This work is supported by the Intramural Research Program of the National Eye Institute (ZIAEY000450 and ZIAEY000546).

All mouse protocols were approved by the Animal Care and Use Committee of the National Eye Institute (NEI ASP# 650). Mice were housed in 12 h light-dark conditions and cared for by following the recommendations of the Guide for the Care and Use of Laboratory Animals, the Institute of Laboratory Animal Resources, and the Public Health Service Policy on Humane Care and Use of Laboratory Animals.
1. Hydrating sensor cartridge and preparation of the assay medium
<ol>
	<li>The day before the experiment, add 1 mL of the calibration medium to each well of the utility plate. Place the Hydro-Booster cover on the top and lower the sensor cartridge through the opening on the cover. Check to ensure that the sensor is submerged in the calibration medium. Incubate the sensor cartridge overnight in a CO2-free incubator at 37 &#730;C to activate the fluorophores. 
	NOTE: To prevent evaporation, the incubator is humidified by keeping a tray of water inside, and the sensor cartridge cassette is wrapped with clear plastic wrap.</li>
	<li>Prepare the assay medium by reconstituting the Seahorse DMEM medium with the addition of glucose, pyruvate, and glutamine to the desired concentrations. In the assays reported in this article, the final concentration of substrates in the assay medium are: 6 mM of glucose, 0.12 mM of pyruvate, and 0.5 mM of glutamine. For each assay plate, 40 mL of the assay medium is prepared fresh on the day of the experiment.</li>
	<li>Set up the assay program in the analyzer following the manufacturer&#39;s instruction26. In the assay demonstrated here, the protocol is set as follows: 5 cycles of measurements for baseline, then inject port A, followed by 4 cycles of measurements, then inject port B and followed by 4 cycles of measurements. Each cycle is composed of mix (3 min), wait (2 min) and measure (3 min).</li>
</ol>
2. Coating mesh inserts of islet capture microplate
<ol>
	<li>Prepare the coating mix by combining 20 &#181;L of the cell attachment medium (e.g., Cell-Tak) with 171 &#181;L of 0.1 M sodium bicarbonate and 9 &#181;L of 1 M NaOH.</li>
	<li>Open the lid of the cassette containing mesh inserts. Pipette 8 &#181;L of the coating mix to each mesh inserts. Use a pipette tip to gently smear/spread the droplet around to distribute the coating mix equally throughout the mesh insert.</li>
	<li>Close the cassette and allow the mesh inserts to incubate at room temperature for at least 25 min for adsorption.</li>
	<li>Wash the mesh insert by pipetting 4 mL of the assay medium directly onto the mesh inserts. Gently shake the cassette to ensure all mesh inserts are washed with the assay medium.</li>
	<li>Keep the mesh insert aside. It is ready to use.</li>
</ol>
3. Preparing injection compounds
<ol>
	<li>Take out stock aliquots of Bam15 (10 mM), Rotenone (10 mM), Antimycin A (10 mM) and 2-DG (500 mM) from -80 &#176;C freezer and thaw at room temperature. 
	NOTE: The 2-DG stock is ready to use. The other drugs need to be diluted to working stock.</li>
	<li>Warm up 10 mL of the assay medium in a 37 &#176;C water bath.</li>
	<li>Dilute 10 mM Bam15 stock to 50 &#181;M working stock using a two-step dilution procedure: mix 20 &#181;L of 10 mM stock with 20 &#181;L of DMSO to get 5 mM intermediate stock. Then mix 10 &#181;L of the above 5 mM intermediate stock with 990 &#181;L of pre-warmed assay medium to get the final 50 &#181;M working stock.</li>
	<li>Dilute and combine 10 mM Rotenone and 10 mM Antimycin A stock to 10 &#181;M Rotenone/Antimycin A (Rot/AA) working stock by two steps of dilutions: mix 10 &#181;L each of 10 mM Rotenone and 10 mM Antimycin A stock with 80 &#181;L of DMSO to get 1 mM Rot/AA intermediate stock. Then mix 10 &#181;L of the above 1 mM intermediate stock with 990 &#181;L of pre-warmed assay medium to get the final 10 &#181;M Rot/AA working stock.</li>
	<li>Freshly prepare the above-mentioned working stocks of injection compounds on the day of the experiment and set them aside at room temperature until loading into injection ports of the sensor cartridge.</li>
</ol>
4. Retinal dissection and retinal punch preparation
<ol>
	<li>Euthanize a mouse by CO2 asphyxiation following AVMA Guidelines on Euthanasia27. 
	NOTE: Do not leave the animal in a CO2 chamber longer than the time needed for euthanasia.</li>
	<li>Enucleate eyes and place into ice-old 1x PBS buffer in a Petri-dish and then place it under a dissection microscope.</li>
	<li>Carefully remove, by cutting with microscissors, the extra rectus muscles attached outside the eyeball and cut off the optic nerve.</li>
	<li>Use a 30 G needle to punch a hole at the edge of the cornea (limbus); this serves as the insertion site for the microscissors. Then, use a fine dissection microscissors to make a circular cut along the edge of the cornea, separating it from the posterior eye cup.</li>
	<li>Use sharp dissection forceps to remove the cornea, lens, and the vitreous humor away from the eye cup.</li>
	<li>Use fine dissection microscissors to make several small cuts on the scleral layer at the rim of eye cup. Avoid cutting the retina layer. Use two sharp dissection forceps to hold on to the scleral tissue at each side of the cut and very carefully pull on the scleral layer to remove it from the neural retina. Repeat this around the eye cup until all sclera is removed and an intact retinal cup is obtained.</li>
	<li>Use dissection microscissors and make radial cuts on the retinal cup to flatten it and generate several distinct sections. 
	NOTE: Depending on the person&#39;s dissection skills and experience in handling fresh retinal tissue, the retinal cup can be cut to generate 3 to 5 distinct sections.</li>
	<li>Use 1 mm diameter biopsy puncher to cut one retinal disk from each section of the flattened retinal cup. 
	NOTE: Care should be taken to get the retinal disks punched at equal distance from the optic nerve head.</li>
	<li>Use forceps to transfer the pre-coated mesh inserts into the dissecting petri-dish. With the help of two superfine eyelash brushes, place the retinal punch disk onto the mesh insert. The retinal punch disk is placed at the center of the mesh insert with ganglion cell layer side down touching the mesh and photoreceptor layer facing up. 
	NOTE: Frequently, some RPE cells remain attached to the photoreceptors, and the pigmentation of these cells can be used as an indicator of the retinal punch disk orientation.</li>
</ol>
5. Loading the sensor cartridge injection ports and calibration
<ol>
	<li>Take the hydrated sensor cartridge plate cassette out of the 37 &#176;C incubator. Remove the Hydro-Booster cover and place the sensor cartridge back on the utility plate.</li>
	<li>Load the desired volume of injection compound working stock solutions into appropriate ports. Hold the pipette tip at 45 &#176;&#160;angle. Insert the pipette tip halfway into an injection port with the bevel of the tip against the opposite wall of the injection port and gently load the compound into each port. Avoid introducing air bubbles.</li>
	<li>Refer to the instrument user guide for the volume of the compound loaded in each injection port for a specific assay. In the experiments presented in this paper, 68 &#181;L of 50 &#181;M Bam15 working stock (for mitochondrial stress assay) or 68 &#181;L of 10 &#181;M Rot/AA working stock (for glycolytic rate assay) is loaded into port A; 75 &#181;L of 10 &#181;M Rot/AA working stock (for mitochondrial stress assay) or 75 &#181;L of 500 mM 2-DG working stock (for glycolytic rate assay) is loaded into port B.</li>
	<li>Load injection ports of all wells of the plate including background correction wells and blank wells to ensure proper injection. Load the respective compound solution in each port for the background correction wells. Assay medium can be substituted, instead of the compound solution, in each of the ports of the bank wells.</li>
	<li>Place the loaded sensor cartridge plate, with lid off, into the analyzer machine to start calibration prior to the assay run. After the calibration is over, the program will automatically pause, waiting for the replacement of the utility plate with the islet capture plate containing retinal punches.</li>
</ol>
6. Loading the islet capture plate and start assay run
<ol>
	<li>Add 607 &#181;L of the assay medium to each well of the islet capture plate</li>
	<li>Use forceps to grab the rim of the mesh insert containing retinal punch disks on top and take it out from the Petri-dish. Lightly tap the bottom of the mesh insert on an absorbing wipe tissue to remove extra liquid and put it into the well of the islet capture plate. Repeat this step until all mesh inserts with retinal punches are placed into the islet capture plate. Fill background correction wells and blank wells with empty mesh inserts.</li>
	<li>Use two Graefe forceps to carefully and gently press the rim of each mesh insert and make sure that these are securely inserted at the bottom of the islet capture plate.</li>
	<li>Place the loaded islet capture plate into a 37 &#176;C incubator for 5 min to warm up.</li>
	<li>Eject the utility plate after the calibration is complete and replace it with an and replace it with the islet capture plate, with lid off, containing&#160;retinal punches.</li>
	<li>Resume the assay run.</li>
</ol>
7. Run termination and data storage
<ol>
	<li>After the run is complete, eject the sensor cartridge and islet capture plate containing retinal punches. The data is automatically saved as .asyr file.</li>
	<li>Use the associated data analysis software to view and analyze the data following the manufacturer&#39;s user guide26.</li>
	<li>Use the Export function to export .xslx file of the data, which can be viewed and analyzed using spreadsheet software.</li>
</ol>
8. Saving the retinal punch sample
<ol>
	<li>After the assay, take out the plate from the machine, remove the sensor cartridge and gently remove the assay medium from each well using a pipette.</li>
	<li>Apply the cover back on and seal the sides of the plate with the parafilm strip.</li>
	<li>Store at -80 &#176;C.</li>
	<li>For normalization, quantify the total DNA or protein content of the punch in each well.</li>
</ol>
9. Data analysis
<ol>
	<li>Mitochondrial stress assay 
	NOTE: The measured OCR value (totalOCR) represents total oxygen consumption by the tissue. After Bam15 (uncoupler) injection, OCR increases from the basal level (totalOCRbasal) to the maximum level (totalOCRmax) and goes down following the Rot/AA injection. The residual OCR value after Rot/AA injection (totalOCRRot/AA) represents non-mitochondrial oxygen consumption.
	<ol>
		<li>Calculate mitochondria-related oxygen consumption as: 
		<img alt="Equation 1" src="/files/ftp_upload/62914/62914eq01.jpg" /> (Eq. 1)28</li>
		<li>Calculate the mitochondrial reserve capacity (MRC) as: 
		<img alt="Equation 2" src="/files/ftp_upload/62914/62914eq2v4.png" /> (Eq. 2)29 
		NOTE: The last reading among the 5 measurements before Bam15 injection is taken as the &#34;basal&#34; value (for totalOCRbasal and mitoOCRbasal). The highest reading among the 4 measurements following Bam15 injection is used as &#34;max&#34; value (for totalOCRmax and mitoOCRmax). The lowest reading among the 4 measurements following the Rot/AA injection is used as totalOCRRot/AA.</li>
	</ol>
	</li>
	<li>Glycolytic rate assay 
	NOTE: The measured ECAR value (totalECAR) represents the total acidification of the medium by the tissue&#39;s metabolic activity. In general, acidification of the extracellular micro-environment results mainly by extrusion of the glycolytic product, lactate. Catabolism of substrates in mitochondrial TCA cycle results in the production of CO2, which also acidifies the extracellular medium through hydration to bicarbonate.
	<ol>
		<li>Substract mitochondrial contributed medium acidification (mitoECAR) from totalECAR to obtain the glycoECAR. 
		<img alt="Equation 3" src="/files/ftp_upload/62914/62914eq03.jpg" /> (Eq. 3)28 
		NOTE: Mitochondrial respiration and TCA cycle are strongly coupled processes. Production of CO2 from mitochondria is a function of the rate of OXPHOS, which is measurable by mitoOCR.</li>
		<li>Calculate the mitoECAR as: 
		<img alt="Equation 4" src="/files/ftp_upload/62914/62914eq04.jpg" /> (Eq. 4)28 
		where, the CCF (CO2 Contribution Factor) is an empirically calculated ratio value, representing the amount of H+ contribution from CO2-mediated acidification vs each O2 consumption from OXPHOS. CCF for this system is pre-determined to be 0.6028. Accurate measurement of medium acidification is determined by the buffer capacity of the medium, the sensitivity of instrument pH sensor, and the effective measurement chamber capacity. Here, the BF (Buffer Factor) is a parameter of the in situ experimental buffer capacity, representing the amount of H+ or OH- added to the effective measurement chamber to change the pH level by 1 unit. When customized assay medium is used, the BF can be determined by titrating known amounts of acid into the assay medium following the Buffer Factor protocol30. The Seahorse DMEM medium pH 7.4 used in this protocol has a pre-determined BF of 2.60 mmol H+/L/pH. The islet capture plate used in this protocol has a Volmicrochamber = 16.6 &#181;L31. The volume scaling factor, Kvol, is an empirically determined constant. Kvol value is not available for the islet capture plate but can be calculated from the value of the microplate28, accounting for the volume difference in their microchambers, to be 0.41. 
		NOTE: Injection of the Rot/AA shuts down mitochondrial respiration and forces the tissue to switch to glycolysis for ATP production, leading to higher lactate extrusion and an increase in ECAR measurement. Glycolysis is ceased with 2-DG injection, and the residual ECAR measurement reveals non-glycolytic and non-mitochondrial acidification of medium.</li>
		<li>Calculate the glycolytic reserve capacity (GRC) as: 
		<img alt="Equation 5" src="/files/ftp_upload/62914/62914eq05.jpg" /> (Eq. 5)32 
		where, the last reading among the 5 measurements before Rot/AA injection is taken as the &#34;basal&#34; value (glycoECARbasal). The highest reading among the 4 measurements following Rot/AA injection is used as &#34;max&#34; value (glycoECARmax). The lowest reading among the 4 measurements following 2-DG injection is used as glycoECAR2-DG.</li>
	</ol>
	</li>
	<li>Normalization 
	NOTE: Normalization is essential when comparing the readings from retinal tissues of different age groups or between wild-type and pathological/degenerative samples, which might differ in cell numbers.
	<ol>
		<li>Use commerically available kits to assess the DNA content in each retinal punch disk33,34.</li>
		<li>Alternatively, use radioimmunoprecipitation assay buffer (RIPA buffer) to extract total protein from the retinal punch and use the protein content for normalization. 
		NOTE: The surface area of an adult mouse retina has been previously determined to be around 20 mm2, and each retina contains ~6.5 million cells35. Hence, each 1 mm diameter retinal punch is ~1/25 of a single retina and contains ~260K cells. One can refer to these numbers when comparing the data from a retinal punch to those from other tissue samples or cultured cells,</li>
	</ol>
	</li>
</ol>

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Report Generator User Guide Agilent Seahorse XF Cell Mito Stress Test Available from: <a target="_blank" href="https://www.agilent.com/cs/library/usermanuals/public/Report_Generator_User_Guide_Seahorse_XF_Cell_Mito_Stress_Test_Single_File.pdf">https://www.agilent.com/cs/library/usermanuals/public/Report_Generator_User_Guide_Seahorse_XF_Cell_Mito_Stress_Test_Single_File.pdf</a> (2021)</li><li>. Agilent Seahorse XF Buffer Factor Protocol Available from: <a target="_blank" href="https://www.agilent.com/cs/library/usermanuals/public/usermanual-xf-buffer-factor-protocol-cell-analysis-S7888-10010en-agilent.pdf">https://www.agilent.com/cs/library/usermanuals/public/usermanual-xf-buffer-factor-protocol-cell-analysis-S7888-10010en-agilent.pdf</a> (2021)</li><li>. Agilent sensor cartridges and cell culture microplates Available from: <a target="_blank" href="https://www.agilent.com/cs/library/brochures/5991-8657EN_seahorse_plastics_brochure.pdf">https://www.agilent.com/cs/library/brochures/5991-8657EN_seahorse_plastics_brochure.pdf</a> (2021)</li><li>. Agilent Seahorse XF Glycolysis Stress Test Kit User Guide Available from: <a target="_blank" href="https://www.agilent.com/cs/library/usermanuals/public/XF_Glycolysis_Stress_Test_Kit_User_Guide.pdf">https://www.agilent.com/cs/library/usermanuals/public/XF_Glycolysis_Stress_Test_Kit_User_Guide.pdf</a> (2021)</li><li>Fan, Y. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+bioassay+to+measure+energy+metabolism+in+mouse+colonic+crypts,+organoids,+and+sorted+stem+cells.">A bioassay to measure energy metabolism in mouse colonic crypts, organoids, and sorted stem cells.</a> American Journal of Physiology-Gastrointestinal and Liver Physiology. 309, 1-9 (2015).</li><li>Huang, 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=Ductal+pancreatic+cancer+modeling+and+drug+screening+using+human+pluripotent+stem+cell-+and+patient-derived+tumor+organoids.">Ductal pancreatic cancer modeling and drug screening using human pluripotent stem cell- and patient-derived tumor organoids.</a> Nature Medicine. 21 (11), 1364-1371 (2015).</li><li>Jeon, C. J., Strettoi, E., Masland, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+major+cell+populations+of+the+mouse+retina.">The major cell populations of the mouse retina.</a> Journal of Neuroscience. 18 (21), 8936-8946 (1998).</li><li>Akimoto, M., 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+of+GFP+to+newborn+rods+by+Nrl+promoter+and+temporal+expression+profiling+of+flow-sorted+photoreceptors.">Targeting of GFP to newborn rods by Nrl promoter and temporal expression profiling of flow-sorted photoreceptors.</a> Proceedings of the National Academy of Science U. S. A. 103 (10), 3890-3895 (2006).</li><li>Kenwood, B. M., 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=Identification+of+a+novel+mitochondrial+uncoupler+that+does+not+depolarize+the+plasma+membrane.">Identification of a novel mitochondrial uncoupler that does not depolarize the plasma membrane.</a> Molecular Metabolism. 3 (2), 114-123 (2014).</li><li>Corso-Diaz, X., 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=Genome-wide+profiling+identifies+DNA+methylation+signatures+of+aging+in+rod+photoreceptors+associated+with+alterations+in+energy+metabolism.">Genome-wide profiling identifies DNA methylation signatures of aging in rod photoreceptors associated with alterations in energy metabolism.</a> Cell Reports. 31 (3), 107525 (2020).</li><li>Berkowitz, B. A., 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=Mitochondrial+respiration+in+outer+retina+contributes+to+light-evoked+increase+in+hydration+in+vivo.">Mitochondrial respiration in outer retina contributes to light-evoked increase in hydration in vivo.</a> Investigative Ophthalmology and Visual Science. 59 (15), 5957-5964 (2018).</li><li>Joyal, J. S., 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=Retinal+lipid+and+glucose+metabolism+dictates+angiogenesis+through+the+lipid+sensor+Ffar1.">Retinal lipid and glucose metabolism dictates angiogenesis through the lipid sensor Ffar1.</a> Nature Medicine. 22 (4), 439-445 (2016).</li></ol>

The authors have nothing to disclose.

Provided here are detailed instructions for performing microplate-based assays of mitochondrial respiration and glycolysis activity using ex vivo, freshly dissected retinal punch disks. The protocol has been optimized to: 1) ensure the use of a suitable assay medium for ex vivo retinal tissue; 2) employ proper size of retinal punch disks to obtain OCR and ECAR readings that fall within the machine&#39;s optimal detecting range; 3) coating mesh inserts to enhance the adhesiveness of retinal punch for stable reading during the measuring cycle; 4) use of optimal concentration of each injected drug compounds; and 5) ensure altered cycle length to reach a plateau of mitochondrial states at each step. The reagents and protocol have been modified from a previously published version23 to adapt to the new generation Seahorse XFe24 machine. Instead of the Ames&#39; buffer used in the previous protocol23, a basic Seahorse DMEM medium is used here to allow the custom constitution of fuel source by adding glucose, glutamine, and pyruvate separately. This also makes it possible to perform various assays where a specific fuel substrate is supplied or deprived from the medium. In the assays presented here, the medium was constituted to the same concentration of glucose (6 mM), glutamine (0.5 mM), and pyruvate (0.12 mM) as in Ames&#39; buffer, which are proven suitable for retinal tissue. Another advantage of this medium (with 5 mM HEPES) over the Ames&#39; buffer (with 22.6 mM NaHCO3) is its low buffer capacity, which ensures sensitive and accurate measurement of ECAR28.
Both mitochondrial stress and glycolytic rate assays can be performed following the protocol described here with high precision, as evidenced by the tight standard error values between replicating wells. However, it is worthwhile to note the factors that can contribute to data variability. Avoid cell death in retinal tissue. The entire dissection process should be performed in ice-cold 1x PBS, and the process from enucleation of eyes to putting islet capture plate containing the retinal punch into the machine should not exceed 2 hours. Caution should be taken during the dissection of the retina cup to avoid any damage to the retinal tissue, and punches should not be taken from areas damaged by dissection. New, sharp biopsy puncher should be used in each experiment, and change the puncher when the edge is dull or bent to ensure consistency and accuracy in cutting retinal punches at 1 mm diameter. Try to get the retinal disks punched at equidistant from the optic nerve head to avoid regional variations (center versus peripheral). After the assay, check each well for any sign of the retinal punch being detached from the mesh insert. When a retinal punch has poor adhesion on mesh insert or detaches during measurement, the distance from senser probe to the tissue will change, affecting the readings. Omit the data from such wells with detached retinal punch.
Measurement of the real-time mitochondrial metabolism in intact retinal tissue has broad applications and can provide useful information for various studies. These assays have been used to measure mitochondrial respiration in retinal tissues from mice of different genetic backgrounds to reveal their intrinsic difference in mitochondrial activity39,40. It was also used to study changes in mitochondrial energy metabolism during aging of the retina38. By providing different fuel substrates and utilizing various inhibitors targeting different metabolic pathways, it provides insights to the preference of the cell/tissue on certain fuel sources22,38. Furthermore, comparison on OCR and MRC between wild-type mouse and mouse models of inherited retinal degeneration can provide evidence of mitochondrial defects in degenerating retina22.
There are limitations of this technique. The islet capture plate used in these assays only contains 24 wells; hence, it is only able to provide mid-throughput analysis. The data quality from this method is contingent upon the quality of retinal punch disks and viability of cells. Also, retinal dissection and retinal punch disks preparation is a time-consuming process, rendering it less feasible to high-throughput analysis on live ex vivo retinal tissues even when 96-well plates are available. Compared to a monolayer of cultured cells, penetration of drug compound into the retinal tissue also affects data readout. In addition, the measured OCR and ECAR values represent the total performance of the entire tissue, which is composed by many different cell types; hence, one needs to consider the relationship and interactions among different neuronal and glial cells in the retina while interpreting the data. Specific experimental designs should be implemented by tailoring to each project. It is recommended that one includes 3 to 5 retinal punches (from same eye or same mouse) as technical replicates and use samples from 3 or more mice as biological replicates.

Mitochondria are essential organelle that regulates cellular metabolism, signaling, homeostasis, and apoptosis by coordinating multiple crucial physiological processes1. Mitochondria serve as the powerhouse in the cell to generate adenosine triphosphate (ATP) through oxidative phosphorylation (OXPHOS) and provide energy that supports almost all cellular events. The majority of cellular oxygen is metabolized in mitochondria, where it serves as the final electron acceptor in the electron transport chain (ETC) during aerobic respiration. Low amounts of ATP can also be produced from glycolysis in the cytosol, where glucose is converted to pyruvate, which can be further converted to lactate or be transported into mitochondria and oxidized to acetyl-CoA, a substrate in the tricarboxylic acid cycle (TCA cycle).
The retina is one of the most metabolically active tissues in mammals2, displaying high levels of mitochondrial respiration and extremely high oxygen consumption3. The rod and cone photoreceptors contain a high density of mitochondria4, and OXPHOS generates most ATP in the retina5. In addition, the retina also relies heavily on aerobic glycolysis6,7 by converting glucose to lactate5. Mitochondrial defects are associated with various neurodegenerative diseases8,9; and with its unique high energy demands, the retina is especially vulnerable to metabolic defects, including those affecting mitochondrial OXPHOS4 and glycolysis10. Mitochondrial dysfunction and defects in glycolysis are implicated in retinal11,12 and macular13 degenerative diseases, age-related macular degeneration10,14,15,16, and diabetic retinopathy17,18. Therefore, accurate measurements of mitochondrial respiration and glycolysis can provide important parameters for assessing the integrity and health of the retina.
Mitochondrial respiration can be measured through the determination of oxygen consumption rate (OCR). Given that the conversion of glucose to pyruvate and subsequently to lactate results in extrusion of protons into and acidification of the extracellular environment, measurements of the extracellular acidification rate (ECAR) provide an indication of glycolysis flux. As the retina is composed of multiple cell types with intimate relationships and active synergy, including the exchange of substrates6, it is imperative to analyze mitochondrial function and metabolism in the context of whole retinal tissue with intact lamination and circuitry. For the past several decades, the Clark type O2 electrodes and other oxygen microelectrodes have been used to measure oxygen consumption in the retina19,20,21. These oxygen electrodes have major limitations in sensitivity, requirement of a large sample volume, and the need for continuous stirring of suspending sample, which usually leads to the disruption of cellular and tissue context. The protocol described here was developed using a microplate-based, fluorescence technique to measure mitochondrial energy metabolism in freshly dissected ex vivo mouse retina tissue. It allows mid-throughput real-time measurements of both OCR and ECAR simultaneously using a small sample (1 mm punch) of ex vivo retinal tissue while avoiding the need for suspension and continuous stirring.
Demonstrated here is the experimental procedure for mitochondrial stress assay and glycolytic rate assay on freshly dissected retinal punch disks. This protocol allows the measurement of mitochondria-related metabolic activities in an ex vivo tissue context. Different from the assays performed using cultured cells, the readings obtained here reflect combined energy metabolism at the tissue level and are influenced by interactions between the different cell types within the tissue. The protocol is modified from a previously published version22,23 to adapt to the new generation of the Agilent Seahorse extracellular flux 24-wells (XFe24) analyzer with Islet Capture plate. The assay medium, injection compound concentrations, and number/duration of assay cycles have also been optimized for retinal tissue. A detailed step-by-step protocol is given for the preparation of retinal punch disks. More information on the program setup and data analysis can be obtained from the manufacturer&#39;s user guide24,25,26.

<table><tbody><tr><td>1X PBS</td><td>Thermo Fisher</td><td>14190-144</td><td></td></tr><tr><td>2-Deoxy glucose (2-DG), 500 mM stock solution</td><td>Sigma</td><td>D6134</td><td>Dissolve in Seahorse XF DMEM medium, prepare ahead of time</td></tr><tr><td>30-gauge needle</td><td>BD Precision Glide</td><td>305106</td><td></td></tr><tr><td>Antimycin A, 10 mM stock solution</td><td>Sigma</td><td>A8674</td><td>Dissolve in DMSO, prepare ahead of time</td></tr><tr><td>Bam15, 10 mM stock solution</td><td>TimTec</td><td>ST056388</td><td>Dissolve in DMSO, prepare ahead of time</td></tr><tr><td>Biopsy puncher, 1 mm</td><td>Integra Miltex</td><td>33-31AA</td><td></td></tr><tr><td>Cell-Tak</td><td>Corning Life Sciences</td><td>CB40240</td><td></td></tr><tr><td>CO2 asphyxiation chamber</td><td></td><td></td><td></td></tr><tr><td>Dissection forceps-Dumont #5</td><td>Fine Science Tools</td><td>11251-10</td><td>Stright tip</td></tr><tr><td>Dissection forceps-Dumont #7</td><td>Fine Science Tools</td><td>11274-20</td><td>Curved tip</td></tr><tr><td>Dissection microscope</td><td></td><td></td><td></td></tr><tr><td>DMSO</td><td>Sigma</td><td>D2438</td><td></td></tr><tr><td>Graefe forceps</td><td>Fine Science Tools</td><td>11051-10</td><td>Curved, Serrated tip</td></tr><tr><td>Microscissors</td><td>Fine Science Tools</td><td>15004-08</td><td>Curved tip</td></tr><tr><td>NaOH solution, 1 M</td><td>Sigma-Aldrich</td><td>S8263</td><td>Aqueous solution, prepare ahead of time</td></tr><tr><td>Rotenone, 10 mM stock solution</td><td>Sigma</td><td>R8875</td><td>Dissolve in DMSO, prepare ahead of time</td></tr><tr><td>Seahorse calibration medium</td><td>Agilent</td><td>100840-000</td><td></td></tr><tr><td>Seahorse XF 1.0 M glucose</td><td>Agilent</td><td>103577-100</td><td></td></tr><tr><td>Seahorse XF 100 mM pyruvate</td><td>Agilent</td><td>103578-100</td><td></td></tr><tr><td>Seahorse XF 200 mM glutamine</td><td>Agilent</td><td>103579-100</td><td></td></tr><tr><td>Seahorse XF DMEM medium</td><td>Agilent</td><td>103575-100</td><td>pH 7.4, with 5 mM HEPES</td></tr><tr><td>Seahorse XFe24 Islet Capture FluxPak</td><td>Agilent</td><td>103518-100</td><td>Containing Sensor Cartridge and Islet Capture microplate</td></tr><tr><td>Seahorse XFe24, Extra Cellular Flux Analyzer</td><td>Agilent</td><td></td><td></td></tr><tr><td>Sodium bicarbonate solution, 0.1 M</td><td>Sigma-Aldrich</td><td>S5761</td><td>Aqueous solution, prepare ahead of time</td></tr><tr><td>Superfine eyelash brush</td><td>Ted Pella</td><td>113</td><td></td></tr></tbody></table>

determination of mitochondrial respiration and glycolysis in ex vivo retinal tissue samples

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. In addition, photoreceptors also exhibit high aerobic glycolysis like cancer cells. Precise measurements of these metabolic activities can provide valuable insights into cellular homeostasis under physiological conditions and in disease states. High throughput microplate-based assays have been developed to measure mitochondrial respiration and various metabolic activities in live cells. However, a vast majority of these are developed for cultured cells and have not been optimized for intact tissue samples and for application ex vivo. Described here is a detailed step-by-step protocol, using microplate-based fluorescence technology, to directly measure oxygen consumption rate (OCR) as an indicator of mitochondrial respiration, as well as extracellular acidification rate (ECAR) as an indicator of glycolysis, in intact ex vivo retinal tissue. This method has been used to successfully assess metabolic activities in adult mouse retina and demonstrate its application in investigating cellular mechanisms of aging and disease.

The data reported here are representative mitochondrial stress assay showing OCR trace (Figure 1) and glycolytic rate assay showing OCR trace and ECAR trace (Figure 2), which were performed using freshly dissected 1 mm retinal punch disks from 4 months old transgenic Nrl-L-EGFP mice36 (C57B/L6 background). These mice express GFP specifically in rod photoreceptors without altering normal retinal development, histology, and physiology and have been widely used as wild-type controls in retinal research. Two Nrl-L-GFP littermate mice were used were used in the assays presented here. GFP expressed in the Nrl-L-GFP mice does not interfere with the measurements of OCR and ECAR in this protocol. Five retinal punches were taken from each retina. Ten of the retinal punches were used for mitochondrial stress assay and the other 10 were used for glycolytic rate assay. Seahorse XF DMEM medium, pH 7.4 (constituted with 6 mM glucose, 0.12 mM pyruvate, and 0.5 mM glutamine) and Seahorse XFe24 Islet Capture plates were used in the experiments. The representative data presented here were obtained using the same 1 mm diameter puncher but was not normalized with respect to the DNA/protein content.
In mitochondrial stress assay, the uncoupler Bam1537 was injected after establishing the OCR baseline, leading to enhanced OCR to the maximal level. Rotenone and Antimycin A were injected to inhibit mitochondria respiration at complex I and complex III, respectively, resulting in OCR to drop to the minimal level (Figure 1). The difference between the maximal level of OCR and the last measurement of the basal OCR level reflects mitochondrial reserve capacity (MRC). The MRC is calculated to be 19.2%&#177;3.4% using Eq. 2, consistent with previously measured MRC values in retinas of ~3 months old Nrl-L-EGFP mice using the previous generation Seahorse XF24 analyzer22,38.
In the glycolytic rate assay, Rotenone and Antimycin A were injected after establishing the baseline for the total ECAR. With the production of ATP from OXPHOS halted, the tissue is forced to rely on glycolysis for energy, and an increase in the extracellular release of lactate drives ECAR to the maximal level. Glycolysis is ceased by injection of 2-DG, which competes with glucose for hexokinase binding, causing ECAR to drop to the minimal level (Figure 2). Mitochondria contributed ECAR (mitoECAR) can be calculated from the mitoOCR value (Eq. 4). Glycolysis contributed ECAR glycoECAR is calculated and plotted by subtracting mitoECAR from totalECAR. The difference between maximal level of glycoECAR and the last measurement of glycoECAR basal level reflects the glycolysis reserve capacity (GRC). Here, the GRC is calculated to be 35.7% &#177; 3.4% using Eq. 5.
As a highly glycolytic tissue, lactate production from the retina accounts for a major source of extracellular acidification, as revealed by the small difference of glycoECAR from the totalECAR. Interestingly, ECAR measurement does not plateau immediately following the Rot/AA injection but drops after the second measurement. The retinal punch disk is an intact ex vivo system composed of different cell types, including the M&#252;ller glia cells, which are known to receive lactate (glycolysis end product) released from the photoreceptors6. Hence, a drop in ECAR measurement following the Rot/AA injection is likely explained by increased removal of lactate from the intercellular space, slowing down/preventing its release into the medium.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62914/62914fig01.jpg" /> 
Figure 1: Mitochondrial stress assay. The plotted graph shows OCR trace from 1 mm retinal punch disks in Seahorse XF DMEM buffer, supplemented with 6 mM of glucose, 0.12 mM of pyruvate and 0.5 mM of glutamine. Each data point represents the average of measurements from 10 wells. Error bar = standard error. MRC is calculated to be 19.2%&#177;3.4%. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62914/62914fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62914/62914fig02.jpg" /> 
Figure 2: Glycolytic rate assay. The plotted graph shows the measured OCR trace, ECAR trace (totalECAR), and the calculated glycolysis contributed ECAR (glycoECAR) from 1 mm retinal punch disks in Seahorse XF DMEM buffer supplemented with 6 mM of glucose, 0.12 mM of pyruvate, and 0.5 mM of glutamine. Each data point represents the average of measurements from 10 wells. Error bar = standard error. GRC is calculated to be 35.7%&#177;3.4% <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62914/62914fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples at JoVE.com

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Described here is a detailed protocol for performing mitochondrial stress assay and glycolytic rate assay in ex vivo retinal tissue samples using a commercial bioanalyzer.

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. ...

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Nanyang Technological University

Research

JoVE Journal

Neuroscience

4.1K Views.  National Institutes of Health. Described here is a detailed protocol for performing mitochondrial stress assay and glycolytic rate assay in ex vivo retinal tissue samples using a commercial bioanalyzer.

Video: Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Measuring Liver Mitochondrial Oxygen Consumption and Proton Leak Kinetics to Estimate Mitochondrial Respiration in Holstein Dairy Cattle

Oxygen consumption, proton motive force (PMF) and proton leak are measurements of mitochondrial respiration, or how well mitochondria are able to convert NADH and FADH into ATP. Since mitochondria are also the primary site for oxygen use and nutrient oxidation to carbon dioxide and water, how efficiently they use oxygen and produce ATP directly relates to the efficiency of nutrient metabolism, nutrient requirements of the animal, and health of the animal. The purpose of this method is to examine mitochondrial respiration, which can be used to examine the effects of different drugs, diets and environmental effects on mitochondrial metabolism. Results include oxygen consumption measured as proton dependent respiration (State 3) and proton leak dependent respiration (State 4). The ratio of State 3 / State 4 respiration is defined as respiratory control ratio (RCR) and can represent mitochondrial energetic efficiency. Mitochondrial proton leak is a process that allows dissipation of mitochondrial membrane potential (MMP) by uncoupling oxidative phosphorylation from ADP decreasing the efficiency of ATP synthesis. Oxygen and TRMP+ sensitive electrodes with mitochondrial substrates and electron transport chain inhibitors are used to measure State 3 and State 4 respiration, mitochondrial membrane PMF (or the potential to produce ATP) and proton leak. Limitations to this method are that liver tissue must be as fresh as possible and all biopsies and assays must be performed in less than 10 h. This limits the number of samples that can be collected and processed by a single person in a day to approximately 5. However, only 1 g of liver tissue is needed, so in large animals, such as dairy cattle, the amount of sample needed is small relative to liver size and there is little recovery time needed.

Here, we share methods for measuring mitochondrial oxygen consumption, a defining concept of nutritional energetics, and proton leak, the primary cause of inefficiency in mitochondrial generation of ATP. These results can account for 30% of the energy lost in nutrient utilization to help evaluate mitochondrial function.

Oxygen consumption, proton motive force (PMF) and proton leak are measurements of mitochondrial respiration, or how well mitochondria are able to convert ...

Biochemistry

Analytical Determination of Mitochondrial Function of Excised Solid Tumor Homogenates

Mitochondria are essential to the onset and progression of cancer through energy production, reactive oxygen species regulation, and macromolecule synthesis. Genetic and functional adaptations of mitochondria to the tumor environment drive proliferative and metastatic potential. The advent of DNA and RNA sequencing removed critical barriers to the evaluation of genetic mediators of tumorigenesis. However, to date, methodological approaches to evaluate tumor mitochondrial function remain elusive and require technical proficiency limiting the feasibility, ultimately diminishing diagnostic and prognostic value in both experimental and clinical settings. Here, we&#160;outline a simple and rapid method to quantify rates of oxidative phosphorylation (OXPHOS) and electron transfer (ET) capacity in freshly excised solid tumor homogenates using high-resolution respirometry. The protocol can be reproducibly applied across species and tumor types as well as adapted to evaluate a diversity of mitochondrial ET pathways. Using this protocol, we demonstrate that mice bearing a luminal B mammary cancer exhibit defective nicotinamide adenine dinucleotide-linked respiration and reliance on succinate to generate adenosine triphosphate via OXPHOS.

We developed a practical protocol and analytical approach to evaluate mitochondrial oxidative phosphorylation and electron transfer capacity in fresh tumor homogenates. This protocol can be easily adapted to survey various mitochondrial functions that contribute to cancer initiation, progression, and treatment response.

Mitochondria are essential to the onset and progression of cancer through energy production, reactive oxygen species regulation, and macromolecule ...

Cancer Research

Preparation and Respirometric Assessment of Mitochondria Isolated from Skeletal Muscle Tissue Obtained by Percutaneous Needle Biopsy

Respirometric profiling of isolated mitochondria is commonly used to investigate electron transport chain function. We describe a method for obtaining samples of human Vastus lateralis, isolating mitochondria from minimal amounts of skeletal muscle tissue, and plate based respirometric profiling using an extracellular flux (XF) analyzer. Comparison of respirometric profiles obtained using 1.0, 2.5 and 5.0 &#956;g of mitochondria indicate that 1.0 &#956;g is sufficient to measure respiration and that 5.0 &#956;g provides most consistent results based on comparison of standard errors. Western blot analysis of isolated mitochondria for mitochondrial marker COX IV and non-mitochondrial tissue marker GAPDH indicate that there is limited non-mitochondrial contamination using this protocol. The ability to study mitochondrial respirometry in as little as 20 mg of muscle tissue allows users to utilize individual biopsies for multiple study endpoints in clinical research projects.

Methods for biopsy of Vastus lateralis, preparation of purified mitochondria, and respirometric profiling are described. The use of small muscle volume makes this technique suitable for clinical research applications.

Respirometric profiling of isolated mitochondria is commonly used to investigate electron transport chain function. We describe a method for obtaining ...

Medicine

Mitochondrial Functional Assessment in Brush Lizard Liver Mitochondria

Assessment of Mitochondrial Oxygen Consumption Using a Plate Reader-based Fluorescent Assay

Mitochondria serve many important functions, including cellular respiration, ATP production, controlling apoptosis, and acting as a central hub of metabolic pathways. Therefore, experimentally assessing mitochondrial functionality can provide insight into variations among different populations or disease states. Additionally, it is valuable to assess whether isolated mitochondria are healthy enough to proceed with experiments. One characteristic often used to compare mitochondrial function in different samples is the rate of oxygen consumption. Oxygen consumption and subsequent calculation of the respiratory control ratio in either intact cells or mitochondria isolated from tissue can serve all three purposes. Using mitochondria isolated from the livers of brush lizards in conjunction with a phosphorescent probe that is sensitive to the fluctuations in oxygen concentration of a solution, we measured oxygen consumption using a fluorescent plate reader. This method is not only quick and efficient but also can be conducted with a small amount of mitochondria and without the need for specialized equipment. The step-by-step protocol described here increases the accessibility of mitochondrial functional assessment to researchers.

Assessment of oxygen consumption provides integral information about mitochondrial function. Using a phosphorescent probe with a fluorescent plate reader, accurate and reproducible data can be obtained easily without specialized equipment. This assay enables any lab to measure the oxygen consumption of isolated mitochondria and calculate respiratory control ratios.

Mitochondria serve many important functions, including cellular respiration, ATP production, controlling apoptosis, and acting as a central hub of ...

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Here, we present a modification of a previously reported method that allows for the isolation of high quality and purified mitochondria from smaller quantities of mouse skeletal muscle. This procedure results in highly coupled mitochondria that respire with high function during microplate based respirometirc assays.

Dysfunctional skeletal muscle mitochondria play a role in altered metabolism observed with aging, obesity and Type II diabetes. Mitochondrial ...

Biology

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 ...

Development of a Mobile Mitochondrial Physiology Laboratory for Measuring Mitochondrial Energetics in the Field

Mitochondrial energetics is a central theme in animal biochemistry and physiology, with researchers using mitochondrial respiration as a metric to investigate metabolic capability. To obtain the measures of mitochondrial respiration, fresh biological samples must be used, and the entire laboratory procedure must be completed within approximately 2 h. Furthermore, multiple pieces of specialized equipment are required to perform these laboratory assays. This creates a challenge for measuring mitochondrial respiration in the tissues of wild animals living far from physiology laboratories as live tissue cannot be preserved for very long after collection in the field. Moreover, transporting live animals over long distances induces stress, which can alter mitochondrial energetics.
This manuscript introduces the Auburn University (AU) MitoMobile, a mobile mitochondrial physiology laboratory that can be taken into the field and used on-site to measure mitochondrial metabolism in tissues collected from wild animals. The basic features of the mobile laboratory and the step-by-step methods for measuring isolated mitochondrial respiration rates are presented. Additionally, the data presented validate the success of outfitting the mobile mitochondrial physiology laboratory and making mitochondrial respiration measurements. The novelty of the mobile laboratory lies in the ability to drive to the field and perform mitochondrial measurements on the tissues of animals captured on site.

We designed and constructed a mobile laboratory to measure respiration rates in isolated mitochondria of wild animals captured at field locations. Here, we describe the design and outfitting of a mobile mitochondrial laboratory and the associated laboratory protocols.

Mitochondrial energetics is a central theme in animal biochemistry and physiology, with researchers using mitochondrial respiration as a metric to ...

Isolation of Mitochondria from Mouse Skeletal Muscle for Respirometric Assays

Most of the cell&#39;s energy is obtained through the degradation of glucose, fatty acids, and amino acids by different pathways that converge on the mitochondrial oxidative phosphorylation (OXPHOS) system, which is regulated in response to cellular demands. The lipid molecule Coenzyme Q (CoQ) is essential in this process by transferring electrons to complex III in the electron transport chain (ETC) through constant oxidation/reduction cycles. Mitochondria status and, ultimately, cellular health can be assessed by measuring ETC oxygen consumption using respirometric assays. These studies are typically performed in established or primary cell lines that have been cultured for several days. In both cases, the respiration parameters obtained may have deviated from normal physiological conditions in&#160;any given organ or tissue.
Additionally, the intrinsic characteristics of cultured single fibers isolated from skeletal muscle impede this type of analysis. This paper presents an updated and detailed protocol for the analysis of respiration in freshly isolated mitochondria from mouse skeletal muscle. We also provide solutions to potential problems that could arise at any step of the process. The method presented here could be applied to compare oxygen consumption rates in diverse transgenic mouse models and study the mitochondrial response to drug treatments or other factors such as aging or sex. This is a feasible method to respond to crucial questions about mitochondrial bioenergetics metabolism and regulation.

Here, we describe a detailed method for mitochondria isolation from mouse skeletal muscle and the subsequent analysis of respiration by Oxygen Consumption Rate (OCR) using microplate-based respirometric assays. This pipeline can be applied to study the effects of multiple environmental or genetic interventions on mitochondrial metabolism.

Most of the cell&#39;s energy is obtained through the degradation of glucose, fatty acids, and amino acids by different pathways that converge on the ...

Real-Time Analysis of Bioenergetics in Primary Human Retinal Pigment Epithelial Cells Using High-Resolution Respirometry

Metabolic dysfunction of retinal pigment epithelial cells (RPE) is a key pathogenic driver of retinal diseases such as age-related macular degeneration (AMD) and proliferative vitreoretinopathy (PVR). Since RPE are highly metabolically-active cells, alterations in their metabolic status reflect changes in their health and function. High-resolution respirometry allows for real-time kinetic analysis of the two major bioenergetic pathways, glycolysis and mitochondrial oxidative phosphorylation (OXPHOS), through quantification of the extracellular acidification rate (ECAR) and oxygen consumption rate (OCR), respectively. The following is an optimized protocol for conducting high-resolution respirometry on primary human retinal pigment epithelial cells (H-RPE). This protocol provides a detailed description of the steps involved in producing bioenergetic profiles of RPE to define their basal and maximal OXPHOS and glycolytic capacities. Exposing H-RPE to different drug injections targeting the mitochondrial and glycolytic machinery results in defined bioenergetic profiles, from which key metabolic parameters can be calculated. This protocol highlights the enhanced response of BAM15 as an uncoupling agent compared to carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) to induce the maximal respiration capacity in RPE. This protocol can be utilized to study the bioenergetic status of RPE under different disease conditions and test the efficacy of novel drugs in restoring the basal metabolic status of RPE.

The metabolic status of human retinal pigment epithelial cells (H-RPE) reflects their health and function. Presented here is an optimized protocol for examining the real-time metabolic flux of H-RPE using high-resolution respirometry.

Metabolic dysfunction of retinal pigment epithelial cells (RPE) is a key pathogenic driver of retinal diseases such as age-related macular degeneration ...

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 ...

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

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Preparation and Respirometric Assessment of Mitochondria Isolated from Skeletal Muscle Tissue Obtained by Percutaneous Needle Biopsy

Assessment of Mitochondrial Oxygen Consumption Using a Plate Reader-based Fluorescent Assay

Isolation of Mitochondria from Minimal Quantities of Mouse Skeletal Muscle for High Throughput Microplate Respiratory Measurements

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

Development of a Mobile Mitochondrial Physiology Laboratory for Measuring Mitochondrial Energetics in the Field

Isolation of Mitochondria from Mouse Skeletal Muscle for Respirometric Assays

Real-Time Analysis of Bioenergetics in Primary Human Retinal Pigment Epithelial Cells Using High-Resolution Respirometry

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