We would like to thank all Ross lab and Murphy lab members. This work is supported by American Heart Association (14SDG17790005) to Y.C. NIH (HL115933, HL127806) and VA Merit (BX003260) to R.S.R.

For work with neonatal mice, please refer to local university/institute guidelines set forth by the animal care programs and adhere to one&#8217;s institutional and other appropriate regulations. All methods described in this protocol have been approved by the UC San Diego Institutional Animal Care and Use Committee (IACUC) and adhere to federal and state regulations.
1. Preparation of Reagents
<ol>
	<li>Prepare 25 mL of pre-digestion solution: HBSS (without Ca2+ and Mg2+) supplemented with trypsin (0.5 mg/mL). Sterilize the solution using a 0.22 &#181;m filter and keep on ice until use. Make pre-digestion solution on the day of experiment. 
	NOTE: It is critical to use HBSS without Ca2+ and Mg2+, as Ca2+ and Mg2+ will cause myocyte contraction and subsequent cell death during isolation.</li>
	<li>Prepare 30 mL of collagenase digestion buffer: collagenase (0.8 mg/mL, approximately 350 U/mL) dissolved in HBSS (without Ca2+ and Mg2+) buffer. Sterilize the solution using a 0.22 &#181;m filter and keep on ice until use. Make collagenase digestion solution on the day of experiment.</li>
	<li>Prepare 500 mL of cardiomyocyte culture media (growth media): Mix 375 mL of DMEM, 125 mL of M-199, 25 mL of Horse serum, and 12.5 mL of FBS. Supplement with 1% penicillin and 1% streptomycin solution.</li>
	<li>Prepare mitochondrial stress test medium: Make 200 mL of DMEM based stress test medium (DMEM without NaHCO3, see Table of Materials) supplemented with 1 mM sodium pyruvate, 2 mM L-glutamine, and 10 mM glucose, and 2 mM Hepes. 
	NOTE: Make 1 L of medium with DMEM without sodium pyruvate, L-glutamine, glucose, and Hepes, filer sterile, and store in 4 &#176;C. Prepare the stress test medium on the day of the assay by adding other reagents. Using DMEM medium without NaHCO3 is critical.</li>
	<li>Adjust pH of mitochondrial stress test media to 7.4 on the day of use. Warm media to 37 &#176;C before use.</li>
	<li>Prepare oligomycin: Prepare 5 mL of a 5 mM stock solution in DMSO, make 250 &#181;L aliquots, and store at -20 &#176;C.</li>
	<li>Prepare FCCP: Prepare 5 mL of a 5 mM stock solution in DMSO, make 250 &#181;L aliquots, and store at -20 &#176;C.</li>
	<li>Prepare antimycin A: Prepare 5 mL of a 5 mM stock solution in DMSO, make 250 &#181;L aliquots, and store at -20 &#176;C.</li>
	<li>Prepare rotenone: Prepare 5 mL of a 5 mM stock solution in DMSO, make 250 &#181;L aliquots, and store at -20 &#176;C. 
	NOTE: All reagents and solutions used in this protocol are listed in Table 1.</li>
</ol>
2. Harvesting and Pre-digestion of Hearts from Neonatal Mice (Day 1)
<ol>
	<li>Autoclave scissors, forceps, and a Moria spoon to sterilize.</li>
	<li>Perform all steps in the cell culture hood for sterility.</li>
	<li>Aliquot 5 mL of HBSS (without Ca2+, Mg2+) to each well of a 6-well cell culture plate; place on ice. Aliquot 10 mL of HBSS into a 10 cm cell culture dish.</li>
	<li>Prepare 20 mL of trypsin pre-digestion solution in a 50 mL sterile conical tube. Keep all solutions on ice.</li>
	<li>Quickly dip newborn (day 0) mice in 70% ethanol solution for sterilization.</li>
	<li>Decapitate pups using sterile scissors (straight) without anesthesia, and then open chest along the sternum to allow access to the chest cavity and the heart. (Figure 1A) 
	NOTE: 1) It is critical to use P0 neonatal mice to achieve high cell viability. 2) This euthanasia method is permitted for neonates in accordance with NIH and American Veterinary Medical Association guidelines 17.</li>
	<li>Extract hearts from the body with a fine scissors and transfer immediately into the sterile cell culture dish containing HBSS (without Ca2+, Mg2+) (Figure 1B).</li>
	<li>Remove any residual lung tissue, larger vessels, etc. (and atria, if desired). Wash hearts in the HBSS solution using gentle agitation.</li>
	<li>Cut each heart with a fine scissors into 8 pieces and transfer the all heart tissue with forceps into one well of a 6-well cell culture plate with HBSS (Figure 1C and 1D).</li>
	<li>Wash the hearts by transferring the hearts from well to well in the 6-well plate filled with HBSS, using a Moria spoon (Figure 1D). 
	NOTE: Transferring the hearts from well to well is enough to wash out the blood. Since blood interferes with enzymatic digestion it is important to wash the hearts with HBSS and remove blood.</li>
	<li>Transfer the hearts with a Moria spoon into a conical tube containing 20 mL of trypsin (0.5 mg/mL) and incubate with gentle agitation at 4&#176;C for 4 h (Figure 1E).</li>
</ol>
3. Prepare a 96-well Culture Plate (Day 1)
<ol>
	<li>Prepare 5 mL of coating solution: PBS containing 0.5% gelatin (autoclave before use) and 1% fibronectin solution. (e.g. 5 mL gelatin solution plus 50 &#181;L of fibronectin solution)</li>
	<li>Aliquot 50 &#181;L of coating solution into each well of the 96-well cell culture plate (see Table of Materials). If bubbles are present, remove them by using a 20 &#181;L pipette to suck bubbles out. 
	NOTE: It is important to cover all the surface area of each well with coating solution.</li>
	<li>Incubate the plate in a 37 &#176;C cell culture incubator for 1 h or more to allow drying of the matrix coating.</li>
	<li>Aspirate any residual coating solution before seeding cardiomyocytes.</li>
</ol>
4. Enzymatic Digestion and Plating of Cells (Day 1)
<ol>
	<li>Pre-warm collagenase digestion solution in a 37 &#176;C water bath. 
	NOTE: This is step is important to achieve efficient enzymatic digestion.</li>
	<li>Move the conical tube containing hearts and pre-digestion solution from 4 &#176;C to a cell culture hood. (Figure 1F)</li>
	<li>Let the hearts sink to the bottom of the tube and remove the pre-digestion solution by using a 10 mL serological pipette (1 to 2 mL of the isolation medium may remain in the tube).</li>
	<li>Add 10 mL of HBSS into the tube. Re-suspend the hearts with HBSS 2-3 times to wash out trypsin using a 10 mL serological pipette. Aspirate HBSS (1 to 2 mL may remain in the tube).</li>
	<li>Add 10 mL of pre-warmed collagenase digestion solution into the tube with hearts. (Figure 1G)</li>
	<li>Incubate the tube with hearts in a 37 &#176;C water bath for 10 min without agitation (1st digestion).</li>
	<li>After 1st digestion, move the tube to the cell culture hood. Gently triturate hearts by re-suspending the hearts within the tube gently 10 times using a 10 mL serological pipette. This will allow hearts to disperse and cells to be released from heart tissue. (Figure 1H) 
	NOTE: Since cardiomyocytes are fragile, gentle trituration is important to achieve high viability.</li>
	<li>Let the undigested tissue sink, transfer digested solution enriched in cardiomyocytes (approximately 9 - 10 mL) to a new conical tube, and immediately add an equal amount of cell culture media to stop the collagenase digestion.</li>
	<li>Add 10 mL of collagenase digestion solution into the tube containing the remaining undigested heart tissue.</li>
	<li>Incubate the tube with heart tissue in a 37 &#176;C water bath for 10 min (2nd digestion).</li>
	<li>Repeat procedure 4.7 and 4.8. 
	NOTE: If there is still much undigested tissue, repeat digestion one more time. However, in most cases, two digestions are enough to disperse most of the cells from the heart tissue.</li>
	<li>Place a sterile cell-strainer (100 &#181;m nylon mesh) in a new sterile 50 mL conical tube. Pre-wet the cell strainer with 2-3 mL of cell culture media and pass cells through the cell-strainer. Rinse the cell-strainer with 2-3 mL of cell culture media. (Figure 1I)</li>
	<li>Centrifuge conical tube containing cardiomyocytes for 5 min at 180 x g (Figure 1J). Aspirate the supernatant (Figure 1K), which will contain cell tissue debris and re-suspend the cell pellet in 10 mL of cell culture media (Figure 1L).</li>
	<li>Gently resuspend the cells and plate cells onto a 10 cm cell culture dish (plastic without any type of coating) and incubate for 1 h in a cell culture incubator (1st pre-plating) (Figure 1M). This pre-plating step allows non-cardiomyocytes, such as fibroblasts and endothelial cells, to adhere to the uncoated cell-culture dish. 
	NOTE: At this point, cardiomyocytes are typically a round shape and appear shiny under the microscope. (Figure 1N).</li>
	<li>After the 1 h incubation, gently agitate the plate, wash non-adherent cells (enriched in cardiomyocytes) from the 10 cm culture dish, and re-suspend cells by repeatedly pipetting the cell culture medium over the dish using a 10 mL serological pipette. Then, transfer non-adherent cells (enriched in cardiomyocytes) into a new 10 cm cell culture dish (plastic without any coating) and incubate for an additional 1 h in a cell culture incubator (2nd pre-plating). 
	NOTE: Cells that attach to the non-coated plate are dominantly non-cardiomyocytes: fibroblasts and endothelial cells, that can be visualized under a microscope (Figure 1O).</li>
	<li>After 2nd pre-plating, gently agitate the plate, wash non-adherent cells (cardiomyocytes) from the 10 cm culture dish, then transfer the cardiomyocytes into a new 50 mL conical tube.</li>
</ol>
5. Counting Cells and Plating Cells into a 96-well Cell Culture Plate (Day 1)
<ol>
	<li>Count the cells using a hemocytometer.</li>
	<li>Plate the cells into an extracellular matrix coated 96-well cell culture plate at a density between&#160;10&#8211;30 x 103&#160;cells/well by using a multi-channel pipette in a final volume of 200 &#181;L (Figure 1P). Use wells A1, A12, H1, and H12 for background: add 200 &#181;L of culture medium as other wells in these wells (no cells). Incubate the plate in a 37 &#176;C cell culture incubator. 
	NOTE: In this study, oxygen consumption was tested by using different cell densities such as 10 x 103, 20 x 103, or 30&#160;x 103 cells/well. (Figure 2A). Also, as above, cardiomyocytes immediately after isolation are typically a round shape and appear shiny under the microscope. Viable cells will flatten out within 16-24 h of culture.</li>
</ol>
6. Oxygen Consumption Assay using a 96-well-format Extracellular Flux Analyzer (Day 2)
NOTE: Oxygen consumption assay can be carried out one day after plating the cells or later. Neonatal cardiomyocytes cultured using this protocol can survive up to 7 days post isolation.
<ol>
	<li>Hydrate a flux analyzer sensor cartridge (see Table of Materials) for at least 3 hours, but ideally for a full day, before the assay. Add 200 &#181;L of Calibrant solution (see Table of Materials) into the each well of the utility plate, put the sensor cartridge back onto the utility plate, and incubate in a 37&#176;C incubator without CO2 or O2 supplementation.</li>
	<li>Change cell culture media to mitochondria stress test medium one hour prior to the assay. Cardiomyocytes are fragile. Therefore, gently remove the cell culture media by using a multi-channel pipette and wash the cells with 200 &#181;L of pre-warmed mitochondrial stress test media twice. After second wash, add 175 &#181;L of pre-warmed mitochondria stress test media and culture the cells in a 37&#176;C incubator without CO2 or O2 supplementation.</li>
	<li>Prepare concentrated test compounds. For mitochondria stress test, prepare 3.0 mL each of 16 &#181;M oligomycin, 9 &#181;M FCCP, and a mixture of 20 &#181;M rotenone and 20 &#181;M antimycin A, all in mitochondrial stress test medium. 
	NOTE: Each compound at the concentration described has been tested. However, titrating the concentration of each compound in one&#8217;s own laboratory is necessary.</li>
	<li>Load 25 &#181;L of each compound into the injector ports of the sensor cartridge using a multichannel pipette (Figure 1Q). The volume and final concentration are described in Table 2.</li>
	<li>Set up extracellular flux assay protocol. The program is described in Table 3.</li>
	<li>Start the program. First, put the sensor cartridge into the machine for calibration (Figure 1R). Replace the calibrant for the assay plate once the calibration step is done. 
	NOTE: Using the software provided by manufacturer, indicate groups of wells and each compound and port.</li>
	<li>If desired, after the assay, carefully discard all assay medium by using a multi-channel pipette and store the cell culture microplate at -20 &#176;C for future cell normalization using protein assay.</li>
	<li>Measure protein content. Add 50 &#181;L of standard RIPA cell lysis solution (see Table of Materials). Incubate the plate on ice for 30 min to fully lyse cells. Transfer all material to a new clear flat bottom 96 well assay plate.</li>
	<li>Measure protein concentration by BCA assay according to manufacturer&#8217;s protocol. 
	NOTE: Coating the well with extracellular matrix results in a high protein concentration in each well. Therefore, subtract the amount of the background well(s) (no cells) to get actual cell protein concentration. The protein concentration derived from cells is low in a 96-well culture plate. In addition, protein concentration can vary from well-to-well due to extracellular matrix coating. Therefore, using cell number to normalize the OCR is suggested.</li>
</ol>

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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+Gene+Expression+Analysis+of+Mouse+and+Human+Cardiac+Maturation.">Comparative Gene Expression Analysis of Mouse and Human Cardiac Maturation.</a> Genomics, Proteomics &amp; Bioinformatics. 14 (4), 207-215 (2016).</li></ol>

The authors declare that they have no competing financial interests.

In this study, we have established a simple protocol for isolating and culturing mouse neonatal cardiomyocytes. By using these cardiomyocytes, we also optimized the conditions to measure oxygen consumption rate by using an extracellular flux analyzer system. The protocol allows one to use mouse neonatal cardiomyocytes as a model system to examine how various factors can alter oxygen consumption in the principal working cells of the heart, akin to what would be measured in the intact organ. Our protocol is different from ...

To sustain a continuous cardiac contractile function, cardiomyocytes must maintain a constant supply of cellular energy primarily in the form of ATP1. In the heart, approximately 95% of ATP is generated by mitochondria, mainly through oxidative phosphorylation, showing that mitochondria play a crucial bioenergetic role in cardiac function2,3. Supporting this notion is that dysregulation of mitochondrial function can lead to cardiomyopathy and heart failure4,5. Conversely, restoring mitochondrial function has been shown to improve cardiac function of the failing heart6,7. Therefore, studying the mechanism of mitochondrial bioenergetics and identifying novel regulators of mitochondrial function in cardiomyocytes will not only reveal mechanistic insights of cardiac energy production but also could provide insight that will lead to development of new therapeutic targets to treat heart diseases6,8.
Compared to the whole heart, which contains a mixture of myocytes and non-myocytes9, cardiomyocyte cultures are extremely pure, with minimal contamination of non-myocytes from the heart, such as fibroblasts and endothelial cells10. In addition, isolating cardiomyocytes from neonatal pups enables culturing a large number of cells in a small amount of time, compared to isolating cells from adult hearts10,11. Most importantly, primary cultured adult mouse cardiomyocytes have short survival times (e.g. 24 h) and at longer time points de-differentiate. Neonatal mouse cardiomyocytes can survive and be manipulated for upwards of 7 days in culture, making them ideal for testing the effects of drug compounds and gene manipulation on the function of mitochondria in cardiomyocytes10. Of course, there are significant biological differences between the adult and neonatal cells, but the longer duration available for culture of neonatal cells makes them appropriate for many different types of studies, including those of mitochondrial function.
To date, primary cultured neonatal mouse and rat cardiomyocytes have been used as models to study cardiac bioenergetics12,13. In recent years, studies used an extracellular flux analyzer to measure oxygen consumption rate (OCR) and evaluate oxidative capacity in mouse and rat neonatal cardiomyocytes14,15. While compared to rats, the cell viability of mouse neonatal cardiomyocytes is lower and has greater variability16. Also, the ability to study cells from genetically engineered mouse models makes the mouse cell model very important. Given that OCR studies are so sensitive to cell number and seeding density, development of a reproducible, reliable, and simple protocol to achieve consistent cell yield and viability is needed.
Here, we report an optimized protocol that has been developed which uses cultured mouse neonatal cardiomyocytes along with a 96-well-format extracellular flux analyzer for OCR analysis. This protocol greatly increases reproducibility of the assay. In addition, the protocol not only provides a novel and reproducible method for OCR analysis, but also could be adapted to a larger size culture for other experimental purposes, such as that which may be needed to study myofibrillar functions and intracellular signaling pathways.
In particular, this protocol describes a one-day procedure for isolation and culture of neonatal mouse cardiomyocytes in a 96-well cell culture plate. In addition, it describes the procedure to measure oxygen consumption using an extracellular flux analyzer. All solutions used are sterile or sterile filtered. All tools are sterilized by 75% ethanol. We provide a&#160;Table of Materials for various parts of the procedure. For culturing cardiomyocytes, all procedures and steps are performed in a standard cell culture hood. This protocol is developed for the isolation of neonatal mouse hearts from one litter (approximately 8-10 pups). However, the protocol can also be adapted for isolating cardiomyocytes from multiple litters.

<table><tbody><tr><td>Antimycin A</td><td>SIGMA</td><td>A8674</td><td>Inhibits complex III of the mitochondria</td></tr><tr><td>Cell strainer 100 &amp;mu;m pores</td><td>FALCON</td><td>352360</td><td>To capture undigested tissue</td></tr><tr><td>Collagenase type II</td><td>Worthington</td><td>LS004176</td><td>To make collagenase digestion solution</td></tr><tr><td>D-Glucose</td><td>SIGMA</td><td>75351</td><td>To make mitochodnrial stress test medium</td></tr><tr><td>DMEM high glucose</td><td>Life technologies</td><td>11965-092</td><td>To make cell culture medium</td></tr><tr><td>DMEM without NaHCO&lt;sub&gt;3&lt;/sub&gt;, Glucose, pyruvate, glumanine, and Hepes</td><td>SIGMA</td><td>D5030-10X1L</td><td>To make mitochodnrial stress test medium</td></tr><tr><td>FCCP (Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone)</td><td>SIGMA</td><td>C2920</td><td>Uncouples mitochondrial respiration</td></tr><tr><td>Fetal bovine serum (FBS)</td><td>Life technologies</td><td>26140-079</td><td>To make cell culture medium</td></tr><tr><td>Fibronectin from bovine plasma</td><td>SIGMA</td><td>F1141-5MG</td><td>To make coating solution for tissue culture plates</td></tr><tr><td>Fine scissors</td><td>Fine Sciences Tools</td><td>14060-10</td><td>For dissection of hearts</td></tr><tr><td>Gelatin from porcine skin</td><td>SIGMA</td><td>G-1890</td><td>To make coating solution for tissue culture plates</td></tr><tr><td>HBSS (Hank&#039;s balanced salt solution, without Ca&lt;sup&gt;2+&lt;/sup&gt;, Mg&lt;sup&gt;2+&lt;/sup&gt;)</td><td>Cellgro</td><td>21-022-CV</td><td>To wash hearts and make pre-digestion and collagnase digestion solution</td></tr><tr><td>HEPES (1 M)</td><td>Fisher scientific</td><td>15630080</td><td>To make mitochodnrial stress test medium</td></tr><tr><td>Horse serum</td><td>Life technologies</td><td>26050-088</td><td>To make cell culture medium</td></tr><tr><td>L-Glutamine</td><td>SIGMA</td><td>G-3126</td><td>To make mitochodnrial stress test medium</td></tr><tr><td>M-199</td><td>Cellgro</td><td>10-060-CV</td><td>To make cell culture medium</td></tr><tr><td>Moria spoon</td><td>Fisher scientific</td><td>NC9190356</td><td>To wash hearts&amp;nbsp;</td></tr><tr><td>Oligomycin</td><td>SIGMA</td><td>75351-5MG</td><td>Inhibits mitochondrial ATP synthase</td></tr><tr><td>RIPA buffer</td><td>Fisher scientific</td><td>89900</td><td>To lyse the cells for protein assay</td></tr><tr><td>Rotenone</td><td>SIGMA</td><td>R8875</td><td>Inhibits complex I of the mitochondria</td></tr><tr><td>Seahorse XFe96 Extracellular Flux&lt;br/&gt; Analyzer</td><td>Agilent</td><td></td><td>Device used to analyze oxygen consumption rate</td></tr><tr><td>Seahorse XFe96 FluxPak</td><td>Agilent</td><td>102601-100</td><td>Package of flux analyzer culture plates, sensor cartridges, and calibrant</td></tr><tr><td>Sodium pyruvate</td><td>SIGMA</td><td>P2256</td><td>To make mitochodnrial stress test medium</td></tr><tr><td>Straight scissors</td><td>Fine Sciences Tools</td><td>91401-12</td><td>For dissection of hearts</td></tr><tr><td>Syringe filter 0.2 &amp;mu;m size</td><td></td><td></td><td>For sterile filtration of digestion medium</td></tr><tr><td>Trypsin</td><td>USB Corporation</td><td>22715 25GM</td><td>To make pre-digestion solution</td></tr></tbody></table>

analyzing oxygen consumption rate in primary cultured mouse neonatal cardiomyocytes using an extracellular flux analyzer

Mitochondria and oxidative metabolism are critical for maintaining cardiac muscle function. Research has shown that mitochondrial dysfunction is an important contributing factor to impaired cardiac function found in heart failure. By contrast, restoring defective mitochondrial function may have beneficial effects to improve cardiac function in the failing heart. Therefore, studying the regulatory mechanisms and identifying novel regulators for mitochondrial function could provide insight which could be used to develop new therapeutic targets for treating heart disease. Here, cardiac myocyte mitochondrial respiration is analyzed using a unique cell culture system. First, a protocol has been optimized to rapidly isolate and culture high viability neonatal mouse cardiomyocytes. Then, a 96-well format extracellular flux analyzer is used to assess the oxygen consumption rate of these cardiomyocytes. For this protocol, we optimized seeding conditions and demonstrated that neonatal mouse cardiomyocytes oxygen consumption rate can be easily assessed in an extracellular flux analyzer. Finally, we note that our protocol can be applied to a larger culture size and other studies, such as intracellular signaling and contractile function analysis.

By using the protocol described, hearts were isolated from day 0 neonatal pups. 5 x 105 cells/pup were obtained, and cardiomyocytes were seeded at densities of 10 x 103, 20 x 103, or 30&#160;x 103 cells/well, in 96 well plates (Figure 2A). After overnight culture, cardiomyocytes were found well-attached to the coated plastic surface and there were very few unattached cells (the unattached cells will still appear as ...

Watch this Scientific Journal Video about Analyzing Oxygen Consumption Rate in Primary Cultured Mouse Neonatal Cardiomyocytes Using an Extracellular Flux Analyzer at JoVE.com

Analyzing Oxygen Consumption Rate in Primary Cultured Mouse Neonatal Cardiomyocytes Using an Extracellular Flux Analyzer

The goal of this protocol is to illustrate how to use mouse neonatal cardiomyocytes as a model system to examine how various factors can alter oxygen consumption in the heart.

Mitochondria and oxidative metabolism are critical for maintaining cardiac muscle function. Research has shown that mitochondrial dysfunction is an ...

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Research

JoVE Journal

Medicine

8.6K Views.  University of California San Diego. Dit protocol stelt ons in staat om te isoleren en cultuur hoge bi-BBT, alleen bekend om de muis cardiomyocyten. Om de mitochondriale functie te evalueren, kan het zuurstofverbruik van cardiomyocyten worden geanalyseerd door een echte zilverfluxanalyzer. Een van de voordelen van deze techniek is dat het zuurstofverbruik van cardiomyocyten gemakkelijk kan worden gemeten in hun aanhangende toestand in 96-well formaat, waardoor het testen van meerdere omstandigheden met een groot aantal replicati...

Video: Analyzing Oxygen Consumption Rate in Primary Cultured Mouse Neonatal Cardiomyocytes Using an Extracellular Flux Analyzer

Exploring Mitochondrial Energy Metabolism of Single 3D Microtissue Spheroids Using Extracellular Flux Analysis

Three-dimensional (3D) cellular aggregates, termed spheroids, have become the forefront of in vitro cell culture in recent years. In contrast to culturing cells as two-dimensional, single-cell monolayers (2D culture), spheroid cell culture promotes, regulates, and supports physiological cellular architecture and characteristics that exist in vivo, including the expression of extracellular matrix proteins, cell signaling, gene expression, protein production, differentiation, and proliferation. The importance of 3D culture has been recognized in many research fields, including oncology, diabetes, stem cell biology, and tissue engineering. Over the last decade, improved methods have been developed to produce spheroids and assess their metabolic function and fate.
Extracellular flux (XF) analyzers have been used to explore mitochondrial function in 3D microtissues such as spheroids using either an XF24 islet capture plate or an XFe96 spheroid microplate. However, distinct protocols and the optimization of probing mitochondrial energy metabolism in spheroids using XF technology have not been described in detail. This paper provides detailed protocols for probing mitochondrial energy metabolism in single 3D spheroids using spheroid microplates with the XFe96 XF analyzer. Using different cancer cell lines, XF technology is demonstrated to be capable of distinguishing between cellular respiration in 3D spheroids of not only different sizes but also different volumes, cell numbers,&#160;DNA content and type.
The optimal mitochondrial effector compound concentrations of oligomycin, BAM15, rotenone, and antimycin A are used to probe specific parameters of mitochondrial energy metabolism in 3D spheroids. This paper also discusses methods to normalize data obtained from spheroids and addresses many considerations that should be considered when exploring spheroid metabolism using XF technology. This protocol will help drive research in advanced in vitro spheroid models.

These protocols will help users probe mitochondrial energy metabolism in 3D cancer cell-line-derived spheroids using Seahorse extracellular flux analysis.

Three-dimensional (3D) cellular aggregates, termed spheroids, have become the forefront of in vitro cell culture in recent years. In contrast to culturing ...

Cancer Research

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 ...

Immunology and Infection

Measurement of Oxygen Consumption Rate in Acute Striatal Slices from Adult Mice

Mitochondria play an important role in cellular ATP production, reactive oxygen species regulation, and Ca2+ concentration control. Mitochondrial dysfunction has been implicated in the pathogenesis of multiple neurodegenerative diseases, including Parkinson&#39;s disease (PD), Huntington&#39;s disease, and Alzheimer&#39;s disease. To study the role of mitochondria in models of these diseases, we can measure mitochondrial respiration via oxygen consumption rate (OCR) as a proxy for mitochondrial function. OCR has already been successfully measured in cell cultures, as well as isolated mitochondria. However, these techniques are less physiologically relevant than measuring OCR in acute brain slices. To overcome this limitation, the authors developed a new method using a Seahorse XF analyzer to directly measure the OCR in acute striatal slices from adult mice. The technique is optimized with a focus on the striatum, a brain area involved in PD and Huntington&#39;s disease. The analyzer performs a live cell assay using a 24-well plate, which allows the simultaneous kinetic measurement of 24 samples. The method uses circular-punched pieces of striatal brain slices as samples. We demonstrate the effectiveness of this technique by identifying a lower basal OCR in striatal slices of a mouse model of PD. This method will be of broad interest to researchers working in the field of PD and Huntington&#39;s disease.

Oxygen consumption rate (OCR) is a common proxy for mitochondrial function and can be used to study different disease models. We developed a new method using a Seahorse XF analyzer to directly measure the OCR in acute striatal slices from adult mice that is more physiologically relevant than other methods.

Mitochondria play an important role in cellular ATP production, reactive oxygen species regulation, and Ca2+ concentration control. Mitochondrial ...

Neuroscience

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.

Metabolic reprogramming is a hallmark of monocyte/macrophage activation and polarization between pro- and anti-inflammatory states. For example, ...

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 ...

Biology

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.

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 ...

Using Real-Time Cell Metabolic Flux Analyzer to Monitor Osteoblast Bioenergetics

Bone formation by osteoblasts is an essential process for proper bone acquisition and bone turnover to maintain skeletal homeostasis, and ultimately, prevent fracture. In the interest to both optimize peak bone mass and combat various musculoskeletal diseases (i.e., post-menopausal osteoporosis, anorexia nervosa, type 1 and 2 diabetes mellitus), incredible efforts have been made in the field of bone biology to fully characterize osteoblasts throughout their differentiation process. Given the primary role of mature osteoblasts to secrete matrix proteins and mineralization vesicles, it has been noted that these processes take an incredible amount of cellular energy, or adenosine triphosphate (ATP). The overall cellular energy status is often referred to as cellular bioenergetics, and it includes a series of metabolic reactions that sense substrate availability to derive ATP to meet cellular needs. Therefore, the current method details the process of isolating primary, murine bone marrow stromal cells (BMSCs) and monitoring their bioenergetic status using the Real-time cell metabolic flux analyzer at various stages in osteoblast differentiation. Importantly, these data have demonstrated that the metabolic profile changes dramatically throughout osteoblast differentiation. Thus, using this physiologically relevant cell type is required to fully appreciate how a cell's bioenergetic status can regulate the overall function.

Real-time cell metabolic flux assay measures the oxygen consumption rate and extracellular acidification rate, which corresponds to mitochondrial and glycolytic adenosine triphosphate production, using pH and oxygen sensors. The manuscript explains a method to understand the energy status of osteoblasts and the characterization and interpretation of the cellular bioenergetic status.

Bone formation by osteoblasts is an essential process for proper bone acquisition and bone turnover to maintain skeletal homeostasis, and ultimately, ...

Novel OCR Monitoring System for Differentiated RPE Cultures

Long-term Monitoring of Oxygen Consumption Rates in Highly Differentiated and Polarized Retinal Pigment Epithelial Cultures

Mitochondrial metabolism is critical for the normal function of the retinal pigment epithelium (RPE), a monolayer of cells in the retina important for photoreceptor survival. RPE mitochondrial dysfunction is a hallmark of age-related macular degeneration (AMD), the leading cause of irreversible blindness in the developed world, and proliferative vitreoretinopathy (PVR), a blinding complication of retinal detachments. RPE degenerative conditions have been well-modeled by RPE culture systems that are highly differentiated and polarized to mimic in vivo RPE. However, monitoring oxygen consumption rates (OCR), a proxy for mitochondrial function, has been difficult in such culture systems because the conditions that promote ideal RPE polarization and differentiation do not allow for easy OCR measurements. 
Here, we introduce a novel system, Resipher, to monitor OCR for weeks at a time in well-differentiated RPE cultures while maintaining the RPE on optimal growth substrates and physiologic culture media in a standard cell culture incubator. This system calculates OCR by measuring the oxygen concentration gradient present in the media above cells. We discuss the advantages of this system over other methods for detecting OCR and how to set up the system for measuring OCR in RPE cultures. We cover key tips and tricks for using the system, caution about interpreting the data, and guidelines for troubleshooting unexpected results. 
We also provide an online calculator for extrapolating the level of hypoxia, normoxia, or hyperoxia RPE cultures experience based on the oxygen gradient in the media above cells detected by the system. Finally, we review two applications of the system, measuring the metabolic state of RPE cells in a PVR model and understanding how the RPE metabolically adapts to hypoxia. We anticipate that the use of this system on highly polarized and differentiated RPE cultures will enhance our understanding of RPE mitochondrial metabolism both under physiologic and disease states.

Author Spotlight: Extended Oxygen Consumption Measurement in Retinal Pigment Epithelium Using Resipher

We introduce a novel device for measuring oxygen consumption rates (OCR) in retinal pigment epithelial (RPE) cultures. The device can measure OCR for weeks at a time on RPE grown on standard cell culture plates with standard media while the plates are in a standard cell culture incubator.

Mitochondrial metabolism is critical for the normal function of the retinal pigment epithelium (RPE), a monolayer of cells in the retina important for ...

Real-Time Quantification of Mitochondrial Respiration in Muscle Fibers

Measurement of Mitochondrial Respiration in Human and Mouse Skeletal Muscle Fibers by High-Resolution Respirometry

Mitochondrial function, a cornerstone of cellular energy production, is critical for maintaining metabolic homeostasis. Its dysfunction in skeletal muscle is linked to prevalent metabolic disorders (e.g., diabetes and obesity), muscular dystrophies, and sarcopenia. While there are many techniques to evaluate mitochondrial content and morphology, the hallmark method to assess mitochondrial function is the measurement of mitochondrial oxidative phosphorylation (OXPHOS) by respirometry. Quantification of mitochondrial OXPHOS provides insight into the efficiency of mitochondrial oxidative energy production and cellular bioenergetics. A high-resolution respirometer provides highly sensitive, robust measurements of mitochondrial OXPHOS in permeabilized muscle fibers by measuring real-time changes in mitochondrial oxygen consumption rate. The use of permeabilized muscle fibers, as opposed to isolated mitochondria, preserves mitochondrial networks, maintains mitochondrial membrane integrity, and ultimately allows for more physiologically relevant measurements. This system also allows for the measurement of fuel preference and metabolic flexibility - dynamic aspects of muscle energy metabolism. Here, we provide a comprehensive guide for mitochondrial OXPHOS measurements in human and mouse skeletal muscle fibers using a high-resolution respirometer. Skeletal muscle groups are composed of different fiber types that vary in their mitochondrial fuel preference and bioenergetics. Using a high-resolution respirometer, we describe methods for evaluating both aerobic glycolytic and fatty acid substrates to assess fuel preference and metabolic flexibility in a fiber-type-dependent manner. The protocol is versatile and applicable to both human and rodent muscle fibers. The goal is to enhance the reproducibility and accuracy of mitochondrial function assessments, which will improve our understanding of an organelle important to muscle health.

Author Spotlight: Unveiling Mitochondrial Function and Cellular Metabolic Adaptation in Metabolic Diseases

Here, we describe a comprehensive method for measuring mitochondrial oxidative phosphorylation in fresh permeabilized skeletal muscle fibers from either human or mouse muscle. This method allows for the real-time quantification of mitochondrial respiration and the assessment of fuel preference and metabolic flexibility while preserving existing mitochondrial networks and membrane integrity.

Mitochondrial function, a cornerstone of cellular energy production, is critical for maintaining metabolic homeostasis. Its dysfunction in skeletal muscle ...

Extracellular Flux Assay: A Method for Measuring Metabolic Profile of Cells

Source: Hlozkova K. et al. <a href="https://www.jove.com/t/58426">Assessment of the Metabolic Profile of Primary Leukemia Cells.</a> J. Vis. Exp. (2018).
This protocol provides a biochemical approach for measuring extracellular acidification rate (ECAR) and oxygen consumption rate (OCR) of primary leukemic cells with extracellular flux assay. The assessment of the metabolic profile of these cells helps to characterize their demands, leading to personalized therapy.

Source: Hlozkova K. et al. Assessment of the Metabolic Profile of Primary Leukemia Cells. J. Vis. Exp. (2018).
This protocol provides a biochemical ...

Analyzing Oxygen Consumption Rate in Primary Cultured Mouse Neonatal Cardiomyocytes Using an Extracellular Flux Analyzer

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