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 different...

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.

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			<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&#160;</td>
			<td>VWR</td>
			<td>1.00579.0500</td>
			<td></td>
		</tr>
		<tr>
			<td>FACSAria Fusion&#160;</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&#160;</td>
			<td>Generon</td>
			<td>26250-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Pannoramic 250 High Throughput Scanner&#160;</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&#160;</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>
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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 matu...

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

Research

JoVE Journal

Biology

352 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

Method for Culture of Early Chick Embryos ex vivo (New Culture)

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  the formation and patterning of brain, neural tube, somite and heart primordia. Applications of chick embryo culture include electroporation of DNA or RNA constructs in order to analyze gene function, grafts of growth factor coated beads such as FGFs and BMPs , as well as whole mount in situ hybridization and immunohistochemistry. This video demonstrates the different steps in chick embryo culture; First, the embryo is explanted in saline. Then, the embryo is centered on a glass ring. The membranes surrounding the embryo are lifted along the walls of the ring. The ring is then placed in a culture dish containing a pool of albumine. The culture dish is sealed and placed in a humid chamber, where the embryo is cultured for up to 24 hrs. Finally, the embryo is removed from the ring, fixed and processed for further applications. A troubleshooting guide is also presented.

Method for Culture of Early Chick Embryos ex vivo New Culture

This video demonstrates New culture, a method by which chick embryos are cultured outside the egg for up to 24 hr. This method enables one to study early development (primitive streak to 14 som.), a period corresponding to E7-9 in mouse. Applications of this technique include electroporation, in situ hybridization and immunohistochemistry.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

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 ...

Ex vivo Culture of Mouse Embryonic Skin and Live-imaging of Melanoblast Migration

Melanoblasts are the neural crest derived precursors of melanocytes; the cells responsible for producing the pigment in skin and hair. Melanoblasts migrate through the epidermis of the embryo where they subsequently colonize the developing hair follicles1,2. Neural crest cell migration is extensively studied in vitro but in vivo methods are still not well developed, especially in mammalian systems. One alternative is to use ex vivo organotypic culture3-6. Culture of mouse embryonic skin requires the maintenance of an air-liquid interface (ALI) across the surface of the tissue3,6. High resolution live-imaging of mouse embryonic skin has been hampered by the lack of a good method that not only maintains this ALI but also allows the culture to be inverted and therefore compatible with short working distance objective lenses and most confocal microscopes. This article describes recent improvements to a method that uses a gas permeable membrane to overcome these problems and allow high-resolution confocal imaging of embryonic skin in ex vivo culture6. By using a melanoblast specific Cre-recombinase expressing mouse line combined with the R26YFPR reporter line we are able to fluorescently label the melanoblast population within these skin cultures. The technique allows live-imaging of melanoblasts and observation of their behavior and interactions with the tissue in which they develop. Representative results are included to demonstrate the capability to live-image 6&#160;cultures in parallel.

Ex vivo Culture of Mouse Embryonic Skin and Live-imaging of Melanoblast Migration

We describe the dissection and ex vivo culture of mouse embryonic skin. The culture system maintains an air-liquid interface across the tissue surface and allows imaging on an inverted microscope. Melanoblasts, a component of the developing skin, are fluorescently labeled allowing their behavior to be observed using confocal microscopy.

Melanoblasts are the neural crest derived precursors of melanocytes; the cells responsible for producing the pigment in skin and hair. Melanoblasts ...

The c-FOS Protein Immunohistological Detection: A Useful Tool As a Marker of Central Pathways Involved in Specific Physiological Responses In Vivo and Ex Vivo

Many studies seek to identify and map the brain regions involved in specific physiological regulations. The proto-oncogene c-fos, an immediate early gene, is expressed in neurons in response to various stimuli. The protein product can be readily detected with immunohistochemical techniques leading to the use of c-FOS detection to map groups of neurons that display changes in their activity. In this article, we focused on the identification of brainstem neuronal populations involved in the ventilatory adaptation to hypoxia or hypercapnia. Two approaches were described to identify involved neuronal populations in vivo in animals and ex vivo in deafferented brainstem preparations. In vivo, animals were exposed to hypercapnic or hypoxic gas mixtures. Ex vivo, deafferented preparations were superfused with hypoxic or hypercapnic artificial cerebrospinal fluid. In both cases, either control in vivo animals or ex vivo preparations were maintained under normoxic and normocapnic conditions. The comparison of these two approaches allows the determination of the origin of the neuronal activation i.e., peripheral and/or central. In vivo and ex vivo, brainstems were collected, fixed, and sliced into sections. Once sections were prepared, immunohistochemical detection of the c-FOS protein was made in order to identify the brainstem groups of cells activated by hypoxic or hypercapnic stimulations. Labeled cells were counted in brainstem respiratory structures. In comparison to the control condition, hypoxia or hypercapnia increased the number of c-FOS labeled cells in several specific brainstem sites that are thus constitutive of the neuronal pathways involved in the adaptation of the central respiratory drive.

The c-FOS Protein Immunohistological Detection: A Useful Tool As a Marker of Central Pathways Involved in Specific Physiological Responses In Vivo and Ex Vivo

Here, we present a protocol based on c-FOS protein immunohistological detection, a classical technique used for the identification of neuronal populations involved in specific physiological responses in vivo and ex vivo.

Many studies seek to identify and map the brain regions involved in specific physiological regulations. The proto-oncogene c-fos, an immediate early gene, ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

Gap Junctional Intercellular Communication: A Functional Biomarker to Assess Adverse Effects of Toxicants and Toxins, and Health Benefits of Natural Products

This protocol describes a scalpel loading-fluorescent dye transfer (SL-DT) technique that measures intercellular communication through gap junction channels, which is a major intercellular process by which tissue homeostasis is maintained. Interruption of gap junctional intercellular communication (GJIC) by toxicants, toxins, drugs, etc. has been linked to numerous adverse health effects. Many genetic-based human diseases have been linked to mutations in gap junction genes. The SL-DT technique is a simple functional assay for the simultaneous assessment of GJIC in a large population of cells. The assay involves pre-loading cells with a fluorescent dye by briefly perturbing the cell membrane with a scalpel blade through a population of cells. The fluorescent dye is then allowed to traverse through gap junction channels to neighboring cells for a designated time. The assay is then terminated by the addition of formalin to the cells. The spread of the fluorescent dye through a population of cells is assessed with an epifluorescence microscope and the images are analyzed with any number of morphometric software packages that are available, including free software packages found on the public domain. This assay has also been adapted for in vivo studies using tissue slices from various organs from treated animals. Overall, the SL-DT assay can serve a broad range of in vitro pharmacological and toxicological needs, and can be potentially adapted for high throughput set-up systems with automated fluorescence microscopy imaging and analysis to elucidate more samples in a shorter time.

This protocol describes a scalpel loading-fluorescent dye transfer technique that measures intercellular communication through gap junction channels. Gap junctional intercellular communication is a major cellular process by which tissue homeostasis is maintained and disruption of this cell signaling has adverse health effects.

This protocol describes a scalpel loading-fluorescent dye transfer (SL-DT) technique that measures intercellular communication through gap junction ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Analysis of SCAP N-glycosylation and Trafficking in Human Cells

Elevated lipogenesis is a common characteristic of cancer and metabolic diseases. Sterol regulatory element-binding proteins (SREBPs), a family of membrane-bound transcription factors controlling the expression of genes important for the synthesis of cholesterol, fatty acids and phospholipids, are frequently upregulated in these diseases. In the process of SREBP nuclear translocation, SREBP-cleavage activating protein (SCAP) plays a central role in the trafficking of SREBP from the endoplasmic reticulum (ER) to the Golgi and in subsequent proteolysis activation. Recently, we uncovered that glucose-mediated N-glycosylation of SCAP is a prerequisite condition for the exit of SCAP/SREBP from the ER and movement to the Golgi. N-glycosylation stabilizes SCAP and directs SCAP/SREBP trafficking. Here, we describe a protocol for the isolation of membrane fractions in human cells and for the preparation of the samples for the detection of SCAP N-glycosylation and total protein by using western blot. We further provide a method to monitor SCAP trafficking by using confocal microscopy. This protocol is appropriate for the investigation of SCAP N-glycosylation and trafficking in mammalian cells.

Analysis of SCAP N-glycosylation and Trafficking in Human Cells

We describe a modified method for membrane fraction isolation from human cells and sample preparation for the detection of SCAP N-glycosylation and total protein by using western blot. We further introduce a GFP-labeling method to monitor SCAP trafficking using confocal microscopy. This protocol can be used in regular biology laboratories.

Elevated lipogenesis is a common characteristic of cancer and metabolic diseases. Sterol regulatory element-binding proteins (SREBPs), a family of ...

In Vitro and In Vivo Approaches to Determine Intestinal Epithelial Cell Permeability

The intestinal barrier defends against pathogenic microorganism and microbial toxin. Its function is regulated by tight junction permeability and epithelial cell integrity, and disruption of the intestinal barrier function contributes to progression of gastrointestinal and systemic disease. Two simple methods are described here to measure the permeability of intestinal epithelium. In vitro, Caco-2BBe cells are plated in tissue culture wells as a monolayer and transepithelial electrical resistance (TER) can be measured by an epithelial (volt/ohm) meter. This method is convincing because of its user-friendly operation and repeatability. In vivo, mice are gavaged with 4 kDa fluorescein isothiocyanate (FITC)-dextran, and the FITC-dextran concentrations are measured in collected serum samples from mice to determine the epithelial permeability. Oral gavage provides an accurate dose, and therefore is the preferred method to measure the intestinal permeability in vivo. Taken together, these two methods can measure the permeability of the intestinal epithelium in vitro and in vivo, and hence be used to study the connection between diseases and barrier function.

In Vitro and In Vivo Approaches to Determine Intestinal Epithelial Cell Permeability

Two methods are presented here to determine intestinal barrier function. An epithelial meter (volt/ohm) is used for measurements of transepithelial electrical resistance of cultured epithelia directly in tissue culture wells. In mice, the FITC-dextran gavage method is used to determine the intestinal permeability in vivo.

The intestinal barrier defends against pathogenic microorganism and microbial toxin. Its function is regulated by tight junction permeability and ...

A Pipeline using Bilateral In Utero Electroporation to Interrogate Genetic Influences on Rodent Behavior

As genome-wide association studies shed light on the heterogeneous genetic underpinnings of many neurological diseases, the need to study the contribution of specific genes to brain development and function increases. Relying on mouse models to study the role of specific genetic manipulations is not always feasible since transgenic mouse lines are quite costly and many novel disease-associated genes do not yet have commercially available genetic lines. Additionally, it can take years of development and validation to create a mouse line. In utero electroporation offers a relatively quick and easy method to manipulate gene expression in a cell-type specific manner in vivo that only requires developing a DNA plasmid to achieve a particular genetic manipulation. Bilateral in utero electroporation can be used to target large populations of frontal cortex pyramidal neurons. Combining this gene transfer method with behavioral approaches allows one to study the effects of genetic manipulations on the function of prefrontal cortex networks and the social behavior of juvenile and adult mice.

The role of recently discovered disease-associated genes in the pathogenesis of neuropsychiatric disorders remains obscure. A modified bilateral in utero electroporation technique allows for the gene transfer in large populations of neurons and examination of the causative effects of gene expression changes on social behavior.

As genome-wide association studies shed light on the heterogeneous genetic underpinnings of many neurological diseases, the need to study the contribution ...