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

Summary

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.

Abstract

Early erythroid progenitors were originally defined by their colony-forming potential in vitro and classified into burst-forming and colony-forming "units" 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⁺ CD49f^med/low CD105^med/high CD71^med/high) contains all of the CFU-e progenitors and may be further subdivided into P1-low (CD71^med CD150^high) and P1-hi (CD71^high CD150^low), corresponding to early and late CFU-e, respectively; Population 2 (P2 or BFU-e, Kit⁺ CD55⁺ CD49f^med/low CD105^med/high CD71^low CD150^high) contains all of the BFU-e progenitors; Population P3 (P3, Kit⁺ CD55⁺ CD49f^med/high CD105^med/low CD150^low CD41^low) is enriched for basophil/mast cell progenitors; Population 4 (P4, Kit⁺ CD55⁺ CD49f^med/high CD105^med/low CD150^high CD41⁺) is enriched for megakaryocytic progenitors; and Population 5 (P5, Kit⁺ CD55⁺ CD49f^med/high CD105^med/low CD150^high 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.

Introduction

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 medium^4,5,6,7,8,9. Broadly, erythroid progenitors are divided into two categories: earlier progenitors that each give rise to a "burst" (a large aggregate of smaller erythroid cell clusters), named "burst-forming unit erythroid" or BFU-e^4,5,6; and their progeny, which each form a single, small erythroid cell cluster or colony, named "colony-forming unit erythroid" or CFU-e^7,8,9. 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 Ter119^11,12,13 (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 vitro^{10,15,16,17,18,19,20}. 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 cultures^22,23. 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 gestation^10,24,25. 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. 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 assay^1,27. 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'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).

Protocol

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

1. Euthanize adult (>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).
2. Tissue harvest:
   1. Prepare 5 mL fluorescence-activated cell sorting (FACS) tubes containing 3 mL of cold (4 °C) staining buffer ("SB5": phosphate-buffered saline (PBS), 0.2% bovine serum albumin (BSA), 0.08% glucose, 5 mM EDTA) on ice. Use this to place harvested tissues.
   2. Prepare separate tubes for each tissue dissected from each mouse. Do not pool tissues from multiple mice.
   3. Harvest both femurs and both tibias. Remove all the muscles from the bones before placing them in the tube with SB5.
   4. Dissect the spleen through an incision on the left side of the abdomen.
3. Bone marrow cell extraction:
   1. 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.
   2. Snip off approximately 1 mm of the ends of each bone with dissection scissors so that the orifice of the bone becomes just visible.
   3. 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.
   4. Filter the flushed marrow through a 100μm mesh filter and collect the cells in a 50 mL centrifuge tube placed on ice.
   5. 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 µm cell strainer and pool with cells collected in step 1.3.4.
4. Spleen cell extraction:
   1. Set up a 100 µm mesh filter on an empty 50 mL centrifuge tube placed on ice. Place a single spleen on the mesh filter.
   2. Add 0.5-1 mL of SB5 to the spleen to ensure it does not dry.
   3. 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.
5. From this point onward, the bone marrow and splenic cells are treated in the same way.
6. Spin the cells down at 900 x g, 4 °C for 10 min. Aspirate the supernatant using a vacuum aspiration setup.
7. 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.
8. 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 μL of the diluted cells with 10 μL of trypan blue and count on a hemocytometer and/or cell counter.
9. Spin the cells down at 900 x g, 4 °C for 10 min. Aspirate the supernatant and resuspend the cells at 33 x 10⁶ live cells/mL in SB5.
10. Prepare the Lin-FITC cocktail by mixing equal volumes of each antibody in Table 3.
11. Prepare the cells for single color controls and "Fluorescence minus one" (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).
    1. Prepare the cells for an unstained cell control by transferring 50 µL (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 µL (1.6 x 10⁶ cells) of the cell suspension into a FACS tube on ice.
    2. Prepare FMO controls for the following colors (Kit, CD41, CD150, CD49f [four tubes]) by transferring 300 µL (10 x 10⁶ cells) of the cell suspension into a separate FACS tube on ice.
    3. Spin down the cells at 900 x g, 4 °C for 10 min and aspirate the supernatant.
12. Antibody staining of FMOs and single color controls
    1. Stain the FMOs at 33 x 10⁶ cells/mL (in a total volume of 300 µL) in FACS tubes. For each control, add all of the antibodies except the FMO'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.
    2. Stain the single color controls at 20 x 10⁶ cells/mL (in a total volume of 50 µL) in FACS tubes; for each control, add a single antibody at the concentration listed in Table 2.
    3. Vortex each control tube for 2-3 s at 3,000 rpm (rotations per minute), and then incubate at 4 °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° and a 60° angle from the horizontal, so that the contents do not touch the cap.
    4. 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 °C for 10 min, and aspirate the supernatant.
13. Cell resuspension:
    1. Resuspend the unstained and all the single color controls (except the DAPI control) in 300 µL of SB5 and filter through a 40 µm filter mesh into a new FACS tube.
    2. Make a DAPI/SB5 solution by diluting the DAPI stock (1 mg/mL in 100% ethanol) at 1:10,000 in SB5.
    3. Resuspend the FMOs in 1.8 mL of DAPI/SB5 and filter through a 40 µm filter mesh into a new FACS tube
    4. Resuspend the DAPI single color control in 300 µL of DAPI/SB5 and filter through a 40 µm filter mesh into a new 4 mL FACS tube.
14. 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.
15. Stain the flow cytometric analysis samples at 33 x 10⁶ cell/mL in a total volume of 300 µL (10 x 10⁶ cells) in FACS tubes:
    1. Prepare the antibody master mix. Determine the volume of the master mix by the number of samples, with 300 µL 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.
    2. Vortex each tube for 2-3 s at 3,000 rpm, and then incubate at 4 °C, rocking for 2.5 h in the dark, protected from light.
    3. 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 °C for 10 min, and aspirate the supernatant.
    4. 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 µm filter mesh into a FACS tube.
    5. 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.
    6. 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.

2. Protocol adjustments for flow cytometric sorting

1. Preparation of Lin^-
   1. 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.
   2. Spin down the cells at 900 x g, 4 °C for 10 min and aspirate the supernatant.
   3. Prepare single color and FMO controls as described in section 1 using un-enriched cells.
   4. Enrich Lin^- cells for sorting using magnetic bead enrichment of Lin^- cells:
   5. 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 °C for 1 h.
   6. Spin the cells down at 900 x g, 4 °C for 10 min and aspirate the supernatant. Resuspend the cells in 15 mL of cold SB5.
   7. Spin the cells down at 900 x g, 4 °C for 10 min and aspirate the supernatant. Resuspend the cells at 1 x 10⁸ cells/mL in cold SB5 and keep them on ice.
   8. Vortex the magnetic nanobeads at 3,000 rpm for 5-10 s. Take 1 mL of magnetic nanobeads for every 10 mL of cells.
   9. Wash the nanobeads in a 5 mL FACS tube. Make up the volume to 4 mL with SB5, spin at 900 x g, 4 °C for 5 min, and aspirate the supernatant.
   10. Resuspend the nanobeads in 500 µL of SB5 and mix with the cells prepared in step 2.1.7. Incubate at 4 °C with rocking for 15 min.
   11. Spin the cells down at 900 x g, 4 °C for 10 min and aspirate the supernatant. Resuspend the cells at 1 x 10⁸ cells/mL with SB5.
   12. Place the tube on a magnet at 4 °C for 5 min.
   13. Carefully pour the supernatant into a new 15 mL tube.
   14. Place the tube containing the supernatant from step 2.1.13 on a magnet at 4 °C for 5 min. Carefully pour the supernatant into a new 15 mL tube.
   15. Spin the cells from the supernatant in step 2.1.14 at 900 x g, 4 °C for 10 min, and aspirate the supernatant.
   16. Resuspend the Lin^--enriched cells in 1 mL of SB5.
   17. Dilute 10 µL of the cells at 1:100 in SB5. Mix 10 μL of the diluted cells with 10 μL of trypan blue and count on a hemocytometer and/or cell counter.
   18. Based on the cell count in step 2.1.17, resuspend the Lin^--enriched cells at 33 x 10⁶ cells/mL.
2. Staining of enriched cells:
   1. 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 °C, rocking for 2.5 h in the dark, protected from light.
   2. 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°C for 10 min, and remove the supernatant.
   3. 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 µm filter mesh into a new 4 mL FACS tube.
3. Sample sorting:
   1. 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 μm nozzle or 12 psi with a 130 μm nozzle. Set up the sorting gates as described under step 2.4 below.
   2. Prepare the collection buffer: Add 20% fetal bovine serum to PBS.
   3. 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%.
4. Use the gating strategy shown in Figure 3 for analysis or setting up the sorting gates.
   1. 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.
   2. Exclude the dead cells by gating out the cells that are positive for the DNA dye DAPI, to which viable cells are impermeable.
   3. Select Lin^-Ter119^-Kit⁺ cells, and from these, select a subpopulation that expresses CD55.
   4. Subdivide the Lin^- Ter119^-Kit⁺CD55⁺ cells into two principal populations, P6 and P7, based on the expression of CD49f and CD105.
   5. 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).
   6. 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).

Representative Results

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 "lineage" 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 cytometric analysis; or magnetic bead enrichment, Table 4, for lineage-negative cells). Cells are also labeled with antibodies against markers that allow us to distinguish a number of early progenitor populations within the lineage-negative cell fraction. Labeling is followed by either flow cytometric sorting or analysis. The approach to flow cytometric sorting is similar to that of flow cytometric analysis but is preceded by magnetic-bead enrichment for the Lin^- cell fraction to reduce the time and expense of sorting large numbers of cells. When sorting progenitors, BM is pooled from multiple mice, the number depending on the downstream application. 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%.

The key difference between the spleen- and BM-derived populations identified by this protocol is the much lower frequency of the P6 population and its subpopulations, P4/P5/P3, in spleen-derived cells.

As an example of flow cytometric analysis, the effect of Epo administration in mice was examined. Epo binds to and activates the Epo receptor (EpoR), which is essential for both basal and accelerated erythropoiesis under stress conditions like hypoxia. Hypoxia stimulates an increase in blood Epo levels^28,18, in turn promoting the expansion of CFU-e^1,29 and erythroid precursors.

Epo (0.25 U/g body weight) or an equal volume of vehicle (saline) were injected subcutaneously into mice once every 24 h for 3 days. The BM and spleen were harvested at 72 h. Epo stimulation led to the expansion of the early and late CFU-e populations (P1-low and P1-hi) in both the BM and spleen (Figure 4 and Figure 5).

Figure 1: Two principal phases of definitive erythropoiesis. Erythropoiesis may be divided into an early phase, where progenitors are defined based on their colony-forming potential, and a later phase known as erythroid terminal differentiation (ETD), where the erythroblast stage is defined based on morphology. Flow cytometric techniques have permitted the prospective isolation of developmentally specific erythroblast stages using FSC, Ter119, and CD71. By contrast, there has not been an equivalent method for isolating BFU-e and CFU-e progenitors directly from tissue with high purity. Note that only a fraction of the BFU-e colony is shown. The scale bar applies to both the CFU-e and BFU-e colonies. Please click here to view a larger version of this figure.

Figure 2: A new flow cytometric strategy for isolating CFU-e and BFU-e directly from tissue. This manuscript describes a protocol for identifying five new flow cytometric populations, P1 to P5, based on an analysis of the scRNAseq data. (A) Force-directed graph projection ("Spring plot") of fresh Balb/C adult bone marrow single-cell transcriptomes. The colors indicate the predicted cell fate potential based on transcriptomic information and the population-balance analysis (PBA) algorithm¹. The regions of the plot that correspond to each of the flow cytometric populations P1 to P5 are indicated. The regions delineated are hand-drawn approximations of the original data, which may be found in Tusi et al.¹. The correspondence was confirmed by undertaking scRNAseq of each of the sorted P1 to P5 populations¹. E = erythroid; Ba = basophilic or mast-cell progenitors; Meg = megakaryocytic progenitors; Ly = Lymphocytic progenitors; D = dendritic progenitors; M = monocytic progenitors; G = granulocytic progenitors. (B) Colony formation assays, replotted from Tusi et al.¹. The P1 to P5 populations and CD55+ cells were sorted as described in this protocol, and each population was plated for the relevant colony formation assays. Erythroid colonies were scored on the indicated days. UF = unifocal; MF = multi-focal; CFU-MK = colony-forming unit megakaryocytic; CFU-GM = colony-forming unit granulocytic/monocytic; CFU-GEMM = colony-forming unit granulocytic, erythroid, monocytic, megakaryocytic. (C) Cell surface markers that define each of the five populations, as well as their cell fate potential. Cell fate potential is both the result of transcriptomic information and of the cell fate assays, both colony formation and single-cell fate assays^1,27. Of note, the P1 population was split based on the expression of the CD71 marker into P1-hi and P1-low¹. Based on data from Tusi et al.¹. Please click here to view a larger version of this figure.

Figure 3: Flow cytometric gating strategy. The complete gating strategy for populations P1 to P5. Freshly harvested bone marrow (BM) or spleen (SP) from a Balb/C mouse subcutaneously injected with 100 µL of 0.9% sterile saline are first gated for the main population, discarding debris; this is followed by selecting singlets, discarding dead cells, and gating on cells that are Ter119-negative, lineage-negative, and express Kit. Further subdivision of Ter119^-Lin^-Kit⁺CD55⁺ into P6 and P7 is followed by the final subdivision into populations P1 to P5 (also see Figure 2). The Ter119 channel allows the analysis of erythroblasts on the same sample (using the Ter119/CD71 approach¹²). However, a separate Ter119 channel is not essential for this protocol, and the Ter119 antibody may be omitted from the master mix and included as part of the Lin-FITC cocktail instead. The exclusion step for Ter119 positive cells is not shown in this gating strategy. Numbers are percentages of each gate within the shown histogram. Please click here to view a larger version of this figure.

Figure 4: Analysis of bone marrow and spleen early erythroid progenitors following Epo stimulation. Representative flow cytometric plots of freshly harvested bone marrow (BM) or spleen (SP). Balb/C mice were injected subcutaneously with either 0.25 U/g body weight of Epo or with an equivalent volume of saline. Please click here to view a larger version of this figure.

Figure 5: The response of BM and spleen progenitors to Epo in vivo. Summary statistics showing changes in the absolute cell number in populations P1 to P5 following Epo injection in freshly harvested (A) bone marrow and (B) spleen cells; *p = 0.02. Experiment as described in Figure 4. n = 4 mice in each group. To calculate the absolute cell number, the frequency of cells in each population in either the BM or spleen (from Figure 4) was multiplied by the total number of BM or spleen cells harvested from each mouse and divided by the mouse weight in grams. Of note, only early CFU-e in the BM reached statistical significance (p = 0.02), likely a combination of a relatively low Epo dose and a small number of mice in each group. Please click here to view a larger version of this figure.

Detector Array	PMT	Dichroic mirror	Bandpass filter	Fluorophore detected
Blue Laser (488 nm)	A	685LP	695/40
	B	505LP	530/30	Lin-FITC (+Ter119-FITC)
	C		488/10
Violet Laser (405 nm)	A	760LP	800/80
	B	630LP	660/20	CD150-BV650
	C	595LP	605/20	CD41-BV605
	D	475LP	525/20
	E		450/50	CD49f-BV421
Red Laser (640 nm)	A	755LP	785/50	cKit-APC-Cy7
	B	700 LP	720/30
	C		675/14	CD55-AF 647
UV Laser (355 nm)	A	505LP	525/20
	B	420LP	450/50	DAPI
	C		379/28	Ter119-BUV 395
YG Laser (561 nm)	A	755LP	780/60	CD71-PE-Cy7
	B	685LP	710/50
	C	635LP	660/20
	D	600LP	610/20
	E	570LP	580/20	CD105-PE

Table 1: Example of the channel layout in an LSRII cytometer. The panel of antibodies used in the described protocol and their arrangement in the LSRII flow cytometer to collect the data.

Reagent or Antibody	Conjugate	Stock conc. (mg/mL)	Dilution factor	Final conc. in cell suspension (µg/mL)	Clone
CD71	PE/Cy7	0.2	2000	0.1	RI7217
TER-119	BUV395	0.2	100	2	TER-119
CD55 (DAF)	AF647	0.5	50	10	RIKO-3
CD105	PE	0.2	50	4	MJ7/18
CD150 (SLAM)	BV650	0.1	50	2	TC15-12F12.2
CD49f	BV421	0.025	50	0.5	GoH3
CD41	BV605	0.2	100	2	MWReg30
CD117 (cKit)	APC/Cy7	0.2	200	1	2B8
Lin cocktail	FITC	0.08 (for each antibody in the cocktail)	83	1 (for each antibody in the cocktail)
Rabbit IgG		10	50	200

Table 2: Antibody master-mix components. Composition of the antibody master mix. The Ter119 channel allows the analysis of erythroblasts on the same sample (using the Ter119/CD71 approach¹²). However, a separate Ter119 channel is not essential for this protocol, and the Ter119 antibody may be omitted from the master mix and included as part of the Lin-FITC cocktail instead.

Reagent or Antibody	Conjugate	Stock conc. (mg/mL)	Clone
Ly-6G and Ly-6C	FITC	0.5	RB6-8C5
CD11b	FITC	0.5	M1/70
CD19	FITC	0.5	1D3
CD4	FITC	0.5	RM4-5
CD8a	FITC	0.5	53-6.7
F4/80	FITC	0.5	BM8

Table 3: Lin cocktail components. Composition of the Lin cocktail used as part of the antibody master mix (see details of the master mix in Table 2).

Reagent or Antibody	Conjugate	Stock conc. (mg/mL)	Dilution factor	Final conc. in cell suspension (µg/mL)	Clone
Ly-6G and Ly-6C	Biotin	0.5	200	2.5	RB6-8C5
CD11b	Biotin	0.5	200	2.5	M1/70
CD19	Biotin	0.5	200	2.5	1D3
CD4	Biotin	0.5	200	2.5	RM4-5
CD8a	Biotin	0.5	200	2.5	53-6.7
F4/80	Biotin	0.5	200	2.5	BM8
TER-119	Biotin	0.5	200	2.5	TER-119
Normal rat serum			50

Table 4: Antibodies for magnetic-bead enrichment. Antibodies used to enrich the Lin^- fraction of either the BM or spleen prior to cell sorting.

Discussion

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 assays^1,27, 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-density cells. Many cells at the CFU-e stage are large, low-density cells that may otherwise be lost. Second, it is necessary to use large orifice tips when pipetting up and down to help break the cell aggregates and prevent the loss of fragile P1 and P2 cells. This step is important since many CFU-e progenitors are associated with erythroblastic-island macrophages (EBIs) and would be lost if not successfully dissociated. Third, the cells are incubated with an EDTA-containing buffer ("SB5") prior to staining, which helps to dissociate the EBIs. Fourth, during flow cytometric data acquisition, the rate of acquisition should not exceed 7,000 events/s or the recommended rate for the flow cytometer instrument that is being used.

The P1 and P2 populations contain all of the CFU-e and BFU-e progenitors in the bone marrow, respectively, and contain no other progenitor cell types¹. The P1 population is further divided into early CFU-e (P1-low) and late CFU-e (P1-hi) based on the expression of CD71¹. These highly pure flow cytometric populations provide a complete picture of the BFU-e and CFU-e progenitor populations in vivo.

One drawback is that the P2 population does contain a small fraction of early CFU-e progenitors. It does, however, contain all of the BFU-e cells in the BM and no other cell types. The P1 population contains all of the CFU-e in the BM, other than the small fraction that is present in P2. Unlike the P1 and P2 populations, P3 to P5 are highly enriched, but not pure, progenitor populations.

Applying this flow cytometric tool to mouse models of erythropoiesis would now allow the determination of cellular and molecular responses during physiological and pathological perturbations to erythropoiesis. In the shown example, mice were injected with the hormone Epo. This approach may be used in genetically modified mice, for example, in mouse models of hemoglobinopathies, for a more accurate molecular analysis of their altered erythroid progenitors in vivo.

DATA AVAILABILITY:
Additional data associated with the experiments in Figure 4 and Figure 5 are available upon request.

Materials

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