This work is supported by NIH grants R01DK130498, R01DK120639, and R01HL141402

All experiments were conducted in accordance with animal protocols A-1586 and 202200017 approved by the University of Massachusetts Chan Medical School Institutional Animal Care and Use Committee.
NOTE: Two protocols are detailed here: first, flow cytometric analysis (section 1), followed by protocol adjustments for flow cytometric sorting (section 2). The protocol below uses a flow cytometer/sorter with 10 channels. An example setup is provided in Table 1, referred to in step 1.14.5. It is also possible to run this protocol with only nine channels; see the legend in Table 2.
1. Flow cytometric analysis
<ol>
	<li>Euthanize adult (&#62;6 weeks old) Balb/C or C57BL6 mice using an appropriate method approved by your Institutional Animal Care and Use Committee (e.g., CO2 inhalation followed by cervical dislocation). For flow cytometric analysis, bone marrow (BM) from a single mouse will be sufficient. For sorting, the number of mice depends on the downstream applications. The cell yield for each population is given below (step 2.3.3).</li>
	<li>Tissue harvest:
	<ol>
		<li>Prepare 5 mL fluorescence-activated cell sorting (FACS) tubes containing 3 mL of cold (4 &#176;C) staining buffer (&#34;SB5&#34;: phosphate-buffered saline (PBS), 0.2% bovine serum albumin (BSA), 0.08% glucose, 5 mM EDTA) on ice. Use this to place harvested tissues.</li>
		<li>Prepare separate tubes for each tissue dissected from each mouse. Do not pool tissues from multiple mice.</li>
		<li>Harvest both femurs and both tibias. Remove all the muscles from the bones before placing them in the tube with SB5.</li>
		<li>Dissect the spleen through an incision on the left side of the abdomen.</li>
	</ol>
	</li>
	<li>Bone marrow cell extraction:
	<ol>
		<li>Place the freshly dissected femurs and tibias directly into cold staining buffer (SB5) and keep the tubes on ice until ready to extract the bone marrow (BM). Place a clean sterilized mortar on ice in a bucket. Transfer the bones to the mortar along with the SB5 from the tube.</li>
		<li>Snip off approximately 1 mm of the ends of each bone with dissection scissors so that the orifice of the bone becomes just visible.</li>
		<li>Use a 26 G needle and a 3 mL syringe filled with SB5 to flush the marrow out of the bones and collect it into the mortar. Repeat the process 3-5 times using fresh buffer each time until the bones appear colorless.</li>
		<li>Filter the flushed marrow through a 100&#956;m mesh filter and collect the cells in a 50 mL centrifuge tube placed on ice.</li>
		<li>Use the mortar and pestle to crush the flushed bones in 5-10 mL of SB5. Filter the mixture of crushed bones and buffer through the 100 &#181;m cell strainer and pool with cells collected in step 1.3.4.</li>
	</ol>
	</li>
	<li>Spleen cell extraction:
	<ol>
		<li>Set up a 100 &#181;m mesh filter on an empty 50 mL centrifuge tube placed on ice. Place a single spleen on the mesh filter.</li>
		<li>Add 0.5-1 mL of SB5 to the spleen to ensure it does not dry.</li>
		<li>Using the rubber side of a 3 mL or 5 mL syringe plunger, mash the spleen through the mesh. Add more SB5, 5 mL at a time, and continue to mash until all the cells have been collected in the tube.</li>
	</ol>
	</li>
	<li>From this point onward, the bone marrow and splenic cells are treated in the same way.</li>
	<li>Spin the cells down at 900 x g, 4 &#176;C&#8194;for 10 min. Aspirate the supernatant using a vacuum aspiration setup.</li>
	<li>To make a single-cell suspension, resuspend the cells in 2 mL of SB5 and pipet up and down 50 times using a P1000 with a large orifice tip (see Table of Materials). These have wide tip openings to help prevent damage to fragile cells.</li>
	<li>To count the cells, resuspend by pipetting up and down 20 times. Dilute the cells at a ratio of 1:100 in SB5. Mix 10 &#956;L of the diluted cells with 10 &#956;L of trypan blue and count on a hemocytometer and/or cell counter.</li>
	<li>Spin the cells down at 900 x g, 4 &#176;C&#8194;for 10 min. Aspirate the supernatant and resuspend the cells at 33 x 106 live cells/mL in SB5.</li>
	<li>Prepare the Lin-FITC cocktail by mixing equal volumes of each antibody in Table 3.</li>
	<li>Prepare the cells for single color controls and &#34;Fluorescence minus one&#34; (FMO) controls. Carry out steps 1.11-1.12 in parallel with the preparation and staining of the sample cells in step 1.14 (samples for flow cytometric analysis) and/or step 2.1 (flow cytometric sorting).
	<ol>
		<li>Prepare the cells for an unstained cell control by transferring 50 &#181;L (1.6 x 106 cells) of the cell suspension into a FACS tube on ice. Prepare the cells for each single color control (11 tubes) by transferring 50 &#181;L (1.6 x 106 cells) of the cell suspension into a FACS tube on ice.</li>
		<li>Prepare FMO controls for the following colors (Kit, CD41, CD150, CD49f [four tubes]) by transferring 300 &#181;L (10 x 106 cells) of the cell suspension into a separate FACS tube on ice.</li>
		<li>Spin down the cells at 900 x g, 4 &#176;C&#8194;for 10 min and aspirate the supernatant.</li>
	</ol>
	</li>
	<li>Antibody staining of FMOs and single color controls
	<ol>
		<li>Stain the FMOs at 33 x 106 cells/mL (in a total volume of 300 &#181;L) in FACS tubes. For each control, add all of the antibodies except the FMO&#39;s namesake antibody. For example, for the Kit FMO, leave out the Kit antibody. For each antibody, use the dilution and final concentration as in Table 2.</li>
		<li>Stain the single color controls at 20 x 106 cells/mL (in a total volume of 50 &#181;L) in FACS tubes; for each control, add a single antibody at the concentration listed in Table 2.</li>
		<li>Vortex each control tube for 2-3 s at 3,000 rpm (rotations per minute), and then incubate at 4 &#176;C, rocking for 2.5 h in the dark, protected from light. Rock the tubes with an axis of rotation parallel to the length of the tube, between approximately a 10&#176; and a 60&#176; angle from the horizontal, so that the contents do not touch the cap.</li>
		<li>Wash the cells with SB5: Make up the volume of the cell suspension to 4 mL with SB5. Spin the cells down at 900 x g, 4 &#176;C&#8194;for 10 min, and aspirate the supernatant.</li>
	</ol>
	</li>
	<li>Cell resuspension:
	<ol>
		<li>Resuspend the unstained and all the single color controls (except the DAPI control) in 300 &#181;L of SB5 and filter through a 40 &#181;m filter mesh into a new FACS tube.</li>
		<li>Make a DAPI/SB5 solution by diluting the DAPI stock (1 mg/mL in 100% ethanol) at 1:10,000 in SB5.</li>
		<li>Resuspend the FMOs in 1.8 mL of DAPI/SB5 and filter through a 40 &#181;m filter mesh into a new FACS tube</li>
		<li>Resuspend the DAPI single color control in 300 &#181;L of DAPI/SB5 and filter through a 40 &#181;m filter mesh into a new 4 mL FACS tube.</li>
	</ol>
	</li>
	<li>Sample preparation: If the cells are being prepared for flow cytometric analysis, go to step 1.14. If preparing the cells for sorting, skip to step 2.1.</li>
	<li>Stain the flow cytometric analysis samples at 33 x 106 cell/mL in a total volume of 300 &#181;L (10 x 106 cells) in FACS tubes:
	<ol>
		<li>Prepare the antibody master mix. Determine the volume of the master mix by the number of samples, with 300 &#181;L per sample. Make the master mix by diluting all of the antibodies listed in Table 2 at the indicated dilution in SB5. The Rabbit IgG in the master mix serves as an Fc block.</li>
		<li>Vortex each tube for 2-3 s at 3,000 rpm, and then incubate at 4 &#176;C, rocking for 2.5 h in the dark, protected from light.</li>
		<li>Wash the cells with SB5: Make up the volume of the cell suspension to 4 mL with SB5. Spin the cells down at 900 x g, 4 &#176;C&#8194;for 10 min, and aspirate the supernatant.</li>
		<li>Resuspend the samples for flow cytometric analysis in 1.8 mL of DAPI/SB5 as prepared in step 1.12.5.2 and filter through a 40 &#181;m filter mesh into a FACS tube.</li>
		<li>Analyze the samples on a cytometer with 10 channels to accommodate the antibody conjugates in Table 2 and the viability dye DAPI. An example channel setup is provided in Table 1. It is also possible to only use nine channels; see the Table 2 legend.</li>
		<li>During the analysis of the flow cytometry data, use standard spectral compensation approaches with the help of the single color controls and FMO controls as detailed in step 2.4.</li>
	</ol>
	</li>
</ol>
2. Protocol adjustments for flow cytometric sorting
<ol>
	<li>Preparation of Lin-
	<ol>
		<li>Pool all the extracted cells from the same tissue (either BM or spleen) from 10 mice into a 50 mL tube. Resuspend each pooled sample by pipetting using a P1000 with a large orifice tip.</li>
		<li>Spin down the cells at 900 x g, 4 &#176;C&#8194;for 10 min and aspirate the supernatant.</li>
		<li>Prepare single color and FMO controls as described in section 1 using un-enriched cells.</li>
		<li>Enrich Lin- cells for sorting using magnetic bead enrichment of Lin- cells:</li>
		<li>Resuspend the cells at 1 x 108 cells/mL with SB5 and transfer them to a 15 mL tube. Add normal rat serum and the biotinylated antibodies listed in Table 4 and incubate with rocking at 4 &#176;C for 1 h.</li>
		<li>Spin the cells down at 900 x g, 4 &#176;C&#8194;for 10 min and aspirate the supernatant. Resuspend the cells in 15 mL of cold SB5.</li>
		<li>Spin the cells down at 900 x g, 4 &#176;C&#8194;for 10 min and aspirate the supernatant. Resuspend the cells at 1 x 108 cells/mL in cold SB5 and keep them on ice.</li>
		<li>Vortex the magnetic nanobeads at 3,000 rpm for 5-10 s. Take 1 mL of magnetic nanobeads for every 10 mL of cells.</li>
		<li>Wash the nanobeads in a 5 mL FACS tube. Make up the volume to 4 mL with SB5, spin at 900 x g, 4 &#176;C&#8194;for 5 min, and aspirate the supernatant.</li>
		<li>Resuspend the nanobeads in 500 &#181;L of SB5 and mix with the cells prepared in step 2.1.7. Incubate at 4 &#176;C with rocking for 15 min.</li>
		<li>Spin the cells down at 900 x g, 4 &#176;C&#8194;for 10 min and aspirate the supernatant. Resuspend the cells at 1 x 108 cells/mL with SB5.</li>
		<li>Place the tube on a magnet at 4 &#176;C for 5 min.</li>
		<li>Carefully pour the supernatant into a new 15 mL tube.</li>
		<li>Place the tube containing the supernatant from step 2.1.13 on a magnet at 4 &#176;C for 5 min. Carefully pour the supernatant into a new 15 mL tube.</li>
		<li>Spin the cells from the supernatant in step 2.1.14 at 900 x g, 4 &#176;C&#8194;for 10 min, and aspirate the supernatant.</li>
		<li>Resuspend the Lin--enriched cells in 1 mL of SB5.</li>
		<li>Dilute 10 &#181;L of the cells at 1:100 in SB5. Mix 10 &#956;L of the diluted cells with 10 &#956;L of trypan blue and count on a hemocytometer and/or cell counter.</li>
		<li>Based on the cell count in step 2.1.17, resuspend the Lin--enriched cells at 33 x 106 cells/mL.</li>
	</ol>
	</li>
	<li>Staining of enriched cells:
	<ol>
		<li>Stain the enriched cell samples at 33 x 106 cells/mL in FACS tubes by adding all the antibodies listed in Table 2. Vortex each tube for 2-3 s at 3,000 rpm, and then incubate at 4 &#176;C, rocking for 2.5 h in the dark, protected from light.</li>
		<li>Wash the cells with SB5: Make up the volume of the FACS tubes to 4 mL with SB5. Spin the cells down at 900 x g, 4&#176;C&#8194;for 10 min, and remove the supernatant.</li>
		<li>Resuspend the samples for flow cytometric analysis in 4-6 mL of DAPI/SB5 as prepared in step 1.12.5.2 and filter through a 40 &#181;m filter mesh into a new 4 mL FACS tube.</li>
	</ol>
	</li>
	<li>Sample sorting:
	<ol>
		<li>Sort the samples with a large nozzle and low pressure to maximize the CFU-e yield since CFU-e cells are easily damaged under high pressure. For the flow cytometer used in this study, use 14 psi and a 100 &#956;m nozzle or 12 psi with a 130 &#956;m nozzle. Set up the sorting gates as described under step 2.4 below.</li>
		<li>Prepare the collection buffer: Add 20% fetal bovine serum to PBS.</li>
		<li>To check the purities of the sorted populations, re-run a small aliquot from each sorted population in a buffer that contains DAPI, as described in step 1.12.5.2. 
		NOTE: Typically, BM from a total of 10 mice (approximately 1 x 109 cells) yields at least 50,000-100,000 cells for each of the P1-hi, P1-low, and P2 populations with a viability of approximately 70%.</li>
	</ol>
	</li>
	<li>Use the gating strategy shown in Figure 3 for analysis or setting up the sorting gates.
	<ol>
		<li>Use the forward-scatter (FSC)parameter to remove the debris based on size. Use the FSC-height versus FSC-area histogram to exclude cell aggregates and select single cells; repeat with the side-scatter (SSC)-height versus SSC-area histogram.</li>
		<li>Exclude the dead cells by gating out the cells that are positive for the DNA dye DAPI, to which viable cells are impermeable.</li>
		<li>Select Lin-Ter119-Kit+ cells, and from these, select a subpopulation that expresses CD55.</li>
		<li>Subdivide the Lin- Ter119-Kit+CD55+ cells into two principal populations, P6 and P7, based on the expression of CD49f and CD105.</li>
		<li>Based on the expression of CD150 and CD41, subdivide P6 further into P3 (basophil or mast cell progenitors), P4 (megakaryocytic progenitors), and P5 (EBMP and erythroid-biased MPP).</li>
		<li>Based on the expression of CD150 and CD71, subdivide P7 further into P2 (BFU-e), early CFU-e (P1-low), and late CFU-e (P1-hi).</li>
	</ol>
	</li>
</ol>

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J., Sigvardsson, M., Bryder, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Deciphering+developmental+stages+of+adult+myelopoiesis.">Deciphering developmental stages of adult myelopoiesis.</a> Cell Cycle. 7 (6), 706-713 (2008).</li><li>Khoramian Tusi, B., Socolovsky, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-throughput+single-cell+fate+potential+assay+of+murine+hematopoietic+progenitors+in+vitro.">High-throughput single-cell fate potential assay of murine hematopoietic progenitors in vitro.</a> Experimental Hematology. 60, 21-29 (2018).</li><li>Erslev, A. J., Caro, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Erythropoietin+titers+in+response+to+anemia+or+hypoxia.">Erythropoietin titers in response to anemia or hypoxia.</a> Blood Cells. 13 (1-2), 207-216 (1987).</li><li>Koury, M. J., Bondurant, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+molecular+mechanism+of+erythropoietin+action.">The molecular mechanism of erythropoietin action.</a> European Journal of Biochemistry. 210 (3), 649-663 (1992).</li></ol>

The authors have no conflicts of interest to declare.

The ability to prospectively isolate BFU-e and CFU-e progenitors directly from fresh tissue with high purity had previously eluded investigators. Our novel approach, validated using scRNAseq and cell fate assays1,27, now offers the tools to do this.
There are a number of key points for successfully executing both the sorting and the analytical protocols. First, the cells need to be spun at 900 x g to prevent the loss of low-de...

Erythropoiesis may be divided into two principal phases: early erythropoiesis and erythroid terminal differentiation (Figure 1)1,2,3. In early erythropoiesis, hematopoietic stem cells commit to the erythroid lineage and give rise to early erythroid progenitors, which were first identified in the 1970s based on their colony-forming potential in semi-solid medium4,5,6,7,8,9. Broadly, erythroid progenitors are divided into two categories: earlier progenitors that each give rise to a &#34;burst&#34; (a large aggregate of smaller erythroid cell clusters), named &#34;burst-forming unit erythroid&#34; or BFU-e4,5,6; and their progeny, which each form a single, small erythroid cell cluster or colony, named &#34;colony-forming unit erythroid&#34; or CFU-e7,8,9. BFU-e and CFU-e do not yet express terminal erythroid genes and are not morphologically recognizable. After a number of self-renewal or expansion cell divisions, the CFU-e undergoes a transcriptional switch in which erythroid genes such as globins are induced, thereby transitioning into erythroid terminal differentiation (ETD)1,10. During ETD, erythroblasts undergo three to five maturational cell divisions before enucleating to form reticulocytes, which mature into red cells.
Erythroblasts during terminal differentiation were originally classified based on their morphology into proerythroblasts, basophilic, polychromatic, and orthochromatic. The advent of flow cytometry allowed their prospective sorting and isolation based on cell size (measured by forward scatter, FSC) and two cell surface markers, CD71 and Ter11911,12,13 (Figure 1). This and similar flow cytometric approaches14 have revolutionized the investigation of the molecular and cellular aspects of ETD, allowing developmental stage-specific analysis of erythroblasts in vivo and in vitro10,15,16,17,18,19,20. The CD71/Ter119 approach is now used routinely in the analysis of erythroid precursors.
Until recently, a similar, accessible flow cytometric approach for direct, high-purity prospective isolation of CFU-e and BFU-e from mouse tissue has eluded investigators. Instead, investigators have used flow cytometric strategies that isolate only a fraction of these progenitors, often in the presence of non-erythroid cells that co-purify within the same flow cytometric subsets21. Consequently, the investigation of BFU-e and CFU-e was limited to in vitro differentiation systems that derive and amplify BFU-e and CFU-e from earlier bone marrow progenitors. It is then possible to apply flow cytometric strategies that distinguish CFU-e from BFU-e in these erythroid progenitor-enriched cultures22,23. An alternative approach makes use of fetal CFU-e and BFU-e, which are highly enriched in the Ter119-negative fraction of the mouse fetal liver at mid gestation10,24,25. Neither of these approaches, however, allow the investigation of adult BFU-e and CFU-e in their physiological state in vivo. The magnitude of the challenge may be appreciated when recalling that, based on colony formation assays, these cells are present in the adult bone marrow at a frequency of only 0.025% and 0.3%, respectively6.
The protocol described here is a novel flow cytometric approach based on single-cell transcriptomic analysis of freshly harvested Kit+ mouse bone marrow cells (Kit is expressed by all of the early progenitor populations of the bone marrow)1. Our approach contains some cell surface markers that were already in use by Pronk et al.21,26. Single-cell transcriptomes were used to determine combinations of cell surface markers that identify erythroid and other early hematopoietic progenitors (Figure 2). Specifically, the CD55+ fraction of lineage-negative (Lin-) Kit+ cells may be subdivided into five populations, three of which yield contiguous segments of the erythroid trajectory (Figure 2). The transcriptomic identities of each of these populations were confirmed by sorting, followed by scRNAseq and projection of the sorted single-cell transcriptomes back onto the original transcriptomic map (the gene expression in each of the five populations and the entire bone-marrow dataset can be explored in https://kleintools.hms.harvard.edu/paper_websites/tusi_et_al/index.html)1.&#160;The cell fate potential of each of the populations was confirmed using traditional colony formation assays (Figure 2), as well as a novel high-throughput single-cell fate assay1,27. These analyses show that the novel flow cytometric approach results in high-purity isolation of all the BFU-e and CFU-e progenitors of fresh adult bone marrow and spleen. Specifically, population 1 (P1) contains only CFU-e and no other hematopoietic progenitors, and population 2 (P2) contains all of the bone marrow&#39;s BFU-e progenitors and a small number of CFU-e but no other progenitors1. The detailed protocol below is further illustrated with an example experiment in mice that were injected with either saline or with the erythropoiesis-stimulating hormone erythropoietin (Epo).

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			<td>0.5 M EDTA, pH 8.0</td>
			<td>Life Technologies</td>
			<td>15575020</td>
			<td></td>
		</tr>
		<tr>
			<td>1000 &#181;L large orifice tips</td>
			<td>USA sceintific</td>
			<td>1011-9000</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 647 anti-mouse CD55 (DAF) Antibody</td>
			<td>BioLegend</td>
			<td>131806</td>
			<td></td>
		</tr>
		<tr>
			<td>APC/Cyanine7 anti-mouse CD117 (c-kit) Antibody</td>
			<td>BioLegend</td>
			<td>105826</td>
			<td></td>
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		<tr>
			<td>Biotin-CD11b</td>
			<td>BD Biosciences</td>
			<td>557395</td>
			<td>M1/70 (clone)</td>
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			<td>Biotin-CD19</td>
			<td>BD Biosciences</td>
			<td>553784</td>
			<td>1D3 (clone)</td>
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			<td>Biotin-CD4</td>
			<td>BD Biosciences</td>
			<td>BDB553045</td>
			<td>RM4-5 (clone)</td>
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		<tr>
			<td>Biotin-CD8a</td>
			<td>BD Biosciences</td>
			<td>BDB553029</td>
			<td>53-6.7 (clone)</td>
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		<tr>
			<td>Biotin-F4/80</td>
			<td>Biolegend</td>
			<td>123106</td>
			<td>BM8 (clone)</td>
		</tr>
		<tr>
			<td>Biotin-Ly-6G and Ly-6C</td>
			<td>BD Biosciences</td>
			<td>553125</td>
			<td>RB6-8C5 (clone)</td>
		</tr>
		<tr>
			<td>Biotin-TER-119</td>
			<td>BD Biosciences</td>
			<td>553672</td>
			<td>TER-119 (clone)</td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Sigma aldritch</td>
			<td>A1470</td>
			<td></td>
		</tr>
		<tr>
			<td>Brilliant Violet 421 anti-human/mouse CD49f Antibody</td>
			<td>BioLegend</td>
			<td>313624</td>
			<td></td>
		</tr>
		<tr>
			<td>Brilliant Violet 605 anti-mouse CD41 Antibody</td>
			<td>BioLegend</td>
			<td>133921</td>
			<td></td>
		</tr>
		<tr>
			<td>Brilliant Violet 650 anti-mouse CD150 (SLAM) Antibody</td>
			<td>BioLegend</td>
			<td>115931</td>
			<td></td>
		</tr>
		<tr>
			<td>BUV395 Rat Anti-Mouse TER-119/Erythroid Cells</td>
			<td>BD Biosciences</td>
			<td>563827</td>
			<td></td>
		</tr>
		<tr>
			<td>ChromPure Rabbit IgG, whole molecule</td>
			<td>Jackson ImmunoResearch Laboratories</td>
			<td>011-000-003</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI (4&#39;,6-Diamidino-2-Phenylindole, Dihydrochloride)</td>
			<td>Life Technologies</td>
			<td>D1306</td>
			<td></td>
		</tr>
		<tr>
			<td>Digital DIVA hardware and software for LSR II</td>
			<td>BD Biosciences</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>FITC anti-mouse F4/80 Antibody</td>
			<td>BioLegend</td>
			<td>123108</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC Rat Anti-CD11b</td>
			<td>BD Biosciences</td>
			<td>557396</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC Rat Anti-Mouse CD19</td>
			<td>BD Biosciences</td>
			<td>553785</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC Rat Anti-Mouse CD4</td>
			<td>BD Biosciences</td>
			<td>553047</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC Rat Anti-Mouse CD8a</td>
			<td>BD Biosciences</td>
			<td>553031</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC Rat Anti-Mouse Ly-6G and LY-6C</td>
			<td>BD Biosciences</td>
			<td>553127</td>
			<td></td>
		</tr>
		<tr>
			<td>FlowJo software&#160;</td>
			<td>FlowJo</td>
			<td>version 10</td>
			<td>Flow cytometer analysis software</td>
		</tr>
		<tr>
			<td>LSR II digital multiparameter flow cytometer analyzer</td>
			<td>BD Biosciences</td>
			<td></td>
			<td>Flow cytometer&#160;</td>
		</tr>
		<tr>
			<td>NewlineNY Stainless Steel Hand Masher &#38; Bowl, Mortar and Pestle Set</td>
			<td>Amazon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Normal rat serum</td>
			<td>Stem Cell Technologies</td>
			<td>13551</td>
			<td></td>
		</tr>
		<tr>
			<td>PE anti-mouse CD105 Antibody</td>
			<td>BioLegend</td>
			<td>120408</td>
			<td></td>
		</tr>
		<tr>
			<td>PE/Cyanine7 anti-mouse CD71 Antibody</td>
			<td>BioLegend</td>
			<td>113812</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline, 10x Solution</td>
			<td>Fisher scientific</td>
			<td>BP3994</td>
			<td></td>
		</tr>
		<tr>
			<td>Streptavidin Nanobeads</td>
			<td>BioLegend</td>
			<td>480016</td>
			<td>Magnetic beads</td>
		</tr>
	</tbody>
</table>

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.

The protocol describes a flow cytometric approach to identify BFU-es and CFU-es in freshly harvested bone marrow and spleen cells. It starts with harvesting fresh BM and spleen from mice and immediately placing the tissue on ice. All procedures are conducted in the cold to preserve cell viability. Cells are labeled with a &#34;lineage&#34; antibody cocktail that allows the exclusion of all cells expressing markers of differentiated blood lineages (the FITC- Lin cocktail, Table 3, in the case of flow cyto...

Watch this Scientific Journal Video about Identification and Isolation of Burst-Forming Unit and Colony-Forming Unit Erythroid Progenitors from Mouse Tissue by Flow Cytometry at JoVE.com

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

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 ...

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2.7K Views.  UMASS Chan Medical School. 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. 

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

Characterization of Thymic Settling Progenitors in the Mouse Embryo Using In Vivo and In Vitro Assays

Characterizing thymic settling progenitors is important to understand the pre-thymic stages of T cell development, essential to devise strategies for T cell replacement in lymphopenic patients. We studied thymic settling progenitors from murine embryonic day 13 and 18 thymi by two complementary in vitro and in vivo techniques, both based on the &#8220;hanging drop&#8221; method. This method allowed colonizing irradiated fetal thymic lobes with E13 and/or E18 thymic progenitors distinguished by CD45 allotypic markers and thus following their progeny. Colonization with mixed populations allows analyzing cell autonomous differences in biologic properties of the progenitors while colonization with either population removes possible competitive selective pressures. The colonized thymic lobes can also be grafted in immunodeficient male recipient mice allowing the analysis of the mature T cell progeny in vivo, such as population dynamics of the peripheral immune system and colonization of different tissues and organs. Fetal thymic organ cultures revealed that E13 progenitors developed rapidly into all mature CD3+ cells and gave rise to the canonical &#947;&#948; T cell subset, known as dendritic epithelial T cells. In comparison, E18 progenitors have a delayed differentiation and were unable to generate dendritic epithelial T cells. The monitoring of peripheral blood of thymus-grafted CD3-/- mice further showed that E18 thymic settling progenitors generate, with time, larger numbers of mature T cells than their E13 counterparts, a feature that could not be appreciated in the short term fetal thymic organ cultures.

Characterization of Thymic Settling Progenitors in the Mouse Embryo Using In Vivo and In Vitro Assays

This article describes in vivo and in vitro methodology to characterize the thymic settling progenitors by the analysis of the kinetics of generation, phenotype and numbers of their T cell progeny.

Characterizing thymic settling progenitors is important to understand the pre-thymic stages of T cell development, essential to devise strategies for T ...

Step-specific Sorting of Mouse Spermatids by Flow Cytometry

The differentiation of mouse spermatids is one critical process for the production of a functional male gamete with an intact genome to be transmitted to the next generation. So far, molecular studies of this morphological transition have been hampered by the lack of a method allowing adequate separation of these important steps of spermatid differentiation for subsequent analyses. Earlier attempts at proper gating of these cells using flow cytometry may have been difficult because of a peculiar increase in DNA fluorescence in spermatids undergoing chromatin remodeling. Based on this observation, we provide details of a simple flow cytometry scheme, allowing reproducible purification of four populations of mouse spermatids fixed with ethanol, each representing a different state in the nuclear remodeling process. Population enrichment is confirmed using step-specific markers and morphological criterions. The purified spermatids can be used for genomic and proteomic analyses.

We describe a sorting strategy for mouse spermatids using flow cytometry. Spermatids are sorted into four highly pure populations, including round (spermiogenesis steps 1-9), early elongating (spermiogenesis steps 10-12), late elongating (spermiogenesis steps 13-14) and elongated spermatids (spermiogenesis steps 15-16). DNA staining, size and granulosity are used as selection parameters.

The differentiation of mouse spermatids is one critical process for the production of a functional male gamete with an intact genome to be transmitted to ...

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 ...

Isolating and Analyzing Cells of the Pancreas Mesenchyme by Flow Cytometry

The pancreas is comprised of epithelial cells that are required for food digestion and blood glucose regulation. Cells of the pancreas microenvironment, including endothelial, neuronal, and mesenchymal cells were shown to regulate cell differentiation and proliferation in the embryonic pancreas. In the adult, the function and mass of insulin-producing cells were shown to depend on cells in their microenvironment, including pericyte, immune, endothelial, and neuronal cells. Lastly, changes in the pancreas microenvironment were shown to regulate pancreas tumorigenesis. However, the cues underlying these processes are not fully defined. Therefore, characterizing the different cell types that comprise the pancreas microenvironment and profiling their gene expression are crucial to delineate the tissue development and function under normal and diseased states. Here, we describe a method that allows for the isolation of mesenchymal cells from the pancreas of embryonic, neonatal, and adult mice. This method utilizes the enzymatic digestion of mouse pancreatic tissue and the subsequent fluorescence-activated cell sorting (FACS) or flow-cytometric analysis of labeled cells. Cells can be labeled by either immunostaining for surface markers or by the expression of fluorescent proteins. Cell isolation can facilitate the characterization of genes and proteins expressed in cells of the pancreas mesenchyme. This protocol was successful in isolating and culturing highly enriched mesenchymal cell populations from the embryonic, neonatal, and adult mouse pancreas.

Here, we describe a method for the isolation of cells in the pancreas microenvironment from embryonic, neonatal and adult mouse tissue, focusing on the isolation of mesenchymal cells. This method allows profiling of cell gene expression and protein secretion in order to elucidate the extrinsic signals that regulate pancreas development, function, and tumorigenesis.

The pancreas is comprised of epithelial cells that are required for food digestion and blood glucose regulation. Cells of the pancreas microenvironment, ...

Expansion and Adipogenesis Induction of Adipocyte Progenitors from Perivascular Adipose Tissue Isolated by Magnetic Activated Cell Sorting

Expansion of Perivascular Adipose Tissue (PVAT), a major regulator of vascular function through paracrine signaling, is directly related to the development of hypertension during obesity. The extent of hypertrophy and hyperplasia depends on depot location, sex, and the type of Adipocyte Progenitor Cell (APC) phenotypes present. Techniques used for APC and preadipocytes isolation in the last 10 years have drastically improved the accuracy at which individual cells can be identified based on specific cell surface markers. However, isolation of APC and adipocytes can be a challenge due to the fragility of the cell, especially if the intact cell must be retained for cell culture applications.
Magnetic-activated Cell Sorting (MCS) provides a method of isolating greater number of viable APC per weight unit of adipose tissue. APC harvested by MCS can be used for in vitro protocols to expand preadipocytes and differentiate them into adipocytes through use of growth factor cocktails allowing for analysis of the prolific and adipogenic potential retained by the cells. This experiment focused on the aortic and mesenteric PVAT depots, which play key roles in the development of cardiovascular disease during expansion. These protocols describe methods to isolate, expand, and differentiate a defined population of APC. This MCS protocol allows isolation to be used in any experiment where cell sorting is needed with minimal equipment or training. These techniques can aid further experiments to determine the functionality of specific cell populations based on the presence of cell surface markers.

Here we report a method for isolation of Adipocyte Progenitor Cell (APC) populations from Perivascular Adipose Tissue (PVAT) using Magnetic-activated Cell Sorting (MCS). This method allows for an increased isolation of APC per gram of adipose tissue when compared to Fluorescence-Activated Cell Sorting (FACS).

Expansion of Perivascular Adipose Tissue (PVAT), a major regulator of vascular function through paracrine signaling, is directly related to the ...

A Standardized Approach for Multispecies Purification of Mammalian Male Germ Cells by Mechanical Tissue Dissociation and Flow Cytometry

Fluorescence-activated cell sorting (FACS) has been one of the methods of choice to isolate enriched populations of mammalian testicular germ cells. Currently, it allows the discrimination of up to 9 murine germ cell populations with high yield and purity. This high-resolution in discrimination and purification is possible due to unique changes in chromatin structure and quantity throughout spermatogenesis. These patterns can be captured by flow cytometry of male germ cells stained with fluorescent DNA-binding dyes such as Hoechst-33342 (Hoechst). Herein is a detailed description of a recently developed protocol to isolate mammalian testicular germ cells. Briefly, single cell suspensions are generated from testicular tissue by mechanical dissociation, double stained with Hoechst and propidium iodide (PI) and processed by flow cytometry. A serial gating strategy, including the selection of live cells (PI negative) with different DNA content (Hoechst intensity), is used during FACS sorting to discriminate up to 5 germ cell types. These include, with corresponding average purities (determined by microscopy evaluation): spermatogonia (66%), primary (71%) and secondary (85%) spermatocytes, and spermatids (90%), further separated into round (93%) and elongating (87%) subpopulations. Execution of the entire workflow is straightforward, allows the isolation of 4 cell types simultaneously with the appropriate FACS machine, and can be performed in less than 2 h. As reduced processing time is crucial to preserve the physiology of ex vivo cells, this method is ideal for downstream high-throughput studies of male germ cell biology. Moreover, a standardized protocol for multispecies purification of mammalian germ cells eliminates methodological sources of variables and allows a single set of reagents to be used for different animal models.

This work describes the standardization of a method to obtain purified germ cell populations from testicular tissue of different mammalian species. It is a straightforward protocol that combines mechanical testis dissociation, staining with Hoechst-33342 and propidium iodide, and FACS sorting, with wide applications in comparative studies of male reproductive biology.

Fluorescence-activated cell sorting (FACS) has been one of the methods of choice to isolate enriched populations of mammalian testicular germ cells. ...

Isolation, Culture, and Differentiation of Bone Marrow Stromal Cells and Osteoclast Progenitors from Mice

Bone marrow stromal cells (BMSCs) constitute a cell population routinely used as a representation of mesenchymal stem cells in vitro. They reside within the bone marrow cavity alongside hematopoietic stem cells (HSCs), which can give rise to red blood cells, immune progenitors, and osteoclasts. Thus, extractions of cell populations from the bone marrow results in a very heterogeneous mix of various cell populations, which can present challenges in experimental design and confound data interpretation. Several isolation and culture techniques have been developed in laboratories in order to obtain more or less homogeneous populations of BMSCs and HSCs invitro. Here, we present two methods for isolation of BMSCs and HSCs from mouse long bones: one method that yields a mixed population of BMSCs and HSCs and one method that attempts to separate the two cell populations based on adherence. Both methods provide cells suitable for osteogenic and adipogenic differentiation experiments as well as functional assays.

In this article, we present methods to isolate and differentiate bone marrow stromal cells and hematopoietic stem cells from mouse long bones. Two different protocols are presented yielding different cell populations suitable for expansion and differentiation into osteoblasts, adipocytes, and osteoclasts.

Bone marrow stromal cells (BMSCs) constitute a cell population routinely used as a representation of mesenchymal stem cells in vitro. They reside within ...

Isolation and Characterization of Cardiac Mesenchymal Stromal Cells from Endomyocardial Bioptic Samples of Arrhythmogenic Cardiomyopathy Patients

A normal adult heart is composed of several different cell types, among which cardiac mesenchymal stromal cells represent an abundant population. The isolation of these cells offers the possibility of studying their involvement in cardiac diseases, and, in addition, provides a useful primary cell model to investigate biological mechanisms.
Here, the method for the isolation of C-MSC from arrhythmogenic cardiomyopathy patients&#39; bioptic samples is described. The endomyocardial biopsy sampling is guided in the right ventricular areas adjacent to the scar visualized by electro-anatomical mapping. The digestion of the biopsies in collagenase and their plating on a plastic dish in culture medium to allow C-MSC growth is described. The isolated cells can be expanded in culture for several passages. To confirm their mesenchymal phenotype, the description of immuno-phenotypical characterization is provided. C-MSC are able to differentiate into several cell types like adipocytes, chondrocytes, and osteoblasts: in the context of ACM, characterized by adipocyte deposits in patients&#39; hearts, the protocols for the adipogenic differentiation of C-MSC and the characterization of lipid droplet accumulation are described.

In this article, a method to isolate cardiac mesenchymal stromal cells from endomyocardial bioptic samples of arrhythmogenic cardiomyopathy patients is provided. Their characterization and the protocol to boost their adipogenic differentiation are described.

A normal adult heart is composed of several different cell types, among which cardiac mesenchymal stromal cells represent an abundant population. The ...

Direct Lineage Reprogramming of Adult Mouse Fibroblast to Erythroid Progenitors

Erythroid cell commitment and differentiation proceed through activation of a lineage-restricted transcriptional network orchestrated by a group of cell fate determining and maturing factors. We previously set out to define the minimal set of factors necessary for instructing red blood cell development using direct lineage reprogramming of fibroblasts into induced erythroid progenitors/precursors (iEPs). We showed that overexpression of Gata1, Tal1, Lmo2, and c-Myc (GTLM) can rapidly convert murine and human fibroblasts directly to iEPs that resemble bona fide erythroid cells in terms of morphology, phenotype, and gene expression. We intend that iEPs will provide an invaluable tool to study erythropoiesis and cell fate regulation. Here we describe the stepwise process of converting murine tail tip fibroblasts into iEPs via transcription factor-driven direct lineage reprogramming (DLR). In this example, we perform the reprogramming in fibroblasts from erythroid lineage-tracing mice that express the yellow fluorescent protein (YFP) under the control of the erythropoietin receptor gene (EpoR) promoter, enabling visualization of erythroid cell fate induction upon reprogramming. Following this protocol, fibroblasts can be reprogrammed into iEPs within five to eight days. 
While improvements can still be made to the process, we show that GTLM-mediated reprogramming is a rapid and direct process, yielding cells with properties of bona fide erythroid progenitor and precursor cells.

Here we present our protocol for producing induced erythroid progenitors (iEPs) from mouse adult fibroblasts using transcription factor-driven direct lineage reprogramming (DLR).

Erythroid cell commitment and differentiation proceed through activation of a lineage-restricted transcriptional network orchestrated by a group of cell ...

Real-time Live-cell Flow Cytometry to Investigate Calcium Influx, Pore Formation, and Phagocytosis by P2X7 Receptors in Adult Neural Progenitor Cells

Live-cell flow cytometry is increasingly used among cell biologists to quantify biological processes in a living cell culture. This protocol describes a method whereby live-cell flow cytometry is extended upon to analyze the multiple functions of P2X7 receptor activation in real-time. Using a time module installed on a flow cytometer, live-cell functionality can be assessed and plotted over a given time period to explore the kinetics of calcium influx, transmembrane pore formation, and phagocytosis. This simple method is advantageous as all three canonical functions of the P2X7 receptor can be assessed using one machine, and the gathered data plotted over time provides information on the entire live-cell population rather than single-cell recordings often obtained using technically challenging patch-clamp methods. Calcium influx experiments use a calcium indicator dye, while P2X7 pore formation assays rely on ethidium bromide being allowed to pass through the transmembrane pore formed upon high agonist concentrations. Yellow-green (YG) latex beads are utilized to measure phagocytosis. Specific agonists and antagonists are applied to investigate the effects of P2X7 receptor activity. Individually, these methods can be modified to provide quantitative data on any number of calcium channels and purinergic and scavenger receptors. Taken together, they highlight how the use of real-time live-cell flow cytometry is a rapid, cost-effective, reproducible, and quantifiable method to investigate P2X7 receptor function.

Providing single-cell sensitivity, real-time flow cytometry is uniquely suited to quantify multimodal receptor functions of live cultures. Using adult neural progenitor cells, the P2X7 receptor function was assessed via calcium influx detected by calcium indicator dye, transmembrane pore formation by ethidium bromide uptake, and phagocytosis using fluorescent latex beads.

Live-cell flow cytometry is increasingly used among cell biologists to quantify biological processes in a living cell culture. This protocol describes a ...