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.

<table><tbody><tr><td>0.5 mL safe-lock Eppendorf tubes</td><td>Eppendorf&amp;nbsp;</td><td>22363611</td><td>0.5 mL microcentrifuge tube</td></tr><tr><td>1.5mL Eppendorf tubes&amp;nbsp;</td><td>Eppendorf&amp;nbsp;</td><td>22364111</td><td>1.5 mL microcentrifuge tube</td></tr><tr><td>15 mL Falcon tubes&amp;nbsp;</td><td>Corning&amp;nbsp;</td><td>352097</td><td>15 mL conical tube</td></tr><tr><td>18 gauge needle</td><td>BD</td><td>305196</td><td></td></tr><tr><td>25 gauge needle</td><td>BD&amp;nbsp;</td><td>305124</td><td></td></tr><tr><td>3 mL syringe</td><td>BD</td><td>309657</td><td></td></tr><tr><td>70 mM MACS SmartStrainer&amp;nbsp;</td><td>Miltenyi Biotec&amp;nbsp;</td><td>130-110-916&amp;nbsp;</td><td>70 mM cell strainer</td></tr><tr><td>96 well U bottom plate&amp;nbsp;</td><td>VWR</td><td>10861-564</td><td></td></tr><tr><td>ACK lysis buffer&amp;nbsp;</td><td>GIBCO&amp;nbsp;</td><td>A1049201</td><td>red blood cell lysis buffer</td></tr><tr><td>Acroprep Advance 96 Well Filter Plate</td><td>Pall Corporation</td><td>8027</td><td>filter plate</td></tr><tr><td>B220</td><td>eBiosciences</td><td>17-0452-82</td><td></td></tr><tr><td>BD CompBead Anti-Mouse Ig/&amp;kappa;</td><td>BD</td><td>552843</td><td>compensation beads</td></tr><tr><td>BD CompBead Anti-Rat Ig/&amp;kappa;</td><td>BD</td><td>552844</td><td>compensation beads</td></tr><tr><td>Bovine Serum Albumin</td><td>Sigma-Aldrich&amp;nbsp;</td><td>A8577</td><td>BSA</td></tr><tr><td>BP-1</td><td>BD</td><td>740882</td><td></td></tr><tr><td>Brilliant Stain Buffer</td><td>BD</td><td>566349</td><td>brilliant stain buffer</td></tr><tr><td>C-Kit</td><td>BD</td><td>564011</td><td></td></tr><tr><td>CD11b</td><td>BD</td><td>563168</td><td></td></tr><tr><td>CD11b</td><td>BioLegend</td><td>101222</td><td></td></tr><tr><td>CD19</td><td>BD</td><td>560143</td><td></td></tr><tr><td>CD21/35</td><td>BD</td><td>562756</td><td></td></tr><tr><td>CD23&amp;nbsp;</td><td>BD</td><td>740216</td><td></td></tr><tr><td>CD24 (HSA)</td><td>BioLegend</td><td>138504</td><td></td></tr><tr><td>CD3</td><td>BD</td><td>561388</td><td></td></tr><tr><td>CD3</td><td>BioLegend</td><td>100214</td><td></td></tr><tr><td>CD43</td><td>BD</td><td>553270</td><td></td></tr><tr><td>CD43</td><td>BioLegend</td><td>121206</td><td></td></tr><tr><td>CD5</td><td>BD</td><td>563194</td><td></td></tr><tr><td>CD93</td><td>BD</td><td>740750</td><td></td></tr><tr><td>CD93</td><td>BioLegend</td><td>136504</td><td></td></tr><tr><td>DPBS (1x)</td><td>ThermoFisher</td><td>14190-144</td><td>DPBS</td></tr><tr><td>eBioscience Fixable Viability Dye eFluor 506</td><td>ThermoFisher</td><td>65-0866-14</td><td>viability dye</td></tr><tr><td>Extended Fine Tip Transfer Pipette</td><td>Samco</td><td>233</td><td>disposable transfer pipette</td></tr><tr><td>FACSymphony A3 flow cytometer</td><td>BD</td><td>custom order</td><td>flow cytometer</td></tr><tr><td>Fc Block, CD16/CD32 (2.4G2)</td><td>BD</td><td>553142</td><td>Fc block</td></tr><tr><td>FlowJo</td><td>Flowjo</td><td></td><td>flow cytometer analysis software</td></tr><tr><td>gentleMACS C Tubes&amp;nbsp;</td><td>Miltenyi Biotec&amp;nbsp;</td><td>130-096-334</td><td>automated dissociation tube&amp;nbsp;</td></tr><tr><td>gentleMACS Octo Dissociator with Heaters&amp;nbsp;</td><td>Miltenyi Biotec&amp;nbsp;</td><td>130-095-937</td><td>tissue dissociator instrument</td></tr><tr><td>GR1 (Ly6C/6G)</td><td>BioLegend</td><td>108422</td><td></td></tr><tr><td>IgD</td><td>BioLegend</td><td>405710</td><td></td></tr><tr><td>IgM</td><td>eBiosciences</td><td>25-5790-82</td><td></td></tr><tr><td>Kappa</td><td>BD</td><td>550003</td><td></td></tr><tr><td>Lambda</td><td>BioLegend</td><td>407308</td><td></td></tr><tr><td>paraformaldehyde, 32% Solution</td><td>Electron Microscopy Sciences</td><td>15714</td><td></td></tr><tr><td>Ter119</td><td>BioLegend</td><td>116220</td><td></td></tr><tr><td>True-Stain Monocyte Blocker</td><td>BioLegend</td><td>426103</td><td>monocyte blocker</td></tr><tr><td>UltraPure EDTA, pH 8.0</td><td>ThermoFisher</td><td>15575038</td><td>EDTA</td></tr><tr><td>Vi-CELL XR</td><td>Beckman Coulter</td><td>731050</td><td>cell counter instrument&amp;nbsp;</td></tr></tbody></table>

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

14.7K 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

Identification Of Erythromyeloid Progenitors And Their Progeny In The Mouse Embryo By Flow Cytometry

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where they act as front line sentinels of infection and tissue damage. While other immune cells are continuously renewed from hematopoietic stem and progenitor cells (HSPC) located in the bone marrow, a lineage of macrophages, known as resident macrophages, have been shown to be self-maintained in tissues without input from bone marrow HSPCs. This lineage is exemplified by microglia in the brain, Kupffer cells in the liver and Langerhans cells in the epidermis among others. The intestinal and colon lamina propria are the only adult tissues devoid of HSPC-independent resident macrophages. Recent investigations have identified that resident macrophages originate from the extra-embryonic yolk sac hematopoiesis from progenitor(s) distinct from fetal hematopoietic stem cells (HSC). Among yolk sac definitive hematopoiesis, erythromyeloid progenitors (EMP) give rise both to erythroid and myeloid cells, in particular resident macrophages. EMP are only generated within the yolk sac between E8.5 and E10.5 days of development and they migrate to the fetal liver as early as circulation is connected, where they expand and differentiate until at least E16.5. Their progeny includes erythrocytes, macrophages, neutrophils and mast cells but only EMP-derived macrophages persist until adulthood in tissues. The transient nature of EMP emergence and the temporal overlap with HSC generation renders the analysis of these progenitors difficult. We have established a tamoxifen-inducible fate mapping protocol based on expression of the macrophage cytokine receptor Csf1r promoter to characterize EMP and EMP-derived cells in vivo by flow cytometry.

While infiltrating macrophages are continuously recruited to adult tissues from circulating precursors, resident macrophages seed their tissue during development, where they are maintained without further input from progenitors. The progenitors for resident macrophages were recently identified. Here, we present methods for the genetic fate mapping of the resident macrophage progenitors.

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where ...

Developmental Biology

Assessing the Development of Murine Plasmacytoid Dendritic Cells in Peyer's Patches Using Adoptive Transfer of Hematopoietic Progenitors

This protocol details a method to analyze the ability of purified hematopoietic progenitors to generate plasmacytoid dendritic cells (pDC) in intestinal Peyer&#39;s patch (PP). Common dendritic cell progenitors (CDPs, lin- c-kitlo CD115+ Flt3+) were purified from the bone marrow of C57BL6 mice by FACS and transferred to recipient mice that lack a significant pDC population in PP; in this case, Ifnar-/- mice were used as the transfer recipients. In some mice, overexpression of the dendritic cell growth factor Flt3 ligand (Flt3L) was enforced prior to adoptive transfer of CDPs, using hydrodynamic gene transfer (HGT) of Flt3L-encoding plasmid. Flt3L overexpression expands DC populations originating from transferred (or endogenous) hematopoietic progenitors. At 7-10 days after progenitor transfer, pDCs that arise from the adoptively transferred progenitors were distinguished from recipient cells on the basis of CD45 marker expression, with pDCs from transferred CDPs being CD45.1+ and recipients being CD45.2+. The ability of transferred CDPs to contribute to the pDC population in PP and to respond to Flt3L was evaluated by flow cytometry of PP single cell suspensions from recipient mice. This method may be used to test whether other progenitor populations are capable of generating PP pDCs. In addition, this approach could be used to examine the role of factors that are predicted to affect pDC development in PP, by transferring progenitor subsets with an appropriate knockdown, knockout or overexpression of the putative developmental factor and/or by manipulating circulating cytokines via HGT. This method may also allow analysis of how PP pDCs affect the frequency or function of other immune subsets in PPs. A unique feature of this method is the use of Ifnar-/- mice, which show severely depleted PP pDCs relative to wild type animals, thus allowing reconstitution of PP pDCs in the absence of confounding effects from lethal irradiation.

This protocol describes experimental procedures to assess the differentiation of plasmacytoid dendritic cells in Peyer&#8217;s patch from common dendritic cell progenitors, using techniques involving FACS-mediated cell isolation, hydrodynamic gene transfer, and flow analysis of immune subsets in Peyer&#8217;s patch.

This protocol details a method to analyze the ability of purified hematopoietic progenitors to generate plasmacytoid dendritic cells (pDC) in intestinal ...

Identification and Isolation of Burst-Forming Unit and Colony-Forming Unit Erythroid Progenitors from Mouse Tissue by Flow Cytometry

Early erythroid progenitors were originally defined by their colony-forming potential in vitro and classified into burst-forming and colony-forming &#34;units&#34; known as BFU-e and CFU-e. Until recently, methods for the direct prospective and complete isolation of pure BFU-e and CFU-e progenitors from freshly isolated adult mouse bone marrow were not available. To address this gap, a single-cell RNA-seq (scRNAseq) dataset of mouse bone marrow was analyzed for the expression of genes coding for cell surface markers. This analysis was combined with cell fate assays, allowing the development of a novel flow cytometric approach that identifies and allows the isolation of complete and pure subsets of BFU-e and CFU-e progenitors in mouse bone marrow or spleen. This approach also identifies other progenitor subsets, including subsets enriched for basophil/mast cell and megakaryocytic potentials. The method consists of labeling fresh bone marrow or spleen cells with antibodies directed at Kit and CD55. Progenitors that express both these markers are then subdivided into five principal populations. Population 1 (P1 or CFU-e, Kit+ CD55+ CD49fmed/low CD105med/high CD71med/high) contains all of the CFU-e progenitors and may be further subdivided into P1-low (CD71med CD150high) and P1-hi (CD71high CD150low), corresponding to early and late CFU-e, respectively; Population 2 (P2 or BFU-e, Kit+ CD55+ CD49fmed/low CD105med/high CD71low CD150high) contains all of the BFU-e progenitors; Population P3 (P3, Kit+ CD55+ CD49fmed/high CD105med/low CD150low CD41low) is enriched for basophil/mast cell progenitors; Population 4 (P4, Kit+ CD55+ CD49fmed/high CD105med/low CD150high CD41+) is enriched for megakaryocytic progenitors; and Population 5 (P5, Kit+ CD55+ CD49fmed/high CD105med/low CD150high CD41-) contains progenitors with erythroid, basophil/mast cell, and megakaryocytic potential (EBMP) and erythroid/ megakaryocytic/ basophil-biased multipotential progenitors (MPPs). This novel approach allows greater precision when analyzing erythroid and other hematopoietic progenitors and also allows for reference to transcriptome information for each flow cytometrically defined population.

Here, we describe a novel flow cytometric method for prospective isolation of early burst-forming unit erythroid (BFU-e) and colony-forming unit erythroid (CFU-e) progenitors directly from fresh mouse bone marrow and spleen. This protocol, developed based on single-cell transcriptomic data, is the first to isolate all the tissue's erythroid progenitors with high purity.

Early erythroid progenitors were originally defined by their colony-forming potential in vitro and classified into burst-forming and colony-forming ...

Identification and Analysis of Mouse Erythroid Progenitors using the CD71/TER119 Flow-cytometric Assay

The study of erythropoiesis aims to understand how red cells are formed from earlier hematopoietic and erythroid progenitors. Specifically, the rate of red cell formation is regulated by the hormone erythropoietin (Epo), whose synthesis is triggered by tissue hypoxia. A threat to adequate tissue oxygenation results in a rapid increase in Epo, driving an increase in erythropoietic rate, a process known as the erythropoietic stress response. The resulting increase in the number of circulating red cells improves tissue oxygen delivery. An efficient erythropoietic stress response is therefore critical to the survival and recovery from physiological and pathological conditions such as high altitude, anemia, hemorrhage, chemotherapy or stem cell transplantation. 
The mouse is a key model for the study of erythropoiesis and its stress response. Mouse definitive (adult-type) erythropoiesis takes place in the fetal liver between embryonic days 12.5 and 15.5, in the neonatal spleen, and in adult spleen and bone marrow. Classical methods of identifying erythroid progenitors in tissue rely on the ability of these cells to give rise to red cell colonies when plated in Epo-containing semi-solid media. Their erythroid precursor progeny are identified based on morphological criteria. Neither of these classical methods allow access to large numbers of differentiation-stage-specific erythroid cells for molecular study. Here we present a flow-cytometric method of identifying and studying differentiation-stage-specific erythroid progenitors and precursors, directly in the context of freshly isolated mouse tissue. The assay relies on the cell-surface markers CD71, Ter119, and on the flow-cytometric 'forward-scatter' parameter, which is a function of cell size. The CD71/Ter119 assay can be used to study erythroid progenitors during their response to erythropoietic stress in vivo, for example, in anemic mice or mice housed in low oxygen conditions. It may also be used to study erythroid progenitors directly in the tissues of genetically modified adult mice or embryos, in order to assess the specific role of the modified molecular pathway in erythropoiesis.

A flow-cytometric method for identification and molecular analysis of differentiation-stage-specific murine erythroid progenitors and precursors, directly in freshly &#x2013;harvested mouse bone marrow, spleen or fetal liver. The assay relies on cell-surface markers CD71, Ter119, and cell size.

The study of erythropoiesis aims to understand how red cells are formed from earlier hematopoietic and erythroid progenitors. Specifically, the rate of ...

Biology

Generation of Large Numbers of Myeloid Progenitors and Dendritic Cell Precursors from Murine Bone Marrow Using a Novel Cell Sorting Strategy

Cultures of monocyte-derived dendritic cells (moDC) generated from mouse bone marrow using Granulocyte-Macrophage Colony Stimulating Factor (GM-CSF) have recently been recognized to be more heterogeneous than previously appreciated. These cultures routinely contain moDC as well monocyte-derived macrophages (moMac), and even some less developed cells such as monocytes. The goal of this protocol is to provide a consistent method for identification and separation of the many cell types present in these cultures as they develop, so that their specific functions may be further investigated. The sorting strategy presented here separates cells first into four populations based on expression of Ly6C and CD115, both of which are expressed transiently by cells as they develop in GM-CSF-driven culture. These four populations include Common myeloid progenitors or CMP (Ly6C-, CD115-), granulocyte/macrophage progenitors or GMP (Ly6C+, CD115-), monocytes (Ly6C+, CD115+), and monocyte-derived macrophages or moMac (Ly6C-, CD115+). CD11c is also added to the sorting strategy to distinguish two populations within the Ly6C-, CD115- population: CMP (CD11c-) and moDC (CD11c+). Finally, two populations may be further distinguished within the Ly6C-, CD115+ population based on the level of MHC class II expression. MoMacs express lower levels of MHC class II, while a monocyte-derived DC precursor (moDP) expresses higher MHC class II. This method allows for the reliable isolation of several developmentally distinct populations in numbers sufficient for a variety of functional and developmental analyses. We highlight one such functional readout, the differential responses of these cell types to stimulation with Pathogen-Associated Molecular Patterns (PAMPs).

Here we provide a method for identifying and isolating large numbers of GM-CSF driven myeloid cells using high speed cell sorting. Five distinct populations (Common myeloid progenitors, granulocyte/macrophage progenitors, monocytes, monocyte-derived macrophages, and monocyte-derived DCs) can be identified based on Ly6C and CD115 expression.

Cultures of monocyte-derived dendritic cells (moDC) generated from mouse bone marrow using Granulocyte-Macrophage Colony Stimulating Factor (GM-CSF) have ...

Identification and Isolation of Oligopotent and Lineage-committed Myeloid Progenitors from Mouse Bone Marrow

Myeloid progenitors that yield neutrophils, monocytes and dendritic cells (DCs) can be identified in and isolated from the bone marrow of mice for hematological and immunological analyses. For example, studies of the cellular and molecular properties of myeloid progenitor populations can reveal mechanisms underlying leukemic transformation, or demonstrate how the immune system responds to pathogen exposure. Previously described flow cytometry strategies for myeloid progenitor identification have enabled significant advances in many fields, but the fractions they identify are very heterogeneous. The most commonly used gating strategies define bone marrow fractions that are enriched for the desired populations, but also contain large numbers of &#34;contaminating&#34; progenitors. Our recent studies have resolved much of this heterogeneity, and the protocol we present here permits the isolation of 6 subpopulations of oligopotent and lineage-committed myeloid progenitors from 2 previously described bone marrow fractions. The protocol describes 3 stages: 1) isolation of bone marrow cells, 2) enrichment for hematopoietic progenitors by magnetic-activated cell sorting (lineage depletion by MACS), and 3) identification of myeloid progenitor subsets by flow cytometry (including fluorescence-activated cell sorting, FACS, if desired). This approach permits progenitor quantification and isolation for a variety of in vitro and in vivo applications, and has already yielded novel insight into pathways and mechanisms of neutrophil, monocyte, and DC differentiation.

We demonstrate how to identify and isolate 6 subsets of myeloid progenitors from murine bone marrow using a combination of magnetic and fluorescence sorting (MACS and FACS). This protocol can be used for in vitro culture assays (methylcellulose or liquid cultures), in vivo adoptive transfer experiments, and RNA/protein analyses.

Myeloid progenitors that yield neutrophils, monocytes and dendritic cells (DCs) can be identified in and isolated from the bone marrow of mice for ...

Analysis of Shear Flow-induced Migration of Murine Marginal Zone B Cells In Vitro

Marginal zone B cells (MZBs) are a population of B cells that reside in the mouse splenic marginal zones that envelop follicles. To reach the follicles, MZBs must migrate up the shear force of blood flow. We present here a method for analyzing this flow-induced MZB migration in vitro. First, MZBs are isolated from the mouse spleen. Second, MZBs are settled on integrin ligands in flow chamber slides, exposed to shear flow, and imaged under a microscope while migrating. Third, images of the migrating MZBs are processed using the MTrack2 automatic cell tracking plugin for ImageJ, and the resulting cell tracks are quantified using the Ibidi chemotaxis tool. The migration data reveal how fast the cells move, how often they change direction, whether the shear flow vector affects their migration direction, and which integrin ligands are involved. Although we use MZBs, the method can easily be adapted for analyzing migration of any leukocyte that responds to the force of shear flow.

Marginal zone B cells (MZBs) respond to the force of shear flow by re-orienting their migration path up the flow. This protocol shows how to record and analyze the migration using a fluidics unit, pump, microscope imaging system, and free software.

Marginal zone B cells (MZBs) are a population of B cells that reside in the mouse splenic marginal zones that envelop follicles. To reach the follicles, ...

Discrimination of Seven Immune Cell Subsets by Two-fluorochrome Flow Cytometry

Immune cell characterization heavily relies on multicolor flow cytometry to identify subpopulations based on differential expression of surface markers. Setup of a classic multicolor panel requires high-end instruments, custom labeled antibodies, and careful study design to minimize spectral overlap. We developed a multiparametric analysis to identify major human immune populations (CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, NK cells and monocytes) in peripheral blood by combining seven lineage markers using only two fluorochromes. Our strategy is based on the observation that lineage markers are constantly expressed in a unique combination by each cell population. Combining this information with a careful titration of the antibodies allows investigators to record five additional markers, expanding the optical limit of most flow cytometers. Head-to-head comparison demonstrated that the vast majority of immune cell populations in peripheral blood can be characterized with comparable accuracy between our method and the classic &#34;one fluorochrome-one marker approach&#34;, although the latter is still more precise for identifying populations such as NKT cells and &#947;&#948; T cells. Combining seven markers using two fluorochromes allows for the analysis of complex immune cell populations and clinical samples on affordable 6-10 fluorochrome flow cytometers, and even on 2-3 fluorochrome field instruments in areas with limited resources. High-end instruments can also benefit from this approach by using extra fluorochromes to accomplish deeper flow cytometry analysis. This approach is also very well suited for screening several cell populations in the case of clinical samples with limited number of cells.

Here, we present a flow cytometric protocol to identify CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, NK cells and monocytes in human peripheral blood by using only two fluorochromes instead of seven. With this approach, five additional markers can be recorded on most flow cytometers.

Immune cell characterization heavily relies on multicolor flow cytometry to identify subpopulations based on differential expression of surface markers. ...

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.

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 Cell Sorting for Different Circulatory B Cell Populations

Source: Tzeng, S. J. <a href="https://app.jove.com/v/54582">The Isolation, Differentiation, and Quantification of Human Antibody-secreting B Cells from Blood: ELISpot as a Functional Readout of Humoral Immunity.</a> J. Vis. Exp. (2016).
In this video, we describe a protocol for the flow cytometry-based sorting of na&#239;ve B cell, memory B cell, and plasmablast plasma cell subpopulations from human peripheral blood B cells. The different B cell subpopulations can be distinguished based on fluorophore-based antibody binding to the CD19, CD27, and CD38 receptors expressed on the B cell surfaces, followed by cell sorting.

Source: Tzeng, S. J. The Isolation, Differentiation, and Quantification of Human Antibody-secreting B Cells from Blood: ELISpot as a Functional Readout of ...