Work in Lombard laboratory is supported by the NIH (NIGMS R01GM101171, NIEHS R21ES032305), DoD (CA190267, CA170628, NF170044, and ME200030), and Glenn Foundation for Medical Research. Work in Li laboratory is supported by NIH (NHLBI 5R01HL150707).

All vertebrate animal experiments were approved by and performed in accordance with the regulations of the University of Michigan Committee on Use and Care of Animals.
1. Day before the assay (Total time: ~10 min)
<ol>
	<li>Hydration of the sensor cartridge (Step time: ~10 min)
	<ol>
		<li>Open the extracellular flux assay kit and remove the sensor cartridge and utility plate assembly. Save loading guide flats for use the next day.</li>
		<li>Manually separate the sensor cartridge (the top green part with lid) from the utility plate (lower 96-well microplate) and place it upside-down next to the utility plate.</li>
		<li>Using a multichannel pipette, fill each well of the utility plate with 200 &#181;L of calibrant, included with flux assay kit.</li>
		<li>Place the sensor cartridge back onto the utility plate, making sure to completely submerge the sensors in the calibrant.</li>
		<li>Place the assembled sensor cartridge and utility plate with calibrant in a non-CO2 37 &#176;C incubator overnight. To prevent evaporation of the calibrant, ensure that the incubator is properly humidified. If using a regular oven incubator for non-CO2 37 &#176;C incubations, place an open beaker containing water inside the incubator to humidify it. If available, use an XF prep station for all non-CO2 37 &#176;C incubations. 
		&#8203;NOTE: Alternatively, the sensor cartridge can be hydrated overnight with 200 &#181;L per well of sterile ultrapure water in a non-CO2 37 &#176;C incubator. Replace water with 200 &#181;L per well of 37 &#176;C prewarmed calibrant, at least 45-60 min before loading the injection ports on the day of assay.</li>
	</ol>
	</li>
</ol>
2. Day of the assay (Total time: ~9 h 30 min)
NOTE: The total time indicated above includes cumulative durations of steps 2.1-2.4 in addition to either step 2.5 or step 2.6.
<ol>
	<li>Preparation of cell-adhesive-coated microplates (Step time: ~1 h 30 min) 
	NOTE: Perform all steps in a biosafety cabinet.
	<ol>
		<li>Prepare 2.5 mL of the cell adhesive solution (22.4 &#181;g/mL, see the Table of Materials) per 96-well cell culture microplate by dissolving 56 &#181;g of the cell adhesive in an appropriate volume of filter-sterilized 0.1 M sodium bicarbonate (pH 8.0) and immediately adding 1 N sodium hydroxide at half the volume of cell adhesive stock used. Vortex or pipette up and down to mix.</li>
		<li>Open the 96-well cell culture microplate (included with the flux assay kit) and dispense 25 &#181;L of the prepared cell adhesive solution at the bottom of each well. Cover the microplate with the lid and incubate for 30 min at room temperature inside the hood.</li>
		<li>After incubation, remove and discard excess cell adhesive solution using a multichannel pipette or aspirator, and wash each well twice with 200 &#181;L of sterile ultrapure water. Air-dry the plate, with the lid removed, inside the hood for 30-45 min. 
		NOTE: Cell-adhesive-coated cell culture microplates may be stored for up to one week at 4 &#176;C. Precoated microplates stored at 4 &#176;C must be allowed to warm to room temperature inside the hood before seeding cells.</li>
	</ol>
	</li>
	<li>Preparation of assay media (Step time: ~30 min)
	<ol>
		<li>Preparation of glycolysis stress test assay medium
		<ol>
			<li>Supplement 100 mL of base medium (see the Table of Materials) with 1 mL of 200 mM L-glutamine. 
			&#8203;NOTE: The final L-glutamine concentration in the assay medium is 2 mM.</li>
			<li>Warm the L-glutamine-supplemented medium to 37 &#176;C in a water bath.</li>
			<li>Adjust the pH of the warm medium to 7.4 with 1 N NaOH.</li>
			<li>Filter-sterilize the medium with a 0.2 &#181;m filter and keep it at 37 &#176;C until ready to use.</li>
		</ol>
		</li>
		<li>Preparation of mitochondrial stress test assay medium
		<ol>
			<li>Supplement 100 mL of the base medium with 0.45 g of glucose, 1 mL of 200 mM L-glutamine, and 1 mL of 100 mM sodium pyruvate. 
			NOTE: The final assay medium contains 25 mM glucose, 2 mM L-glutamine, and 1 mM sodium pyruvate.</li>
			<li>Warm glucose, L-glutamine, and the sodium pyruvate-supplemented medium to 37 &#176;C in a water bath.</li>
			<li>Adjust the pH of the warm medium to 7.4 with 1 N NaOH.</li>
			<li>Filter-sterilize medium with a 0.2 &#181;m filter and keep at 37 &#176;C until ready to use.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Harvest mouse lineage-negative HSPCs (Step time: ~3 h)
	<ol>
		<li>Prepare the assay buffer by supplementing Hank&#39;s balanced salt solution with 4% fetal bovine serum. Prepare 1x enrichment buffer by diluting the 5x stock with sterile distilled water. Keep both buffers on ice until use.</li>
		<li>Humanely euthanize mice using CO2 followed by cervical dislocation and remove the hindlimbs with scissors. Carefully remove all tissues from the bones, and cut the ends from both sides of the femur and tibia using scissors. Flush out bone marrow cells from the femur and tibia using the assay buffer. Pass the flushed cells through a sterile 70 &#181;m filter, centrifuge the filtrate at 200 &#215; g for 5 min at 4 &#176;C, and discard the supernatant.</li>
		<li>Lyse the red blood cells by resuspending the cell pellet in 500 &#181;L of ACK buffer (150 mM NH4Cl, 10 mM KHCO3, 0.1 mM EDTA, pH 7.2). Following lysis for 1 min on ice, wash the samples by adding 1 mL of assay buffer and centrifuge at 200 &#215; g for 5 min at 4 &#176;C. Discard the supernatant.</li>
		<li>After washing, incubate the samples with a cocktail of biotinylated antibodies directed against CD2 (1:200 dilution), CD3 (1:200 dilution), CD5 (1:200 dilution), CD8 (1:200 dilution), Ter-119 (1:200 dilution), B220 (1:200 dilution), and Gr-1 (1:800 dilution) for 45 min on ice in a total volume of 500 &#181;L of assay buffer.</li>
		<li>Wash the samples with 1 mL of assay buffer with centrifugation as above.</li>
		<li>Wash the samples with 1 mL of 1x enrichment buffer with centrifugation as above.</li>
		<li>Incubate the samples with 20 &#181;L of streptavidin nanobeads and 100 &#181;L of 1x enrichment buffer on ice for 15 min.</li>
		<li>Wash the samples with 1 mL of 1x enrichment buffer with centrifugation as above.</li>
		<li>Resuspend the samples in 2.5 mL of 1x enrichment buffer. Place them on a separation magnet for 5 min. Collect the supernatants separately in 15 mL conical tubes.</li>
		<li>Resuspend magnet-separated samples in an additional 2.5 mL of 1x enrichment buffer, and place them again on the separation magnet for 5 min. Collect and combine supernatants to the prior collections in 15 mL conical tubes from step 2.3.9. 
		NOTE: The supernatants contain the lineage-negative cell fractions.</li>
		<li>Centrifuge the 15 mL conical tubes containing the negatively selected cells for 5 min at 200 &#215; g at 4 &#176;C.</li>
		<li>Resuspend the cell pellets in 1 mL of assay buffer and count the cell number using trypan blue manually or via an automated cell counter such as a Countess 3. 
		NOTE: Following the above-described approach, ~6 &#215; 106 lineage-negative HSPCs should be readily purified from the hindlimbs of a single mouse. If reagents, such as assay buffer, 1x enrichment buffer, and lineage-specific antibody cocktail, are prepared on the day prior to the experiment, it takes approximately 2.5-3 h to enrich HSPCs from 4 mice. It would take an additional 10 min for each mouse beyond this number.</li>
	</ol>
	</li>
	<li>Seeding cells in cell-adhesive-coated microplates (Step time: ~1 h)
	<ol>
		<li>Centrifuge cells from step 2.3.12 at 200 &#215; g for 5 min at room temperature.</li>
		<li>Remove the supernatant and resuspend the cell pellet in the appropriate assay medium (glycolysis stress test medium or mitochondrial stress test medium). Centrifuge at 200 &#215; g for 5 min at room temperature.</li>
		<li>Repeat step 2.4.2. Remove the supernatant and resuspend the cells in the appropriate warmed assay medium to the concentration of 2.5 &#215; 105 cells per 50 &#181;L or 5 &#215; 106 per mL.</li>
		<li>Pipette 50 &#181;L of the cell suspension along the side of each well of the room-temperature cell-adhesive-coated 96-well cell culture microplate. Pipette 50 &#181;L of the assay medium into the corner background measurement wells. Use a multichannel pipette for cell plating to ensure consistency.</li>
		<li>Create a centrifuge balance plate by adding 50 &#181;L of water per well of a non-coated 96-well cell culture microplate.</li>
		<li>Centrifuge the cells in the plate at 200 &#215; g for 1 min at room temperature. Do not apply brake. Transfer the plate to a non-CO2 37 &#176;C incubator for 25-30 min to ensure the cells have completely attached. Visually confirm under a microscope that the cells are stably adhered to the microplate surface. 
		NOTE: As ~6 &#215; 106 lineage-negative HSPCs can be harvested from the hindlimbs of a single mouse, HSPCs isolated from 4 mice (~2.4 &#215; 107 cells)&#160;are needed to fill an entire 96-well plate if the aim is to screen multiple interventions ex vivo in parallel. However, if the aim is to investigate the impact of a genetic alteration or an intervention on the metabolic parameters of HSPCs in vivo, then HSPCs isolated from multiple pairs of control and test mice can be analyzed in multiple technical replicates in parallel using a single plate.</li>
	</ol>
	</li>
	<li>Performing glycolysis stress test using the extracellular flux analyzer (Step time: ~3 h 30 min)
	<ol>
		<li>Loading sensor cartridge with injecting compounds
		<ol>
			<li>Prepare 100 mM glucose, 20 &#181;M oligomycin, and 500 mM 2-DG solutions in prewarmed glycolysis stress test assay medium (pH 7.4). 
			NOTE: All injecting compound solutions are made at 10x concentration. The final well concentrations are 10 mM for glucose, 2 &#181;M for oligomycin, and 50 mM for 2-DG (see Table 1).</li>
			<li>Remove the hydrated sensor cartridge from the non-CO2 37 &#176;C incubator from step 1.1.5. Remove air bubbles from the calibrant in the utility plate by lifting the sensor cartridge out of the calibrant and replacing it on to the same utility plate with the calibrant.</li>
			<li>Place the A/D loading guide flat (included in the extracellular flux assay kit) on the top of the sensor cartridge, orienting it so that the letter &#39;A&#39; is located on the upper left-hand corner to ensure that only ports A are available for loading. Using a multichannel pipette, dispense 20 &#181;L of 100 mM glucose solution in ports A.</li>
			<li>Replace the A/D loading guide flat with the B/C loading guide flat, orienting so that the letter &#39;B&#39; is located on the upper left-hand corner to ensure that only ports B are available for loading. Using a multichannel pipette, dispense 22 &#181;L of 20 &#181;M oligomycin solution in ports B.</li>
			<li>Re-orient the B/C loading guide flat to locate the letter &#39;C&#39; on the upper left-hand corner for loading ports C. Using a multichannel pipette, dispense 25 &#181;L of 500 mM 2-DG solution in ports C.</li>
			<li>Remove and discard the loading guide flats. 
			NOTE: It is important that each port series of all 96 wells, including those of the background wells, be loaded completely with the same volume of the injecting compound.</li>
		</ol>
		</li>
		<li>Template creation, calibration, and measurements
		<ol>
			<li>Create or load the template for the glycolysis stress test on the controller. Enter details regarding injection strategies, treatment conditions, and cell types, and press generate groups. Go to the plate map and assign wells to each group to be analyzed. Assign the 4 corner wells for background measurements.</li>
			<li>In protocol, make sure calibration and equilibration are checked in the initialization step.</li>
			<li>For baseline measurement and measurements after each injection (glucose, oligomycin, and 2-DG), set the number of measurement cycles to 3 and Mix - Wait - Measure times to 3 min - 0 min - 3 min&#160;(see Table 2).</li>
			<li>Remove the lid from the compounds loaded and hydrated sensor cartridge, submerged in calibrant in the utility plate. Place it on the work tray of the extracellular flux analyzer and begin the run. 
			NOTE: The first step is the calibration, which usually takes ~20 min.</li>
			<li>Take out the microplate containing the cells from the non-CO2 37 &#176;C incubator from step 2.4.6 after 25-30 min of incubation is over. Without disturbing the cells, slowly add 130 &#181;L of the prewarmed glycolysis stress test assay medium (pH 7.4) per well to make up the medium volume in each well to 180 &#181;L and return the plate to the non-CO2 37 &#176;C incubator for additional 15-20 min. 
			NOTE: A total incubation time of 45-60 min after centrifugation is preferred.</li>
			<li>After the calibration is over, replace the utility plate with the assay microplate (without lid) containing the cells. Press Load cell plate&#160;to start the measurements, which will take ~1.5 h for completion of the assay.</li>
			<li>After the measurements are over, collect the assay microplate containing the cells and remove the assay medium without disturbing the cells. Wash once gently with 250 &#181;L of 1x phosphate-buffered saline without dislodging the cells.</li>
			<li>Add 10 &#181;L of RIPA lysis buffer (50 mM Tris-HCl, pH 7.4, 1% NP-40, 0.5 % Na deoxycholate, 0.1% sodium dodecylsulfate, 150 mM NaCl, 2 mM EDTA, 50 mM NaF), supplemented with 1x protease inhibitor cocktail, to each well and agitate the plate on a shaker for 10 min. Freeze the whole plate at -80 &#176;C.</li>
			<li>Thaw the plate, and perform a protein measurement assay for data normalization following the manufacturer&#39;s instructions.</li>
			<li>Retrieve and analyze the data. Open the data file using Wave Desktop Software. Click on Normalize and paste the normalization values for each well. Click on apply to normalize the data. Click on Export and select glycolysis stress test report generator to export the analyzed data to a report generator. Alternatively, export the data to an external application. 
			NOTE: Alternatively, the total nuclear DNA content in each well can be used to normalize the data. A fluorescent dye, such as CyQuant that binds to nucleic acid, can be used to assess the total nuclear DNA content per well.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Performing a mitochondrial stress test using the extracellular flux analyzer (Step time: ~3 h 30 min)
	<ol>
		<li>Loading sensor cartridge with injecting compounds
		<ol>
			<li>Prepare 20 &#181;M oligomycin, 20 &#181;M FCCP, and 5 &#181;M rotenone + 5 &#181;M antimycin A solutions in prewarmed mitochondrial stress test assay medium (pH 7.4). 
			NOTE: All injecting compound solutions are made at 10x concentration. The final well concentrations are 2 &#181;M for oligomycin, 2 &#181;M for FCCP, and 0.5 &#181;M each for rotenone and antimycin A (see Table 3).</li>
			<li>Remove the hydrated sensor cartridge from the 37 &#176;C incubator from step 1.1.5 and remove air bubbles as described previously in step 2.5.1.2.</li>
			<li>By following steps 2.5.1.3 to 2.5.1.6, dispense 20 &#181;L of 20 &#181;M oligomycin in ports A, 22 &#181;L of 20 &#181;M FCCP in ports B, and 25 &#181;L of a mixture of 5 &#181;M rotenone and 5 &#181;M antimycin A in ports C. 
			NOTE: It is important that each port series of all 96 wells, including those of the background wells, be loaded completely with the same volume of injecting compound.</li>
		</ol>
		</li>
		<li>Template creation, calibration, and measurements
		<ol>
			<li>Create or load the template for the mitochondrial stress test on the controller. Enter details regarding injection strategies, treatment conditions, and cell types, and press generate groups. Go to the plate map and assign wells to each group to be analyzed. Assign the 4 corner wells for background measurements.</li>
			<li>In protocol, make sure calibration and equilibration are checked in the initialization step.</li>
			<li>For baseline measurement and measurements after each injection (oligomycin, FCCP, and rotenone + antimycin A), set the number of measurement cycles to 3 and Mix - Wait - Measure times to 3 min - 0 min - 3 min (see Table 4).</li>
			<li>Start the run to perform the calibration as in step 2.5.2.4.</li>
			<li>Add 130 &#181;L of the prewarmed mitochondrial stress test assay medium (pH 7.4) per well as in step 2.5.2.5.</li>
			<li>Start the measurements; retrieve and analyze the data as in steps 2.5.2.6 to 2.5.2.10. Export the analyzed data to the mitochondrial stress test report generator or an external application.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors declare that there is no conflict of interest.

This method paper describes an optimized protocol for the assessment of cellular bioenergetics (glycolysis and OxPHOS) in mouse HSPCs using the Seahorse extracellular flux analyzer. This device is a powerful tool that simultaneously measures the ECAR and OCR of live cells, which are metrics of glycolysis and mitochondrial respiration, respectively. Thus, it can be used to assess the cellular bioenergetics in real time. Further, the 96-well microplate-based platform offers high-throughput quantification with high sensitiv...

Hematopoiesis is the process by which various types of mature blood cells with highly specialized functions are formed from HSCs1. HSCs are capable of self-renewal and differentiation into various multipotent and lineage-specific progenitor populations. These progenitors ultimately produce cells of lymphoid, myeloid, erythroid, and megakaryocyte lineages. To maintain their self-renewal capacity, HSCs remain largely quiescent and, like other tissue stem cells, are believed to rely on glycolysis rather than mitochondrial OxPHOS for ATP production2,3. Entry into the cell cycle leads to enhanced respiration and OxPHOS, resulting in elevated levels of reactive oxygen species (ROS), which are detrimental to HSC maintenance and function3. Repeated cell division thus may lead to reduced self-renewal capacity of HSCs and ultimately to their exhaustion.
In adult hematopoiesis, HSCs primarily undergo asymmetric cell division, during which one of the daughter cells retains HSC potential and continues to rely on glycolytic metabolism. The other daughter cell becomes a primitive progenitor cell that loses self-renewal capacity but proliferates and eventually gives rise to differentiated functional hematopoietic cells4. When HSCs differentiate to produce downstream progenitors, a switch from glycolysis to mitochondrial metabolism is thought to occur to supply the energy and building blocks needed to support this rapid transition5, as suggested by the observations that HSCs possess inactive mitochondrial mass6,7,8,9. In contrast, mitochondrial activity (indicated by linked ROS levels) is much higher in lineage-committed progenitors than in HSCs9,10,11. Metabolic changes that occur during the earliest step of hematopoiesis thus suggest a direct and crucial role of mitochondria in regulating HSC fate.
Dysfunctional mitochondria present under various stress conditions, such as aging, cancer, diabetes, and obesity12, can interfere with HSC self-renewal capacity, inducing an imbalance in HSC/progenitor differentiation by producing excessive amounts of ROS, impairing ATP production, and/or by altering other metabolic processes9,12,13. Perturbations in metabolic homeostasis in HSC/progenitor differentiation could significantly impact hematopoiesis, potentially contributing to the development of hematologic abnormalities13. Given the critical influences of glycolysis and mitochondrial OxPHOS on HSC stemness and differentiation, it is of interest to investigate both metabolic parameters under normal and stress conditions. Valuable insights into the glycolytic and mitochondrial function of HSCs and progenitor cells can be gained by assessing their ECAR and OCR, which are indicators of cellular glycolysis and mitochondrial respiration, respectively.
The Seahorse extracellular flux analyzer is a powerful apparatus equipped with two probes per well to simultaneously measure ECAR and OCR in live cells and thus can be used to assess cellular bioenergetics in real time, in response to various substrates or inhibitors. The assay cartridge used with the analyzer contains injection ports to hold up to four drugs for automated injection during the assay. A scheme of a typical glycolysis stress test is shown in Figure 1A. The assay starts with the measurement of ECAR of cells, incubated in glycolysis stress test medium containing glutamine but not glucose or pyruvate. This represents acidification occurring due to non-glycolytic activities of the cells and is reported as non-glycolytic acidification. This is followed by the injection of glucose at a saturating concentration. Via glycolysis, glucose in the cell is converted into pyruvate, which is further metabolized in the cytoplasm to produce lactate, or in mitochondria to produce CO2 and water.
Conversion of glucose to lactate causes net production and subsequent release of protons into the extracellular medium, resulting in a rapid increase in the ECAR14,15,16. This glucose-stimulated change in ECAR is reported as glycolysis under basal conditions. The second injection consists of oligomycin (an inhibitor of ATP synthase, a.k.a. complex V17), which inhibits mitochondrial ATP production. During oligomycin-mediated OxPHOS inhibition, cells maximally upregulate glycolysis to meet their energetic demands. This causes a further increase in ECAR, revealing the maximum glycolytic capacity of the cells. The difference between the maximum glycolytic capacity and basal glycolysis is referred to as glycolytic reserve. Finally, 2-DG is injected, which causes a significant drop in ECAR, usually close to non-glycolytic acidification levels. 2-DG is a glucose analog that competitively binds to hexokinase, resulting in inhibition of glycolysis18. Thus, the 2-DG-induced decrease in ECAR further confirms that glycolysis is indeed the source of ECAR observed after glucose and oligomycin injections.
Figure 1B displays the schematic for a typical mitochondrial stress test. The assay starts with baseline OCR measurement of the cells, incubated in mitochondrial stress test medium containing glucose, glutamine, and pyruvate. Following basal OCR measurements, oligomycin is injected in this assay, which inhibits complex V, thereby reducing electron flow through the electron transport chain (ETC)17. Consequently, OCR is reduced in response to oligomycin injection, and this decrease in OCR is linked to mitochondrial ATP production. The second injection consists of carbonyl cyanide-4 (trifluoromethoxy) phenylhydrazone (FCCP), a protonophore and an uncoupler of mitochondrial OxPHOS17. FCCP collapses the mitochondrial proton gradient by allowing the flow of protons across the mitochondrial inner membrane. Because of the FCCP injection, electron flow through the ETC is derepressed, and complex IV consumes oxygen at the maximal level. The difference between maximal OCR and basal OCR is referred to as the spare respiratory capacity, which is a measure of the cell&#39;s ability to respond to increased energy demand under stress conditions. Finally, a mixture of two ETC inhibitors (rotenone, a complex I inhibitor, and antimycin A, a complex III inhibitor17) is injected, which completely shuts down electron flow, and OCR decreases to a low level. OCR measured after rotenone and antimycin A injection corresponds to non-mitochondrial OCR driven by other processes inside the cells. Non-mitochondrial OCR enables the calculation of basal respiration, proton leak, and maximal respiration.
Basal respiration represents the difference between baseline OCR and non-mitochondrial OCR. Proton leak refers to the difference between OCR after oligomycin injection and non-mitochondrial OCR. Maximal respiration represents the difference between OCR after FCCP injection and non-mitochondrial OCR. Coupling efficiency is calculated as the percentage of ATP production rate to basal respiration rate. This method paper provides a detailed protocol to measure ECAR and OCR in lineage-negative HSPCs using the Seahorse XFe96 extracellular flux analyzer. This protocol describes approaches to isolate mouse lineage-negative HSPCs, explains the optimization of cell-seeding density and concentrations of various drugs used in extracellular flux assays, and describes drug treatment strategies.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.2 &#956;m filter</td>
			<td>Corning</td>
			<td>430626</td>
			<td>Used to filter-sterilize the assay media</td>
		</tr>
		<tr>
			<td>100 mM sodium pyruvate</td>
			<td>Life Technologies</td>
			<td>11360-070</td>
			<td>Component of mitochondrial stress test assay medium</td>
		</tr>
		<tr>
			<td>15 mL conical Falcon tubes</td>
			<td>Corning</td>
			<td>352096</td>
			<td>Used during HSPCs harvest and to prepare assay drug solutions</td>
		</tr>
		<tr>
			<td>200 mM L-glutamine</td>
			<td>Life Technologies</td>
			<td>25030-081</td>
			<td>Component of glycolysis stress test and mitochondrial stress test assay media</td>
		</tr>
		<tr>
			<td>2-Deoxy-D-glucose (2-DG)</td>
			<td>Sigma-Aldrich</td>
			<td>D8375</td>
			<td>3rd drug injection during glycolysis stress test</td>
		</tr>
		<tr>
			<td>5x Enrichment buffer (MojoSort)</td>
			<td>Biolegend</td>
			<td>480017</td>
			<td>Used for washings during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Ammonium chloride (NH4Cl)</td>
			<td>Fisher Scientific</td>
			<td>A661-3</td>
			<td>Component of ACK lysis buffer</td>
		</tr>
		<tr>
			<td>Antimycin A</td>
			<td>Sigma-Aldrich</td>
			<td>A8674</td>
			<td>3rd drug injection during mitochondrial stress test</td>
		</tr>
		<tr>
			<td>Bio-Rad DC protein assay kit</td>
			<td>Bio-Rad</td>
			<td>500-0112</td>
			<td>Used as per manufacturer&#39;s instructions</td>
		</tr>
		<tr>
			<td>Carbonyl cyanide-4 (trifluoromethoxy) phenylhydrazone (FCCP)</td>
			<td>Sigma-Aldrich</td>
			<td>C2920</td>
			<td>2nd drug injection during mitochondrial stress test</td>
		</tr>
		<tr>
			<td>Cell-Tak</td>
			<td>Corning</td>
			<td>354240</td>
			<td>Cell adhesive. Used for coating cell microplates</td>
		</tr>
		<tr>
			<td>Countes 3 Automated Cell Counter</td>
			<td>ThermoFisher Scientific</td>
			<td></td>
			<td>For cell counting</td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Fisher Scientific</td>
			<td>O2793-500</td>
			<td>Component of ACK lysis buffer and RIPA lysis buffer</td>
		</tr>
		<tr>
			<td>Falcon 70 &#956;m filter</td>
			<td>Fisher Scientific</td>
			<td>08-771-2</td>
			<td>Used as cells strainer during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Gibco Fetal bovine serum (FBS)</td>
			<td>Fisher Scientific</td>
			<td>26400044</td>
			<td>Used to prepare assay buffer during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Gibco HBSS</td>
			<td>Fisher Scientific</td>
			<td>14175095</td>
			<td>Used to prepare assay buffer during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma-Aldrich</td>
			<td>G7528</td>
			<td>Component of mitochondrial stress test assay medium and first injection of glycolysis stress test</td>
		</tr>
		<tr>
			<td>Oligomycin</td>
			<td>Sigma-Aldrich</td>
			<td>O4876</td>
			<td>2nd drug injection during glycolysis stress test and 1st drug injection during mitochondrial stress test</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Life Technologies</td>
			<td>10010-049</td>
			<td>Used to wash cells after assay for protein quantification</td>
		</tr>
		<tr>
			<td>Potassium bicarbonate (KHCO3)</td>
			<td>Fisher Scientific</td>
			<td>P235-500</td>
			<td>Component of ACK lysis buffer</td>
		</tr>
		<tr>
			<td>Protease Inhibitor Cocktail (PIC)</td>
			<td>Roche</td>
			<td>11836170001</td>
			<td>Supplied as tablets. One tablet was dissolved in 10 mL of RIPA buffer to make 1x PIC.</td>
		</tr>
		<tr>
			<td>Rat biotin antimouse-B220, Clone ID: RA3-6B2</td>
			<td>Biolegend</td>
			<td>103203</td>
			<td>Used for lineage depletion during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Rat biotin antimouse-CD2, Clone ID: RM2-5</td>
			<td>Biolegend</td>
			<td>100103</td>
			<td>Used for lineage depletion during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Rat biotin antimouse-CD3, Clone ID: 17A2</td>
			<td>Biolegend</td>
			<td>100243</td>
			<td>Used for lineage depletion during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Rat biotin antimouse-CD5, Clone ID: 53-7.3</td>
			<td>Biolegend</td>
			<td>100603</td>
			<td>Used for lineage depletion during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Rat biotin antimouse-CD8, Clone ID: 53-6.7</td>
			<td>Biolegend</td>
			<td>100703</td>
			<td>Used for lineage depletion during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Rat biotin antimouse-Gr-1, Clone ID: RB6-8C5</td>
			<td>Biolegend</td>
			<td>108403</td>
			<td>Used for lineage depletion during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Rat biotin antimouse-Ter-119, Clone ID: TER-119</td>
			<td>Biolegend</td>
			<td>116203</td>
			<td>Used for lineage depletion during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Rotenone</td>
			<td>Sigma-Aldrich</td>
			<td>R8875</td>
			<td>3rd drug injection during mitochondrial stress test</td>
		</tr>
		<tr>
			<td>Seahorse XFe96 extracellular flux analyzer</td>
			<td>Seahorse Biosciences now Agilent</td>
			<td></td>
			<td>For ECAR and OCR measurments in real time.</td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Sigma-Aldrich</td>
			<td>S5761</td>
			<td>Used to make Cell-adhesive solution for microplate coating</td>
		</tr>
		<tr>
			<td>Sodium chloride (NaCl)</td>
			<td>Fisher</td>
			<td>BP358</td>
			<td>Component of RIPA lysis buffer</td>
		</tr>
		<tr>
			<td>Sodium deoxycholate</td>
			<td>Sigma-Aldrich</td>
			<td>D6750</td>
			<td>Component of RIPA lysis buffer</td>
		</tr>
		<tr>
			<td>Sodium Fluoride (NaF)</td>
			<td>Sigma-Aldrich</td>
			<td>S7920</td>
			<td>Component of RIPA lysis buffer</td>
		</tr>
		<tr>
			<td>Sodium hydroxide (NaOH)</td>
			<td>Sigma-Aldrich</td>
			<td>S8045</td>
			<td>Prepared 1 N solution. Used for pH normalization</td>
		</tr>
		<tr>
			<td>Streptavidin Nanobeads (MojoSort)</td>
			<td>Biolegend</td>
			<td>480015</td>
			<td>Used for lineage depletion during HSPCs harvest</td>
		</tr>
		<tr>
			<td>Tris-HCl</td>
			<td>Fisher</td>
			<td>BP153</td>
			<td>Component of RIPA lysis buffer</td>
		</tr>
		<tr>
			<td>XF base medium</td>
			<td>Agilent</td>
			<td>102353-100</td>
			<td>base medium used to prepare glycolysis stress test and mitochondrial stress test assay media</td>
		</tr>
		<tr>
			<td>XF prep station</td>
			<td>Seahorse Biosciences</td>
			<td></td>
			<td>Used for non-CO2 37 &#176;C incubations</td>
		</tr>
		<tr>
			<td>XFe96 extracellular FluxPak</td>
			<td>Agilent</td>
			<td>102416-100 or 102601-100</td>
			<td>Includes assay cartridges with utility plates, loading guide flats for loading 
			drugs onto the assay cartridge, XF calibrant solution, and XF cell culture microplate</td>
		</tr>
	</tbody>
</table>

assessment of cellular bioenergetics in mouse hematopoietic stem and primitive progenitor cells using the extracellular flux analyzer

Under steady state, hematopoietic stem cells (HSCs) remain largely quiescent and are believed to be predominantly reliant on glycolysis to meet their energetic needs. However, under stress conditions such as infection or blood loss, HSCs become proliferative and rapidly produce downstream progenitor cells, which in turn further differentiate, ultimately producing mature blood cells. During this transition and differentiation process, HSCs exit from quiescence and rapidly undergo a metabolic switch from glycolysis to oxidative phosphorylation (OxPHOS). Various stress conditions, such as aging, cancer, diabetes, and obesity, can negatively impact mitochondrial function and thus can alter the metabolic reprogramming and differentiation of HSCs and progenitors during hematopoiesis. Valuable insights into glycolytic and mitochondrial functions of HSCs and progenitors under normal and stress conditions can be gained through the assessment of their extracellular acidification rate (ECAR) and oxygen consumption rate (OCR), which are indicators of cellular glycolysis and mitochondrial respiration, respectively. 
Here, a detailed protocol is provided to measure ECAR and OCR in mouse bone marrow-derived lineage-negative cell populations, which include both hematopoietic stem and primitive progenitor cells (HSPCs), using the extracellular flux analyzer. This protocol describes approaches to isolate lineage-negative cells from mouse bone marrow, explains optimization of cell seeding density and concentrations of 2-deoxy-D-glucose (2-DG, a glucose analog that inhibits glycolysis) and various OxPHOS-targeted drugs (oligomycin, FCCP, rotenone, and antimycin A) used in these assays, and describes drug treatment strategies. Key parameters of glycolytic flux, such as glycolysis, glycolytic capacity, and glycolytic reserve, and OxPHOS parameters, such as basal respiration, maximal respiration, proton leak, ATP production, spare respiratory capacity, and coupling efficiency, can be measured in these assays. This protocol allows ECAR and OCR measurements on non-adherent HSPCs and can be generalized to optimize analysis conditions for any type of suspension cells.

Using this protocol, the cell number and the concentrations of various OxPHOS-targeting drugs (used in the extracellular flux assays) were optimized to measure ECAR and OCR of HSPCs isolated from 24-week-old female C57BL/6 mice. First, the glycolysis stress test was performed to optimize cell number and oligomycin concentration. A varying number of HSPCs per well ranging from 5 &#215; 104 to 2.5 &#215; 105 were used in this assay. As shown in Figure 2A and <strong class...

Watch this Scientific Journal Video about Assessment of Cellular Bioenergetics in Mouse Hematopoietic Stem and Primitive Progenitor Cells using the Extracellular Flux Analyzer at JoVE.com

Assessment of Cellular Bioenergetics in Mouse Hematopoietic Stem and Primitive Progenitor Cells using the Extracellular Flux Analyzer

The method presented here summarizes optimized protocols for assessing cellular bioenergetics in non-adherent mouse hematopoietic stem and primitive progenitor cells (HSPCs) using the extracellular flux analyzer to measure the extracellular acidification rate (ECAR) and oxygen consumption rate (OCR) of HSPCs in real time.

Under steady state, hematopoietic stem cells (HSCs) remain largely quiescent and are believed to be predominantly reliant on glycolysis to meet their ...

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JoVE Journal

Biology

2.5K Views.  University of Michigan, Ann Arbor. The method presented here summarizes optimized protocols for assessing cellular bioenergetics in non-adherent mouse hematopoietic stem and primitive progenitor cells (HSPCs) using the extracellular flux analyzer to measure the extracellular acidification rate (ECAR) and oxygen consumption rate (OCR) of HSPCs in real time.

Video: Assessment of Cellular Bioenergetics in Mouse Hematopoietic Stem and Primitive Progenitor Cells using the Extracellular Flux Analyzer

Isolation and Transplantation of Hematopoietic Stem Cells (HSCs)

Isolation and Transplantation of Hematopoietic Stem Cells HSCs

I am Christina Ceso. I am a postdoc in David's Den lab and I work on the hematopoietic stem cells. Today we are going to purify long-term recuperating hematopoietic stem cells and we'll transplant them into a mouse.We start by spraying the mice with alcohol so that first of all, the mind starts sterilized and also the hair doesn't interfere with the dissection. We use PBS with 2%serum for the dissection. I cut the back skin and I pull the skin apart in order to expose the tissue.In order to isolate the tibia, I hold the tibia with tweezers. I cut the tendon on top of the knee and I cut the muscles below the knee and after that I cut through the knee so that the TBI is not connected to the rest of the limb anymore and I pull in order to isolate it. I then hold the TB upside down and with the scissors I separate the foot from the tibia In order to obtain the femur.I cut the muscles around the femur and then I dislocate the femur from the hip. In order to isolate the hip, I cut the muscle around the hip bone on the side of the tail of the mouse and then I pull the hip away from the body of the mouth In order to isolate the spine. I cut through the tail as close as I can to the beginning of the back and then I cut on the two sides of the spine and below the spine, trying to stay as close as possible to the spine so that the peritoneum is not cut and the intestine is left untouched.A first round of cleaning of the bones is done with a scalpel, which is used to scrape the bone, the muscles away from the bone. The scalpel is good to clean the femurs, the hip and the spine. In order to clean the tibia, I hold them with forceps and I use the forceps to remove as much tissue as possible.A second round of cleaning is done using clean wipe to completely clean the bones outta the muscle, and I use the F wipe to clean the tibia, the femurs and the hips. So this is how bones look. After cleaning, I prefer 50 amount tubes with filters on top and I transfer the bones into the mortar.I add the section media and I crush the bones ask of the first round of crashing of the bones. I piped the mixture in order to release as many styles as possible into solution and reduce clumps. I transferred the mixture on the tubes through the filters and this is what the bone fragments look like after the first round of crushing.So I add more dissection medium to the bone fragments and I flash them again. I transferred the seline mixture to the tubes through the filters and at this point the remaining bone fragments are white and there is no bone marrow left. The tubes are filled with PBS 2%serum and in order to have them all with the same weight so that they're balanced in the centrifuge, but also in order to wash the filters so that we obtain as many cells as possible, I then centrifuge the cell mixture and after centrifugation, the palt includes the bone marrow cells.I aspirate the medium and I move back and forward the tube on the rack so that the pelle is becomes loose and fewer clamps are formed. When I add medium, at this point I add one ml of medium per tube and I ics since I'm sorting hematopoietic stem cells. Later on, I'm going to store an ALI output of total bone marrow sizes at this point, which I will use later for the fact single color control.I add lineage antibody cocktail. I ordered the samples to mix them well and I invaded four degrees for 15 minutes. In the meantime, I start preparing the columns for the lineage division.I put the columns on the magnet and I prepare Ian tubes under the columns. I also prepared the gas PBS using aster flip filter and attaching it to the vacuum so that all the bubbles of air move in the top part of the filter. I then add three ml of the gas DBS to each column in order for the column to become wet.After the incubation with the antibodies, I add medium to the cell samples and I centrifuge them in order to remove the excess of antibodies. After the centrifugation, the stain cells are in the pallet. I aspirate the medium and I suspend the pallet in one mile of the gas PBSI Add the strip tab beans, I mix the cells and I incubate for 15 minutes at four degrees.After the incubation, I put new elucian tubes under the columns. I add two ml of PBS to each tube with the cells and I transferred the cell mixture to the column through a 40 micron filter. I wash the tubes with more medium in order to collect all the cells remaining and I apply the remaining medium through the filters to the columns.The progenitor cells lineage low are alluded into the collection views. I pulled the luted samples and I sent tofu them. So after the centrifugation, the depleted cells are in a white pate.I aspirate the medium and re suspend the cells. I store an adequate of the depleted cells and I add the antibodies to the cell mixture for the sorting and also to all the single color controls. The antibodies are incubated in that with the size for 15 minutes in the fridge.After the ation, I add medium to the cells and and centrifuge them to remove the ex excess of antibodies. After the centrifugation, the cells are in the pt, I aspirate the media that I suspend the cells and I transfer the cells into fox tubes with a blue filter. At this point, the cells are ready to be sorted.I place, I place the mouse under a heating lamp for a few minutes so that the veins dilate and they are more easily seen. When preparing the syringe, you have to be careful to eliminate all the bubbles from the, the mixture that is in the syringe and then you bend the needle at 90 degrees. I put the mouse under a glass cup with a weight on top so that it can run away.I sterilize the tail with the alcohol pad so that the site of injection is sterile, but also it's easier to identify the vein. If you look at the tail of the mouse from the top in the middle and top of the tail, there is an artery while on the two sides there are the two veins and we inject the cells in one of the two veins. I insert the needle into the vein and I inject following injection.It is to wait for a couple of seconds before removing the needle, the needle, and after that I clean the side of injection. This is a representation of the syringe with the ben needle bent at 90 degrees with the knee, the at this ankle compared to the tail, which makes the injection the easiest. When you injecting the tail of the mouse, you have to keep in mind that the tail vein is extremely superficial, so really this is the orientation of the needle compared to the tail.And it looks like you will just keep sliding on top of the tail, but actually you will get into the tail then. I am Christina Ceso. I am a post in David Kaden lab, and I work on the hematopoietic stem cells.I usually use for animals to start with, and I had more variability at the beginning, but once you become consistent with your methods, I consistently get about 50, 000 to 60, 000 stem cells out for mice. I did my PhD in London with Dr.Fiona Wat working on derma stem cells. After that, I just wanted to have a wider knowledge of adult stem cells in various tissues, and so I decided to move to David Kaden lab to work on hematopoietic stem cells, mainly because the things that are known about hematopoietic stem cells are the things that are not known for the epidermic stem cells and a number of assays that can be done with epi.Epidermal stem cells are not really done yet with the hematopoietic stem cells, and so at this point I'm trying to put together the, the knowledge from the two different fields and ultimately I'm interested in the study of the mechanism to regulate adult tissue sensors in general, so not just the epidermis or the stem cells, but also different tissues because there are, there are similar mechanism that is, it's always the same genes that are involved in the regulation that just interact with each other in different ways, and it's just interesting to understand what makes each stem cell be a stem, but then ultimately a stem cell in a different tissue. So giving rise to a different protein.

Isolation and Analysis of Hematopoietic Stem Cells from the Placenta

Hematopoietic stem cells (HSCs) have the ability to self-renew and generate all cell types of the blood lineages throughout the lifetime of an individual. All HSCs emerge during embryonic development, after which their pool size is maintained by self-renewing cell divisions. Identifying the anatomical origin of HSCs and the critical developmental events regulating the process of HSC development has been complicated as many anatomical sites participate during fetal hematopoiesis. Recently, we identified the placenta as a major hematopoietic organ where HSCs are generated and expanded in unique microenvironmental niches (Gekas, et al 2005, Rhodes, et al 2008). Consequently, the placenta is an important source of HSCs during their emergence and initial expansion.
In this article, we show dissection techniques for the isolation of murine placenta from E10.5 and E12.5 embryos, corresponding to the developmental stages of initiation of HSCs and the peak in the size of the HSC pool in the placenta, respectively. In addition, we present an optimized protocol for enzymatic and mechanical dissociation of placental tissue into single-cell suspension for use in flow cytometry or functional assays. We have found that use of collagenase for single-cell suspension of placenta gives sufficient yields of HSCs. An important factor affecting HSC yield from the placenta is the degree of mechanical dissociation prior to, and duration of, enzymatic treatment.
We also provide a protocol for the preparation of fixed-frozen placental tissue sections for the visualization of developing HSCs by immunohistochemistry in their precise cellular niches. As hematopoietic specific antigens are not preserved during preparation of paraffin embedded sections, we routinely use fixed frozen sections for localizing placental HSCs and progenitors.

We have identified the placenta as a major hematopoietic organ during development. We found that hematopoietic stem cells (HSCs) are both generated and expanded in the placenta in unique microenvironmental niches. Here, we describe experimental techniques required for isolation and visualization of HSCs in the mouse placenta.

Hematopoietic stem cells (HSCs) have the ability to self-renew and generate all cell types of the blood lineages throughout the lifetime of an individual. ...

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 ...

Phenotypic Analysis and Isolation of Murine Hematopoietic Stem Cells and Lineage-committed Progenitors

The bone marrow is the principal site where HSCs and more mature blood cells lineage progenitors reside and differentiate in an adult organism. HSCs constitute a minute cell population of pluripotent cells capable of generating all blood cell lineages for a life-time1. The molecular dissection of HSCs homeostasis in the bone marrow has important implications in hematopoiesis, oncology and regenerative medicine. We describe the labeling protocol with fluorescent antibodies and the electronic gating procedure in flow cytometry to score hematopoietic progenitor subsets and HSCs distribution in individual mice (Fig. 1). In addition, we describe a method to extensively enrich hematopoietic progenitors as well as long-term (LT) and short term (ST) reconstituting HSCs from pooled bone marrow cell suspensions by magnetic enrichment of cells expressing c-Kit. The resulting cell preparation can be used to sort selected subsets for in vitro and in vivo functional studies (Fig. 2).
Both trabecular osteoblasts2,3 and sinusoidal endothelium4 constitute functional niches supporting HSCs in the bone marrow. Several mechanisms in the osteoblastic niche, including a subset of N-cadherin+ osteoblasts3 and interaction of the receptor tyrosine kinase Tie2 expressed in HSCs with its ligand angiopoietin-15 concur in determining HSCs quiescence. "Hibernation" in the bone marrow is crucial to protect HSCs from replication and eventual exhaustion upon excessive cycling activity6. Exogenous stimuli acting on cells of the innate immune system such as Toll-like receptor ligands7 and interferon-&alpha;6 can also induce proliferation and differentiation of HSCs into lineage committed progenitors. Recently, a population of dormant mouse HSCs within the lin- c-Kit+ Sca-1+ CD150+ CD48- CD34- population has been described8. Sorting of cells based on CD34 expression from the hematopoietic progenitors-enriched cell suspension as described here allows the isolation of both quiescent self-renewing LT-HSCs and ST-HSCs9. A similar procedure based on depletion of lineage positive cells and sorting of LT-HSC with CD48 and Flk2 antibodies has been previously described10. In the present report we provide a protocol for the phenotypic characterization and ex vivo cell cycle analysis of hematopoietic progenitors, which can be useful for monitoring hematopoiesis in different physiological and pathological conditions. Moreover, we describe a FACS sorting procedure for HSCs, which can be used to define factors and mechanisms regulating their self-renewal, expansion and differentiation in cell biology and signal transduction assays as well as for transplantation.

A method to analyse the distribution of bone marrow hematopoietic progenitors in flow cytometry as well as to efficiently isolate highly purified hematopoietic stem cells (HSCs) is described. The isolation procedure is essentially based on magnetic enrichment of c-Kit+ cells and cell sorting to purify HSCs for cellular and molecular studies.

The bone marrow is the principal site where HSCs and more mature blood cells lineage progenitors reside and differentiate in an adult organism. HSCs ...

Generation of Mice Derived from Induced Pluripotent Stem Cells

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also produce a source of patient-matched cells for regenerative therapies. iPSCs may be generated using multiple protocols and derived from multiple cell sources. Once generated, iPSCs are tested using a variety of assays including immunostaining for pluripotency markers, generation of three germ layers in embryoid bodies and teratomas, comparisons of gene expression with embryonic stem cells (ESCs) and production of chimeric mice with or without germline contribution2. Importantly, iPSC lines that pass these tests still vary in their capacity to produce different differentiated cell types2. This has made it difficult to establish which iPSC derivation protocols, donor cell sources or selection methods are most useful for different applications.
The most stringent test of whether a stem cell line has sufficient developmental potential to generate all tissues required for survival of an organism (termed full pluripotency) is tetraploid embryo complementation (TEC)3-5. Technically, TEC involves electrofusion of two-cell embryos to generate tetraploid (4n) one-cell embryos that can be cultured in vitro to the blastocyst stage6. Diploid (2n) pluripotent stem cells (e.g. ESCs or iPSCs) are then injected into the blastocoel cavity of the tetraploid blastocyst and transferred to a recipient female for gestation (see Figure 1). The tetraploid component of the complemented embryo contributes almost exclusively to the extraembryonic tissues (placenta, yolk sac), whereas the diploid cells constitute the embryo proper, resulting in a fetus derived entirely from the injected stem cell line.
Recently, we reported the derivation of iPSC lines that reproducibly generate adult mice via TEC1. These iPSC lines give rise to viable pups with efficiencies of 5-13%, which is comparable to ESCs3,4,7 and higher than that reported for most other iPSC lines8-12. These reports show that direct reprogramming can produce fully pluripotent iPSCs that match ESCs in their developmental potential and efficiency of generating pups in TEC tests. At present, it is not clear what distinguishes between fully pluripotent iPSCs and less potent lines13-15. Nor is it clear which reprogramming methods will produce these lines with the highest efficiency. Here we describe one method that produces fully pluripotent iPSCs and "all- iPSC" mice, which may be helpful for investigators wishing to compare the pluripotency of iPSC lines or establish the equivalence of different reprogramming methods.

Generating induced pluripotent stem cell (iPSC) lines produces lines of differing developmental potential even when they pass standard tests for pluripotency. Here we describe a protocol to produce mice derived entirely from iPSCs, which defines the iPSC lines as possessing full pluripotency1.

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Isolation, Characterization, And High Throughput Extracellular Flux Analysis of Mouse Primary Renal Tubular Epithelial Cells

Mitochondrial dysfunction in the renal tubular epithelial cells (TECs) can lead to renal fibrosis, a major cause of&#160;chronic kidney disease (CKD). Therefore, assessing mitochondrial function in primary TECs may provide valuable insight into the bioenergetic status of the cells, providing insight into the pathophysiology of CKD. While there are a number of complex protocols available for the isolation and purification of proximal tubules in different species, the field lacks a cost-effective method optimized for&#160;tubular cell isolation without the need for purification. Here, we provide an isolation protocol that allows for studies focusing on both primary mouse proximal and distal renal TECs. In addition to cost-effective reagents and minimal animal procedures required in this protocol, the isolated cells maintain high energy levels after isolation and can be sub-cultured up to four&#160;passages, allowing for continuous studies. Furthermore, using a high throughput extracellular flux analyzer, we assess the mitochondrial respiration directly in the isolated TECs in a 96-well plate for which we provide recommendations for the optimization of cell density and compound concentration. These observations suggest that this protocol can be used for renal tubular ex vivo studies with a consistent, well-standardized production of renal TECs. This protocol may have broader future applications to study mitochondrial dysfunction associated with renal disorders for drug discovery or drug characterization purposes.

This protocol provides a cost-effective approach to isolate and characterize mouse primary renal tubular cells that can subsequently be sub-cultured to assess renal biological functions ex vivo, including mitochondrial bioenergetics.

Mitochondrial dysfunction in the renal tubular epithelial cells (TECs) can lead to renal fibrosis, a major cause of&#160;chronic kidney disease (CKD). ...

Measurement of Energy Metabolism in Explanted Retinal Tissue Using Extracellular Flux Analysis

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while maintaining transparency of the visual axis. Perturbations to this delicate balance cause blinding illnesses, such as diabetic retinopathy. Therefore, the understanding of energy metabolism changes in the retina during disease is imperative to the development of rational therapies for various causes of vison loss. The recent advent of commercially-available extracellular flux analyzers has made the study of retinal energy metabolism more accessible. This protocol describes the use of such an analyzer to measure contributions to retinal energy supply through its two principle arms - oxidative phosphorylation and glycolysis - by quantifying changes in oxygen consumption rates (OCR) and extracellular acidification rates (ECAR) as proxies for these pathways. This technique is readily performed in explanted retinal tissue, facilitating assessment of responses to multiple pharmacologic agents in a single experiment. Metabolic signatures in retinas from animals lacking rod photoreceptor signaling are compared to wild-type controls using this method. A major limitation in this technique is the lack of ability to discriminate between light-adapted and dark-adapted energy utilization, an important physiologic consideration in retinal tissue.

This technique describes real time recording of oxygen consumption and extracellular acidification rates in explanted mouse retinal tissues using an extracellular flux analyzer.

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while ...

Improving the Accuracy of Flow Cytometric Assessment of Mitochondrial Membrane Potential in Hematopoietic Stem and Progenitor Cells Through the Inhibition of Efflux Pumps

As cellular metabolism is a key regulator of hematopoietic stem cell (HSC) self-renewal, the various roles played by the mitochondria in hematopoietic homeostasis have been extensively studied by HSC researchers. Mitochondrial activity levels are reflected in their membrane potentials (&#916;&#936;m), which can be measured by cell-permeant cationic dyes such as TMRM (tetramethylrhodamine, methyl ester). The ability of efflux pumps to extrude these dyes from cells can limit their usefulness, however. The resulting measurement bias is particularly critical when assessing HSCs, as xenobiotic transporters exhibit higher levels of expression and activity in HSCs than in differentiated cells. Here, we describe a protocol utilizing Verapamil, an efflux pump inhibitor, to accurately measure &#916;&#936;m across multiple bone marrow populations. The resulting inhibition of pump activity is shown to increase TMRM intensity in hematopoietic stem and progenitor cells (HSPCs), while leaving it relatively unchanged in mature fractions. This highlights the close attention to dye-efflux activity that is required when &#916;&#936;m-dependent dyes are used, and as written and visualized, this protocol can be used to accurately compare either different populations within the bone marrow, or the same population across different experimental models.

Xenobiotic efflux pumps are highly active in hematopoietic stem and progenitor cells (HSPCs) and cause extrusion of TMRM, a mitochondrial membrane potential fluorescent dye. Here, we present a protocol to accurately measure mitochondrial membrane potential in HSPCs by TMRM in the presence of Verapamil, an efflux pump inhibitor.

As cellular metabolism is a key regulator of hematopoietic stem cell (HSC) self-renewal, the various roles played by the mitochondria in hematopoietic ...