Name: an optimized protocol to analyze glycolysis and mitochondrial respiration in lymphocytes
Uploaded: 2016-11-21T15:00:00.000+00:00
Duration: 8 min 40 s
Description: Watch this Scientific Journal Video about An Optimized Protocol to Analyze Glycolysis and Mitochondrial Respiration in Lymphocytes at JoVE.com

The authors thank Dr. Michael N. Sack (National Heart, Lung, and Blood Institute) for support and discussion, Ms. Ann Kim for optimizing the B cell isolation protocol, Dr. Joseph Brzostowski for his help with microscopy, and Ms. Mirna Pe&#241;a for maintaining the animals used. This study was supported by the Intramural Research Programs of the National Institutes of Health, National Institute of Allergy and Infectious Diseases, and National Heart, Lung, and Blood Institute.

All animal experiments were carried out in accordance with the LIG-4 animal protocol, which was approved by the NIAID Animal Care and Use Committee.
1. Preparation of Reagents
<ol>
	<li>Prepare magnetic separation (MS) buffer: PBS (pH 7.2) supplemented with 0.5% BSA and 2 mM EDTA or automatic separation solution supplemented with 0.5% BSA. Sterile filter (0.22 &#181;m) and store at 2-8 &#176;C.</li>
	<li>Prepare RF10 medium: RPMI 1640 medium supplemented with 10% heat-inactivated fetal bovine serum, 50 U/ml penicillin, 50 &#181;M streptomycin, 1 mM sodium pyruvate, 2 mM L-glutamine, 0.1 mM non-essential amino acids, 50 &#181;M 2-mercaptoethanol, 10 mM HEPES. Use sterile reagents. Sterile filter and store at 2-8 &#176;C.</li>
	<li>Prepare mitochondrial stress test medium: XF base medium supplemented with 1 mM sodium pyruvate, 2 mM L-glutamine and 25 mM glucose.</li>
	<li>Prepare glycolysis stress test medium: XF base medium supplemented with 2 mM L-glutamine.</li>
	<li>Adjust the pH of both the mitochondrial and glycolysis stress test media to 7.4, sterile filter and store at 2-8 &#176;C. Warm media to 37 &#176;C and re-adjust the pH before use (carry out pH measurements at 37 &#176;C). 
	NOTE: Using XF medium, which lacks bicarbonate, is critical. Since the atmosphere in the extracellular flux analyzer contains only ambient CO2, any bicarbonate in the medium, upon binding to protons released as a byproduct of glycolysis, will dissociate to form CO2 and water. CO2 will degas during the experiment, causing drifts in pH that would obscure the glycolytic acidification signal.</li>
	<li>Prepare oligomycin: Prepare 10 mM stock solution in DMSO; store at -20 &#176;C.</li>
	<li>Prepare 2,4-DNP: Prepare 1 M stock solution in DMSO; store at -20 &#176;C.</li>
	<li>Prepare antimycin A: Prepare 10 mM stock solution in DMSO; store at -20 &#176;C.</li>
	<li>Prepare rotenone: Prepare 10 mM stock solution in DMSO; store at -20 &#176;C.</li>
	<li>Prepare glucose: Dissolve glucose in glycolysis stress test medium to form a 250 mM solution; prepare fresh before each experiment and do not freeze.</li>
	<li>Prepare 2-DG: Dissolve solid 2-DG in glycolysis stress test medium to form a 500 mM solution; prepare fresh before each experiment and do not freeze.</li>
</ol>
2. Harvesting Spleen and Lymph Nodes from Mice
<ol>
	<li>Euthanize the mouse by CO2 asphyxiation followed by cervical dislocation.</li>
	<li>Spray the mouse with 70% ethanol.</li>
	<li>For subsequent T cell isolation, harvest the spleen, superficial cervical, superficial/deep axillary, mandibular, inguinal, and mesenteric lymph nodes.</li>
	<li>For subsequent B cell isolation, harvest only the spleen. 
	NOTE: Use a biosafety cabinet and keep the harvested organs in PBS or RF10 media on ice until processing. Use sterile labware and sterile technique for all processing and isolation steps. Keep all buffers and media on ice to increase isolation efficiency and to reduce cell death.</li>
</ol>
3. Tissue Processing
<ol>
	<li>Place a 70 &#181;m cell strainer over a 50 ml conical tube and transfer the organs onto the strainer. For T cell isolation, combine all the organs and for B cell isolation transfer only the spleen.</li>
	<li>Using the plunger from a sterile 3 ml syringe, mash the tissue through the strainer. During mashing, make sure that the filter and the organs are moist at all times and wet them as required by adding MS buffer. This will both keep cell viability high and prevent clogging of the filter.</li>
	<li>After mashing, apply 5-10 ml MS buffer to the filter to increase cell recovery. Centrifuge the suspension at 400 x g for 5 min at 4 &#176;C. (Carry out all further centrifugations under these conditions, unless specified).</li>
	<li>Decant the liquid and resuspend the pellet in 5 ml ACK red blood cell lysis buffer. Incubate for 5 min on ice to lyse red blood cells (red blood cells need to be removed because they interfere with the magnetic separation). Add 10-20 ml MS buffer and centrifuge.</li>
	<li>Place a 30 &#181;m cell filter over a 15 ml conical tube and prime it with 1 ml MS buffer (this filtration will remove debris from the erythrocyte lysis). Decant the liquid from the centrifuged tube, resuspend the pellet in 5 ml MS buffer and pass it through the filter. Wash the initial tube with 4 ml MS buffer to increase cell recovery, and use this to rinse the filter.</li>
	<li>Using a hemocytometer or an automated cell counter, determine the total number of cells. This cell number will be used to determine the amount of magnetic separation reagents required in Section 4. If splenocytes are to be used in the subsequent extracellular flux assay, set aside the appropriate number of cells at this time.</li>
	<li>Centrifuge the suspension and proceed to magnetic separation. Resuspend splenocytes not used for B cell isolation in 5 ml RF10 and keep on ice until the extracellular flux assay.</li>
</ol>
4. Magnetic Separation of B Cells and Na&#239;ve CD4+ T Cells
<ol>
	<li>Determine the necessary volumes of biotin-antibody cocktail, anti-biotin microbeads, and MS buffer. Use the following volumes for 108 cells as a guide and scale up or down as necessary. 
	NOTE: Incubate samples in a refrigerator rather than on ice since incubating on ice may reduce antibody binding efficiency and longer incubation times might be required.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td rowspan="2">Reagents for na&#239;ve CD4+ T cell separation</td>
			<td>Volume for 108 cells</td>
		</tr>
		<tr>
			<td>&#160;(scale appropriately)</td>
		</tr>
		<tr>
			<td>Biotin-antibody cocktail</td>
			<td>100 &#956;l</td>
		</tr>
		<tr>
			<td>MS buffer for first incubation</td>
			<td>400 &#956;l</td>
		</tr>
		<tr>
			<td>Anti-biotin microbeads</td>
			<td>200 &#956;l</td>
		</tr>
		<tr>
			<td>CD44 microbeads</td>
			<td>100 &#956;l</td>
		</tr>
		<tr>
			<td>MS buffer for second incubation</td>
			<td>200 &#956;l</td>
		</tr>
		<tr>
			<td colspan="2">Incubate the biotin-antibody cocktail and cells at 4 &#176;C for 5 min. Add anti-biotin microbeads, CD44 microbeads, MS buffer and incubate at 4 &#176;C for 15 min.</td>
		</tr>
	</tbody>
</table>
Table 1: T cell isolation volumes and instructions.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td rowspan="2">Reagents for B cell separation</td>
			<td>Volume for 108 cells&#160;</td>
		</tr>
		<tr>
			<td>(scale appropriately)</td>
		</tr>
		<tr>
			<td>Biotin-antibody cocktail</td>
			<td>100 &#956;l</td>
		</tr>
		<tr>
			<td>MS buffer for first incubation</td>
			<td>400 &#956;l</td>
		</tr>
		<tr>
			<td>Anti-biotin microbeads</td>
			<td>200 &#956;l</td>
		</tr>
		<tr>
			<td>MS buffer for second incubation</td>
			<td>300 &#956;l</td>
		</tr>
		<tr>
			<td colspan="2">Incubate the biotin-antibody cocktail and cells at 4 &#176;C for 15 min. Add the appropriate volume of biotin-antibody cocktail and MS buffer, and incubate the mixture at 4 &#176;C for 15 min. Add the appropriate volume of anti-biotin microbeads and MS buffer and incubate the mixture at 4 &#176;C for 15 min. After the second incubation, fill the tube with MS buffer and centrifuge.</td>
		</tr>
	</tbody>
</table>
Table 2: B cell isolation volumes and instructions.
<ol start="2">
	<li>Resuspend the pellet in MS buffer: 500 &#181;l for manual separation, 3-4 ml for automatic separation.</li>
	<li>Continue with either manual or automated separations as follows:
	<ol>
		<li>Manual Separation:
		<ol style="margin-left: 40px;">
			<li>Insert an LS column into the separation magnet, place a sterile 15 ml conical tube below to collect the flow through, and prime the column by washing with 3 ml MS buffer. Discard the flow through.</li>
			<li>Transfer the cell suspension to the primed LS column and allow it to pass through; wash the tube used during incubation with 3 ml MS buffer and pass this through the column as well. The eluate will contain the purified cells. For B cell isolation, pass the purified cells through the column a second time into a new 15 ml tube, and repeat the washing step. This will increase purity and yield.</li>
		</ol>
		</li>
		<li>Automated Separation:
		<ol>
			<li>Switch on the automatic separator and check the levels of running buffer, rinsing solution and 70% Ethanol. Ensure that the waste is empty before starting the separation.</li>
			<li>Select the appropriate chilled rack depending on the tube size of the unpurified sample (e.g., 15 ml). In order to keep the cells viable throughout the separation process, use racks that have been previously chilled at 2-8 &#176;C for 3-4 hr, or until the coolant becomes solid.</li>
			<li>Mount the chilled rack on the sample stage of the automatic separator.</li>
			<li>Place the sample tube in slot A1, a tube for the negative fraction in slot B1, and a tube for the positive fraction in C1. If collecting more than one sample, place additional tubes in the next columns (A2, B2, C2, etc.).</li>
			<li>Select the separation tab on the touchscreen, and indicate the arrangement of tubes on the rack. From the separation menu, select the &#34;Depletes&#34; program and complete the program cycle with the appropriate type of wash (see note below). 
			NOTE: The &#34;Depletes&#34; program carries out depletion using the machine&#39;s most sensitive mode, which prioritizes purity. Additionally, there are three different wash options: quick rinse, rinse, and sleep. If the samples are from the same source and are to be combined, select quick rinse. If the samples are to be kept separate, select rinse. Select the sleep option if no further isolations will be carried out that day and if the machine will be shut down following isolation</li>
			<li>Start the isolation by pressing &#34;Run,&#34; and confirm the buffer levels when prompted. After the program has ended, the negative fraction will contain purified cells.</li>
		</ol>
		</li>
		<li>Fill the conical tube up to 15 ml with MS buffer and centrifuge.</li>
		<li>Decant the MS buffer and resuspend cells in 5 ml RF10 media. Keep cells on ice until the extracellular flux assay. 
		NOTE: Ideally, the extracellular flux assay should be carried out immediately after isolation. However, in preliminary experiments no significant differences in assay outcomes were observed in cells used within 3 hr of isolation versus freshly-isolated ones.</li>
	</ol>
	</li>
</ol>
5. Extracellular Flux Assay
NOTE: The protocol outlined here is for the 96-well format of the instrument. Volumes will need to be adjusted if another format is used.
<ol>
	<li>Hydration of Sensor Cartridge
	<ol>
		<li>Lift up the sensor cartridge, fill each well of the utility plate with 200 &#181;l of calibrant solution and lower the sensor cartridge onto the utility plate, submerging the sensors in the calibrant solution. 
		NOTE: Verify that the calibrant solution level is high enough to keep the sensors submerged. The cartridge harbors fluorophores associated to O2 and H+ for each well; these need to be hydrated in order for them to work properly.</li>
		<li>Place the cartridge in a 37 &#176;C incubator not supplemented with CO2 or oxygen/nitrogen. Carry out all further incubations under these conditions, unless specified. Incubate the cartridge overnight to hydrate.</li>
	</ol>
	</li>
	<li>Preparation of Adhesive-coated Plates
	<ol>
		<li>Prepare 2.5 ml of a 22.4 &#181;g/ml adhesive solution in 0.1 M sodium bicarbonate, pH 8.0 (bicarbonate provides the optimum pH for adhesive adsorption).</li>
		<li>Apply 25 &#181;l of the solution to each well of the assay plate, and incubate on the bench at room temperature for 20 min.</li>
		<li>Aspirate adhesive and wash each well twice using 200 &#181;l of sterile water. Wait until wells are dry before seeding the cells.</li>
	</ol>
	</li>
	<li>Seeding Cells in Adhesive-coated Plates
	<ol>
		<li>Centrifuge cells at room temperature at 200 x g for 5 min. Resuspend cells in 5 ml of the appropriate assay medium (see above for formulation of mitochondrial stress and glucose stress media). Centrifuge cells again and resuspend at the desired concentration in assay medium (resuspension volume will vary based on concentration; each well will contain 180 &#181;l of cell suspension). 
		NOTE: As long as the appropriate assay media and compounds are used, the extracellular flux analyzer can simultaneously run the mitochondrial and glycolysis stress tests on the same plate.</li>
		<li>Plate 180 &#181;l of cell suspension in each well. Use wells A1, A12, H1, and H12 for background temperature correction: add 180 &#181;l of assay medium in these wells (no cells).</li>
		<li>Incubate plates for 25 min at 37 &#176;C.</li>
		<li>Centrifuge the plate at 200 x g for 5 min with no brake to ensure that all cells have completely attached. Visually confirm that the cells are stably adhered to the culture surface, forming a monolayer, by viewing under the microscope. Transfer the plates back to the incubator for an additional 30 min.</li>
	</ol>
	</li>
	<li>Preparation of 10x Concentrations of Compounds, and Loading of Injector Ports
	<ol>
		<li>For the mitochondrial stress test, prepare 2.5 ml each of 10 &#181;M oligomycin, 1 mM DNP, and a mixture of 10 &#181;M rotenone and 10 &#181;M antimycin A, all in mitochondrial stress assay medium. 
		NOTE: The concentration of 2,4-DNP is based on previously-published studies;21 this is a general guideline but the concentration of uncoupler for each cell type used should be determined by the researcher.</li>
		<li>For the glycolysis stress test, prepare 2.5 ml each of 250 mM glucose, 10 &#181;M oligomycin, and 500 mM 2-deoxyglucose (2-DG) in glycolysis stress assay medium.</li>
		<li>Warm 10x solutions to 37 &#176;C, and adjust pH to 7.4.</li>
		<li>Load the compounds into the appropriate injector ports of the cartridge using a multichannel micropipettor and the loading guides, as follows:
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
			<tbody>
				<tr>
					<td>Port</td>
					<td>Mitochondrial Stress Assay</td>
					<td>Glycolysis Stress Assay</td>
				</tr>
				<tr>
					<td>A</td>
					<td>20 &#181;l oligomycin</td>
					<td>20 &#181;l glucose</td>
				</tr>
				<tr>
					<td>B</td>
					<td>22 &#181;l 2,4-DNP</td>
					<td>22 &#181;l oligomycin</td>
				</tr>
				<tr>
					<td>C</td>
					<td>25 &#181;l antimycin A/rotenone</td>
					<td>25 &#181;l 2-DG</td>
				</tr>
			</tbody>
		</table>
		Table 3: Compound volumes. 
		 
		NOTE: Each series of ports (e.g., all ports A) must contain the same volume. It is critical that all wells in a given series are loaded, even those not used in the experiment, otherwise the compounds will not be injected (ports in unused wells can be loaded with the same volume of assay medium).</li>
		<li>Incubate the cartridge while setting up the program.</li>
	</ol>
	</li>
	<li>Setting Up Extracellular Flux Assay Protocols
	<ol>
		<li>Set up the following program (Table 4).
		<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
			<tbody>
				<tr>
					<td>Step</td>
					<td>Loop</td>
				</tr>
				<tr>
					<td>Calibration</td>
					<td>&#8211;&#8211;</td>
				</tr>
				<tr>
					<td>Equilibration</td>
					<td>&#8211;&#8211;</td>
				</tr>
				<tr>
					<td>Baseline readings</td>
					<td>3 times: Mix 3 min, wait 0 min, measure 3 min</td>
				</tr>
				<tr>
					<td>End loop</td>
					<td>&#8211;&#8211;</td>
				</tr>
				<tr>
					<td>Inject Port A</td>
					<td>&#8211;&#8211;</td>
				</tr>
				<tr>
					<td>Measurements</td>
					<td>3 times: Mix 3 min, wait 0 min, measure 3 min</td>
				</tr>
				<tr>
					<td>End loop</td>
					<td>&#8211;&#8211;</td>
				</tr>
				<tr>
					<td>Inject Port B</td>
					<td>&#8211;&#8211;</td>
				</tr>
				<tr>
					<td>Measurements</td>
					<td>3 times: Mix 3 min, wait 0 min, measure 3 min</td>
				</tr>
				<tr>
					<td>End loop</td>
					<td>&#8211;&#8211;</td>
				</tr>
				<tr>
					<td>Inject Port C</td>
					<td>&#8211;&#8211;</td>
				</tr>
				<tr>
					<td>Measurements</td>
					<td>3 times: Mix 3 min, wait 0 min, measure 3 min</td>
				</tr>
				<tr>
					<td>End loop</td>
					<td>&#8211;&#8211;</td>
				</tr>
				<tr>
					<td>End Program</td>
					<td>&#8211;&#8211;</td>
				</tr>
			</tbody>
		</table>
		Table 4: Program layout. 
		NOTE: In certain conditions, e.g., when using activated cells, the acidification may continue for longer than in na&#239;ve cells. For this reason, it may be necessary to extend the &#34;wait&#34; times in the above program or repeat each measurement cycle more times.</li>
		<li>Begin the program. After the calibration step, replace the calibrant plate for the assay plate (when prompted). 
		NOTE: Using the software, it is possible to indicate groups of wells that have similar conditions. Additionally, it is crucial to indicate which wells are the control wells (in this protocol, they are the four corners of the plate) and to indicate which wells are empty. For a more detailed, step-by-step, protocol for setting up the machine and its software, the manufacturer&#39;s website should be consulted.</li>
	</ol>
	</li>
	<li>Measuring Protein Content
	<ol>
		<li>Remove the remaining assay medium from each well without disturbing the cells. 
		NOTE: If it is not convenient to proceed with the protein concentration assay, it is possible to freeze the whole plate at -20 &#176;C until analysis. If frozen, thaw before continuing.</li>
		<li>Prepare a 1x solution of protease inhibitors (100x stock) in RIPA lysis medium (sufficient for 50 &#181;l/well).</li>
		<li>Add 50 &#181;l RIPA lysis medium supplemented with protease inhibitors to each well. Agitate plate on a shaker for 5 min, and then incubate plate on ice for 30 min to fully lyse cells.</li>
		<li>Spin the plate at 200 x g for 5 min at room temperature to bring lipids and other molecules to the bottom of the plate so that they do not interfere with the bicinchoninic acid (BCA) assay.</li>
		<li>Measure protein concentration by BCA assay according to manufacturer&#39;s recommendations.</li>
	</ol>
	</li>
</ol>

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L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeting+rare+populations+of+murine+antigen-specific+T+lymphocytes+by+retroviral+transduction+for+potential+application+in+gene+therapy+for+autoimmune+disease.">Targeting rare populations of murine antigen-specific T lymphocytes by retroviral transduction for potential application in gene therapy for autoimmune disease.</a> J Immunol. 164 (7), 3581-3590 (2000).</li><li>Franz, B., May, K. F., Dranoff, G., Wucherpfennig, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ex+vivo+characterization+and+isolation+of+rare+memory+B+cells+with+antigen+tetramers.">Ex vivo characterization and isolation of rare memory B cells with antigen tetramers.</a> Blood. 118 (2), 348-357 (2011).</li><li>Schneider, D. F., Glenn, C. H., Faunce, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Innate+lymphocyte+subsets+and+their+immunoregulatory+roles+in+burn+injury+and+sepsis.">Innate lymphocyte subsets and their immunoregulatory roles in burn injury and sepsis.</a> J Burn Care Res. 28 (3), 365-379 (2007).</li><li>Reeves, J. P., Reeves, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Removal+of+lymphoid+organs.">Removal of lymphoid organs.</a> Curr Protoc Immunol. Chapter 1, Unit 1 9 (2001).</li><li>Akkaya, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Simple,+Versatile+Antibody-Based+Barcoding+Method+for+Flow+Cytometry.">A Simple, Versatile Antibody-Based Barcoding Method for Flow Cytometry.</a> J Immunol. 197 (5), 2027-2038 (2016).</li><li>Traba, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fasting+and+refeeding+differentially+regulate+NLRP3+inflammasome+activation+in+human+subjects.">Fasting and refeeding differentially regulate NLRP3 inflammasome activation in human subjects.</a> J. Clin. Invest. 125 (12), 4592-4600 (2015).</li><li>Wick, A. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Localization+of+the+Primary+Metabolic+Block+Produced+by+2-Deoxyglucose.">Localization of the Primary Metabolic Block Produced by 2-Deoxyglucose.</a> J. Biol. Chem. 224 (2), 963-969 (1957).</li><li>Linnett, P. E., Beechey, R. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inhibitors+of+the+ATP+synthetase+system.">Inhibitors of the ATP synthetase system.</a> Methods Enzymol. 55, 472-518 (1979).</li><li>Terada, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Uncouplers+of+oxidative+phosphorylation.">Uncouplers of oxidative phosphorylation.</a> Environmental Health Perspectives. 87, 213-218 (1990).</li><li>Mookerjee, S. A., Nicholls, D. G., Brand, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determining+Maximum+Glycolytic+Capacity+Using+Extracellular+Flux+Measurements.">Determining Maximum Glycolytic Capacity Using Extracellular Flux Measurements.</a> PLOS ONE. 11 (3), e0152016 (2016).</li><li>Lai, L., Alaverdi, N., Maltais, L., Morse, H. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+cell+surface+antigens:+nomenclature+and+immunophenotyping.">Mouse cell surface antigens: nomenclature and immunophenotyping.</a> J Immunol. 160 (8), 3861-3868 (1998).</li><li>Gerberick, G. F., Cruse, L. W., Miller, C. M., Sikorski, E. E., Ridder, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Selective+modulation+of+T+cell+memory+markers+CD62L+and+CD44+on+murine+draining+lymph+node+cells+following+allergen+and+irritant+treatment.">Selective modulation of T cell memory markers CD62L and CD44 on murine draining lymph node cells following allergen and irritant treatment.</a> Toxicol Appl Pharmacol. 146 (1), 1-10 (1997).</li><li>Mookerjee, S. A., Goncalves, R. L. S., Gerencser, A. A., Nicholls, D. G., Brand, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+contributions+of+respiration+and+glycolysis+to+extracellular+acid+production.">The contributions of respiration and glycolysis to extracellular acid production.</a> Biochim. Biophys. Acta - Bioenergetics. 1847 (2), 171-181 (2015).</li><li>Buck, M. D., O'Sullivan, D., Pearce, E. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=T+cell+metabolism+drives+immunity.">T cell metabolism drives immunity.</a> J Exp Med. 212 (9), 1345-1360 (2015).</li></ol>

The authors declare that they have no competing financial interests. Open Access fees for this article were provided by Agilent Technologies.

The protocol we developed allowed for efficient isolation of pure and viable lymphocyte subsets, which were subsequently used in extracellular flux analysis at different concentrations to evaluate the differences in the glycolysis and mitochondrial respiration performance. This protocol is designed specifically for lymphocytes and addresses the special considerations related to these cell types, such as low basal metabolic activity, fragility, low frequency, and their inability to adhere to assay plates. Therefore, compared to previously published protocols11,12, our method offers a more convenient and better-optimized guide for researchers working in the field of immunology. There are several critical steps in this protocol, including getting pure and viable cell populations, plating the cells at optimal confluence, and standardizing the extracellular flux assay measurements to the protein concentration in each well.
The initial number of cells plated could both be visualized by light microscopy and also be quantified using protein concentration measurements. The linear correlation between cell number and protein concentration confirmed that the protein concentration can indeed be used as a way to standardize the effects of changes in cell numbers between different wells.
By standardizing the ECAR and OCR results to the protein concentrations, we showed that, despite lower cell confluence, as few as 2.5 x 105 cells/well could be used in most assays without compromising the quality of the data. However, the lower limit of cells that can be used varied between cell types and assays. For example, while as few as 1.25 x 105 cells could be used for B cell OCR measurements, even 2.5 x 105 cells/well were not enough to assess the glycolytic performance of the same cell type. Therefore, as long as cell number is not a limiting factor, plating at over 90% confluence, which approximately corresponds to 5 x 105 lymphocytes/well, is preferable. Further optimization might be required when cells of different sizes-such as previously activated and partially differentiated lymphocytes-are used. Additionally, when using previously cultured primary lymphocytes, there might be a variation in cell viability between different treatment conditions, which would decrease the reliability of protein concentration as a measure of the cell number since dead or dying cells can also contribute to the measured protein levels. In such instances, it might be helpful to sort live cells by flow cytometry before carrying out the extracellular flux assays.
In our assays using freshly isolated and highly viable lymphocyte populations, we obtained reliable functional data for both OCR and ECAR measurements. While all cell types behaved similarly in the mitochondrial stress test, striking differences between cell types were observed in the glycolysis stress test. For instance, the glycolytic performance of na&#239;ve T cells was low compared to splenocytes or B cells, and it did not change with the addition of oligomycin. This observation is in line with previously published studies7,30, confirming the validity of our protocol.
In conclusion, our method offers an efficient and convenient way of testing the metabolic activity of lymphocytes using an extracellular flux analyzer, and it can be useful in a wide range of immunological studies exploring the metabolic changes in immune cells upon activation, cell differentiation or due to disease phenotypes such as infection, autoimmunity and hematologic malignancies.

The immune response against antigens is a tightly regulated balance between immune activation and immune suppression. Immune activation drives rapid cell proliferation and migration, as well as cytokine production, antibody secretion and increased phagocytosis in response to the stimulant, whereas immune suppression simply inhibits these events and therefore is important in preventing unnecessary immune responses1-6. Recent studies have shown that a direct link exists between the activation status of immune cells and the activity of various metabolic pathways7. Immune cells can shift between resting and activated states by switching energy producing pathways on and off. Furthermore, it has been observed that different immune cell types may use different metabolic strategies to fuel their increased energy needs during activation. For example, while activation of T lymphocytes directs cells into an almost completely glycolytic state8, activated B lymphocytes use a balance of glycolysis and oxidative phosphorylation9,10. These studies point out the importance of investigating the effects of immune cell activation on cellular metabolism.
Real time, simultaneous measurements of oxygen consumption rate (OCR) and extracellular acidification rate (ECAR), as indicators of oxidative phosphorylation and glycolysis, is a common strategy to address the states of energy producing pathways11-13. In order to achieve this, an extracellular flux analyzer, such as the Seahorse XF 96, is routinely used. Such an instrument can rapidly compare changes in OCR and ECAR across cell types or upon different stimulation conditions. So far, various cell types, including immune cells, have been studied using these devices. However, an optimized protocol specifically designed for immune cells is not available.
Immune cells, particularly lymphocytes, differ from other cell types in several critical ways. Lymphocytes are fragile cells that do not survive for long durations in ex vivo conditions14-16. This is an even greater issue when they are cultured in suboptimal growth media lacking essential nutrients, such as those used in extracellular flux analysis. Unlike macrophages and many cell lines, lymphocytes do not adhere to plastic surfaces; therefore it is critical to attach them to the analysis plate without generating stress. Finally, some lymphocyte subpopulations may be extremely rare and harvesting them at the required, optimal amounts might be challenging17-19.
Here, we provide an optimized protocol that is specifically developed for lymphocytes. Using splenocytes, B lymphocytes and na&#239;ve CD4+ T lymphocytes isolated from mouse spleen and lymph nodes20, we show the characteristics of their resting state glycolysis and oxidative phosphorylation at different cell densities. Data were normalized to account for the differences in the initial cell numbers for each well by measuring the end assay cell lysate protein concentrations, which were directly proportional to the cell numbers. Our protocol not only provides guidelines for the rapid isolation of viable lymphocytes for extracellular flux assays, but it also allows for working at suboptimal cell concentrations without compromising the data quality.

<table><tbody><tr><td>PBS (pH 7.2)</td><td>Gibco-ThermoFisher</td><td>20012-050</td><td>To make MACS buffer</td></tr><tr><td>Bovine Serum Albumin BSA</td><td>Sigma-Aldrich</td><td>A3803</td><td>To make MACS buffer</td></tr><tr><td>0.5 M EDTA, pH 8</td><td>Quality Biological</td><td>10128-446</td><td>To make MACS buffer</td></tr><tr><td>autoMACS rinsing solution</td><td>Miltenyi Biotec</td><td>130-091-222</td><td>Instead of PBS + EDTA to make MS buffer. Also used in autoMACS Pro Separator.</td></tr><tr><td>RPMI-1640</td><td>Gibco-ThermoFisher</td><td>11875-093</td><td>Contains phenol red and L-glutamine</td></tr><tr><td>Fetal Bovine Serum</td><td>Gibco-ThermoFisher</td><td>10437-028</td><td>For heat-inactivation, thaw frozen stock bottle in a 37 &amp;deg;C water bath and then inactivate at 56 &amp;deg;C for 30 min. Aliquot heat-inactivated serum for storage (&lt;em&gt;e.g.&lt;/em&gt;, in 50 ml conical tubes) and freeze at -20 &amp;deg;C until needed.&amp;nbsp;</td></tr><tr><td>Penicillin-Streptomycin (Pen Strep)</td><td>Gibco-ThermoFisher</td><td>15140-122</td><td>Combine Pen Strep with L-Glut 1:1 (if making 500 ml media, make a total of 12 ml Pen Strep/L-Glut); keep aliquots at -20 &amp;deg;C until ready to make media.</td></tr><tr><td>L-Glutamine 200 mM (L-Glut)</td><td>Gibco-ThermoFisher</td><td>25030-081</td><td>Component of RF10 and stress test media</td></tr><tr><td>Sodium pyruvate 100 mM</td><td>Gibco-ThermoFisher</td><td>11360-070</td><td>Component of RF10 and stress test media</td></tr><tr><td>HEPES 1 M</td><td>Gibco-ThermoFisher</td><td>15630-080</td><td>Component of RF10</td></tr><tr><td>MEM Non-Essential Amino Acids Solution (100x)</td><td>Gibco-ThermoFisher</td><td>11140-050</td><td>Component of RF10</td></tr><tr><td>2-Mercaptoethanol (55 mM)</td><td>Gibco-ThermoFisher</td><td>21985-023</td><td>Component of RF10</td></tr><tr><td>Falcon 70 &amp;mu;m cell strainer</td><td>Falcon-Fischer Scientific</td><td>87712</td><td>Used in cell isolation</td></tr><tr><td>Monoject 3 ml Syringe, Luer-Lock Tip</td><td>Covidien</td><td>8881513934</td><td>Used in cell isolation</td></tr><tr><td>Falcon 15 ml Conical Centrifuge Tubes&amp;nbsp;</td><td>Falcon-Fischer Scientific</td><td>14-959-53A</td><td>Used in cell isolation</td></tr><tr><td>Falcon 50 ml Conical Centrifuge Tubes&amp;nbsp;</td><td>Falcon-Fischer Scientific</td><td>14-432-22</td><td>Used in cell isolation</td></tr><tr><td>ACK Lysing Buffer</td><td>Lonza</td><td>10-548E</td><td>Used in cell isolation</td></tr><tr><td>CellTrics 30 &amp;mu;m cell filter, sterile, single-packed</td><td>Partec CellTrics</td><td>04-004-2326</td><td>Used in cell isolation</td></tr><tr><td>B Cell Isolation Kit, mouse</td><td>Miltenyi Biotec</td><td>130-090-862</td><td>Used in cell isolation</td></tr><tr><td>Na&amp;iuml;ve CD4&lt;sup&gt;+&lt;/sup&gt; T cell isolation kit, mouse</td><td>Miltenyi Biotec</td><td>130-104-453</td><td>Used in cell isolation</td></tr><tr><td>LS Column</td><td>Miltenyi Biotec</td><td>130-042-401</td><td>Used in cell isolation</td></tr><tr><td>MidiMACS separator</td><td>Miltenyi Biotec</td><td>130-042-302</td><td>Magnet for separation</td></tr><tr><td>MACS MultiStand</td><td>Miltenyi Biotec</td><td>130-042-303</td><td>Holder for magnets</td></tr><tr><td>autoMACS Rinsing Solution</td><td></td><td>130-091-222</td><td>Rinsing solution for autoMACS Pro Separator</td></tr><tr><td>autoMACS Pro Separator Instrument</td><td>Miltenyi Biotec</td><td>N/A</td><td></td></tr><tr><td>2-Deoxy-D-glucose (2-DG)</td><td>Sigma-Aldrich</td><td>D8375-5G</td><td></td></tr><tr><td>Cell-Tak</td><td>Corning</td><td>354240</td><td>Cell adhesive. Take care to note concentration, as each lot is different (package is 1 mg).</td></tr><tr><td>Oligomycin</td><td>Sigma-Aldrich</td><td>75351</td><td></td></tr><tr><td>2,4-Dinotrophenol (2,4-DNP)</td><td>Sigma-Aldrich</td><td>D198501</td><td></td></tr><tr><td>Antimycin A</td><td>Sigma-Aldrich</td><td>A8674</td><td></td></tr><tr><td>Rotenone</td><td>Sigma-Aldrich</td><td>R8875</td><td></td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>G8270</td><td></td></tr><tr><td>Halt Protease Inhibitor Cocktail</td><td>ThermoFisher Scientific</td><td>78429</td><td>Supplied as 100x cocktail, combine with RIPA to form 1x solution for lysis.</td></tr><tr><td>RIPA Buffer</td><td>Boston BioProducts</td><td>BP-115</td><td></td></tr><tr><td>Pierce BCA Protein Assay Kit</td><td>ThermoFisher Scientific</td><td>23225</td><td>Follow manufacturer&#039;s instructions</td></tr><tr><td>Seahorse XFe96 FluxPak</td><td>Seahorse Bioscience</td><td>101085-004</td><td>Includes assay plates, cartridges, loading guides for transferring compounds to the assay cartridge, and calibrant solution.</td></tr><tr><td>Seahorse XF Base Medium</td><td>Seahorse Bioscience</td><td>102353-100</td><td>Used to prepare stress test media</td></tr><tr><td>Seahorse XFe96 Extracellular Flux Analyzer</td><td>Seahorse Bioscience</td><td></td><td></td></tr><tr><td>Stericup Sterile Vacuum Filter Units, 0.22 &amp;mu;m</td><td>EMD Millipore-Fisher Scientific</td><td>SCGPU10RE</td><td>Used to sterile filter media.</td></tr><tr><td>Sodium bicarbonate</td><td>Sigma-Aldrich</td><td>487031</td><td>Dissolved to 0.1 M and used to dilute Cell-Tak.</td></tr></tbody></table>

an optimized protocol to analyze glycolysis and mitochondrial respiration in lymphocytes

Lymphocytes respond to a variety of stimuli by activating intracellular signaling pathways, which in turn leads to rapid cellular proliferation, migration and differentiation, and cytokine production. All of these events are tightly linked to the energy status of the cell, and therefore studying the energy-producing pathways may give clues about the overall functionality of these cells. The extracellular flux analyzer is a commonly used device for evaluating the performance of glycolysis and mitochondrial respiration in many cell types. This system has been used to study immune cells in a few published reports, yet a comprehensive protocol optimized particularly for lymphocytes is lacking. Lymphocytes are fragile cells that survive poorly in ex vivo conditions. Oftentimes lymphocyte subsets are rare, and working with low cell numbers is inevitable. Thus, an experimental strategy that addresses these difficulties is required. Here, we provide a protocol that allows for rapid isolation of viable lymphocytes from lymphoid tissues, and for the analysis of their metabolic states in the extracellular flux analyzer. Furthermore, we provide results of experiments in which the metabolic activities of several lymphocyte subtypes at different cell densities were compared. These observations suggest that our protocol can be used to achieve consistent, well-standardized results even at low cell concentrations, and thus it may have broad applications in future studies focusing on the characterization of metabolic events in immune cells.

Eukaryotic cells use an integrated network of glycolysis, the tricarboxylic acid (TCA) cycle, and oxidative phosphorylation to meet the majority of their energy demands and to provide intermediates required for cell growth and proliferation. These pathways start with the intracellular trapping of free glucose in the form of glucose-6-phosphate, which subsequently gets processed into pyruvate. Pyruvate is either reduced to lactate or transported into the mitochondria, where it forms acetyl coenzyme A (CoA). Acetyl CoA then enters into the TCA cycle. High-energy intermediates of the TCA cycle propel the movement of electrons in the electron transport chain (ETC), which in turn expels H+ from the mitochondrial matrix to generate a H+ gradient across the inner mitochondrial membrane. Oxygen acts as the final electron acceptor, and H+ return back to the mitochondrial matrix through the FO/F1 complex, where their potential energy is used to generate ATP (Figure 1A).
The working principle of the extracellular flux assay depends on interfering with glycolysis and oxidative phosphorylation at specific points, and assessing the resulting effects. For this purpose, we used glucose to propagate glycolysis in starved cells, and 2-deoxyglucose (2-DG), which is converted to 2-deoxyglucose-6-phosphate, a competitive inhibitor of phosphoglucoisomerase23, by hexokinase, in order to block glycolysis. Rotenone (a complex I-specific inhibitor of the ETC), antimycin A (a complex III-specific inhibitor of the ETC), oligomycin (inhibitor of ATP synthase24), and the uncoupling agent 2,4-dinotrophenol (DNP)25 were used to intervene specific events related to electron transport, proton gradient and ATP synthesis (Figure 1A). Instead of 2,4-DNP, carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone (FCCP) can also be used, but further optimization may be required. In either case, uncouplers should always be titrated for a specific cell type before use to determine the optimal concentration necessary.
The glycolysis stress test (Figure 1B) starts with a baseline measurement of the extracellular acidification rate (ECAR) in the previously starved cells. Since these cells are at their basal minimum, and can therefore be practically considered as non-glycolytic, the ECAR measured at this point is referred to as the non-glycolytic acidification. This acidification likely corresponds to respiratory CO2 generated in the TCA cycle being converted to HCO3- and H+. This is followed by the injection of glucose to activate glycolysis, which presents as an increase in the extracellular acidification due to the formation of lactate. This increase represents the normal rate of glycolysis. The cells are then challenged with the injection of oligomycin, which blocks the generation of ATP through oxidative phosphorylation. Cells respond to this dramatic decrease in ATP production by activating glycolysis to its maximum level, and that results in a secondary increase in the ECAR level (glycolytic reserve). The test is terminated by total inhibition of glycolysis using the glucose analog 2-DG, which returns the ECAR to its non-glycolytic level. Interestingly, it has been proposed that, contrary to the manufacturer's recommendations, maximal glycolysis is not necessarily attained by the injection of oligomycin26. In cells with a high glycolytic capacity, if there is not a significant increase in ATP demand, glycolysis may be perfectly able to cope with the loss of mitochondrial ATP without it needing to be up-regulated. The glycolytic rate with oligomycin could be surpassed by adding respiratory inhibitors, such as rotenone and myxothiazol, which provide a few benefits over oligomycin: 1) They increase the ATP demand, as they cause the reversal of ATP Synthase, with ATP hydrolysis, to pump protons in an attempt to recover the mitochondrial membrane potential26; 2) They prevent respiratory acidification of the medium, which can confound the ECAR results (see below). Other ways to increase ATP demand include the addition of compounds that stimulate the hydrolysis of ATP by the plasma membrane ATPases26. All this should be considered carefully by researchers when planning a glycolysis stress test.
The mitochondrial stress test (Figure 1C) starts with a baseline measurement of the oxygen consumption rate (OCR) in non-starved cells. This is followed by the injection of oligomycin, which inhibits the return of protons through the FO/F1 complex and thus rapidly hyperpolarizes the mitochondrial membrane. Hyperpolarization prevents further proton pumping through respiratory complexes, and the respiratory rate decreases. The remaining respiration is called proton leak, which represents the flow of protons through lipids or other channels. This hyperpolarized state is rapidly reversed by the addition of the uncoupling agent 2,4-DNP, which acts as a proton ionophore. In response, cells try to recover the membrane potential in a futile attempt by increasing the electron transport rate to its maximum, and this in turn increases the OCR. Finally, with the addition of two ETC inhibitors (antimycin A and rotenone), mitochondrial respiration completely stops and OCR decreases to its lowest level. At this level, oxygen consumption is not due to mitochondrial activity (non-mitochondrial). The difference in the OCR generated by these inhibitors is called the maximum mitochondrial respiration, which is the sum of the baseline respiration and the spare capacity.
B and T cell isolations yield highly viable pure lymphocyte populations.
B cells and naïve CD4+ T cells were isolated as outlined in Section 4 of the protocol, and splenocytes were simply obtained by lysing the red blood cells as described in Section 3.3.
The sensitivity of cell type-specific metabolic assays depends on the viability and the purity of the starting cell population. Therefore, in order to verify viability of the isolated mouse T cells, splenocytes, and B cells, and the purity of T and B cells, small aliquots of cells were stained for flow cytometric analysis (Figure 2). In the forward scatter-area vs. side scatter-area (FSC-A vs. SSC-A) plot, lymphocytes were gated, and within this gate, the population along the diagonal in the forward scatter-height (FSC-H) vs. FSC-A plot were determined as the singlets. Within the singlet population, viability was measured by gating the cells that stained negative for the live/dead marker (Figure 2A); T cell viability was 97.9%, splenocyte viability was 92%, and B cell viability was 94%. B cell purity, as measured by the B220+ CD19+ population26, was 99%, while CD4+ T cell purity, as measured by the CD44- CD4+ population27, was 98.3% (Figure 2B).
Protein concentration of the cell lysate can be used as a direct indicator of plated cell number.
Isolated lymphocytes and splenocytes were plated in an adhesive-coated 96-well assay plate at 5 x 105 cells/well, 2.5 x 105 cells/well, 1.25 x 105 cells/well, and 0.625 x 105 cells/well. The confluence at each plating density of the three cell types was visualized under the light microscope (Figure 3A). As expected, confluence correlated with the initial plating densities. Upon completion of the extracellular flux assay, plated cells were lysed and their protein concentrations were quantified using the BCA assay. For all cell types, lysate protein concentrations were shown to be linearly correlated with the initial plating densities (Figure 3B), which confirms that lysate protein concentrations can be used as an accurate measure for the normalization of cell numbers when interpreting the extracellular flux data. The plating densities used in this experiment have been optimized for naïve, unstimulated lymphocytes. If stimulated or previously cultured cells are to be used, further optimization may be required.
Mitochondrial and glycolytic stress assays are dependent on plated cell number.
The OCR was measured for each cell type and plating density in the extracellular flux analyzer. As expected, higher cell numbers have a higher measured OCR, as well as more dramatic responses to oligomycin, 2,4-DNP, and antimycin A/rotenone (Figure 4A). Standardizing OCR measurements to each sample's protein concentration reveals that in general, larger numbers of cells lead to more accurate OCR measurements. Plating at 5 and 2.5 x 105 cells/well resulted in similar normalized OCR measurements in the T cells and splenocytes, and 5 and 1.25 x 105 cells/well resulted similar normalized OCR measurements in B cells (Figure 4B). The slight differences in the normalized B cell OCR measurements might be an indirect indication that B cells perform better at 2.5 x 105 cells/well compared to other cell densities. By all measures, 0.625 x 105 cells/well gave suboptimal results, demonstrating that this plating density is insufficient. Baseline respiration, proton leak, maximum mitochondrial respiration, non-mitochondrial respiration, and ATP production linearly correlated with plating densities for all cell types (Figure 4C). Additionally, most of the baseline respiration is used towards synthesizing ATP, as indicated by the low proton leak in the three cell types. While the primary focus of the mitochondrial stress experiment is to measure changes in OCR, the ECAR is still useful to record to ensure that the assay was successfully carried out. Similar to the OCR, the two higher plating densities resulted in higher ECAR and more dramatic responses to oligomycin, 2,4-DNP, and antimycin A/rotenone (Figure 4D). The dramatic changes in ECAR upon addition of 2,4-DNP, and antimycin A/rotenone may be due to changes in respiration in addition to changes in glycolysis, since CO2 generated in the TCA cycle is converted to HCO3 - and H+. This issue has been recently addressed, and there exists a simple method for correcting the total extracellular acidification signal using oxygen consumption data, to obtain the real glycolytic rate12,29.
The glycolysis stress assay was most successful at the highest plating density (Figure 5A, B). In all cell types, the 5 x 105 cells/well samples had the greatest changes in ECAR after the addition of glucose, oligomycin, and 2-DG (Figure 5B). Additionally, normalizing to protein concentration demonstrated that for all cell types, the 5 x 105 cells/well samples yielded optimal results, while lower concentrations-especially 0.625 x 105 cells/well-did not display a robust glycolytic reserve capacity. Non-glycolytic acidification, glycolysis, and apparent glycolytic capacity linearly correlated with plating densities for all cell types (Figure 5C). For the glycolysis stress experiment, the OCR graph shows that glucose slightly stimulates mitochondrial respiration, which is then inhibited by oligomycin treatment; the addition of 2-DG did not affect the OCR (Figure 5D). The OCR graph can be used as a further indicator that the glycolysis stress experiment was successfully carried out. Also, the OCR graph can be used to correct the ECAR graph, if required, as explained above in the case of the mitochondrial stress test12,29.
<img alt="Glycolysis and oxidative phosphorylation pathways diagram; ECAR and OCR graphs depict metabolic analysis." src="/files/ftp_upload/54918/54918fig1.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Outline of extracellular flux assays. (A) Illustration of glycolysis (left) and oxidative phosphorylation (right) showing the action of the metabolic drugs used in the extracellular flux assays. (B) Schematic of the extracellular acidification rate (ECAR) graph; schematic of the oxygen consumption rate (OCR) graph (C). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54918/54918fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Flow cytometry graphs for T cell, splenocyte, and B cell analysis; includes CD4, CD44, CD19 metrics." src="/files/ftp_upload/54918/54918fig2.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Stepwise gating of T cells, B cells, and splenocytes to determine viability and purity. Flow cytometry plots showing the viability of T cells, splenocytes, and B cells (A); and purity of T and B cells (B). Results are representative of at least three independent experiments. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54918/54918fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="T cell, splenocyte, B cell densities; microscopy, well plate, protein levels; correlation graphs." src="/files/ftp_upload/54918/54918fig3.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: Cell confluence correlates with the lysate protein concentrations of each cell type at different plating densities. (A) Light micrographs of cells in assay plate wells at plating densities ranging from 5 x 105 cells/well to 0.625 x 105 cells/well. Scale bars denote 50 µm. (B) Lysate protein concentrations at different plating densities, as measured by the BCA assay. Results are representative of at least three independent experiments. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54918/54918fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="T cells, splenocytes, B cells OCR/ECAR graph; shows respiration, proton leak, ATP production analysis." src="/files/ftp_upload/54918/54918fig4.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4: Mitochondrial stress assay. Raw (A) and standardized (B) OCR for each cell type and plating density are shown. (C) Cell number-dependent changes in the levels of baseline, maximum mitochondrial and non-mitochondrial respiration, as well as the oxygen consumptions linked to proton leak or ATP production, are shown for each cell types. (D) Raw ECAR values obtained from the mitochondrial stress tests are shown. Each data point represents the mean of 7-8 wells with standard deviation. Labeled arrows denote injections of oligomycin (O), 2,4-DNP (D), and rotenone/antimycin A (R/A). Results are representative of at least three independent experiments. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54918/54918fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="T cell metabolism analysis; ECAR vs time plots; glycolysis and non-glycolytic acidification; graph." src="/files/ftp_upload/54918/54918fig5.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5: Glycolytic stress assay. Raw (A) and standardized (B) ECAR for each cell type and plating density are shown. (C) Cell number-dependent changes in ECAR for non-glycolytic acidification, glucose-induced glycolysis and total glycolytic capacity are shown. (D) Raw OCR values obtained from the glycolytic stress tests are shown. Each data point represents the mean of 7-8 wells with standard deviation. Labeled arrows denote injections of glucose (G), oligomycin (O), and 2-deoxyglucose (2-DG). Results are representative of at least three independent experiments. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54918/54918fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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An Optimized Protocol to Analyze Glycolysis and Mitochondrial Respiration in Lymphocytes

The study of metabolism is becoming increasingly relevant to immunological research. Here, we present an optimized method for measuring glycolysis and mitochondrial respiration in mouse splenocytes, and T and B lymphocytes.

Lymphocytes respond to a variety of stimuli by activating intracellular signaling pathways, which in turn leads to rapid cellular proliferation, migration ...

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29.6K Views.  National Institutes of Health. The study of metabolism is becoming increasingly relevant to immunological research. Here, we present an optimized method for measuring glycolysis and mitochondrial respiration in mouse splenocytes, and T and B lymphocytes.

Video: An Optimized Protocol to Analyze Glycolysis and Mitochondrial Respiration in Lymphocytes

Assessment of the Metabolic Profile of Primary Leukemia Cells

The metabolic requirement of cancer cells can negatively influence survival and treatment efficacy. Nowadays, pharmaceutical targeting of metabolic pathways is tested in many types of tumors. Thus, characterization of cancer cell metabolic setup is inevitable in order to target the correct pathway to improve the overall outcome of patients. Unfortunately, in a majority of cancers, the malignant cells are quite difficult to obtain in higher numbers and the tissue biopsy is required. Leukemia is an exception, where a sufficient number of leukemic cells can be isolated from the bone marrow. Here, we provide a detailed protocol for the isolation of leukemic cells from leukemia patients bone marrow and subsequent analysis of their metabolic state using extracellular flux analyzer. Leukemic cells are isolated by the density gradient, which does not affect their viability. The next cultivation step helps them to regenerate, thus the metabolic state measured is the state of cells in optimal conditions. This protocol allows achieving consistent, well-standardized results, which could be used for the personalized therapy.

Here we present a protocol for the isolation of leukemic cells from leukemia patients bone marrow and analysis of their metabolic state. Assessment of the metabolic profile of primary leukemia cells could help to better characterize the demand of primary cells and could lead up to more personalized medicine.

The metabolic requirement of cancer cells can negatively influence survival and treatment efficacy. Nowadays, pharmaceutical targeting of metabolic ...

Cancer Research

Analysis of Hematopoietic Stem Progenitor Cell Metabolism

Hematopoietic stem progenitor cells (HSPCs) have distinct metabolic plasticity, which allows them to transition from their quiescent state to a differentiation state to sustain demands of the blood formation. However, it has been difficult to analyze the metabolic status (mitochondrial respiration and glycolysis) of HSPCs due to their limited numbers and lack of optimized protocols for non-adherent, fragile HSPCs. Here, we provide a set of clear, step-by-step instructions to measure metabolic respiration (oxygen consumption rate; OCR) and glycolysis (extracellular acidification rate; ECAR) of murine bone marrow-LineagenegSca1+c-Kit+ (LSK) HSPCs. This protocol provides a higher amount of LSK HSPCs from murine bone marrow, improves the viability of HSPCs during incubation, facilitates extracellular flux analyses of non-adherent HSPCs, and provides optimized injection protocols (concentration and time) for drugs targeting oxidative phosphorylation and glycolytic pathways. This method enables the prediction of the metabolic status and the health of HSPCs during blood development and diseases.

Hematopoietic stem progenitor cells (HSPCs) transition from a quiescent state to a differentiation state due to their metabolic plasticity during blood formation. Here, we present an optimized method for measuring mitochondrial respiration and glycolysis of HSPCs.

Hematopoietic stem progenitor cells (HSPCs) have distinct metabolic plasticity, which allows them to transition from their quiescent state to a ...

Developmental Biology

Metabolic Characterization of Polarized M1 and M2 Bone Marrow-derived Macrophages Using Real-time Extracellular Flux Analysis

Specific metabolic pathways are increasingly being recognized as critical hallmarks of macrophage subsets. While LPS-induced classically activated M1 or M(LPS) macrophages are pro-inflammatory, IL-4 induces alternative macrophage activation and these so-called M2 or M(IL-4) support resolution of inflammation and wound healing. Recent evidence shows the crucial role of metabolic reprogramming in the regulation of M1 and M2 macrophage polarization. 
In this manuscript, an extracellular flux analyzer is applied to assess the metabolic characteristics of naive, M1 and M2 polarized mouse bone marrow-derived macrophages. This instrument uses pH and oxygen sensors to measure the extracellular acidification rate (ECAR) and oxygen consumption rate (OCR), which can be related to glycolytic and mitochondrial oxidative metabolism. As such, both glycolysis and mitochondrial oxidative metabolism can be measured in real-time in one single assay.
Using this technique, we demonstrate here that inflammatory M1 macrophages display enhanced glycolytic metabolism and reduced mitochondrial activity. Conversely, anti-inflammatory M2 macrophages show high mitochondrial oxidative phosphorylation (OXPHOS) and are characterized by an enhanced spare respiratory capacity (SRC). 
The presented functional assay serves as a framework to investigate how particular cytokines, pharmacological compounds, gene knock outs or other interventions affect the macrophage&#8217;s metabolic phenotype and inflammatory status.

Metabolic reprogramming is a characteristic and prerequisite for M1 and M2 macrophage polarization. This manuscript describes an assay for the measurement of fundamental parameters of glycolysis and mitochondrial function in mouse bone marrow-derived macrophages. This tool can be applied to investigate how particular factors affect the macrophage&#8217;s metabolism and phenotype.

Specific metabolic pathways are increasingly being recognized as critical hallmarks of macrophage subsets. While LPS-induced classically activated M1 or ...

Measuring Yeast Mitochondrial Respiration While Maintaining Cellular Context

Mitochondrial Respiration Quantification in Yeast Whole Cells

Metabolism is mainly coordinated by cellular energy availability and environmental conditions. Assays for knowing how cells adapt energetic metabolism to different nutritional and environmental conditions give valuable information to elucidate molecular mechanisms. Oxidative phosphorylation is the primary source of ATP in most cells, and mitochondrial respiration activity is a key component of oxidative phosphorylation, maintaining mitochondrial membrane potential for ATP synthesis. Mitochondrial respiration is often studied in isolated mitochondria that are missing the cellular context. Here, we present a method for quantifying mitochondrial respiration in yeast-intact cells. This method applies to any yeast species, although it has been generally used for Saccharomyces cerevisiae cells. First, the yeast growth in specific conditions is tested. Then, cells are washed and resuspended in deionized water with a 1:1 ratio (w/v). Cells are then placed in an oximeter chamber with constant stirring, and a Clark electrode is used to quantify oxygen consumption. Some molecules are sequentially placed into the chamber and selected according to this effect on the electron transport chain or ATP synthesis. ATPase inhibitor oligomycin is first added to measure respiration coupled to ATP synthesis. Afterward, an uncoupler is used to measure the maximal respiratory capacity. Finally, a mix of electron transport chain inhibitors is added to discard oxygen consumption unrelated to mitochondrial respiration. Data are analyzed using a linear regression to obtain the slope, representing the oxygen consumption rate. The advantage of this method is that it is specific for yeast mitochondrial respiration, maintaining the cellular context. It is essential to highlight that inhibitors used in mitochondrial respiration quantification could vary between yeast species.

Mitochondrial respiration in yeast whole cells is a valuable indicator of cell bioenergetics. Here, we present a protocol to quantify this phenotype applicable to different yeast species.

Metabolism is mainly coordinated by cellular energy availability and environmental conditions. Assays for knowing how cells adapt energetic metabolism to ...

Biochemistry

Multicolor Flow Cytometry-based Quantification of Mitochondria and Lysosomes in T Cells

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular components responsible for supplying cell energy; however, excess mitochondria also produce reactive oxygen species (ROS) that could cause cell death. Therefore, the number of mitochondria must constantly be adjusted to fit the needs of the cells. This dynamic regulation is achieved in part through the function of lysosomes that remove surplus/damaged organelles and macromolecules. Hence, cellular mitochondrial and lysosomal contents are key indicators to evaluate the metabolic adjustment of cells. With the development of probes for organelles, well-characterized lysosome or mitochondria-specific dyes have become available in various formats to label cellular lysosomes and mitochondria. Multicolor flow cytometry is a common tool to profile cell phenotypes, and has the capability to be integrated with other assays. Here, we present a detailed protocol of how to combine organelle-specific dyes with surface markers staining to measure the amount of lysosomes and mitochondria in different T cell populations on a flow cytometer.

This article illustrates a powerful method to quantify mitochondria or lysosomes in living cells. The combination of lysosome- or mitochondria-specific dyes with fluorescently conjugated antibodies against surface markers allows the quantification of these organelles in mixed cell populations, like primary cells harvested from tissue samples, by using multicolor flow cytometry.

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular ...

Analysis of Human Natural Killer Cell Metabolism

Natural Killer (NK) cells mediate mainly innate anti-tumor and anti-viral immune responses and respond to a variety of cytokines and other stimuli to promote survival, cellular proliferation, production of cytokines such as interferon gamma (IFN&#947;) and/or cytotoxicity programs. NK cell activation by cytokine stimulation requires a substantial remodeling of metabolic pathways to support their bioenergetic and biosynthetic requirements. There is a large body of evidence that suggests that impaired NK cell metabolism is associated with a number of chronic diseases including obesity and cancer, which highlights the clinical importance of the availability of a method to determine NK cell metabolism. Here we describe the use of an extracellular flux analyzer, a platform that allows real-time measurements of glycolysis and mitochondrial oxygen consumption, as a tool to monitor changes in the energy metabolism of human NK cells. The method described here also allows for the monitoring of metabolic changes after stimulation of NK cells with cytokines such as IL-15, a system that is currently being investigated in a wide range of clinical trials.

In this paper, we describe a method to measure glycolysis and mitochondrial respiration in primary human Natural Killer (NK) cells isolated from peripheral blood, at rest or following IL15-induced activation. The protocol described could be easily extended to primary human NK cells activated by other cytokines or soluble stimuli.

Natural Killer (NK) cells mediate mainly innate anti-tumor and anti-viral immune responses and respond to a variety of cytokines and other stimuli to ...

A Comprehensive High-Efficiency Protocol for Isolation, Culture, Polarization, and Glycolytic Characterization of Bone Marrow-Derived Macrophages

Macrophages are among the most important antigen-presenting cells. Many subsets of macrophages have been identified with unique metabolic signatures. Macrophages are commonly classified as M1-like (inflammatory) and M2-like (anti-inflammatory) subtypes. M1-like macrophages are pro-inflammatory macrophages that get activated by LPS and/or pro-inflammatory cytokines such as INF-&#947;, IL-12 &#38; IL-2. M1-like polarized macrophages are involved in various diseases by mediating the host&#39;s defense to a variety of bacteria and viruses. That is very important to study LPS induced M1-like macrophages and their metabolic states in inflammatory diseases. M2-like macrophages are considered anti-inflammatory macrophages, activated by anti-inflammatory cytokines and stimulators. Under the pro-inflammatory state, macrophages show increased glycolysis in glycolytic function. The glycolytic function has been actively investigated in the context of glycolysis, glycolytic capacity, glycolytic reserve, compensatory glycolysis, or non-glycolytic acidification using extracellular flux (XF) analyzers.
This paper demonstrates how to assess the glycolytic states in real-time with easy-to-follow steps when the bone marrow-derived macrophages (BMDMs) are respiring, consuming, and producing energy. Using specific inhibitors and activators of glycolysis in this protocol, we show how to obtain a systemic and complete view of glycolytic metabolic processes in the cells and provide more accurate and realistic results. To be able to measure multiple glycolytic phenotypes, we provide an easy, sensitive, DNA-based normalization method for polarization assessment of BMDMs. Culturing, activation/polarization and identification of the phenotype and metabolic state of the BMDMs are crucial techniques that can help to investigate many different types of diseases.
In this paper, we polarized the na&#239;ve M0 macrophages to M1-like and M2-like macrophages with LPS and IL4, respectively, and measured a comprehensive set of glycolytic parameters in BMDMs in real-time and longitudinally over time, using extracellular flux analysis and glycolytic activators and inhibitors.

This protocol provides detailed and comprehensive methods for the isolation, culture, polarization, and measurement of the glycolytic metabolic state of live bone marrow-derived macrophages (BMDMs). This paper provides step-by-step instructions with realistic visual illustrations for workflow and glycolytic assessment of BMDMs in real-time.

Macrophages are among the most important antigen-presenting cells. Many subsets of macrophages have been identified with unique metabolic signatures. ...

Real-time Monitoring of Mitochondrial Respiration in Cytokine-differentiated Human Primary T Cells

During activation, the metabolism of T cells adapts to changes that impact their fate. An increase in mitochondrial oxidative phosphorylation is indispensable for T cell activation, and the survival of memory T cells is dependent on mitochondrial remodeling. Consequently, this affects the long-term clinical outcome of cancer immunotherapies. Changes in T cell quality are often studied by flow cytometry using well-known surface markers and not directly by their metabolic state. This is an optimized protocol for measuring real-time mitochondrial respiration of primary human T cells using an Extracellular Flux Analyzer and the cytokines IL-2 and IL-15, which differently affect T cell metabolism. It is shown that the metabolic state of T cells can clearly be distinguished by measuring the oxygen consumption when inhibiting key complexes in the metabolic pathway and that the accuracy of these measurements is highly dependent on optimal inhibitor concentration and inhibitor injection strategy. This standardized protocol will help implement mitochondrial respiration as a standard for T cell fitness in monitoring and studying cancer immunotherapies.

Metabolic adaptation is fundamental for T cells as it dictates differentiation, persistence, and cytotoxicity. Here, an optimized protocol for monitoring mitochondrial respiration in ex vivo cytokine-differentiated human primary T cells is presented.

During activation, the metabolism of T cells adapts to changes that impact their fate. An increase in mitochondrial oxidative phosphorylation is ...

Real-Time Measurement of the Mitochondrial Bioenergetic Profile of Neutrophils

Neutrophils are the first line of defense and the most abundant leukocytes in humans. These effector cells perform functions such as phagocytosis and oxidative burst, and create neutrophil extracellular traps (NETs) for microbial clearance. New insights into the metabolic activities of neutrophils challenge the early concept that they primarily rely on glycolysis. Precise measurement of metabolic activities can unfold different metabolic requirements of neutrophils, including the tricarboxylic acid (TCA) cycle (also known as the Krebs cycle), oxidative phosphorylation (OXPHOS), pentose phosphate pathway (PPP), and fatty acid oxidation (FAO) under physiological conditions and in disease states. This paper describes a step-by-step protocol and prerequirements to measure oxygen consumption rate (OCR) as an indicator of mitochondrial respiration on mouse bone marrow-derived neutrophils, human blood-derived neutrophils, and the neutrophil-like HL60 cell line, using metabolic flux analysis on a metabolic extracellular flux analyzer. This method can be used for quantifying the mitochondrial functions of neutrophils under normal and disease conditions.

We describe stepwise protocols measuring the mitochondrial respiration of mouse and human neutrophils and HL60 cells using the metabolic extracellular flux analyzer.

Neutrophils are the first line of defense and the most abundant leukocytes in humans. These effector cells perform functions such as phagocytosis and ...

Analyzing Ex Vivo Metabolic Flux in Splenic and Cardiac Macrophages and Bone Marrow Monocytes

Metabolic reprogramming is a hallmark of monocyte/macrophage activation and polarization between pro- and anti-inflammatory states. For example, pro-inflammatory (i.e., M1-like) monocytes/macrophages display more reliance on anaerobic glycolysis and less reliance on mitochondrial oxidative phosphorylation, whereas anti-inflammatory (M2-like) macrophages display more reliance on glucose and fatty acid oxidation in the mitochondria. Here, we describe in-depth protocols for extracting macrophages from the two major monocyte/macrophage reservoirs in the body, the spleen and bone marrow, as well as injured tissues such as the heart following myocardial infarction.
Macrophages or monocytes are extracted by immunomagnetic sorting by using antibody-tagged microbeads, which easily bind to cells without compromising their phenotypes. The extracted cells are then cultured in 96-well plates, followed by extracellular flux analysis using a metabolic flux analyzer. Both glycolysis and mitochondrial oxidative phosphorylation can be measured simultaneously in small numbers of cells (as little as 2-3 &#215; 105 cells). This method can easily be performed in 1 day and produces reliable and repeatable results. Ultimately, these methods help to enhance our understanding of metabolic changes during immune and inflammatory responses to injury and disease, which could lead to the development of novel therapeutic targets for immunometabolic pathways.

Here, we describe in detail methods for extracting macrophages from the bone marrow, spleen, and infarcted heart, and subsequently assessing metabolic flux in live cells.

An Optimized Protocol to Analyze Glycolysis and Mitochondrial Respiration in Lymphocytes

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