We thank Matthew Sleeman for critical reading of the manuscript. We also thank the Vivarium Operations and Flow Cytometry Core departments at Regeneron for supporting this research.

All mouse studies were overseen and approved by Regeneron&#39;s Institutional Animal Care and Use Committee (IACUC). The experiment was conducted on tissues from three C57BL/6J female mice (17 weeks of age) from Jackson Laboratories. Titrate all antibodies prior to starting the experiment to determine ideal concentration. When using compensation beads for single-color compensation, ensure they stain as bright or brighter than your samples. Keep all buffers, antibodies, and cells on ice or at 4 &#176;C. After the addition of viability dye, perform all steps and incubations at 4&#176;C in either low light or in the dark.
1. Peritoneal cell harvest and single cell isolation
<ol>
	<li>Euthanize the mouse using CO2 or according to approved protocol.</li>
	<li>Lay the mouse on its back, spray with 70% ethanol, and cut outer abdominal skin with scissors, being careful not to cut the peritoneum.</li>
	<li>Inject 3 mL of ice-cold wash buffer (0.5% bovine serum albumin (BSA) in DPBS [vol/vol]) into the peritoneal cavity with a 3 mL syringe fitted with a 25 gauge needle.</li>
	<li>Gently massage the peritoneum with fingertips.</li>
	<li>Repeat steps 1.3 and 1.4.</li>
	<li>Insert a 3 mL syringe fitted with an 18 G needle through the peritoneum, being careful to avoid organs and fat.</li>
	<li>Extract the wash buffer, now containing peritoneal cells, and transfer to 15 mL conical tube on ice.</li>
	<li>Repeat steps 1.3 and 1.4.</li>
	<li>Cut a small hole in the peritoneum while holding up with tweezers.</li>
	<li>Insert a disposable transfer pipette into the hole and collect the remaining wash buffer, once again avoiding fat and organs.</li>
	<li>Transfer the collected remaining peritoneal cells to the 15 mL conical tube on ice. 
	NOTE: Discard sample if blood contamination is evident.</li>
	<li>Incubate the cells on ice until spleen and bone extraction are complete.</li>
	<li>Centrifuge the cells at 300 x g for 8 min at 4 &#730;C. Aspirate the supernatant.</li>
	<li>Resuspend the cell pellet in 1 mL of wash buffer.</li>
	<li>Filter the cells through a 70 &#181;M cell strainer into a clean 15 mL conical tube on ice.</li>
	<li>Determine the cell concentration using a cell counter instrument or hemocytometer.</li>
</ol>
2. Spleen harvest and single cell isolation
<ol>
	<li>Lay the mouse on its belly and cut through the peritoneum on the left backside using clean scissors. Cut out the spleen, removing fat and connective tissue.</li>
	<li>Transfer the spleen to a 1.5 mL microcentrifuge tube containing 1 mL of wash buffer on ice.</li>
	<li>Incubate the spleen on ice until bone extraction is complete.</li>
	<li>Move the spleen to automated dissociation tube with 5 mL of red blood cell lysis buffer. Place the tube on the tissue dissociator instrument and dissociate for 60 s to create a single cell suspension. 
	NOTE: It is also permissible to use other routine methods of obtaining single-cell spleen suspensions such as smashing between frosted glass slides in wash buffer. If another method of dissociation is used, follow the dissociation with centrifugation, aspiration, and then resuspension in 5 mL of red blood cell lysis buffer before continuing to step 2.5.</li>
	<li>Incubate the cells at room temperature for 3 min.</li>
	<li>Add 10 mL of 4 &#176;C wash buffer containing 2mM EDTA.</li>
	<li>Transfer to a clean 15 mL conical tube.</li>
	<li>Centrifuge the cells at 300 x g for 8 min at 4 &#176;C. Aspirate the supernatant.</li>
	<li>Resuspend the cell pellet in 5 mL of 4 &#176;C wash buffer.</li>
	<li>Filter the cells through a 70 &#181;M cell strainer into a clean 15 mL conical tube on ice.</li>
	<li>Determine the cell concentration using a cell counter instrument or hemocytometer.</li>
</ol>
3. BM harvest and single cell isolation
<ol>
	<li>Remove the skin from the lower half of the mouse body. Trim the excess muscle from leg. Remove the entire leg with scissors, being careful not to cut the femur. Clean the femur and tibia by removing remaining muscle, fat, and feet.</li>
	<li>Transfer the bones to a 1.5 mL microcentrifuge tube containing 1 mL of wash buffer on ice.</li>
	<li>Perforate the bottom of a 0.5 mL microcentrifuge tube, leaving a hole small enough for leg bones not to protrude. Insert the 0.5 mL tube into a clean 1.5 mL microcentrifuge tube. Snip off the end of the femur and tibia proximal to the knee and place the cut ends facing down into the 0.5 mL tube.</li>
	<li>Centrifuge the cells at 6,780 x g for 2 min at 4 &#176;C.</li>
	<li>Resuspend the cell pellet in 1 mL of red blood cell lysis buffer and transfer to a 15 mL conical tube containing an additional 3 mL of red blood cell lysis buffer.</li>
	<li>Incubate at room temperature for 3 min.</li>
	<li>Add 10 mL of 4 &#176;C wash buffer containing 2mM EDTA.</li>
	<li>Centrifuge the cells at 300 x g for 8 min at 4 &#176;C. Aspirate the supernatant.</li>
	<li>Resuspend the cell pellet in 3 mL of 4 &#176;C wash buffer.</li>
	<li>Filter cells through a 70 &#181;M cell strainer into a clean 15 mL conical tube on ice.</li>
	<li>Determine the cell concentration using a cell counter instrument or hemocytometer.</li>
</ol>
4. Stain cells and prepare compensation
<ol>
	<li>Aliquot 106 cells of each cell type from each animal to a 96 well U bottom plate.
	<ol>
		<li>Make sure to include enough wells for all samples and controls, including full stain, fluorescence-minus-one (FMO), unstained, and finally the optional single-color compensation for each fluorophore used.</li>
		<li>For the BM maturation panel and the spleen maturation panel, aliquot cells into 2 wells, 106 cells per well, for each full stain sample. For the single-color compensation viability controls, add 2 x 106 cells of each cell type to individual wells.</li>
	</ol>
	</li>
	<li>Centrifuge the plate at 845 x g for 2 min at 4 &#176;C. Decant the supernatant by quickly inverting and flicking the plate over a sink, being careful not to cross-contaminate wells.</li>
	<li>Resuspend the cells in 200 &#181;L of DPBS (without BSA or FBS). This step is important to remove protein before staining with amine-reactive viability dye.</li>
	<li>Repeat steps 4.2 and 4.3.</li>
	<li>Repeat step 4.2.</li>
	<li>Resuspend the cells in 100 &#181;L viability dye diluted 1:1,000 in DPBS. 
	&#8203;NOTE: If using cells for single-color compensation, do not add viability dye to those wells.
	<ol>
		<li>For each stain set, leave several unstained wells for a completely unstained sample and any other controls you might need.</li>
		<li>For each stain set, leave an additional unstained well for the viability FMO control.</li>
		<li>For the single-color viability compensation controls: Resuspend the 2 x 106 cells, aliquoted in step 4.1, in 200 &#181;L of diluted viability dye. Transfer 100 &#181;L of cells to a 1.5 mL microcentrifuge tube, heat cells for 5 min at 65 &#176;C, and transfer the 100 &#181;L of heat-killed cells back to the original well with the 100 &#181;L remaining live cells.</li>
	</ol>
	</li>
	<li>Incubate cells at 4 &#176;C, protected from light, for 30 min.</li>
	<li>Centrifuge the plate at 845 x g for 2 min at 4 &#176;C. Decant the supernatant by quickly inverting and flicking the plate over a sink, being careful not to cross-contaminate wells.</li>
	<li>Resuspend the cells in 200 &#181;L of DPBS (without BSA or FBS).</li>
	<li>Repeat steps 4.8 and 4.9.</li>
	<li>Repeat step 4.8.</li>
	<li>Resuspend the cells in 50 &#181;L of Fc block diluted 1:50 (final concentration=10 &#181;g/mL) in stain buffer (0.5% BSA in DPBS [vol/vol]).
	<ol>
		<li>For peritoneal cells - also add 5 &#181;L of monocyte blocker to reduce non-specific staining.</li>
	</ol>
	</li>
	<li>Incubate the cells at 4 &#176;C, protected from light, for 15 min.</li>
	<li>Prepare full stain master mixes and FMOs in stain buffer for a final volume of 100 &#181;l per 106 cells.Refer to Table 1-Table 4 for the antibody lists. 
	&#8203;NOTE: FMOs are made by including all antibodies in a stain set except one. Prepare an FMO for each antibody in a stain set. When a stain set contains multiple brilliant dyes, substitute 50 &#181;L of brilliant stain buffer for stain buffer per sample</li>
	<li>Without removing Fc block, add 100 &#181;L of full stain mixes and FMOs to selected wells.</li>
	<li>Prepare single-color compensation controls for each antibody in a stain set.
	<ol>
		<li>If using compensation beads follow the manufacture&#39;s directions for use.</li>
		<li>If using cells, add titrated antibody to 106 cells, reserved previously in step 4.6.1 without viability dye, in 100 &#181;L stain buffer. If all cells in the sample are positive for a particular marker, set aside unstained cells to be used when acquiring compensation data on the flow cytometer.</li>
	</ol>
	</li>
	<li>Incubate the cells and beads at 4 &#176;C, protected from light, for 30 min.</li>
	<li>Centrifuge the plate at 845 x g for 2 min at 4 &#176;C. Decant the supernatant by quickly inverting and flicking the plate over a sink, being careful not to cross-contaminate wells.</li>
	<li>Resuspend the cells and beads in 200 &#181;L of stain buffer.</li>
	<li>Repeat steps 4.18 and 4.19 two times.</li>
	<li>Repeat step 4.18.</li>
	<li>To fix the samples for analysis within 48 h, resuspend cells and beads in 200 &#181;L of 2% paraformaldehyde in DPBS. 
	CAUTION: Parafomaldehyde is a serious health hazard and flammable. Refer to the Safty Data Sheet before use.</li>
	<li>Incubate the cells and beads at 4 &#176;C, protected from light, for 30 min.</li>
	<li>Repeat steps 4.18 and 4.19 two times.</li>
	<li>Place a filter plate over a clean 96 well U-bottom plate. Using a multi-pipette, transfer each sample to a well of the filter plate.</li>
	<li>Centrifuge the filter plate-96 well U-bottom plate setup at 845 x g for 2 min at 4 &#176;C. Remove the filter plate and decant the supernatant by quickly inverting and flicking the plate over a sink, being careful not to cross-contaminate wells.</li>
	<li>For the BM and spleen maturation panels resuspend the fully stained cells in 100 &#181;L of stain buffer. Combine the 2 wells for each animal into 1 well. Resuspend the remaining panels, FMOs, and controls in 200 &#181;L of stain buffer.</li>
	<li>Incubate fixed cells and beads at 4 &#176;C, protected from light, overnight.</li>
</ol>
5. Flow cytometric data acquisition
<ol>
	<li>Initialize and QC the flow cytometer as per manufacturer instructions.</li>
	<li>Load the template specific for each panel.</li>
	<li>Prior to recording data, ensure all events for each sample are on scale and visible on the dot plots.</li>
	<li>Record compensation controls for each stain panel using single stain compensations prepared in step 4.16. Set positive and negative gates for each sample. Have the software calculate the compensation matrix.</li>
	<li>Start acquiring the first sample, and ensure gates are set appropriately.</li>
	<li>Set the machine to record at least 50,000 B cell events for the peritoneal B cell panel and spleen Ig&#954; and Ig&#955;&#160;panel; 150,000 B cell events for the BM maturation panel; and 300,000 B cell events for the spleen maturation panel.</li>
	<li>For each stain panel, run and record the fully stained samples for each animal, an unstained sample, and the FMOs.</li>
</ol>
6. Analyze data
<ol>
	<li>Proceed with data analysis using flow cytometry analysis software. Follow gating strategies outlined in Figure 1, Figure 2, Figure 3,&#160;Figure 4.</li>
</ol>

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P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flow+Cytometry.">Flow Cytometry.</a> Encyclopedia of Biomaterials and Biomedical Engineering. , 630-640 (2004).</li><li>Lugli, E., Roederer, M., Cossarizza, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Data+analysis+in+flow+cytometry:+the+future+just+started.">Data analysis in flow cytometry: the future just started.</a> Cytometry A. 77 (7), 705-713 (2010).</li><li>Cossarizza, A., 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=Guidelines+for+the+use+of+flow+cytometry+and+cell+sorting+in+immunological+studies.">Guidelines for the use of flow cytometry and cell sorting in immunological studies.</a> European Journal of Immunology. 47 (10), 1584 (2017).</li><li>Bigos, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Separation+index:+an+easy-to-use+metric+for+evaluation+of+different+configurations+on+the+same+flow+cytometer.">Separation index: an easy-to-use metric for evaluation of different configurations on the same flow cytometer.</a> Current Protocols in Cytometry. , 21 (2007).</li><li>Pillai, S., Mattoo, H., Cariappa, A. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=B+cells+and+autoimmunity.">B cells and autoimmunity.</a> Current Opinion in Immunology. 23 (6), 721-731 (2011).</li></ol>

All authors are employees and shareholders of Regeneron Pharmaceuticals, Inc.

Flow cytometric analysis of lymphoid and non-lymphoid tissues has enabled simultaneous identification and enumeration of B cell sub-populations in mice and humans since the 1980&#39;s. It has been used as a measure of humoral immunity and can be applied further to evaluate B cell functionality. This method takes advantage of reagent availability to assess different stages of B cell maturation in mice and humans, by way of simultaneous analysis of multiple parameters enabling the assessment of B cell heterogeneity, even i...

Monoclonal antibodies have increasingly become the choice therapy for many human diseases as they become part of mainstream medicine1,2. We have previously described genetically engineered mice which efficiently produce antibodies harboring fully human variable regions with mouse IgH constants3,4. Most recently, we have described genetically engineered mice that produce antibody-like molecules that have distinct antigen-binding5. Antibodies are secreted by B cells and form the basis of adaptive humoral immunity. There are two distinct types of B cells, B-1 and B-2. In mammals, B-1 cells originate in the fetal liver and are enriched in mucosal tissues and the pleural and peritoneal cavities after birth, while B-2 cells originate in the fetal liver prior to birth and thereafter in the bone marrow (BM). B-2 cells are enriched in secondary lymphoid organs including the spleen and blood6,7,8. In the BM, B-2 hematopoietic progenitors start to differentiate to pro-B cells upon the initiation of Ig mu heavy chain rearrangement9,10. Successful rearrangement of Ig heavy chain and its assembly into the pre-B cell receptor (pre-BCR), along with signaling and proliferative expansion, leads to differentiation to pre-B cells. After pre-B cells rearrange their Ig kappa (Ig&#954;), or if unproductive, Ig lambda (Ig&#955;) light chains, they pair with &#956; heavy chain, resulting in surface IgM BCR expression. It is important to point out that IgM surface expression is known to be reduced under conditions of autoreactivity, thus contributing to self tolerance in functionally unresponsive or anergic B cells11,12. Immature B cells then enter a transitional stage, where they begin to co-express IgD and migrate from the BM to the spleen. In the spleen, IgD expression increases further and the cells mature into a second stage of transitional B cells, followed by completion of their maturation status and development into either marginal zone (MZ) or follicular (Fol) cells13,14,15. In adult mice, in a non-diseased setting, the number of mature B cells remains constant despite 10-20 million immature B cells being generated daily in the BM. Of these, only three percent enter the pool of mature B cells. The size of the peripheral B cell compartment is constrained by cell death, due in part to several factors including self-reactivity and incomplete maturation16,17,18. Flow cytometric analysis has been extensively used to characterize and enumerate many immune cell sub-compartments in humans and mice. While there are some similarities between human and murine B cell compartments, this protocol applies only to the analysis of murine B cells. This protocol was developed with the purpose of phenotyping genetically engineered mice, to determine whether genetic manipulation would alter B cell development. Flow cytometry has also been hugely popular in many additional applications, including in measuring cell activation, function, proliferation, cycle analysis, DNA content analysis, apoptosis and cell sorting 19,20.
Flow cytometry is the tool of choice to characterize various lymphocyte compartments in mice and humans, including in complex organs such as the spleen, BM and blood. Due to widely available mouse-specific antibody reagents for flow cytometry, this technique can be used to investigate not only cell surface proteins but also intracellular phosphoproteins and cytokines, as well as functional readouts21. Herein we demonstrate how flow cytometry reagents can be used to identify B cells subsets as they mature and differentiate in secondary lymphoid organs. After optimization of staining conditions, sample handling, correct instrument set up and data acquisition, and finally data analysis, a protocol for comprehensive flow cytometric analysis of the B cell compartment in mice can be utilized. Such comprehensive analysis is based on a decades old nomenclature devised by Hardy and colleagues, where developing BM B-2 cells can be divided into different fractions (Fraction) depending on their expression of B220, CD43, BP-1, CD24, IgM and IgD22. Hardy et al., showed that B220+ CD43 BM B cells can be subdivided into four subsets (Fraction A-C&#39;) on the basis of BP-1 and CD24 (30F1) expression, while B220+ CD43-(dim to neg) BM B cells can be resolved into three subsets (Fraction D-F) based on differential expression of IgD and surface IgM23. Fraction A (pre-pro-B cells) are defined as BP-1- CD24 (30F1)-, Fraction B (early pro-B cells) are defined as BP-1- CD24 (30F1)+, Fraction C (late pro-B cells) are defined as BP-1+ CD24 (30F1)+, and Fraction C&#39; (early pre-B cells) are defined as BP-1+ and CD24high. Furthermore, Fraction D (pre-B cells) are defined as B220+ CD43- IgM- B cells, and Fraction E (newly generated B cells, combination of immature and transitional) are defined as B220+ CD43- IgM+ B cells and Fraction F (mature, recirculting B cells) are defined as B220high CD43- IgM+ B cells. In contrast, the majority of na&#239;ve B cells found in the spleen can be divided into mature (B220+ CD93-) B cells and transitional (T1, T2, T3) cells depending on expression of CD93, CD23 and IgM. Mature B cells can be resolved into marginal zone and follicular subsets based on expression of IgM and CD21/CD35, and follicular subsets can be further divided into mature follicular type I and follicular type II B cell subsets depending on the level of their IgM and IgD surface expression24. These splenic B cell populations express predominantly Ig&#954; light chain. Finally, B-1 B cell populations, which originate in the fetal liver and are mainly found in the peritoneal and pleural cavities of adult mice, have been described in the literature. These peritoneal B cells can be distinguished from the previously described B-2 B cells by their lack of CD23 expression. They are then further subdivided into B-1a or B-1b populations, with the former defined by the presence of CD5 and the latter by its absence25. B-1 cell progenitors are abundant in the fetal liver, but are not found in adult BM. While B-1a and B-1b cells originate from different progenitors, they both seed the peritoneal and pleural cavities24. In contrast to B-2 cells, B-1 cells are uniquely capable of self-renewal and are responsible for production of natural IgM antibodies.
Defects in B cell development can arise in many instances, including deficiencies in the components of the BCR26,27, perturbations of signaling molecules that impact BCR signaling strength14,28,29, or disruption of cytokines that modulate B cell survival30,31. Flow cytometry analysis of the lymphoid compartments has contributed to the characterization of the B cell development blocks in these mice and many others. One advantage of flow cytometric analysis of lymphoid compartments is that it offers the ability to make measurements on individual cells obtained from live dissociated tissue. The availability of reagents in an ever-expanding range of fluorophores allows for the simultaneous analysis of multiple parameters and enables the assessment of B cell heterogeneity. Furthermore, enumeration of B cells by flow cytometric analysis complements other immunological assays such as immunohistochemistry methods that visualize cell localization within lymphoid organs, detection of circulating antibody levels as a measure of humoral immunity, as well as two photon microscopy to measure B cell responses in real space and time32.

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			<td>1.5mL Eppendorf tubes&#160;</td>
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			<td>Corning&#160;</td>
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			<td>15 mL conical tube</td>
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			<td>18 gauge needle</td>
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			<td>BD&#160;</td>
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			<td>3 mL syringe</td>
			<td>BD</td>
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			<td>70 mM MACS SmartStrainer&#160;</td>
			<td>Miltenyi Biotec&#160;</td>
			<td>130-110-916&#160;</td>
			<td>70 mM cell strainer</td>
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			<td>96 well U bottom plate&#160;</td>
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			<td>10861-564</td>
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			<td>ACK lysis buffer&#160;</td>
			<td>GIBCO&#160;</td>
			<td>A1049201</td>
			<td>red blood cell lysis buffer</td>
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			<td>Acroprep Advance 96 Well Filter Plate</td>
			<td>Pall Corporation</td>
			<td>8027</td>
			<td>filter plate</td>
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			<td>B220</td>
			<td>eBiosciences</td>
			<td>17-0452-82</td>
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			<td>BD CompBead Anti-Mouse Ig/&#954;</td>
			<td>BD</td>
			<td>552843</td>
			<td>compensation beads</td>
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			<td>BD CompBead Anti-Rat Ig/&#954;</td>
			<td>BD</td>
			<td>552844</td>
			<td>compensation beads</td>
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			<td>Bovine Serum Albumin</td>
			<td>Sigma-Aldrich&#160;</td>
			<td>A8577</td>
			<td>BSA</td>
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			<td>BP-1</td>
			<td>BD</td>
			<td>740882</td>
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			<td>Brilliant Stain Buffer</td>
			<td>BD</td>
			<td>566349</td>
			<td>brilliant stain buffer</td>
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			<td>C-Kit</td>
			<td>BD</td>
			<td>564011</td>
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			<td>CD11b</td>
			<td>BD</td>
			<td>563168</td>
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			<td>CD11b</td>
			<td>BioLegend</td>
			<td>101222</td>
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			<td>CD19</td>
			<td>BD</td>
			<td>560143</td>
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			<td>CD21/35</td>
			<td>BD</td>
			<td>562756</td>
			<td></td>
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			<td>CD23&#160;</td>
			<td>BD</td>
			<td>740216</td>
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			<td>CD24 (HSA)</td>
			<td>BioLegend</td>
			<td>138504</td>
			<td></td>
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			<td>BD</td>
			<td>561388</td>
			<td></td>
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			<td>CD3</td>
			<td>BioLegend</td>
			<td>100214</td>
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			<td>CD43</td>
			<td>BD</td>
			<td>553270</td>
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			<td>CD43</td>
			<td>BioLegend</td>
			<td>121206</td>
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			<td>BD</td>
			<td>563194</td>
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			<td>CD93</td>
			<td>BD</td>
			<td>740750</td>
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			<td>BioLegend</td>
			<td>136504</td>
			<td></td>
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			<td>DPBS (1x)</td>
			<td>ThermoFisher</td>
			<td>14190-144</td>
			<td>DPBS</td>
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			<td>eBioscience Fixable Viability Dye eFluor 506</td>
			<td>ThermoFisher</td>
			<td>65-0866-14</td>
			<td>viability dye</td>
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			<td>Extended Fine Tip Transfer Pipette</td>
			<td>Samco</td>
			<td>233</td>
			<td>disposable transfer pipette</td>
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			<td>FACSymphony A3 flow cytometer</td>
			<td>BD</td>
			<td>custom order</td>
			<td>flow cytometer</td>
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			<td>Fc Block, CD16/CD32 (2.4G2)</td>
			<td>BD</td>
			<td>553142</td>
			<td>Fc block</td>
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			<td>FlowJo</td>
			<td>Flowjo</td>
			<td></td>
			<td>flow cytometer analysis software</td>
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			<td>gentleMACS C Tubes&#160;</td>
			<td>Miltenyi Biotec&#160;</td>
			<td>130-096-334</td>
			<td>automated dissociation tube&#160;</td>
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			<td>gentleMACS Octo Dissociator with Heaters&#160;</td>
			<td>Miltenyi Biotec&#160;</td>
			<td>130-095-937</td>
			<td>tissue dissociator instrument</td>
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			<td>GR1 (Ly6C/6G)</td>
			<td>BioLegend</td>
			<td>108422</td>
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			<td>BioLegend</td>
			<td>405710</td>
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			<td>IgM</td>
			<td>eBiosciences</td>
			<td>25-5790-82</td>
			<td></td>
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			<td>Kappa</td>
			<td>BD</td>
			<td>550003</td>
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			<td>Lambda</td>
			<td>BioLegend</td>
			<td>407308</td>
			<td></td>
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			<td>paraformaldehyde, 32% Solution</td>
			<td>Electron Microscopy Sciences</td>
			<td>15714</td>
			<td></td>
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			<td>Ter119</td>
			<td>BioLegend</td>
			<td>116220</td>
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			<td>True-Stain Monocyte Blocker</td>
			<td>BioLegend</td>
			<td>426103</td>
			<td>monocyte blocker</td>
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			<td>UltraPure EDTA, pH 8.0</td>
			<td>ThermoFisher</td>
			<td>15575038</td>
			<td>EDTA</td>
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			<td>Vi-CELL XR</td>
			<td>Beckman Coulter</td>
			<td>731050</td>
			<td>cell counter instrument&#160;</td>
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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.

Here we present the gating strategy for characterizing B cell development in mouse peritoneum, BM and spleen. The basis of the analysis is formed around the concept of staining with viability dye, then gating out doublets based on the Forward-Scatter-Area (FSC-A) and Forward-Scatter-Height (FSC-H), and finally gating out debris by selecting cells according to their FSC-A and Side-Scatter-Area (SSC-A) characteristics, referred to here as the size gate, which are reflective of relative cell...

Watch this Scientific Journal Video about Flow Cytometric Characterization of Murine B Cell Development at JoVE.com

Flow Cytometric Characterization of Murine B Cell Development

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 ...

flow-cytometric-characterization-of-murine-b-cell-development

Research

JoVE Journal

Immunology and Infection

14.5K Views.  Regeneron Pharmaceuticals, Inc.. 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.

Video: Flow Cytometric Characterization of Murine B Cell Development

Murine Model of CD40-activation of B cells

Research on B cells has shown that CD40 activation improves their antigen presentation capacity. When stimulated with interleukin-4 and CD40 ligand (CD40L), human B cells can be expanded without difficulties from small amounts of peripheral blood within 14 days to very large amounts of highly-pure CD40-B cells (&gt;109 cells per patient) from healthy donors as well as cancer patients1-4. CD40-B cells express important lymph node homing molecules and can attract T cells in vitro5. Furthermore they efficiently take up, process and present antigens to T cells6,7. CD40-B cells were shown to not only prime na&iacute;ve, but also expand memory T cells8,9. Therefore CD40-activated B cells (CD40-B cells) have been studied as an alternative source of immuno-stimulatory antigen-presenting cells (APC) for cell-based immunotherapy1,5,10. In order to further study whether CD40-B cells induce effective T cell responses in vivo and to study the underlying mechanism we established a cell culture system for the generation of murine CD40-activated B cells. Using splenocytes or purified B cells from C57BL/6 mice for CD40-activation, optimal conditions were identified as follows: Starting from splenocytes of C57BL/6 mice (haplotype H-2b) lymphocytes are purified by density gradient centrifugation and co-cultured with HeLa cells expressing recombinant murine CD40 ligand (tmuCD40L HeLa)11. Cells are recultured every 3-4 days and key components such as CD40L, interleukin-4, -Mercaptoethanol and cyclosporin A are replenished. In this protocol we demonstrate how to obtain fully activated murine CD40-B cells (mCD40B) with similar APC-phenotype to human CD40-B cells (Fig 1a,b). CD40-stimulation leads to a rapid outgrowth and expansion of highly pure (&gt;90%) CD19+ B cells within 14 days of cell culture (Fig 1c,d). To avoid contamination with non-transfected cells, expression of the murine CD40 ligand on the transfectants has to be controlled regularly (Fig 2). Murine CD40-activated B cells can be used to study B-cell activation and differentiation as well as to investigate their potential to function as APC in vitro and in vivo. Moreover, they represent a promising tool for establishing therapeutic or preventive vaccination against tumors and will help to answer questions regarding safety and immunogenicity of this approach12.

In this video, we demonstrate the procedure of CD40-activation and expansion of murine B cells from splenocytes of C57BL/6 mice, which can be used as a model antigen-presenting cell (APC) to study induction of immunity.

Research on B cells has shown that CD40 activation improves their antigen presentation capacity. When stimulated with interleukin-4 and CD40 ligand ...

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 ...

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 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 ...

Isolation and Flow Cytometric Analysis of Glioma-infiltrating Peripheral Blood Mononuclear Cells

Our laboratory has recently demonstrated that natural killer (NK) cells are capable of eradicating orthotopically implanted mouse GL26 and rat CNS-1 malignant gliomas soon after intracranial engraftment if the cancer cells are rendered deficient in their expression of the &#946;-galactoside-binding lectin galectin-1 (gal-1). More recent work now shows that a population of Gr-1+/CD11b+ myeloid cells is critical to this effect. To better understand the mechanisms by which NK and myeloid cells cooperate to confer gal-1-deficient tumor rejection we have developed a comprehensive protocol for the isolation and analysis of glioma-infiltrating peripheral blood mononuclear cells (PBMC). The method is demonstrated here by comparing PBMC infiltration into the tumor microenvironment of gal-1-expressing GL26 gliomas with those rendered gal-1-deficient via shRNA knockdown. The protocol begins with a description of&#160;how to culture and prepare GL26 cells for inoculation into the syngeneic C57BL/6J mouse brain. It then explains the steps involved in the isolation and flow cytometric analysis of glioma-infiltrating PBMCs from the early brain tumor microenvironment. The method is adaptable to a number of in vivo experimental designs in which temporal data on immune infiltration into the brain is required. The method is sensitive and highly reproducible, as glioma-infiltrating PBMCs can be isolated from intracranial tumors as soon as 24 hr post-tumor engraftment with similar cell counts observed from time point matched tumors throughout independent experiments. A single experimentalist can perform the method from brain harvesting to flow cytometric analysis of glioma-infiltrating PBMCs in roughly 4-6 hr depending on the number of samples to be analyzed. Alternative glioma models and/or cell-specific detection antibodies may also be used at the experimentalists&#8217; discretion to assess the infiltration of several other immune cell types of interest without the need for alterations to the overall procedure.

Presented here is a straightforward method for the isolation and flow cytometric analysis of glioma-infiltrating peripheral blood mononuclear cells that yields time-dependent quantitative data on the number and activation status of immune cells entering the early brain tumor microenvironment.

Our laboratory has recently demonstrated that natural killer (NK) cells are capable of eradicating orthotopically implanted mouse GL26 and rat CNS-1 ...

Isolation and Flow Cytometric Characterization of Murine Small Intestinal Lymphocytes

The intestines &#8212; which contain the largest number of immune cells of any organ in the body &#8212; are constantly exposed to foreign antigens, both microbial and dietary. Given an increasing understanding that these luminal antigens help shape the immune response and that education of immune cells within the intestine is critical for a number of systemic diseases, there has been increased interest in characterizing the intestinal immune system. However, many published protocols are arduous and time-consuming. We present here a simplified protocol for the isolation of lymphocytes from the small-intestinal lamina propria, intraepithelial layer, and Peyer&#39;s patches that is rapid, reproducible, and does not require laborious Percoll gradients. Although the protocol focuses on the small intestine, it is also suitable for analysis of the colon. Moreover, we highlight some aspects that may need additional optimization depending on the specific scientific question. This approach results in the isolation of large numbers of viable lymphocytes that can subsequently be used for flow cytometric analysis or alternate means of characterization.

There is growing interest in the quantitative characterization of intestinal lymphocytes owing to increasing recognition that these cells play a critical role in a variety of intestinal and systemic diseases. In this protocol, we describe how to isolate single cell populations from different small-intestinal compartments for subsequent flow cytometric characterization.

The intestines &#8212; which contain the largest number of immune cells of any organ in the body &#8212; are constantly exposed to foreign antigens, both ...

A Murine Model of Group B Streptococcus Vaginal Colonization

Streptococcus agalactiae (group B Streptococcus, GBS), is a Gram-positive, asymptomatic colonizer of the human gastrointestinal tract and vaginal tract of 10 - 30% of adults. In immune-compromised individuals, including neonates, pregnant women, and the elderly, GBS may switch to an invasive pathogen causing sepsis, arthritis, pneumonia, and meningitis. Because GBS is a leading bacterial pathogen of neonates, current prophylaxis is comprised of late gestation screening for GBS vaginal colonization and subsequent peripartum antibiotic treatment of GBS-positive mothers. Heavy GBS vaginal burden is a risk factor for both neonatal disease and colonization. Unfortunately, little is known about the host and bacterial factors that promote or permit GBS vaginal colonization. This protocol describes a technique for establishing persistent GBS vaginal colonization using a single &#946;-estradiol pre-treatment and daily sampling to determine bacterial load. It further details methods to administer additional therapies or reagents of interest and to collect vaginal lavage fluid and reproductive tract tissues. This mouse model will further the understanding of the GBS-host interaction within the vaginal environment, which will lead to potential therapeutic targets to control maternal vaginal colonization during pregnancy and to prevent transmission to the vulnerable newborn. It will also be of interest to increase our understanding of general bacterial-host interactions in the female vaginal tract.

A Murine Model of Group B Streptococcus Vaginal Colonization

The purpose of this protocol is to imitate human group B Streptococcus (GBS) vaginal colonization in a murine model. This method may be used to investigate host immune responses and bacterial factors contributing to GBS vaginal persistence, as well as to test therapeutic strategies.

Streptococcus agalactiae (group B Streptococcus, GBS), is a Gram-positive, asymptomatic colonizer of the human gastrointestinal tract and vaginal tract of ...

Flow Cytometric Analysis of Particle-bound Bet v 1 Allergen in PM10

Flow cytometry is a method widely used to quantify suspended solids such as cells or bacteria in a size range from 0.5 to several tens of micrometers in diameter. In addition to a characterization of forward and sideward scatter properties, it enables the use of fluorescent labeled markers like antibodies to detect respective structures. Using indirect antibody staining, flow cytometry is employed here to quantify birch pollen allergen (precisely Bet v 1)-loaded particles of 0.5 to 10 &#181;m in diameter in inhalable particulate matter (PM10, particle size &#8804;10 &#181;m in diameter). PM10 particles may act as carriers of adsorbed allergens possibly transporting them to the lower respiratory tract, where they could trigger allergic reactions.
So far the allergen content of PM10 has been studied by means of enzyme linked immunosorbent assays (ELISAs) and scanning electron microscopy. ELISA measures the dissolved and not the particle-bound allergen. Compared to scanning electron microscopy, which can visualize allergen-loaded particles, flow cytometry may additionally quantify them. As allergen content of ambient air can deviate from birch pollen count, allergic symptoms might perhaps correlate better with allergen exposure than with pollen count. In conjunction with clinical data, the presented method offers the opportunity to test in future experiments whether allergic reactions to birch pollen antigens are associated with the Bet v 1 allergen content of PM10 particles &#62;0.5 &#181;m.

Here, we present a protocol to quantify allergen-loaded particles by flow cytometry. Ambient particulate matter particles may act as carriers of adsorbed allergens. We show here that flow cytometry, a method widely used to characterize suspended solids &#62;0.5 &#181;m in diameter, can be used to measure these allergen-loaded particles.

Flow cytometry is a method widely used to quantify suspended solids such as cells or bacteria in a size range from 0.5 to several tens of micrometers in ...

Flow Cytometric Analysis of Natural Killer Cell Lytic Activity in Human Whole Blood

NK cell cytotoxicity is a widely used measure to determine the effect of outside intervention on NK cell function. However, the accuracy and reproducibility of this assay can be considered unstable, either because of user&#39;s errors or because of the sensitivity of NK cells to experimental manipulation. To eliminate these issues, a workflow that reduces them to a minimum was established and is presented here. To illustrate, we obtained blood samples, at various time points, from runners (n = 4) that were submitted to an intense bout of exercise. First, NK cells are simultaneously identified and isolated through CD56 tagging and magnetic-based sorting, directly from whole blood and from as little as one milliliter. The sorted NK cells are removed of any reagent or capping antibodies. They can be counted in order to establish an accurate NK cell count per milliliter of blood. Secondly, the sorted NK cells (effectors cells or E) can be mixed with 3,3&#39;-Diotadecyloxacarbocyanine Perchlorate (DiO) tagged K562 cells (target cells or T) at an assay-optimal 1:5 T:E ratio, and analyzed using an imaging flow-cytometer that allows for the visualization of each event and the elimination of any false positive or false negatives (such as doublets or effector cells). This workflow can be completed in about 4 h, and allows for very stable results even when working with human samples. When available, research teams can test several experimental interventions in human subjects, and compare measurements across several time points without compromising the data&#39;s integrity.

This work describes an advanced workflow for the accurate and fast determination of NK (Natural Killer) cell count and NK cell cytotoxicity in human blood samples.

NK cell cytotoxicity is a widely used measure to determine the effect of outside intervention on NK cell function. However, the accuracy and ...

Isolation and Flow Cytometric Analysis of Human Endocervical Gamma Delta T Cells

The female reproductive tract (FRT) mucosal immune system serves as the first line of defense. Better knowledge of the genital mucosa is therefore essential for understanding pathogenicity of different pathogens including HIV. Gamma delta (GD) T cells are the prototype of &#39;unconventional&#39; T cells and represent a relatively small subset of T cells defined by their expression of heterodimeric T-cell receptors (TCRs) composed of gamma and delta chains. This sets them apart from the classical and much better known CD4+ helper T cells and CD8+ cytotoxic T cells that are defined by alpha-beta TCRs. GD T cells often show tissue-specific localization and are enriched in epithelium. GD T cells orchestrate immune responses in inflammation, tumor surveillance, infectious disease, and autoimmunity.
Here, we present a method to reproducibly isolate and analyze human endocervical intraepithelial GD T lymphocytes. We have used endocervical cytobrush samples from women participating in the Women&#39;s Interagency HIV Infection Study (WIHS). Knowledge about GD T cells interactions during conditions in which there is an insult to the vaginal mucosal could be applied to any clinical study in which mucosal vulnerability is addressed, including the development of vaginal microbicides.In addition, knowledge about mucosal GD T cell responses has potential for application of GD T cell-based immune therapy in treating infectious diseases.

We describe here an isolation method to obtain human endocervical intraepithelial lymphocytes for the analysis of intraepithelial gamma delta T cells. This protocol can be extended for the purification of endocervical gamma delta T cells by magnetic beads or by cell sorting.