai, G., Lavine, K. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+of+macrophage+subsets+and+stromal+cells+from+human+and+mouse+myocardial+specimens.">Isolation of macrophage subsets and stromal cells from human and mouse myocardial specimens.</a> Journal of Visualized Experiments. (154), e60015 (2019).</li><li>Anderson, K. G., 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=Intravascular+staining+for+discrimination+of+vascular+and+tissue+leukocytes.">Intravascular staining for discrimination of vascular and tissue leukocytes.</a> Nature Protocols. 9 (1), 209-222 (2014).</li><li>Coffman, R. L., Weissman, I. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=B220:+a+B+cell-specific+member+of+th+T200+glycoprotein+family.">B220: a B cell-specific member of th T200 glycoprotein family.</a> Nature. 289 (5799), 681-683 (1981).</li><li>Montecino-Rodriguez, E., Dorshkind, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=B-1+B+cell+development+in+the+fetus+and+adult.">B-1 B cell development in the fetus and adult.</a> Immunity. 36 (1), 13-21 (2012).</li><li>Bermea, K., Bhalodia, A., Huff, A., Rousseau, S., Adamo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+B+cells+in+cardiomyopathy+and+heart+failure.">The role of B cells in cardiomyopathy and heart failure.</a> Current Cardiology Reports. , 01722-01724 (2022).</li><li>Zhao, T. 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=Rituximab+in+patients+with+acute+ST-elevation+myocardial+infarction:+an+experimental+medicine+safety+study.">Rituximab in patients with acute ST-elevation myocardial infarction: an experimental medicine safety study.</a> Cardiovascular Research. 118 (3), 872-882 (2022).</li><li>Kushnir, N., 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=B2+but+not+B1+cells+can+contribute+to+CD4++T-cell-mediated+clearance+of+rotavirus+in+SCID+mice.">B2 but not B1 cells can contribute to CD4+ T-cell-mediated clearance of rotavirus in SCID mice.</a> Journal of Virology. 75 (12), 5482-5490 (2001).</li></ol>

Luigi Adamo is co-founder of i-Cordis, LLC, a company focused on the development of immunomodulatory molecules for the treatment of heart failure, with a specific interest in derivatives of the B-cell modulating drug pirfenidone. The remaining authors have no conflicts to declare.

A growing body of evidence indicates that B-cells play an important role in the context of myocardial physiology and myocardial remodeling/adaptation to injury7,8,9,10,11,12,13,36. Flow cytometry is an excellent tool to study immune cell populations in any tissue and almost a mandatory tool for any study that focuses on B-cells in the heart.
The emerging literature on B-cells and the heart already reports contrasting findings on very basic aspects, such as the number of B-cells in the heart and changes in myocardial B-cell number with injury. B-cells have been reported as the most prevalent immune cell in rodent hearts7. However, other studies evaluated their relative prevalence compared to other myeloid cells and showed markedly lower numbers26,27. Pinto et al.26 found that B-cells comprised around 9% of the total leukocytes, while Yu et al.27 reported 10%. Adamo et al.7 reported that the average percentage of B-cells relative to leukocyte numbers was around 20%, which is very close to the 18.1% reported in this manuscript. Other studies have reported that B-cells have a very low prevalence in the myocardium28.
Several critical experimental steps could explain the variabilities in B-cell numbers reported in the literature. Namely, differences in the perfusion of the heart, tissue mincing, and digestion can significantly affect the number of B-cells recovered per mg of heart tissue. Perfusion with buffers containing calcium chelating agents and no calcium, or perfusion with a higher volume or perfusion rate, could detach the B-cells attached to the endothelium and reduce the recovery yield. Excessively fine tissue mincing could lead to over-digestion of the tissue and loss of cell viability. A reduction in cell viability could also be the result of digesting for a prolonged time or with higher enzyme concentrations.
The percentage of CD45+ cells that correspond to B-cells could be in a range between 15%-30% in a wild-type mouse. Depending on the mouse strain used, the age, and the genetic background, this value can vary. However, the percentage should remain similar in the studied group. Large variability in the number and percentage of B-cells between mice belonging to the same group suggests that an experimental error may be occurring. Low percentages could indicate an excess in perfusion strength and time. Global reduction in CD45+ cells could indicate that the tissue is being over-digested.
The protocol described here has been optimized to provide a consistent yield of B-cells from the digested murine heart and to allow the discrimination of intravascular vs. extravascular B-cells6. Discrimination of intravascular vs. extravascular B-cells is achieved via intravascular injection of an antibody immediately prior to sacrifice. The antibody injected intravascularly readily diffuses throughout the vasculature and encounters all the intravascular B-cells. If the animal is sacrificed soon enough and the heart is promptly harvested, the antibody does not have time to diffuse in the extravascular space and stain extravascular B-cells. Since the time between the intravascular injection of the antibody and organ harvest is critical, it is recommended to inoculate with the antibody and collect one individual mouse heart at a time. Intravascular injection of antibodies can be omitted if the discrimination of intravascular vs. extravascular B-cells is not of interest. The gating strategy displayed in Figure 1 would remain the same except for the last step, which would not be possible due to the lack of B220 labeling. Depending on the experimental design, any fluorophore from the panel shown can be modified to a color that fits with the panel used. Also, it is important to consider that the selection of the surface markers could be modified according to the needs of the group performing it; for instance, if in the experimental design it is believed that CD11b expression on B1 cells could be reduced, the addition of different markers is recommended, such as IgM, IgD, and CD537.
Please note that it might be necessary to process a heart just to run compensation controls for flow cytometry. This is recommended the first time the experiment is optimized. In the following experiments, saving some tissue at the moment of heart collection could be sufficient, as it would provide sufficient cells to run a negative &#34;unstained&#34; control and a positive &#34;live-dead&#34; stain control. Individual controls for the other colors could be either imported from prior experiments or generated using beads.
Some steps of the protocol can be modified without altering the outcomes of the experiment. For instance, the anesthesia method could be changed for a preferred one depending on the researcher&#39;s experience; the use of isoflurane instead of 2,2,2-tribromoethanol could reduce the time spent between anesthesia administration and heart collection. Also, the perfusion of the heart could be done manually without the use of a syringe pump as long as it is performed slowly, gently, and consistently.
Despite the accurate execution of the protocol, inexperienced operators could encounter high variability between samples within the same experimental session or between different experimental sessions. Based on experience, this variability disappears once the scientist has had the opportunity to become fully familiar with the workflow. It is therefore recommended that anyone who wants to implement this protocol assess its reproducibility in two to three trial runs&#160;before collecting crucial data on their research projects.
In summary, existing methods for isolating immune cells from the heart and analyzing them via flow cytometry typically do not consider the ability of immune cells to attach to the endothelium, and either do not standardize perfusion or implement extensive perfusion, with the assumption that any immune cell in the intravascular space is a contaminant that must be purged. This ignores the notion that intravascular immune cells adhered to the endothelium have biological relevance. The protocol here presented has been optimized to maximize the recovery of myocardial immune cells with specific attention to myocardial B-cells, intravascular and extravascular. We expect that this protocol will be of interest to all those scientists who work in the field of cardiac immunology and are interested in exploring the function that B-cells might play in healthy and injured hearts.

B-lymphocytes are highly specialized immune cells that play an important role in both adaptive and innate immune responses1. There are two main populations of B-cells: a smaller population of B1 cells that are mostly produced during embryonic life, and a preponderant population of B2 cells that are produced in adult life in the bone marrow1. After maturing in the bone marrow, B-cells migrate to primary and secondary lymphoid organs. From there they continuously recirculate between lymphoid organs traveling through blood vessels and lymphatic vessels2. B-cells express specific antibodies on their surface, which function as receptors. When B-cells encounter an antigen that binds to their receptor, an activating signal can be triggered. Activated B-cells either migrate to the tissue where the antigen was found or go back to the bone marrow where they can mature into antibody-producing plasma cells3,4.
Recently, it has been appreciated that the heart harbors a sizeable population of B-cells. Studies in rodents have shown that B-cells colonize the heart early during embryonic development5, and that myocardial-associated B-cells are mostly intravascular, naive B2 cells adhered to the endothelium6,7, with a small percentage of B1 cells7. There are still many areas of uncertainty, but the available data indicates that B-cells play an important role both in the naive heart and in the context of myocardial adaptation to injury.
Studies in the naive murine heart have shown that at baseline myocardial B-cells are mostly located in the intravascular space, adhered to the endothelium (&#62;95% of murine cardiac B-cells were found to be located in the intravascular space). These B cells were found to have gene expression patterns different from those of circulating B cells isolated from the peripheral blood. Analysis of naive hearts from B-cell-deficient animals and syngeneic controls found that animals lacking B-cells had smaller hearts and higher ejection fraction6. All this evidence suggests that B-cells might modulate myocardial growth and/or myocardial function, and that not only interstitial but also intravascular B-cells could be responsible for such observations. B-cells were also found to modulate the phenotype of myocardial resident macrophages8.
Several studies have shown that B-cells play an important role in the context of myocardial adaptation to injury8,9,10,11,12,13. B-cells accumulate transiently in the injured heart, likely through a CXCL13-CXCR5 dependent mechanism11,13. From there, B-cells promote adverse cardiac remodeling through several mechanisms that include cytokine-mediated monocyte recruiting9,12. In addition, B-cells can produce antibodies against cardiac proteins that can promote the extension of cardiac damage and adverse cardiac remodeling through several mechanisms14,15,16,17,18,19,20,21,22,23,24,25. B-cells can also exert protective effects on the injured heart through the secretion of IL-1010.
As the number of groups investigating the role of B-cells in the naive and injured heart grows, it is becoming more and more important to define shared protocols to properly quantify and assess myocardial B-cells and thus avoid inconsistencies that have already started to appear in the literature. So far B-cells have in fact both been reported to be one of the most prevalent immune cells in the rodent heart7 and to be present at a markedly lower prevalence than myeloid cells26,27, or to be quite rare28. Similarly, several groups have described that the number of myocardial B-cells increases after acute ischemic myocardial injury7,9,13, but one group reported no changes in the number of B-cells of the injured myocardium29. Studies on cardiac immune cells rarely give details on perfusion conditions and there is no consensus on the digestion conditions. Since in the rodent heart a large proportion of B-cells are intravascular and extraction of immune cells from the myocardium is highly dependent on the digestion method used, the differences reported in the literature might be the result of differences in organ perfusion and tissue digestion.
Presented here is a detailed method for flow cytometry-based quantification of murine myocardial B-cells that maximizes the yield of B-cell recovery by optimizing perfusion and digestion conditions and allows the discrimination of intravascular vs. extravascular myocardial B-cells6. This protocol is an adaptation and optimization of other similar protocols that distinguish between intravascular and interstitial immune cells28,30,31.
In this protocol, we standardize myocardial perfusion to eliminate B-cells floating&#160;in the intravascular space without removing biologically relevant B-cells adhered to the microvascular endothelium. Moreover, building on prior protocols that have described the use of intravenous injection of antibodies to distinguish intravascular from interstitial immune cells32, and taking advantage of the fact that B-cells express the surface marker B22033, we demonstrate how to distinguish intravascular vs. extravascular myocardial B-cells through intravascular injection of a B220-specific antibody immediately before the animal sacrifice and cardiac perfusion. This protocol is relevant to the research of any scientist interested in including the analysis of myocardial B-cells in the naive and injured heart. The widespread implementation of this protocol will reduce inconsistencies between research groups, will allow the analysis of changes in the intravascular and extravascular myocardial B-cell pools, and thus will bolster the advancement of discoveries in the field of cardiac immunology.
In summary, the protocol represents an optimized workflow to quantify and analyze myocardial B-cells via flow cytometry, and at the same time distinguish between cells located in the extravascular space and the intravascular space.

<table><tbody><tr><td>Alexa Fluor 700 anti-mouse/human CD11b Antibody</td><td>101222</td><td>BioLegend</td><td>100 &amp;micro;g 200 &amp;micro;L</td></tr><tr><td>(CellTreat 29481) Cell Strainer, 40 &amp;micro;m, Blue</td><td>QBIAP303</td><td>Southern Labware</td><td></td></tr><tr><td>0.5 mL Natural Microcentrifuge Tube</td><td>1605-0000</td><td>SealRite, USA Scientific</td><td></td></tr><tr><td>0.9% Sodium Chloride Injection, USP</td><td>114-055-101</td><td>Quality Biological</td><td>0.90%</td></tr><tr><td>1.5 mL Natural Microcentrifuge Tube</td><td>1615-5500</td><td>SealRite, USA Scientific</td><td></td></tr><tr><td>10 &amp;micro;L Graduated TipOne Filter Tips</td><td>11213810</td><td>USA Scientific</td><td></td></tr><tr><td>1000 &amp;micro;L Graduated TipOne Filter Tips</td><td>11267810</td><td>USA Scientific</td><td></td></tr><tr><td>15 mL Centrifuge Tube, Plug Seal Cap, Polypropylene, RNase-/DNase-free</td><td>430052</td><td>Corning</td><td></td></tr><tr><td>1-Way Stop Valve, Polycarbonate</td><td>SVPT951</td><td>ECT Manufacturing</td><td></td></tr><tr><td>2,2,2-Tribromoethanol</td><td>T48402</td><td>Sigma-Aldrich</td><td></td></tr><tr><td>200 &amp;micro;L Graduated TipOne Filter Tips</td><td>11208810</td><td>USA Scientific</td><td></td></tr><tr><td>3-Way Stop Valve, Polycarbonate</td><td>SVPT953</td><td>ECT Manufacturing</td><td></td></tr><tr><td>5 mL Polystyrene Round-Bottom Tube, 12 x 75 mm style</td><td>352054</td><td>Falcon, a Corning Brand</td><td></td></tr><tr><td>50 mL Centrifuge Tube, Plug Seal Cap, Polypropylene, RNase-/DNase-free</td><td>430290</td><td>Corning</td><td></td></tr><tr><td>ACK (Ammonium-Chloride-Potassium) Lysing Buffer</td><td>118-156-101</td><td>Quality Biological</td><td>Osmolality: 290 + or -5% mOsm/Kg H20</td></tr><tr><td>Adapter 4x50ml, for 250 mL rectangular bucket in Rotor A-4-63</td><td>5810759005</td><td>Eppendorf</td><td></td></tr><tr><td>Adapter for 15 mL Centrifuge Tubes, 9 Tubes per Adapter, Conical Bottom for use with Rotor Model A-4-62</td><td>22638289</td><td>Eppendorf</td><td></td></tr><tr><td>Adapter for 15 round-bottom tubes 2.6 &amp;ndash; 7 mL, for 250 mL rectangular bucket in Rotor A-4-62</td><td>22638246</td><td>Eppendorf</td><td></td></tr><tr><td>Aluminum Foil 12 in x 75&#039; Roll .0007</td><td>UPC 109153</td><td>Reynolds Wrap</td><td></td></tr><tr><td>Anesthesia Induction Chamber - Mouse</td><td>RWD-AICMV-100</td><td>Conduct Science</td><td></td></tr><tr><td>BD Luer Slip Tip Syringe with attached needle 25 G x 5/8 in., sterile, single use, 1 mL</td><td>309626</td><td>BD Becton, Dickinson and Company</td><td></td></tr><tr><td>Brandzig Ultra-Fine Insulin Syringes 29G 1cc 1/2&quot; 100-Pack</td><td>CMD 2613</td><td>Brandzig</td><td></td></tr><tr><td>Brilliant Violet 421 anti-mouse CD19 Antibody</td><td>115537</td><td>BioLegend</td><td>50 &amp;micro;g/mL</td></tr><tr><td>CAPS for Flow Tubes w/strainer mesh 35 &amp;micro;m, Dual position for 12 x 75 mm tubes, sterile</td><td>T9009</td><td>Southern Labware</td><td></td></tr><tr><td>Carbon Dioxide USP E CGA 940&amp;nbsp;</td><td>CD USPE</td><td>AirGas USA</td><td></td></tr><tr><td>Cole-Parmer Essentials Low-Form Beaker, Glass, 500 mL</td><td>UX-34502-46</td><td>Cole-Parmer</td><td></td></tr><tr><td>Collagenase 2</td><td>LS004176</td><td>Sigma-Aldrich</td><td></td></tr><tr><td>Connector brass chrome plated 1/4&quot; female NPT x 1/4&quot; barb</td><td>Y992611-AG</td><td>AirGas USA</td><td></td></tr><tr><td>Cytek Aurora Flow Cytometer</td><td></td><td>Cytek Biosciences</td><td></td></tr><tr><td>Diss 1080 Nipple 1/4 BARB CP</td><td>M-08-12</td><td>AirGas USA</td><td></td></tr><tr><td>DNase I - 40,000 U</td><td>D4527</td><td>Sigma-Aldrich</td><td></td></tr><tr><td>Easypet&amp;nbsp;3 - Electronic Pipette Controller</td><td>4430000018</td><td>Eppendorf</td><td></td></tr><tr><td>Electronic Balance, AX223/E</td><td>30100606</td><td>Ohaus Corp.</td><td></td></tr><tr><td>Eppendorf 5810R centrifuge</td><td>5810R</td><td>Eppendorf</td><td></td></tr><tr><td>Eppendorf Research plus 1-channel variable pipettes</td><td></td><td>Eppendorf</td><td></td></tr><tr><td>FlowJo 10.8.1</td><td></td><td>BD Becton, Dickinson and Company</td><td></td></tr><tr><td>GLACIERbrand, triple density Ice Pan (IPAN-3100)</td><td>Z740287</td><td>Heathrow Scientific</td><td></td></tr><tr><td>HBSS (1x) - Ca2+ [+] Mg2+ [+]</td><td>14025076</td><td>gibco</td><td>1x</td></tr><tr><td>Hyaluronidase</td><td>H3506</td><td>Sigma-Aldrich</td><td></td></tr><tr><td>Kelly Hemostats, Straight</td><td>13018-14</td><td>Fine Science Tools</td><td></td></tr><tr><td>Luer Slip Syringe sterile, single use, 20 mL</td><td>302831</td><td>BD Becton, Dickinson and Company</td><td></td></tr><tr><td>M1 Adj. Reg 0-100 PSI/CGA940</td><td>M1-940-PG</td><td>AirGas USA</td><td></td></tr><tr><td>McKesson Underpads, Moderate</td><td>4033-CS150</td><td>McKesson</td><td></td></tr><tr><td>Navigator Multi-Purpose Portable Balance</td><td>NV2201</td><td>Ohaus Corp.</td><td></td></tr><tr><td>PBS pH 7.4 (1X) Ca2+ [-] Mg2+ [-]</td><td>10010023</td><td>gibco</td><td>1x</td></tr><tr><td>PE anti-mouse/human CD45R/B220 Antibody</td><td>103208</td><td>BioLegend</td><td>200 &amp;micro;g/mL</td></tr><tr><td>PerCP/Cyanine5.5 anti-mouse CD45 Antibody</td><td>103132</td><td>BioLegend</td><td>100 &amp;micro;g 500 uL</td></tr><tr><td>Petri dish, Stackable 35 mm x 10 mm Sterile Polystyrene</td><td>FB0875711YZ</td><td>Fisher Scientific</td><td></td></tr><tr><td>Pkgd: Diss 1080 Nut/CO2/CO2-02</td><td>M08-1</td><td>AirGas USA</td><td></td></tr><tr><td>Powerful 6 Watt LED Dual Goose-Neck Illuminator</td><td>LED-6W</td><td>AmScope</td><td></td></tr><tr><td>PrecisionGlide Needle 25 G x 5/8 (0.5 mm x 16 mm)</td><td>305122</td><td>BD Becton, Dickinson and Company</td><td></td></tr><tr><td>Purified Rat Anti-Mouse CD16/CD32 (Mouse BD Fc Block) Clone 2.4G2 (RUO)</td><td>553141</td><td>BD Becton, Dickinson and Company Biosciences</td><td>0.5 mg/mL</td></tr><tr><td>R 4.1.1</td><td></td><td>The R Foundation</td><td></td></tr><tr><td>Razor Blades</td><td>9501250000</td><td>Accutec Blades Inc</td><td></td></tr><tr><td>Regulator analytical two stage 0-25 psi delivery CGA320 3500 psi inlet</td><td>Y12244A320-AG</td><td>AirGas USA</td><td></td></tr><tr><td>Rotor A-4-62, incl. 4 x 250 mL rectangular buckets</td><td>Rotor A-4-62</td><td>Eppendorf</td><td></td></tr><tr><td>Serological pipette, plugged, 10 mL, sterile, non-pyrogenic/endotoxin-free, non-cytotoxic, 1 piece(s)/blister</td><td>86.1254.001</td><td>Sarstedt AG &amp;amp; Co KG</td><td></td></tr><tr><td>Sigma label tape</td><td>L8394</td><td>Sigma-Aldrich</td><td></td></tr><tr><td>SpectroFlo 3.0.0</td><td></td><td>Cytek Biosciences</td><td></td></tr><tr><td>Spex VapLock Luer Fitting, PP, Straight, Male Luer Lock x 1/8&quot; Hose Barb; 1/EA</td><td>MTLL230-6005</td><td>Spex</td><td></td></tr><tr><td>Std Wall Lab Tubing, Size S2, Excelon, 1/8&quot; ID x 3/16&quot; OD x 1/32&quot; Wall x 50&#039; Long</td><td>CG-730-003</td><td>Excelon Laboratory</td><td></td></tr><tr><td>Syringe PP/PE without needle, 3 mL</td><td>Z683566</td><td>Millipore Sigma</td><td></td></tr><tr><td>Syringe pump</td><td>55-1199 (95-240)</td><td>Harvard Apparatus</td><td></td></tr><tr><td>Thomas 3-Channel Alarm Timer TM10500</td><td>9371W13</td><td>Thomas Scientific</td><td></td></tr><tr><td>Tube Rack, 12&amp;nbsp;positions, 6 for 5.0&amp;nbsp;mL and 15&amp;nbsp;mL tubes and 6 for 25&amp;nbsp;mL and 50&amp;nbsp;mL tubes, polypropylene, numbered positions, autoclavable</td><td>30119835</td><td>Eppendorf</td><td></td></tr><tr><td>Tube Rack, 12&amp;nbsp;positions, for 5.0&amp;nbsp;mL and 15&amp;nbsp;mL tubes, polypropylene, numbered positions, autoclavable</td><td>30119827</td><td>Eppendorf</td><td></td></tr><tr><td>TYGON R-3603 Laboratory Tubing, I.D. &amp;times; O.D. 1/4 in. &amp;times; 3/8 in.</td><td>T8913 (Millipore Sigma)</td><td>Tygon, Saint-Gobain</td><td></td></tr><tr><td>Vortex-Genie 2</td><td>SI-0236</td><td>Scientific Industries, Inc.</td><td></td></tr><tr><td>VWR Dissecting Forceps with Guide Pin with Curved Tips</td><td>89259-946</td><td>Avantor, by VWR</td><td></td></tr><tr><td>VWR Dissecting Scissors, Sharp Tip, 4&amp;frac12;&quot;</td><td>82027-578</td><td>Avantor, by VWR</td><td></td></tr><tr><td>VWR Incubating Orbital Shaker, Model 3500I</td><td>12620-946</td><td>Avantor, by VWR</td><td></td></tr><tr><td>Zombie Aqua Fixable Viability Kit</td><td>423102</td><td>BioLegend</td><td></td></tr></tbody></table>

flow cytometry-based quantification and analysis of myocardial b-cells

A growing body of evidence shows that B-lymphocytes play an important role in the context of myocardial physiology and myocardial adaptation to injury. However, the literature reports contrasting data on the prevalence of myocardial B-cells. B-cells have been reported to be both among the most prevalent immune cells in the rodent heart or to be present, but at a markedly lower prevalence than myeloid cells, or to be quite rare. Similarly, several groups have described that the number of myocardial B-cells increases after acute ischemic myocardial injury, but one group reported no changes in the number of B-cells of the injured myocardium. Implementation of a shared, reproducible method to assess the prevalence of myocardial B-cells is critical to harmonize observations from different research groups and thus promote the advancement of the study of B-cell myocardial interactions. Based on our experience, the seemingly contrasting observations reported in the literature likely stem from the fact that murine myocardial B-cells are mostly intravascular and connected to the microvascular endothelium. Therefore, the number of B-cells recovered from a murine heart is exquisitely sensitive to the perfusion conditions used to clean the organ and to the method of digestion used. Here we report an optimized protocol that accounts for these two critical variables in a specific way. This protocol empowers reproducible, flow cytometry-based analysis of the number of murine myocardial B-cells and allows researchers to distinguish extravascular vs. intravascular myocardial B-cells.

Once the acquisition is complete and all the events are collected, the data should be analyzed according to standard flow cytometry practice. The focus of the analysis will vary depending on the individual goal of each experiment. In this case, quantification of intravascular and extravascular B-cells was pursued, and it was expressed as the number of cells per mg of tissue.
When using a spectral cytometer, it is recommended to start the analysis with an initial gating to remove debris with sizes that are not in a range corresponding to immune cells, followed by removal of the live-dead stain+ signal. This allows the detection of a clean population of CD45 positive cells corresponding to leukocytes (Figure 1). After the removal of doublets, in a naive uninjured heart, it is expected that about 20% of CD45+ cells will&#160;be CD19+ B-cells (Figure 2A, 18.1% B-cells in this representative experiment). Of these cells, on average 5.5% are located in the interstitial space, while 94.5% are located in the intravascular space (Figure 2B). From the whole B-cell population found in the heart, only about 1.6% corresponds to B1 cells based on the surface marker CD11b (which is typically considered sufficient to classify a murine CD19+ cell as a B1 cell with a high degree of confidence1,34,35), and the other 98.4% corresponds&#160;to B2 cells (Figure 2C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64344/64344fig01.jpg" /> 
Figure 1: Flow cytometry gating strategy used to detect and characterize myocardial-associated B-cells. (A) FSC-A vs. SSC-A: The plot shows all the events collected, and the area selected is drawn to enrich the analysis with events that are in the range of size of leukocytes. (B) Autofluorescence vs. live-dead staining: The selected area corresponds to the lower live-dead signal. The gate drawn removes dead cells and some other cell debris that bind to the live-dead staining. (C) CD45 signal vs. SSC-A: The selected area corresponds to the CD45+ cells (i.e., myocardial leukocytes). (D) FSC-A vs. FSC-H: This gate is drawn to remove cell doublets (unselected events). (E) CD19 signal vs. SSC-A: The selected area corresponds to myocardial B-cells. (F) B220 vs. CD19: The area inside the black square corresponds to the intramyocardial B-cells (CD19+, B220-), and the area inside the red square corresponds to the intravascular B-cell population. (G) CD11b vs. SSC-A: The area inside the black circle corresponds to the CD11b+ (B1 cells), and the area inside the red circle corresponds to the CD11b- (B2 cells). The percentages displayed on each panel represent the average percentage that the population displayed represents from the parental population. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64344/64344fig01large.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/64344/64344fig02.jpg" /> 
Figure 2: Representative plots reporting the average number of B-cells per milligram of myocardial tissue. (A) Average number of CD45+ and CD19+ cells/mg of murine myocardial tissue. The percentage displayed corresponds to the percentage of cells from the parental population (CD45+). (B) Average number of CD19+ cells and the percentage of the B-cells that are in the interstitial (extravascular) space of intravascular spaces. (C) Absolute number and percentage of myocardial B-cells that express markers of B1 or B2 cells. Error bars correspond to the standard error of the mean. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64344/64344fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: This table contains the solutions and reagents needed for the procedure. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64344/64344 Table 1.xlsx" target="_blank">Please click here to download this Table.</a>

Watch this Scientific Journal Video about Flow Cytometry-Based Quantification and Analysis of Myocardial B-Cells at JoVE.com

Flow Cytometry-Based Quantification and Analysis of Myocardial B-Cells

Here we report a protocol for the quantification and differentiation of myocardial B-lymphocytes based on their location in the intravascular or endothelial space using flow cytometry.

A growing body of evidence shows that B-lymphocytes play an important role in the context of myocardial physiology and myocardial adaptation to injury. ...

flow-cytometry-based-quantification-and-analysis-of-myocardial-b-cells

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

2.4K Views.  The Johns Hopkins University School of Medicine. Here we report a protocol for the quantification and differentiation of myocardial B-lymphocytes based on their location in the intravascular or endothelial space using flow cytometry.

Video: Flow Cytometry-Based Quantification and Analysis of Myocardial B-Cells

Flow Cytometry Analysis of Immune Cells Within Murine Aortas

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

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

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

Isolation of Cardiomyocyte Nuclei from Post-mortem Tissue

Identification of cardiomyocyte nuclei has been challenging in tissue sections as most strategies rely only on cytoplasmic marker proteins1. Rare events in cardiac myocytes such as proliferation and apoptosis require an accurate identification of cardiac myocyte nuclei to analyze cellular renewal in homeostasis and in pathological conditions2. Here, we provide a method to isolate cardiomyocyte nuclei from post mortem tissue by density sedimentation and immunolabeling with antibodies against pericentriolar material 1 (PCM-1) and subsequent flow cytometry sorting. This strategy allows a high throughput analysis and isolation with the advantage of working equally well on fresh tissue and frozen archival material. This makes it possible to study material already collected in biobanks. This technique is applicable and tested in a wide range of species and suitable for multiple downstream applications such as carbon-14 dating3, cell-cycle analysis4, visualization of thymidine analogues (e.g. BrdU and IdU)4, transcriptome and epigenetic analysis.

Cardiac nuclei are isolated via density sedimentation and immunolabeled with antibodies against pericentriolar material 1 (PCM-1) to identify and sort cardiomyocyte nuclei by flow cytometry.

Identification of cardiomyocyte nuclei has been challenging in tissue sections as most strategies rely only on cytoplasmic marker proteins1. Rare events ...

Medicine

Measurement of Antibody Effects on Cellular Function of Isolated Cardiomyocytes

Dilated cardiomyopathy (DCM) is one of the main causes for heart failure in younger adults1. Although genetic disposition and exposition to toxic substances are known causes for this disease in about one third of the patients, the origin of DCM remains largely unclear. In a substantial number of these patients, autoantibodies against cardiac epitopes have been detected and are suspected to play a pivotal role in the onset and progression of the disease2,3. The importance of cardiac autoantibodies is underlined by a hemodynamic improvement observed in DCM patients after elimination of autoantibodies by immunoadsorption3-5. A variety of specific antigens have already been identified2,3 and antibodies against these targets may be detected by immunoassays. However, these assays cannot discriminate between stimulating (and therefore functionally effective) and blocking autoantibodies. There is increasing evidence that this distinction is crucial6,7. It can also be assumed that the targets for a number of cardiotropic antibodies are still unidentified and therefore cannot be detected by immunoassays. Therefore, we established a method for the detection of functionally active cardiotropic antibodies, independent of their respective antigen. The background for the method is the high homology usually observed for functional regions of cardiac proteins in between mammals8,9. This suggests that cardiac antibodies directed against human antigens will cross-react with non-human target cells, which allows testing of IgG from DCM patients on adult rat cardiomyocytes. Our method consists of 3 steps: first, IgG is isolated from patient plasma using sepharose coupled anti-IgG antibodies obtained from immunoadsorption columns (PlasmaSelect, Teterow, Germany). Second, adult cardiomyocytes are isolated by collagenase perfusion in a Langendorff perfusion apparatus using a protocol modified from previous works10,11. The obtained cardiomyocytes are attached to laminin-coated chambered coverglasses and stained with Fura-2, a calcium-selective fluorescent dye which can be easily brought into the cell to observe intracellular calcium (Ca2+) contents12. In the last step, the effect of patient IgG on the cell shortening and Ca2+ transients of field stimulated cardiomyocytes is monitored online using a commercial myocyte calcium and contractility monitoring system (IonOptix, Milton, MA, USA) connected to a standard inverse fluorescent microscope.

The presented method offers a way to detect functional effective cardiotropic autoantibodies in the plasma of patients with dilated cardiomyopathy, irrespective of the specific antigen, by analysing the impact of isolated patient immunoglobulin on cellular shortening and intracellular calcium transients in isolated rat cardiomyocytes.

Dilated cardiomyopathy  (DCM) is one of the main causes for heart failure in younger adults1.  Although genetic disposition and exposition to toxic ...

Assessing Cardiac Reprogramming using High Content Imaging Analysis

The goal of this protocol is to describe a method for quantifying induced cardiomyocyte-like cells (iCMs), which are directly reprogrammed in vitro by a reprogramming technique. Cardiac reprogramming provides a strategy to generate new cardiomyocytes. By introducing core cardiogenic transcription factors into fibroblasts; fibroblasts can be converted to iCMs without transition through the pluripotent stem cell state. However, the conversion rate of fibroblasts to iCMs still remains low. Accordingly, there have been numerous additional approaches to enhance cardiac reprogramming efficiency. Most of these studies assessed cardiac reprogramming efficiency using flow cytometry, while at the same time performed immunocytochemistry to visualize iCMs. Thus, at least two separate sets of reprogramming experiments are required to demonstrate the success of iCM reprogramming. In contrast, automated high content imaging analysis will provide both quantification and qualification of iCM reprogramming with a relatively small number of cells. With this method, it is possible to directly assess the quantity and quality of iCMs with a single reprogramming experiment. This approach will be able to facilitate future cardiac reprogramming studies that require large-scale reprogramming experiments such as screening genetic or pharmacological factors for enhancing reprogramming efficiency. In addition, the application of high content imaging analysis protocol is not limited to cardiac reprogramming. It can be applied to reprogramming of other cell lineages as well as any immunostaining experiments which need both quantification and visualization of immunostained cells.

We present a protocol to quantify directly reprogrammed induced cardiomyocyte-like cells (iCMs) in vitro using high content imaging analysis. This method allows us to quantify the efficiency of cardiac reprogramming in an automated manner and to directly visualize iCMs.

The goal of this protocol is to describe a method for quantifying induced cardiomyocyte-like cells (iCMs), which are directly reprogrammed in vitro by a ...

Developmental Biology

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.

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 ...

Multicolor Flow Cytometry-based Quantification of Mitochondria and Lysosomes in T Cells

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular components responsible for supplying cell energy; however, excess mitochondria also produce reactive oxygen species (ROS) that could cause cell death. Therefore, the number of mitochondria must constantly be adjusted to fit the needs of the cells. This dynamic regulation is achieved in part through the function of lysosomes that remove surplus/damaged organelles and macromolecules. Hence, cellular mitochondrial and lysosomal contents are key indicators to evaluate the metabolic adjustment of cells. With the development of probes for organelles, well-characterized lysosome or mitochondria-specific dyes have become available in various formats to label cellular lysosomes and mitochondria. Multicolor flow cytometry is a common tool to profile cell phenotypes, and has the capability to be integrated with other assays. Here, we present a detailed protocol of how to combine organelle-specific dyes with surface markers staining to measure the amount of lysosomes and mitochondria in different T cell populations on a flow cytometer.

This article illustrates a powerful method to quantify mitochondria or lysosomes in living cells. The combination of lysosome- or mitochondria-specific dyes with fluorescently conjugated antibodies against surface markers allows the quantification of these organelles in mixed cell populations, like primary cells harvested from tissue samples, by using multicolor flow cytometry.

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular ...

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 ...

Flow Cytometric Characterization of Murine B Cell Development

Extensive studies have characterized the development and differentiation of murine B cells in secondary lymphoid organs. Antibodies secreted by B cells have been isolated and developed into well-established therapeutics. Validation of murine B cell development, in the context of autoimmune prone mice, or in mice with modified immune systems, is a crucial component of developing or testing therapeutic agents in mice and is an appropriate use of flow cytometry. Well established B cell flow cytometric parameters can be used to evaluate B cell development in the murine peritoneum, bone marrow, and spleen, but a number of best practices must be adhered to. In addition, flow cytometric analysis of B cell compartments should also complement additional readouts of B cell development. Data generated using this technique can further our understanding of wild type, autoimmune prone mouse models as well as humanized mice that can be used to generate antibody or antibody-like molecules as therapeutics.

We describe herein a simple analysis of the heterogeneity of the murine immune B cell compartment in the peritoneum, spleen, and bone marrow tissues by flow cytometry. The protocol can be adapted and extended to other mouse tissues.

Extensive studies have characterized the development and differentiation of murine B cells in secondary lymphoid organs. Antibodies secreted by B cells ...

A Sensitive and Specific Quantitation Method for Determination of Serum Cardiac Myosin Binding Protein-C by Electrochemiluminescence Immunoassay

Biomarkers are becoming increasingly more important in clinical decision-making, as well as basic science. Diagnosing myocardial infarction (MI) is largely driven by detecting cardiac-specific proteins in patients' serum or plasma as an indicator of myocardial injury. Having recently shown that cardiac myosin binding protein-C (cMyBP-C) is detectable in the serum after MI, we have proposed it as a potential biomarker for MI. Biomarkers are typically detected by traditional sandwich enzyme-linked immunosorbent assays. However, this technique requires a large sample volume, has a small dynamic range, and can measure only one protein at a time.	
Here we show a multiplex immunoassay in which three cardiac proteins can be measured simultaneously with high sensitivity. Measuring cMyBP-C in uniplex or together with creatine kinase MB and cardiac troponin I showed comparable sensitivity. This technique uses the Meso Scale Discovery (MSD) method of multiplexing in a 96-well plate combined with electrochemiluminescence for detection. While only small sample volumes are required, high sensitivity and a large dynamic range are achieved. Using this technique, we measured cMyBP-C, creatine kinase MB, and cardiac troponin I levels in serum samples from 16 subjects with MI and compared the results with 16 control subjects. We were able to detect all three markers in these samples and found all three biomarkers to be increased after MI. This technique is, therefore, suitable for the sensitive detection of cardiac biomarkers in serum samples.

Measuring biomarkers in complex biological samples is increasingly guiding clinical decision-making. We describe a highly sensitive method to simultaneously measure cardiac myosin binding protein-C, creatine kinase MB, and cardiac troponin I in serum samples from subjects with myocardial infarction and healthy control subjects.

Biomarkers are becoming increasingly more  important in clinical decision-making, as well as basic science. Diagnosing  myocardial infarction (MI) is ...

Biology

Analysis of Cell Suspensions Isolated from Solid Tissues by Spectral Flow Cytometry

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to the analysis of a few fluorochromes, currently there are dozens of different fluorescent dyes, and up to 14-18 different dyes can be combined at a time. However, several limitations still impair the analytical capabilities. Because of the multiplicity of fluorescent probes, data analysis has become increasingly complex due to the need of large, multi-parametric compensation matrices. Moreover, mutant mouse models carrying fluorescent proteins to detect and trace specific cell types in different tissues have become available, so the analysis (by flow cytometry) of auto-fluorescent cell suspensions obtained from solid organs is required. Spectral flow cytometry, which distinguishes the shapes of emission spectra along a wide range of continuous wavelengths, addresses some of these problems. The data is analyzed with an algorithm that replaces compensation matrices and treats auto-fluorescence as an independent parameter. Thus, spectral flow cytometry should be capable of discriminating fluorochromes with similar emission peaks and can provide a multi-parametric analysis without compensation requirements.
This protocol describes the spectral flow cytometry analysis, allowing for a 21-parameter (19 fluorescent probes) characterization and the management of an auto-fluorescent signal, providing high resolution in minor population detection. The results presented here show that spectral flow cytometry presents advantages in the analysis of cell populations from tissues difficult to characterize in conventional flow cytometry, such as the heart and the intestine. Spectral flow cytometry thus demonstrates the multi-parametric analytical capacity of high-performing conventional flow cytometry without the requirement for compensation and enables auto-fluorescence management.

This article describes spectral cytometry, a new approach in flow cytometry that uses the shapes of emission spectra to distinguish fluorochromes. An algorithm replaces compensations and can treat auto-fluorescence as an independent parameter. This new approach allows for the proper analysis of cells isolated from solid organs.

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to ...