Calibrating and Using a Polarographic System

Before you can conduct experiments and write the research paper (recommendations follow this page), you have some learning to do. Is it not difficult to use a polarographic system to study mitochondria, but several essential practices must be learned if you are to be successful. A training session on polarography will precede the actual study on isolated mitochondria.

The voltage output of a Yellow Springs Instruments biological oxygen monitor is proportional to the concentration of oxygen measured by the system. Thus we need only calibrate to the end points, namely saturation of the chamber with oxygen and total depletion of oxygen. With depletion of oxygen the monitor puts out zero volts, so we need only set the zero position of the recorder with the input from the monitor grounded. To get the upper point we equilibrate the chamber medium with room air, set the oxygen monitor to read 100% saturation, then set the linear recorder to read 100% full scale. Thereafter, as oxygen is consumed the amount consumed is read as a percentage of full saturation. From a chart record we can determine exactly what molar amount of oxygen was consumed, knowing the temperature and chamber volume. Our chambers hold about 2 ml, but volume varies depending on how the probe is inserted, and of course with the height to which the chamber is filled.

A carefully conducted experiment on isolated mitochondria using a polarographic system can yield a great deal of information. We have developed a methodology for calibration and use of our polarographic systems so as to obtain the most information with a minimum of problems. These methods were developed for the polarographic system described on the previous page.

Calibration

Prior to conducting an experiment we clean out the chamber with several rinses using a squirt bottle and plastic transfer pipet.

When we conduct experiments a glass stopper with capillary bore is inserted into the top of the chamber so that respiration medium just enters the bore. No air bubbles should be trapped in the chamber, nor should there be a vortex when stirring. The chamber must be filled correctly and the stopper inserted gently, otherwise an unstable record will result. We use an automatic pipettor set to a half ml or so to nearly fill the chamber, recording total volume of respiration medium. We then "sneak up" on the final volume by adding volumes of 100-200 µl until the fluid just enters the bottom of the glass stopper when inserted. The stopper must be inserted gently. Forcing the stopper may push out the Clark electrode, causing the chamber to leak.

The filling volume will change when electrodes are changed.  Once the chamber is filled and volume recorded we leave out the glass stopper, while running the stirrer to allow the respiration medium to equilibrate with room air. To facilitate rapid equilibration we keep a working quantity of respiration medium at room temperature. The Clark electrode itself consumes some oxygen, so to obtain 100% saturation with room air it is essential that the medium be stirred and the vortex exposed to room air. The chamber is ready to be sealed when the reading on the oxygen monitor is stable. There may be some drift resulting from temperature variation. If drift is not tolerable, then the water jacket should be used.

Adding a reagent dissolved in ethanol raises the total oxygen in the chamber. For studies using succinate as substrate an investigator may include rotenone in the respiration medium to block the NADH pathway. Bacause rotenone is dissolved in ethanol one may wish to include it in the respiration medium during the equilibration instead of adding it later.

After equilibration and with the chamber still open, we set the oxygen monitor to AIR mode and calibrate to 100% (+/- 1%). The value 100% represents the amount of oxygen per unit volume (oxygen concentration) when the chamber is completely equilibrated (saturated) at the current temperature and pressure.  The percentage will drop as oxygen is consumed.

The zero suppress button on a chart recorder grounds the input signal so that the recorder receives zero volts.  With the input grounded we set the record to physically read zero, then release the zero suppress for recording. Our oxygen monitors deliver a 1 volt signal at 100% saturation, therefore we set the recorder to read 1 volt full scale.

Some investigators prefer to set 100% saturation at 95% of full scale, so that the recording doesn't "peg" should oxygen content drift up. The calibrated setting must be defeated in that case by depressing the "variable" button so that pen position can be set manually. Once the recorder is calibrated the system is ready for an experiment.

Conducting an experiment

For training/practice sessions we use previously frozen liver mitochondria suspensions. Thawed mitochondria cannot produce state III respiration, but they conduct electron transport. We conduct a brief experiment in which we initiate NADH-supported respiration then block electron transport using rotenone. We then re-initiate respiration by adding succinate then block the succinate pathway using antimycin. We finish with ascrobate/TMPD followed by KCN.

A paper speed to 0.2 mm/sec gives us measurable slopes. We start the chart running just prior to adding mitochondria (or other type suspension) so that all of the events can be observed. Twenty to forty microliters of mitochondria suspension usually give us good results. When we use smaller volumes we may fail to see a response because our chambers do not seal perfectly, and with low oxygen consumption rates oxygen can re-enter the chamber as fast as it is consumed. Mitochondria suspensions are quite viscous. We add material using an automatic pipettor. Some investigators prefer to use plunger type pipettors with capillary tubes rather than air displacement, to obtain precise volumes for quantitative work. After adding mitochondria we triturate using a 1 ml automatic pipettor for a few seconds to disperse any particles. Triuration should not be necessary if the suspension has been homogenized sufficiently. Care must be taken not to introduce any air into the chamber.

Immediately after trituration we place the glass stopper gently into the opening, allowing all air to escape through the opening and so that medium enters the bore, stabilizing the vortex. The record should show a rapid drop in oxygen content of 5% or so, followed by a steady state of little or no oxygen consumption. Mitochondria are stored as a very concentrated, oxygen poor suspension.  Upon addition to an oxygen rich medium they take up oxygen into the matrix, taking it out of solution. This uptake accounts for the initial drop in oxygen content. The slow linear decline in oxygen content that follows is actual oxygen consumption, supported by fatty acids remaining in the suspension.

We add substrates and reagents through the capillary bore in the stopper using Hamilton syringes. We rinse our syringes thoroughly using deionized water before drawing up reagent. However, to minimize the risk of poisoning our substrates we use one syringe exclusively for substrates (and later for ADP) and the other for poisons. We identify them using colored tape.

To add a reagent we draw up an appropriate volume, usually 20 µl of substrate or 10 µl of a poison that is dissolved in ethanol. We wipe the tip before inserting it into the stopper, and before ejecting material make sure that the tip of the needle is below the bottom of the stopper. We eject material into the chamber fairly assertedly, so that the material is mixed immediately. After adding reagent we wait until we obtain a measurable steady state, then go to the next reagent. It should not take more than a few seconds to see a response.

For the electron transport study we add, in order, 20 µl 0.1M NADH, 10 µl 10 mM rotenone, 20 µl 0.5M succinate, 10 µl 10 mM antimycin, 20 µl ascorbate and 10 µl 30 mM TMPD (added together) and 20 µl 0.5M KCN. Rotenone and antimycin are dissolved in ethanol. We adjust the pH of the other reagents to 7 (except for KCN). The substrates used to support electron transport are acids. Adding substrates that are not pH neutral will uncouple mitochondria.

After any experiment it is necessary to thoroughly rinse out both the chamber and the stopper. When a chamber with electrode will not be used for some time it should be filled with deionized water to prevent the electrode membrane from drying out.