C.P., S.B., and K.RP. were supported by the HARP program, the Barts Charity (G-002167), the Kay Kendall Leukaemia fund (KKL1149) and the Academy of Medical Sciences (SBF004\1099). We thank Dr. Pantelitsa Protopapa for the MGG staining and microscopy acquisition. We also acknowledge the flow cytometry and microscopy facilities of the Barts Cancer Institute, Queen Mary University of London.&#160;We thank the UK Charity Anthony Nolan for providing us with the umbilical cord blood units used in this manuscript.

Research on primary samples was performed in compliance with institutional guidelines (REC reference: 21/EE/0133; IRAS project ID: 283103).
1. Lentivirus production
NOTE Lentiviruses must be produced in a laboratory of biosafety level L2 under sterile conditions. All required PPE for this lab must be worn, including double gloves. Contaminated material is decontaminated with virucide disinfectant prior to disposal in an autoclaved bin. Here, a shRNA scramble (control) harboring an EGFP reporter gene is used to illustrate the process.
Day 1
<ol>
	<li>Warm gelatin at 37 &#176;C in a water bath until it completely dissolves. Dilute gelatin at 0.1% in phosphate-buffered saline (PBS).</li>
	<li>Coat a 150 mm dish with 2 mL of 0.1% gelatin. Use 2 dishes per virus. Incubate the dishes for 30 min at 37 &#176;C.</li>
	<li>Seed 5 million HEK293T cells per gelatin-coated dish in 20 mL of 10% Dulbecco&#8242;s Modified Eagle&#8242;s Medium (DMEM) and place the cells in an incubator at 37 &#176;C.</li>
</ol>
Day 2
<ol start="4">
	<li>Check that the cells have reached about 50% confluency. Change the medium with 22 mL of 10% DMEM without disturbing the cell layer 2 h prior to transfection.</li>
	<li>Prepare the transfection mix. 
	NOTE: One transfection mix is prepared per dish, as a larger volume might impact the calcium-phosphate precipitate formation.
	<ol>
		<li>Prepare the reaction mix in a 15 mL centrifuge tube as follows: 32 &#181;g of plasmid, 20 &#181;g of PAX (pCMVdR8.74) plasmid, 9 &#181;g of VSVG (pMD.G2) plasmid, and bring the volume up to 1125 &#181;L using H2O.</li>
		<li>Add 125 &#181;L of CaCl2 to the reaction and incubate 5 min at room temperature (RT) on a roller.</li>
		<li>Prepare 1.25 mL of 2x HEPES in a separate 15mL centrifuge tube.</li>
		<li>Add 2x HEPES drop by drop to the reaction mixture while gently vortexing, then place the mixture on a roller for 10 min at RT.</li>
	</ol>
	</li>
	<li>Dispense the reaction drop by drop on the cells in a spiral pattern and place the cells in an incubator at 37 &#176;C after gently moving the plate in a cross-shape manner to homogenize the medium.</li>
</ol>
Day 3
<ol start="7">
	<li>Change the medium with 16 mL of 10% DMEM without disturbing the cell&#39;s layer.</li>
</ol>
Day 4
<ol start="8">
	<li>Collect the supernatant and store it at 4 &#176;C overnight. Add 16 mL of fresh DMEM to each plate without disturbing the cell layer.</li>
</ol>
Day 5
<ol start="9">
	<li>Collect the supernatant and pool with the supernatant from day 4. Filter to eliminate debris with a 0.45 &#181;M filter.</li>
	<li>Transfer each 30 mL (day 4 + day 5) supernatant in a conical tube for ultracentrifugation. Add the tubes in a swing bucket, with adapters for the conical bottom if required.</li>
	<li>Balance the tubes face to face at 0.02 g accuracy and ultracentrifuge at 90,000 x g for 2 h at 4 &#176;C.</li>
	<li>Discard the supernatant and add 200 &#181;L of stem cell medium per tube. Close tubes with parafilm and incubate the tube in a cold room for 2-4 h under agitation.</li>
	<li>Make 20 &#181;L aliquot of each virus and store the vials at -80 &#176;C. Keep 2 &#181;L of each virus and add 400 &#181;L of DMEM. 
	NOTE: This dilution will be used for titration.</li>
</ol>
2. Lentivirus titration
NOTE Lentiviruses must be handled in a laboratory of biosafety level L2 under sterile conditions. All required PPE for this lab must be worn, including double gloves. Contaminated material is decontaminated with virucide disinfectant prior to disposal in an autoclaved bin.
<ol>
	<li>Seed HEK293T in a 24 well plate. Add 50,000 cells/well in 500 &#181;L of 10% DMEM. 
	NOTE: Use 6 wells per virus and keep one well for untransduced cells.</li>
	<li>Perform the transduction. Add in the different wells: 2 &#181;L, 4 &#181;L, 8 &#181;L, 16 &#181;L, 32 &#181;L, and 64 &#181;L of the diluted virus from step 1.13. Homogenize gently.</li>
	<li>Change the medium 24 h later by adding 500 &#181;L of 10% DMEM.</li>
	<li>At day 4 post transduction, prepare cells for flow cytometry.
	<ol>
		<li>Prepare FACS buffer with PBS, 2% fetal bovine serum (FBS), 1% penicillin-streptomycin (P/S), 2 mM ethylenediaminetetraacetic acid (EDTA) and 1/2,000 4&#8242;,6-diamidino-2-phenylindole (DAPI).</li>
		<li>Remove the supernatant and add 500 &#181;L of FACS buffer directly with EDTA to each well.</li>
		<li>Detach the cells by pipetting up and down.</li>
	</ol>
	</li>
	<li>Filter the cells through a 40 &#181;m strainer and transfer them in FACS tubes.</li>
	<li>Determine the percentage of EGFP cells by flow cytometry.</li>
	<li>Set up the following gates:
	<ol>
		<li>Set up DAPI versus side scatter (SSC-A) to exclude dead cells.</li>
		<li>Set up SSC height (SSC-H) versus SSC area (SSC-A) to exclude cell aggregates and select single cells.</li>
		<li>Set up forward-scatter area (FSC-A) versus SSC-A to gate the cells and remove the debris based on size.</li>
		<li>Set up green fluorescence protein (GFP) versus SSC-A to select GFP+ cells.</li>
	</ol>
	</li>
	<li>Titer the virus.
	<ol>
		<li>Calculate the particle concentration (number of particles/mL) for each condition:</li>
		<li>EGFP cells x Dilution factor virus (200) x Number of seeded cells x1000)/ volume of virus. 
		NOTE: For example, cells were treated with 64 &#181;L of virus diluted at 1/200. Following flow cytometry analysis, 54% of cells were EGFP+. In those conditions, the titer will be (54% x 200 x 50 000 x 1000)/64 = 8.44 x 107 particles/mL.</li>
	</ol>
	</li>
</ol>
3. Transducing CD34+ cells isolated from umbilical cord blood
NOTE This part is performed under sterile conditions. Here, CD34+ UCB cells (purity &#62;85%) are transduced with one shRNA (control) harboring an eGFP reporter. The experiment is performed with three technical replicates. Proportion must be adjusted to the number of samples.
Day 1
<ol>
	<li>Prepare the following media and buffer:
	<ol>
		<li>Prepare defrosting medium: FBS supplemented with 1% P/S and 10 &#181;g/mL DNAse</li>
		<li>Prepare FACS buffer: PBS supplemented with 1% P/S and 2% FBS (&#177; 10 &#181;g/mL DNAse)</li>
		<li>Prepare stimulation media (in stem cell medium) by adding 150 ng/mL SCF, 150 ng/mL FLT-3, 10 ng/mL IL-6, 25 ng/mL G-CSF, 20 ng/mL TPO, and 1% HEPES.</li>
	</ol>
	</li>
	<li>Thaw 100,000 CD34+ UCB cells by dipping for 30 s in the water bath at 37 &#176;C. Add 1 mL of defrosting medium dropwise to each vial and then resuspend gently by pipetting up and down.</li>
	<li>Transfer the cell suspension drop by drop in a total of 5 mL defrosting medium gently. 
	NOTE: To avoid losing cells, keep a 2 mL aliquot of defrosting medium aside that will be used at the end to wash the tip and vial.</li>
	<li>Centrifuge for 8 min at 320 x g at RT. Wash cells in 10 mL of FACS buffer + DNAse.</li>
	<li>Centrifuge for 8 min at 320 x g at RT. Resuspend the cell pellet in 2 mL of stimulation media (washing tip and tube with stimulation medium).</li>
	<li>Count the cells using 5 &#181;L of cell suspension and 5 &#181;L of trypan blue. 
	NOTE: Aim for 75,000 cells total minimum (25,000 cells/condition).</li>
	<li>Transfer the 2 mL of cell suspension into a 24 well plate and incubate for &#8805;4-6 h at 37 &#176;C. 
	NOTE: All the cells (75,000-100,000) are seeded in one well at this step.</li>
	<li>Collect the cells in FACS tubes and top up with 1 mL of stem cell medium. Centrifuge for 8 min at 320 x g at RT.</li>
	<li>Thaw the virus and add the required volume to stimulation media at a multiplicity of infection (MOI) of 30.</li>
	<li>Resuspend the pelleted cells in virus-containing media at 100,000 cells per 100 &#181;L virus into 1 well of a 96-well plate. Add PBS to empty wells and incubate overnight at 37 &#176;C.</li>
</ol>
Day 2
<ol start="11">
	<li>Prepare expansion medium (in stem cell medium) by adding 150 ng/mL SCF, 150 ng/mL FLT3-L, and 20 ng/mL TPO.</li>
	<li>Add the full volume of cell suspension from the 96-well plate well to FACS tubes with 1 mL of stem cell medium. Wash the well and the tip with stem cell medium, resulting in the addition of 2 mL stem cell medium to the cell suspension in the FACS tube.</li>
	<li>Centrifuge for 8 min at 320 x g at RT.</li>
	<li>Discard the supernatant and resuspend cells in 2 mL of expansion media. Add the cell suspension into a 24 well plate well (wash the well and tip in the process to obtain a total of 2 mL). Incubate the cells for 4 days at 37 &#176;C. 
	NOTE: Cell sorting is then performed on day 7.</li>
</ol>
4. CD34+ EGFP+ Cell sorting and erythroid differentiation
NOTE This part is performed under sterile conditions.
<ol start="1">
	<li>Prepare the following media and buffer. 
	NOTE: Media with cytokines have to be prepared freshly:
	<ol>
		<li>Prepare FACS buffer by supplementing PBS with 1% P/S and 2% FBS.</li>
		<li>Prepare expansion media (in stem cell medium) by adding 150 ng/mL SCF, 150 ng/mL FLT3-L, and 20 ng/mL TPO.</li>
		<li>Prepare erythrocyte differentiation media (in stem cell medium) by adding 25 ng/mL SCF, 3 U/mL EPO, and 50 ng/mL IGF1.</li>
		<li>Prepare 2x Erythrocyte differentiation media (in stem cell medium) by adding 50 ng/mL SCF, 6 U/mL EPO, and 100 ng/mL IGF1.</li>
	</ol>
	</li>
	<li>Add the full volume of cells to a separate FACS tube (wash the well and tip in the process with stem cell medium). Centrifuge for 8 min at 320 x g at RT. Resuspend the pelleted cells in 100 &#181;L of FACS buffer.</li>
	<li>Add 5 &#181;L of CD34 Ab/tube and incubate in the dark at 4 &#176;C for 20 min.</li>
	<li>Resuspend the cells in 1 mL of FACS buffer with DAPI (1/2000 dilution in FACS buffer) and centrifuge for 8 min at 320 x g at RT.</li>
	<li>Resuspend cells in 200 &#181;L of FACS buffer with DAPI (1/2000 dilution in FACS buffer) and filter cells through a 40 &#181;M strainer prior to sorting.</li>
	<li>Prepare the beads for control:
	<ol>
		<li>Add 0.5 &#181;L of CD34 antibody (Ab) to 1 drop of beads in 100 &#181;L of PBS and incubate in the dark at 4 &#176;C for 20 min.</li>
		<li>Add 1 mL of PBS. Centrifuge for 8 min at 320 x g at RT. Discard the supernatant and resuspend the cells in 500 &#181;L of PBS.</li>
	</ol>
	</li>
	<li>Prepare leukemic cell lines for GFP control. 
	NOTE: Use any available GFP+ cell line as control; beads can be used instead.
	<ol>
		<li>Add GFP- cells (2 FACS tubes with 100,000 cells) and GFP+(1 FACS tube with 100,000 cells) in the FACS tube. 
		NOTE: A total of 3 tubes.</li>
		<li>Wash cells in 1 mL of FACS buffer. Centrifuge for 8 min at 320 x g at RT.</li>
		<li>Resuspend 100,000 GFP- cells in 500 &#181;L of FACS buffer (Unstained), 100,000 GFP- cells in 500 &#181;L of FACS buffer with DAPI (1/2000 dilution in FACS buffer) (DAPI stained), and 100,000 of GFP+ cells in 500 &#181;L of FACS buffer (GFP+ cells). 
		NOTE: Control tubes will be used for gating and compensation settings at the flow cytometer. If the cells used are 100% GFP, GFP- cells can be added to the tube (50,000 GFP- cells and 50,000 GFP+ cells).</li>
		<li>Filter the cells through a 40 &#181;m strainer prior to sorting.</li>
	</ol>
	</li>
	<li>Set up the following gates:
	<ol>
		<li>Set up DAPI versus side scatter (SSC-A) to exclude dead cells.</li>
		<li>Set up SSC height versus SSC area (SSC-A) to exclude cell aggregates and select single cells.</li>
		<li>Set up forward-scatter area (FSC-A) versus SSC-A to remove the debris based on size.</li>
		<li>Set up GFP versus SSC-A to select GFP+ cells.</li>
		<li>Set up CD34 (PerCP-Cy5.5) versus SSC-A to select CD34+ cells.</li>
	</ol>
	</li>
	<li>Flip the collection FACS tube prior to cell collection to coat the walls of the tube with FBS.</li>
	<li>Collect cells in the stem cell medium. 
	NOTE: Aim for 75,000-80,000 cells (25,000 cells x 3)</li>
	<li>Centrifuge for 8 min at 320 x g at RT and resuspend cells in 3 mL of erythroid differentiation media.</li>
	<li>Resuspend and then dispense 1 mL per well in 3 wells of a 24 well plate. Fill up all outer wells with PBS.</li>
	<li>Perform a half depletion of media every other day and replace it with a 2x erythrocyte differentiation media.
	<ol>
		<li>To refresh the media, make sure the cells are settled at the bottom of the plate. Gently lean the plate and take out 500 &#181;L of media.</li>
		<li>Refresh by adding 500 &#181;L with a 2x cytokine erythroid differentiation media.</li>
		<li>Transfer the cells on day 6 to a 12 well plate and add 1 mL of media to make a final volume of 2 mL.</li>
	</ol>
	</li>
</ol>
5. Analyzing erythroid differentiation by flow cytometry twice a week from day 6 until day 14
<ol>
	<li>Prepare the following media and buffer.
	<ol>
		<li>Prepare FACS buffer by supplementing PBS with 1% P/S and 2% FBS</li>
		<li>Prepare erythrocyte differentiation media (in stem cell medium) by adding 25 ng/mL SCF, 3 U/mL EPO, and 50 ng/mL IGF1.</li>
	</ol>
	</li>
	<li>Collect 200 &#181;L cells from each well of the 12 well plate and replace them with 250 &#181;L of 2x erythroid differentiation media. Add 1 mL of FACS buffer to each tube.</li>
	<li>Centrifuge for 8 min at 320 x g at RT. Resuspend the pelleted cells in 100 &#181;L of FACS buffer.</li>
	<li>Add 1 &#181;L of CD71 Ab (1/100), 1 &#181;L of CD34 Ab (1/100), and 1 &#181;L of CD235a Ab (1/100) and incubate in the dark at 4 &#176;C for 20 min.</li>
	<li>Resuspend cells in 1 mL of FACS buffer with DAPI (1/2000 dilution in FACS buffer). Centrifuge for 8 min at 320 x g at RT.</li>
	<li>Resuspend cells in 200 &#181;L of FACS buffer with DAPI (1/2000 dilution in FACS buffer).</li>
	<li>Prepare the beads for control:
	<ol>
		<li>Add 1 drop compensation beads to two tubes in 100 &#181;L of PBS and then add 0.55 &#181;L of a single antibody to each tube to make single stain controls (one for CD34, one for CD71, and one for CD235a). 
		NOTE: 1 drop is sufficient for both controls.</li>
		<li>Incubate in the dark at 4 &#176;C for 20 min. Add 1 mL of PBS.</li>
		<li>Centrifuge for 8 min at 320 x g at RT. Discard the supernatant and resuspend the cells in 500 &#181;L of PBS.</li>
	</ol>
	</li>
	<li>Prepare a leukemic cell line for control. 
	NOTE: K562 cells express the CD71 marker and can be used as a positive control to set up the FACS.
	<ol>
		<li>Add GFP- cells (2 FACS tubes with 100,000 cells) and GFP+ (1 FACS tube with 100,000 cells) cells in the FACS tube.</li>
		<li>Wash cells in 1 mL of FACS buffer. Centrifuge for 8 min at 320 x g at RT.</li>
		<li>Resuspend 100,000 GFP- cells in FACS buffer (unstained), 100,000 GFP- cells in FACS buffer with DAPI (1/2000 dilution in FACS buffer) (DAPI stained), and 100,000 GFP+ cells in FACS buffer (GFP+ cells).</li>
	</ol>
	</li>
	<li>Set up the following gates:
	<ol>
		<li>Set up DAPI versus side scatter (SSC-A) to exclude dead cells.</li>
		<li>Set up SSC height versus SSC area (SSC-A) to exclude cell aggregates and select single cells.</li>
		<li>Set up forward-scatter area (FSC-A) versus SSC-A to remove the debris based on size.</li>
		<li>Set up CD235a (APC-Cy7) versus CD71 (PE).</li>
		<li>Set up CD34 (PerCP-Cy5.5) versus SSC-A.</li>
	</ol>
	</li>
	<li>Run a sample of unstained GFP- cells and full-stained cells in order to adjust voltages so that cells sit in the correct place.</li>
	<li>Run beads, GFP+ cells, and DAPI-stained cells. Compensate and apply compensation to the cytometer settings.</li>
	<li>Run the samples and record the events.</li>
</ol>
6. Analysis of erythroid differentiation by Giemsa staining
<ol>
	<li>Prepare 5 slide staining jars:
	<ol>
		<li>Prepare a slide staining jar with May-Gr&#252;nwald&#39;s eosin-methylene blue solution 1:1 stock and PBS.</li>
		<li>Prepare a slide staining jar with 2.5 mL of Giemsa stock + 50 mL of PBS.</li>
		<li>Prepare a slide staining jar with distilled water.</li>
		<li>Prepare a slide staining jar with PBS.</li>
		<li>Prepare a slide staining jar with iced cold methanol.</li>
	</ol>
	</li>
	<li>Collect 100,000 cells for optimum staining, add 1 mL of PBS, and centrifuge 300 x g for 5 min.</li>
	<li>Discard the medium and resuspend in 150 &#181;L of PBS.</li>
	<li>Prepare slides, filter paper, and conical adaptor into a cytocentrifuge and drop cell preparation slowly into the conical adaptor.</li>
	<li>Spin at 800 x g for 3 min. Remove the slides and place them in the methanol slide jar in ice for 15 min.</li>
	<li>Immerse the slides in the May-Gr&#252;nwald solution for 5 min and agitate occasionally to ensure proper staining.</li>
	<li>Then, immerse the slides in the PBS jar until no stain runs off.</li>
	<li>Immerse the slides in the Giemsa solution for 20 min and agitate occasionally to ensure proper staining.</li>
	<li>Carefully immerse slides with PBS until no stain runs off.</li>
	<li>Allow the slides to remain in PBS for an additional 3 min.</li>
	<li>Immerse the slides quickly in distilled water and air dry at RT.</li>
	<li>Add a drop of synthetic resin on each spot of cells and gently add a coverslip on top.</li>
	<li>Let the slides dry for 1 h at RT and seal the coverslip with nail polish to prevent drying slip.</li>
</ol>

Modeling Erythropoiesis Using HSPCs from Cord Blood and Bone Marrow

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The authors have nothing to disclose.

Here, we described an efficient and reliable method to induce erythroid differentiation of CD34+ hematopoietic stem cells (HSPCs) isolated either from adult bone marrow (BM) or from umbilical cord blood (UCB). This pipeline includes the editing/modification of primary cells to investigate the function of any gene of interest during the erythroid differentiation process. This method has been successfully used to investigate the functional relevance of genes that were not previously linked to erythroid differentiation12,13,14,15 in healthy CD34+ HSPCs and hematopoietic malignancies such as acute myeloid leukemia and myelodysplastic syndrome. Modeling erythroid differentiation is an asset to understanding the biological relevance of genes in this process and testing new therapeutic strategies that could alleviate differentiation defects.
With this protocol, we also provided robust and reproducible read-outs that can be used during the differentiation process and at endpoints. This includes the CD71/CD235a phenotyping by flow cytometry and a May-Grunwald Giemsa (MGG) staining that allows further characterization of the erythroid differentiation stages. Profiling of CD71/CD235a, revealed that BM CD34+ cells are differentiating faster than the UCB CD34+ cells and suggest a bias towards erythroid lineage in adult HSPCs. Our approach allows the production of mature red blood cells, characterized by nucleus extrusion, visualized with an MGG staining on UCB CD34+ cells undergoing erythroid differentiation.
While other methods require many changes in cytokines during the differentiation process5, 3D modeling using scaffold6, or even IPSCs generation8, we offer an alternative simplified method, requiring only changes in cytokines between expansion and differentiation, and that allows the production of mature red blood cells in less than 20 days. Critical steps include an optimal CD34+ purity (&#62;80%-90%), efficient transduction of the cells, the sort of CD34+GFP+ cells post-transduction, and proper maintenance of the differentiation medium with fresh cytokines added every other day. A possible limitation is the maintenance of differentiated red blood cells that start dying soon after 14 days in culture. This duration is, for example, not compatible with the time necessary to observe ring sideroblasts&#39; formation, which are abnormal erythroid progenitors with iron accumulation that characterize myelodysplastic syndrome with SF3B1 mutations8. The heterogeneity among primary samples is likely to affect the kinetic of differentiation and needs to be carefully monitored when using new samples. Finally, the addition of a step of transduction requires the production of viral particles, a greater number of primary cells at the start, and access to the FACS facility to sort the cells.
Overall, this pipeline offers a simplified, robust, and reproducible assay to assess human erythropoiesis in vitro and ex vivo.

Every day about 280 billion red blood cells (RBCs) are produced in the bone marrow to ensure oxygen transport throughout the entire body1. Erythropoiesis is a finely tuned differentiation process that produces mature, functional red blood cells from hematopoietic stem cells. Many diseases, both acquired and congenital, can affect this process, including myelodysplastic syndrome, aplastic anemia, thalassemia, and congenital dyserythropoietic anemia2.
The clinical manifestation of disrupted erythropoiesis is anemia, a major cause of morbidity worldwide. Symptoms of anemia range from fatigue, shortness of breath, and dizziness to life-threatening conditions, such as heart failure and cardiac arrest3.With anemia affecting 1.8 billion people worldwide4, modeling erythropoiesis ex vivo in order to investigate these conditions and their underlying mechanisms remains a global priority. The method described here aims to provide a simple and robust model for studying both healthy and diseased erythroid differentiation. This model allows for the dissection of the roles of specific genes in this process and enables the testing of compounds that could benefit anemic patients. Historically, modeling ineffective erythropoiesis has had many challenges, including a paucity of stem cells in some diseased bone marrows and difficulties mimicking the bone marrow niche.
Previous studies have used similar or more complex protocols; for instance, Bondu et al. used a cocktail of cytokines including erythropoietin (EPO), stem cell factor (SCF), and interleukin-6 (IL6) for 4 days before removing IL6 from the cocktail of erythroid differentiation, in contrast, Yip et al. only added EPO to their cytokines cocktail after 7 days of liquid culture. Elvarsd&#243;ttir et al. developed a three-dimensional (3D) culture model for CD34+ cells to facilitate the highest expansion and maturation of erythroid cells, including the generation of erythroblastic islands and enucleated erythrocytes, which are important to study phenomena such as ring sideroblasts5,6,7,8. This protocol uses only three cytokines when expanding the cells (i.e., SCF, thrombopoietin [TPO], and FMS-like tyrosine kinase 3 [FLT3]) and three cytokines maintained throughout the whole process of differentiation (i.e., EPO, IGF1, and SCF). One of the key criteria shared across these different studies is the use of CD34+ cells. Although the origin of these cells varies, they are consistently employed across these different protocols. The sources might range from non-invasive collection of UCB to more invasive procedures to obtain adult bone marrow samples that can be harvested either from granulocyte colony-stimulating factor (G-CSF) mobilized patients or bone marrow punctures. In general, liquid culture models are carried over 14 days, while 3D models can maintain cells over longer periods. These protocols are essential for modeling genetic diseases and disruptions in erythropoiesis. Genetic approaches, such as short hairpin RNA (shRNA) knockdowns, can be employed to silence genes of interest and study their roles in erythropoiesis. The impact of gene silencing can then be monitored at various stages of differentiation, providing an ex vivo human model of red blood cell production to study anemia in both health and disease. This protocol includes the transduction of CD34+ cord blood cells with shRNA that constitutively express a reporter gene, such as enhanced green fluorescence protein (EGFP), allowing the samples to be sorted by fluorescence-activated cell sorting (FACS). Cells are then incubated in differentiation media for 2 weeks and monitored by flow cytometry analysis and immunochemistry staining.

<table><tbody><tr><td>2x HEPES</td><td>Merck</td><td>51558</td><td></td></tr><tr><td>Beads</td><td>Thermo Scientific</td><td>01-2222-42</td><td></td></tr><tr><td>CD235a APC Cy7</td><td>Biolegend</td><td>349115</td><td></td></tr><tr><td>CD34 PerCP Cy5.5 antibody</td><td>BD Biosciences</td><td>347222</td><td></td></tr><tr><td>CD71 PE antibody</td><td>Biolegend</td><td>334106</td><td></td></tr><tr><td>DAPI</td><td>Merck</td><td>D9542</td><td></td></tr><tr><td>DMEM</td><td>Thermo Scientific</td><td>41966-029</td><td></td></tr><tr><td>DNAse</td><td>Merck</td><td>D4527-200KU</td><td></td></tr><tr><td>DPX mounting medium&amp;nbsp;</td><td>VWR</td><td>1.00579.0500</td><td></td></tr><tr><td>FACSAria Fusion&amp;nbsp;</td><td>BD Biosciences</td><td>NA</td><td></td></tr><tr><td>FBS</td><td>Merck</td><td>F9665</td><td></td></tr><tr><td>Gelatin</td><td>Merck</td><td>G1393</td><td></td></tr><tr><td>Giemsa solution</td><td>Abcam</td><td>ab150670</td><td></td></tr><tr><td>HEPES</td><td>Merck</td><td>H0887-100mL</td><td></td></tr><tr><td>Human EPO</td><td>PeproTech</td><td>100-64</td><td></td></tr><tr><td>Human Flt-3 ligand</td><td>PeproTech</td><td>300-19</td><td></td></tr><tr><td>Human G-CSF</td><td>PeproTech</td><td>300-23</td><td></td></tr><tr><td>Human IGF1</td><td>PeproTech</td><td>100-11</td><td></td></tr><tr><td>Human IL-6</td><td>PeproTech</td><td>200-06</td><td></td></tr><tr><td>Human SCF</td><td>PeproTech</td><td>300-07</td><td></td></tr><tr><td>Human TPO</td><td>PeproTech</td><td>AF-300-18</td><td></td></tr><tr><td>LSRFortessa Cell Analyzer</td><td>BD Biosciences</td><td>NA</td><td></td></tr><tr><td>May-Grunwald solution&amp;nbsp;</td><td>Generon</td><td>26250-01</td><td></td></tr><tr><td>Pannoramic 250 High Throughput Scanner&amp;nbsp;</td><td>3DHISTECH</td><td>NA</td><td></td></tr><tr><td>Penicillin/Streptomycin</td><td>Thermo Scientific</td><td>15140-122</td><td></td></tr><tr><td>Shandon Cytospin 3&amp;nbsp;</td><td>Thermo Scientific</td><td>NA</td><td></td></tr><tr><td>Stem Cell Medium medium</td><td>Stem Cell Technologies</td><td>9655</td><td></td></tr></tbody></table>

a comprehensive pipeline to assess the efficiency of human erythropoiesis in vitro and ex vivo

Erythropoiesis, a remarkably dynamic and efficient process responsible for generating the daily quota of red blood cells (approximately 280 &plusmn; 20 billion cells per day), is crucial for maintaining individual health. Any disruption in this pathway can have significant consequences, leading to health issues. According to the World Health Organization, an estimated 25% of the global population presents symptoms of anemia. This protocol describes how to generate human erythroid cells both in vitro using hematopoietic stem and progenitor cells (HSPCs) from sources such as umbilical cord blood (UCB) or blood taken from healthy donors and ex vivo with HSPCs isolated from patients' bone marrow. Using genetic approach, genes of interest can be modulated in HSPCs, and their impact on erythropoiesis can be monitored at various stages of the differentiation process. This method allows for the screening of compounds perturbing, enhancing, or rescuing the capacity of HSPCs to differentiate into mature erythroid cells and to investigate the role of genes of interest during the erythroid differentiation process.

CD34+ cells from umbilical cord blood or bone marrow are firstly thawed, stimulated and transduced with a GFP-expressing shRNA (Figure 1A). CD34+GFP+ cells are sorted 4 days following transduction, according to the gating strategy illustrated in Figure 1B. Representative results showed the maturation of CD34+ cells after transduction and FACS sorting. Cells are analyzed (1) by flow cytometry at different stages of maturation, using CD71/CD235a9,10,11 staining (Figure 1A); (2) by immunochemistry following a Giemsa staining visualized by microscope.
Erythroid differentiation is characterized by the sequential acquisition of CD71 and CD235a, with the final steps of maturation requiring the loss of CD71. These are well-established, reproducible, and robust markers of erythroid differentiation9,10,11. The CD71/CD235a profile is established by flow cytometry (Figure 1). Here, CD34+ UBC cells were transduced with a shRNA (scramble) expressing GFP. Cells were sorted on CD34+/GFP+ and then cultured in erythroid differentiation media for 14 days. Cells were analyzed by flow cytometry via gating successively on live cells (DAPI-), single cell (SSC-A-SSC-H), cells excluding debris (FSC-A, SSC-A), GFP+ populations, and CD71 (Y axis)/CD235a (X-axis) profile.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67123/67123fig01.jpg" /> 
Figure 1:&#160;Representative workflow and gating strategy. (A) Schematics of the workflow used for erythroid differentiation. Isolated CD34+ cells are transduced with lentiviral particles and cultured for 4 days prior to FACS sorting. CD34+GFP+ cells are then grown in an erythroid differentiation medium supplemented with SCF, EPO, and IGF1. Flow cytometry is performed to follow the differentiation progress over time. (B) Representation of the gating strategy used for CD34+GFP+ cell sorting. Live cells were gated (DAPI-), single cells (SSC-A, SSC-H), cells excluding debris (FSC-A, SSC-A), CD34+, and then GFP+ cells. (C) Representation of the gating strategy used for the CD71/CD235a analysis by flow cytometry. We successively gated on DAPI- cells, which were gated as single cells (SSC-A-SSC-H). We then cleared the population from small debris (FSC-A, SSC-A) and selected the GFP+ population to assess the CD71/CD235a profile. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67123/67123fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The erythroid differentiation kinetic, with CD34+ cells isolated from the umbilical cord or adult healthy donor bone marrow, was compared at days 4, 7, 14, and 17 (Figure 2A,B). A faster differentiation process is usually observed in CD34+ cells from adult bone marrow. The variability between samples is displayed in Figure 2B and is represented by the average and standard deviation of three independent experiments.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67123/67123fig02.jpg" /> 
Figure 2:&#160;Representative CD71/CD235a flow cytometry plots. (A)&#160;The CD71/CD235a profile is assessed at days 4, 7, 14, and 17 of the erythroid differentiation protocol for CD34+ cells isolated from umbilical cord blood (upper panel) or for CD34+ cells isolated from adult bone marrow (bottom panel). (B) CD71+/CD235a+ and CD71-/CD235a+ cells percentage, at day 14 of erythroid differentiation, for 3 individual cord blood samples (CB) and 3 individual adult healthy donors (HD). The average and standard deviation are displayed on the graph. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67123/67123fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
As differentiated cells eventually undergo apoptosis, we assessed the percentage of Annexin V cells at different time points (Figure 3). CD34+ cells from umbilical cord blood showed an increase in apoptosis by day 17.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67123/67123fig03.jpg" /> 
Figure 3:&#160;Annexin V flow cytometry plots and quantification. (A) Representative Annexin V flow cytometry plots.(B) The Annexin V percentage of cells is monitored at days 4, 7, 14, and 17 of the erythroid differentiation protocol for CD34+ cells isolated from umbilical cord blood. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67123/67123fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In order to determine and compare the different stages of differentiation, we performed a May Grunwald-Giemsa staining (MGG) on erythroid differentiated samples. Here, the staining displayed has been performed at day 14 on cells initially isolated from umbilical cord blood (Figure 4). The staining allows for discrimination of the early basophilic and polychromatic erythroblasts (progenitors) with blue cytoplasm and nucleus from more differentiated orthochromatic erythroblasts that are smaller with a purple cytoplasm and a blue nucleus. The late stage of differentiation is associated with nucleus extrusion and reticulocyte formation, which appear as light pink cells without a nucleus.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/67123/67123fig04.jpg" /> 
Figure 4: May-Grunwald Giemsa staining. Representative pictures of the May-Grunwald Giemsa staining performed at day 14 of erythroid differentiation with CD34+ cells isolated from umbilical cord blood. Baso/Poly: Basophilic/Polychromatic erythroblasts. Ortho: Orthochromatic erythroblasts. RBC: Red blood cells. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67123/67123fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Author Spotlight: Advancing Erythropoiesis Research - A Simplified Pipeline for Assessing Hematopoietic Stem Cell Function in Myelodysplastic Syndromes

Modeling erythropoiesis provides a valuable chance to comprehend the biological significance of actors in this process and to evaluate novel therapeutic approaches that may alleviate differentiation defects. Here, we describe a simple and reliable method to efficiently differentiate CD34+ hematopoietic stem and progenitor cells ex vivo.

A Comprehensive Pipeline to Assess the Efficiency of Human Erythropoiesis In Vitro and Ex Vivo

Erythropoiesis, a remarkably dynamic and efficient process responsible for generating the daily quota of red blood cells (approximately 280 &plusmn; 20 ...

a-comprehensive-pipeline-to-assess-efficiency-human-erythropoiesis

Nanyang Technological University

Research

JoVE Journal

Biology

383 Views.  INSERM U1242, University of Rennes. Modeling erythropoiesis provides a valuable chance to comprehend the biological significance of actors in this process and to evaluate novel therapeutic approaches that may alleviate differentiation defects. Here, we describe a simple and reliable method to efficiently differentiate CD34+ hematopoietic stem and progenitor cells ex vivo.

Video: Author Spotlight: Advancing Erythropoiesis Research - A Simplified Pipeline for Assessing Hematopoietic Stem Cell Function in Myelodysplastic Syndromes

Ex vivo Mimicry of Normal and Abnormal Human Hematopoiesis

Hematopoietic stem cells require a unique microenvironment in order to sustain blood cell formation1; the bone marrow (BM) is a complex three-dimensional (3D) tissue wherein hematopoiesis is regulated by spatially organized cellular microenvironments termed niches2-4. The organization of the BM niches is critical for the function or dysfunction of normal or malignant BM5. Therefore a better understanding of the in vivo microenvironment using an ex vivo mimicry would help us elucidate the molecular, cellular and microenvironmental determinants of leukemogenesis6.
Currently, hematopoietic cells are cultured in vitro in two-dimensional (2D) tissue culture flasks/well-plates7 requiring either co-culture with allogenic or xenogenic stromal cells or addition of exogenous cytokines8. These conditions are artificial and differ from the in vivo microenvironment in that they lack the 3D cellular niches and expose the cells to abnormally high cytokine concentrations which can result in differentiation and loss of pluripotency9,10.
Herein, we present a novel 3D bone marrow culture system that simulates the in vivo 3D growth environment and supports multilineage hematopoiesis in the absence of exogenous growth factors. The highly porous scaffold used in this system made of polyurethane (PU), facilitates high-density cell growth across a higher specific surface area than the conventional monolayer culture in 2D11. Our work has indicated that this model supported the growth of human cord blood (CB) mononuclear cells (MNC)12 and primary leukemic cells in the absence of exogenous cytokines. This novel 3D mimicry provides a viable platform for the development of a human experimental model to study hematopoiesis and to explore novel treatments for leukemia.

Ex vivo Mimicry of Normal and Abnormal Human Hematopoiesis

A 3D culture system for hematopoiesis is described using human cord blood and leukemic bone marrow cells. The method is based on the use of a porous synthetic polyurethane scaffold coated with extracellular matrix proteins. This scaffold is adaptable to accommodate a wide range of cells.

Hematopoietic stem cells require a unique microenvironment in order to sustain blood cell formation1; the bone marrow (BM) is a complex three-dimensional ...

Bioengineering

Detection of Residual Donor Erythroid Progenitor Cells after Hematopoietic Stem Cell Transplantation for Patients with Hemoglobinopathies

The presence of incomplete chimerism is noted in a large proportion of patients following bone marrow transplant for thalassemia major or sickle cell disease. This observation has tremendous implications, as subsequent therapeutic immunomodulation strategies can improve clinical outcome. Conventionally, polymerase chain reaction-based analysis of short tandem repeats is used to identify chimerism in donor-derived blood cells. However, this method is restricted to nucleated cells and cannot distinguish between dissociated single-cell lineages. We applied the analysis of short tandem repeats to flow cytometric-sorted hematopoietic progenitor cells and compared this with the analysis of short tandem repeats obtained from selected burst-forming unit - erythroid colonies, both collected from the bone marrow. With this method we are able to demonstrate the different proliferation and differentiation of donor cells in the erythroid compartment. This technique is eligible to complete current monitoring of chimerism in the stem cell transplant setting and thus may be applied in future clinical studies, stem cell research and design of gene therapy trials.

Quantification of donor-derived cells is required to monitor engraftment after stem cell transplantation in patients with hemoglobinopathies. A combination of flow cytometry-based cell sorting, colony formation assay, and subsequent analysis of short tandem repeats may be used to assess the proliferation and differentiation of progenitors in the erythroid compartment.

The presence of incomplete chimerism is noted in a large proportion of patients following bone marrow transplant for thalassemia major or sickle cell ...

Developmental Biology

Mouse Fetal Liver Culture System to Dissect Target Gene Functions at the Early and Late Stages of Terminal Erythropoiesis

Erythropoiesis involves a dynamic process that begins with committed erythroid burst forming units (BFU-Es) followed by rapidly dividing erythroid colony forming units (CFU-Es). After CFU-Es, cells are morphologically recognizable and generally termed terminal erythroblasts. One of the challenges for the study of terminal erythropoiesis is the lack of experimental approaches to dissect gene functions in a chronological manner. In this protocol, we describe a unique strategy to determine gene functions in the early and late stages of terminal erythropoiesis. In this system, mouse fetal liver TER119 (mature erythroid cell marker) negative erythroblasts were purified and transduced with exogenous expression of cDNAs or small hairpin RNAs (shRNAs) for the genes of interest. The cells were subsequently cultured in medium containing growth factors other than erythropoietin (Epo) to maintain their progenitor stage for 12 hr while allowing the exogenous cDNAs or shRNAs to express. The cells were changed to Epo medium after 12 hr to induce cell differentiation and proliferation while the exogenous genetic materials were already expressed. This protocol facilitates analysis of gene functions in the early stage of terminal erythropoiesis. To study late stage terminal erythropoiesis, cells were immediately cultured in Epo medium after transduction. In this way, the cells were already differentiated to the late stage of terminal erythropoiesis when the transduced genetic materials were expressed. We recommend a general application of this strategy that would help understand detailed gene functions in different stages of terminal erythropoiesis.

We present an in vitro mouse fetal liver erythroblast culture system that dissects the early and late stages of terminal erythropoiesis. This system facilitates functional analysis of specific genes in different developmental stages.

Erythropoiesis involves a dynamic process that begins with committed erythroid burst forming units (BFU-Es) followed by rapidly dividing erythroid colony ...

Immunology and Infection

Hemogenic Endothelium Differentiation from Human Pluripotent Stem Cells in A Feeder- and Xeno-free Defined Condition

Deriving functional hematopoietic stem and progenitor cells (HSPCs) from human pluripotent stem cells (PSCs) have been an elusive goal for autologous transplants to treat hematological and malignant disorders. Hemogenic endothelium (HE) is an amenable platform to produce functional HSPCs by transcription factors or to induce lymphocytes in a robust manner. However, current methods to derive HE are either costly or variable in yield. In this protocol, we established a cost-effective and precise method to derive HE within 4 days in a feeder- and xeno-free defined condition. The resultant HE underwent an endothelial-to-hematopoietic transition and produced CD34+CD45+ clonogenic hematopoietic progenitors. The potential capacity of our platform for generating functional HSPCs will have broad applications in disease modeling and screening for therapeutic compounds.

Here, we present a protocol to precisely form hemogenic endothelium from human pluripotent stem cells in a feeder- and xeno-free condition. This method will have broad applications in disease modeling and screening for therapeutic compounds.

Deriving functional hematopoietic stem and progenitor cells (HSPCs) from human pluripotent stem cells (PSCs) have been an elusive goal for autologous ...

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 ...

Lentiviral-mediated Knockdown During Ex Vivo Erythropoiesis of Human Hematopoietic Stem Cells

Erythropoiesis is a commonly used model system to study cell differentiation. During erythropoiesis, pluripotent adult human hematopoietic stem cells (HSCs) differentiate into oligopotent progenitors, committed precursors and mature red blood cells 1. This process is regulated for a large part at the level of gene expression, whereby specific transcription factors activate lineage-specific genes while concomitantly repressing genes that are specific to other cell types 2. Studies on transcription factors regulating erythropoiesis are often performed using human and murine cell lines that represent, to some extent, erythroid cells at given stages of differentiation 3-5. However transformed cell lines can only partially mimic erythroid cells and most importantly they do not allow one to comprehensibly study the dynamic changes that occur as cells progress through many stages towards their final erythroid fate. Therefore, a current challenge remains the development of a protocol to obtain relatively homogenous populations of primary HSCs and erythroid cells at various stages of differentiation in quantities that are sufficient to perform genomics and proteomics experiments.
Here we describe an ex vivo cell culture protocol to induce erythroid differentiation from human hematopoietic stem/progenitor cells that have been isolated from either cord blood, bone marrow, or adult peripheral blood mobilized with G-CSF (leukapheresis). This culture system, initially developed by the Douay laboratory 6, uses cytokines and co-culture on mesenchymal cells to mimic the bone marrow microenvironment. Using this ex vivo differentiation protocol, we observe a strong amplification of erythroid progenitors, an induction of differentiation exclusively towards the erythroid lineage and a complete maturation to the stage of enucleated red blood cells. Thus, this system provides an opportunity to study the molecular mechanism of transcriptional regulation as hematopoietic stem cells progress along the erythroid lineage. 
Studying erythropoiesis at the transcriptional level also requires the ability to over-express or knockdown specific factors in primary erythroid cells. For this purpose, we use a lentivirus-mediated gene delivery system that allows for the efficient infection of both dividing and non-dividing cells 7. Here we show that we are able to efficiently knockdown the transcription factor TAL1 in primary human erythroid cells. In addition, GFP expression demonstrates an efficiency of lentiviral infection close to 90%. Thus, our protocol provides a highly useful system for characterization of the regulatory network of transcription factors that control erythropoiesis.

Lentiviral-mediated Knockdown During Ex Vivo Erythropoiesis of Human Hematopoietic Stem Cells

An ex vivo protocol to generate mature human red blood cells from hematopoietic stem/progenitors is described. Additionally we describe an efficient lentiviral-delivery method to knockdown the transcription factor TAL1 in primary erythroid cells. The efficiency of lentivirus mediated gene delivery is demonstrated using GFP expressing viruses.

Erythropoiesis is a commonly used model system to study cell differentiation. During erythropoiesis, pluripotent adult human hematopoietic stem cells ...

Measuring Deformability and Red Cell Heterogeneity in Blood by Ektacytometry

Decreased red cell deformability is characteristic of several disorders. In some cases, the extent of defective deformability can predict severity of disease or occurrence of serious complications. Ektacytometry uses laser diffraction viscometry to measure the deformability of red blood cells subject to either increasing shear stress or an osmotic gradient at a constant value of applied shear stress. However, direct deformability measurements are difficult to interpret when measuring heterogenous blood that is characterized by the presence of both rigid and deformable red cells. This is due to the inability of rigid cells to properly align in response to shear stress and results in a distorted diffraction pattern marked by an exaggerated decrease in apparent deformability. Measurement of the degree of distortion provides an indicator of the heterogeneity of the erythrocytes in blood. In sickle cell anemia, this is correlated with the percentage of rigid cells, which reflects the hemoglobin concentration and hemoglobin composition of the erythrocytes. In addition to measuring deformability, osmotic gradient ektacytometry provides information about the osmotic fragility and hydration status of erythrocytes. These parameters also reflect the hemoglobin composition of red blood cells from sickle cell patients. Ektacytometry measures deformability in populations of red cells and does not, therefore, provide information on the deformability or mechanical properties of individual erythrocytes. Regardless, the goal of the techniques described herein is to provide a convenient and reliable method for measuring the deformability and cellular heterogeneity of blood. These techniques may be useful for monitoring temporal changes, as well as disease progression and response to therapeutic intervention in several disorders. Sickle cell anemia is one well-characterized example. Other potential disorders where measurements of red cell deformability and/or heterogeneity are of interest include blood storage, diabetes, Plasmodium infection, iron deficiency, and the hemolytic anemias due to membrane defects.

Here we present techniques to measure red cell deformability and cellular heterogeneity by ektacytometry. These techniques are applicable to general investigations of red cell deformability and specific investigations of blood diseases characterized by the presence of both rigid and deformable red cells in circulation, such as sickle cell anemia.

Decreased red cell deformability is characteristic of several disorders. In some cases, the extent of defective deformability can predict severity of ...

Identification of a Murine Erythroblast Subpopulation  Enriched in Enucleating Events by Multi-spectral Imaging Flow Cytometry

Erythropoiesis in mammals concludes with the dramatic process of enucleation that results in reticulocyte formation. The mechanism of enucleation has not yet been fully elucidated. A common problem encountered when studying the localization of key proteins and structures within enucleating erythroblasts by microscopy is the difficulty to observe a sufficient number of cells undergoing enucleation.&#160;We have developed a novel analysis protocol using multiparameter high-speed cell imaging in flow (Multi-Spectral Imaging Flow Cytometry), a method that combines immunofluorescent microscopy with flow cytometry, in order to identify efficiently a significant number of enucleating events, that allows to obtain measurements and perform statistical analysis.
We first describe here two in vitro erythropoiesis culture methods used in order to synchronize murine erythroblasts and increase the probability of capturing enucleation at the time of evaluation.&#160;Then, we describe in detail the staining of erythroblasts after fixation and permeabilization in order to study the localization of intracellular proteins or lipid rafts during enucleation by multi-spectral imaging flow cytometry.&#160;Along with size and DNA/Ter119 staining which are used to identify the orthochromatic erythroblasts, we utilize the parameters &#8220;aspect ratio&#8221; of a cell in the bright-field channel that aids in the recognition of elongated cells and &#8220;delta centroid XY Ter119/Draq5&#8221; that allows the identification of cellular events in which the center of Ter119 staining (nascent reticulocyte) is far apart from the center of Draq5 staining (nucleus undergoing extrusion), thus indicating a cell about to enucleate.&#160;The subset of the orthochromatic erythroblast population with high delta centroid and low aspect ratio is highly enriched in enucleating cells.

The present protocol describes a novel method of identifying a population of enucleating orthochromatic erythroblasts by multi-spectral imaging flow cytometry, providing a visualization of the erythroblast enucleation process.

Erythropoiesis in mammals concludes with the dramatic process of enucleation that results in reticulocyte formation. The mechanism of enucleation has not ...

Simultaneous Assessment of Kinship, Division Number, and Phenotype via Flow Cytometry for Hematopoietic Stem and Progenitor Cells

Few techniques can assess phenotype and fate for the same cell simultaneously. Most of the current protocols used to characterize phenotype, although able to generate large datasets, necessitate the destruction of the cell of interest, making it impossible to assess its functional fate. Heterogeneous biological differentiating systems like hematopoiesis are therefore difficult to describe. Building on cell division tracking dyes, we further developed a protocol to simultaneously determine kinship, division number, and differentiation status for many single hematopoietic progenitors. This protocol allows the assessment of the ex vivo differentiation potential of murine and human hematopoietic progenitors, isolated from various biological sources. Moreover, as it is based on flow cytometry and a limited number of reagents, it can quickly generate a large amount of data, at the single-cell level, in a relatively inexpensive manner. We also provide the analytical pipeline for single-cell analysis, combined with a robust statistical framework. As this protocol allows the linking of cell division and differentiation at the single-cell level, it can be used to quantitatively assess symmetric and asymmetric fate commitment, the balance between self-renewal and differentiation, and the number of divisions for a given commitment fate. Altogether, this protocol can be used in experimental designs aiming to unravel the biological differences between hematopoietic progenitors, from a single-cell perspective.

Simultaneous Assessment of Kinship, Division Number, and Phenotype via Flow Cytometry for Hematopoietic Stem and Progenitor Cells

Presented here is a flow cytometry-based technique that allows for simultaneously measuring the number of cell divisions, surface cell phenotype, and cellular kinship. Those properties can be tested statistically using a permutation-based framework.

Few techniques can assess phenotype and fate for the same cell simultaneously. Most of the current protocols used to characterize phenotype, although able ...

A Comprehensive Pipeline to Assess the Efficiency of Human Erythropoiesis In Vitro and Ex Vivo

Summary

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Chapters in this video

Ex vivo Mimicry of Normal and Abnormal Human Hematopoiesis

Detection of Residual Donor Erythroid Progenitor Cells after Hematopoietic Stem Cell Transplantation for Patients with Hemoglobinopathies

Mouse Fetal Liver Culture System to Dissect Target Gene Functions at the Early and Late Stages of Terminal Erythropoiesis

Hemogenic Endothelium Differentiation from Human Pluripotent Stem Cells in A Feeder- and Xeno-free Defined Condition

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

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

Lentiviral-mediated Knockdown During Ex Vivo Erythropoiesis of Human Hematopoietic Stem Cells

Measuring Deformability and Red Cell Heterogeneity in Blood by Ektacytometry

Identification of a Murine Erythroblast Subpopulation Enriched in Enucleating Events by Multi-spectral Imaging Flow Cytometry

Simultaneous Assessment of Kinship, Division Number, and Phenotype via Flow Cytometry for Hematopoietic Stem and Progenitor Cells