Assessing oxidative phosphorylation using high-resolution respirometers has become an integral part of the functional analysis of mitochondria and cellular energy metabolism. Here, we present protocols for the analysis of cellular energy metabolism using chamber and microplate-based high-resolution respirometers and discuss the key benefits of each device.
Mitochondria provide cellular energy through oxidative phosphorylation contributing to almost all processes within the cell. Consequently, mitochondrial dysfunctions are associated with a wide spectrum of diseases. The main advantage of this technique is the realtime measurement of mitochondrial bioenergetics by oxygen consumption, and thus, an accurate assessment of total cellular energy metabolism.
High-resolution respirometry allows direct assessment of what in the oxidative phosphorylation system is failing, potentially providing diagnostic clues in primary mitochondrial diseases and secondary mitochondrial dysfunctions associated with many disorders. Demonstrating the procedure will be Ryan Awadhpersad, PhD student in my lab. To begin, perform the oxygen calibration by running the respirometers in 2.1 milliliters of mitochondrial respiration medium at 37 degrees Celsius for more than 45 minutes and proceed if baseline variation is within four picomoles per second.
Culture HEK293 cells in high-glucose DMEM supplemented with 10%heat-activated FBS, GlutaMAX, non-essential amino acids, sodium pyruvate, and uridine in an incubator at 37 degrees Celsius at 5%carbon dioxide. On the day of the experiment, collect and count the cells and resuspend 2.5 million cells in 2.5 milliliters respiration medium. To assess routine respiration, add 2.3 milliliters of warm respiration medium cell suspension to the chamber.
Run the chambers at 37 degrees Celsius and a stirring speed of 700 RPM. Wait for at least three minutes to allow media to de-gas, then close the chambers by twisting the stopper in a rotating motion. Next, aspirate the excess liquid on top of the stopper.
After 10 minutes, obtain a stable oxygen flux signal to record routine or state I respiration. To perform OXPHOS analysis in intact cells, add two microliters of 0.01-millimolar oligomycin for a final concentration of 10-nanomolar. Titrate FCCP from two-millimolar stock by adding 0.6 microliters at 0.2-microliter steps until no increase in respiration is observed and respiration is maximally uncoupled.
Next, add one microliter of one-millimolar rotenone for a final concentration of 0.5-micromolar. Then, add two microliters of one milligram per milliliter antimycin stock for a final concentration of one microgram per milliliter. Reoxygenate the chamber to the same oxygen level, approximately 150-micromolar, by lifting the plunger.
Afterward, add five microliters of 0.8-molar ascorbate for a final concentration of two-millimolar. Then, immediately add five microliters of 0.2-molar TMPD for a final concentration of 0.5-millimolar and evaluate complex IV activity. As the peak oxygen flux is reached with TMPD, add five microliters or four-molar azide for a final concentration of 10-millimolar.
Then, continue the run for at least five minutes to assay auto-oxidation of TMPD for complex IV base level calculation. After the run, collect one milliliter of sample suspension from each chamber and centrifuge at 1, 000 G for permeabilized cells or at 20, 000 G for tissue lysate. Then, discard the supernatant and freeze the pellet at minus 80 degrees Celsius for further downstream analysis.
First, seed the cells according to the growth rates of individual cell lines. Dilute oligomycin, FCCP, rotenone, and antimycin in three microliters of assay medium to a final concentration of 1.5-micromolar, 1.125-micromolar, and one-micromolar, respectively. Subsequently, fill them into separate ports.
Observe the injection ports to ensure an even loading of the samples. Turn on the microplate-based system and computer and equilibrate them at 37 degrees Celsius for at least three hours. On the day of the microplate-based assay, verify the confluency of the cell culture plate, the morphology of cells, and ensure that the background wells are empty.
Then, remove everything except 20 microliters of the culture medium from each well. Then, wash each well with 90 microliters of assay medium and add 100 microliters of assay medium to make a final volume of up to 120 microliters. Next, incubate the plate at 37 degrees Celsius in an incubator without carbon dioxide for 60 minutes.
After retrieving the plate from the incubator, remove the lid and place the microplate in the slot. Click on Continue to start the run. After the run is finished, take the plate out and remove the remaining assay media without disturbing the cells.
Then, freeze the entire plate at minus 80 degrees Celsius. After performing the digitonin permeabilization and respiration experiments, raw oxygen consumption traces of wild-type HEK293 cells and HEK293 cells with a CRISPR-mediated gene knockout resulting in multiple OXPHOS deficiencies were recorded. Overlaid cell input-normalized oxygen consumption traces were obtained.
CRISPR knockout 1 shows impaired respiration and CRISPR knockout 2 shows no respiration compared to wild-type when normalized to cell number. Protein amounts were quantified and respiration values were normalized to protein amount in order to determine absolute values of respiratory states and respective flux control ratios. Extracellular acidification rates were found to increase for OXPHOS deficiency in CRISPR knockout 2, suggesting compensation of mitochondrial oxidative phosphorylation deficiency in HEK293 cells through increased glycolysis.
In addition, the respiration values were determined in the presence of oligomycin, FCCP, and rotenone, and the values obtained were normalized to the protein amount. Protein-normalized oxygen consumption traces of wild-type HEK293 cells and HEK293 cells with CRISPR-mediated mitochondrial translation deficiency causing multiple OXPHOS deficiency was studied. Wet weight tissue-normalized oxygen consumption traces of mouse cerebellum and mouse soleus muscle were obtained.
The soleus muscles show three times higher OXPHOS and respiration capacity than the cerebellum. With chamber-based respirometry, it is important to work fast, as mitochondrial function will decline the moment you collect your samples, and with plate-based respirometry, it is crucial to acquire an optimal seeding density to minimize variability. For further analysis, protein quantification or immuno-blotting is possible.
This could determine whether an alteration in mitochondrial respiratory function was due to abundance of OXPHOS complexes or mitochondrial amount. Already a century ago, cancer cells were found to perform anaerobic glycolysis in addition to mitochondrial OXPHOS. This underlines the need to assay mitochondrial bioenergetics.
Here we demonstrated two respirometers considered today's gold standard.
