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

This method can help answer key questions such as how energy metabolism in retinal tissue differs between biologic samples, or after exposure to pharmacologic agents. The main advantage of this technique is that it utilizes explanted organotypic retinal tissue to measure rates of both glycolysis, and oxidated phosphorylation and allows assessments of real time interventions in vitro. To calibrate the instrumentation for an extracellular flux assay first add one milliliter of calibration solution to each well of the 24 well sensor cartridge, and incubate at 37 degrees Celsius in a Co2 free incubator overnight.

Load additives for the mitochondrial stress protocol, or the glycolysis protocol into each injector port on the sensor cartridge, adjusting for volume changes in the well. Assuming an initial well volume of 450 microliters, a typical assay would include 45 microliters of an additive into the first injector port, 49.5 microliters in to the second, 54.5 microliters into the third, and 60 microliters in to the last. During the first several experiments, load base medium into port A to gauge how much tissue movement artifact occurs after a port injection.

Allow the loaded sensor plate to incubate at 37 degrees Celsius in a non Co2 incubator for at least 60 minutes. Begin isolation of fresh mouse retinal tissue with a freshly euthanized mouse. Use a pair of medium sized curved forceps to grasp the posterior globe at the optic nerve, and gently apply forward pressure to proptose the eye.

With a clean razor blade, create a limbus to limbus incision across the cornea using a single, deliberate pass of the blade. Using fine angled McPherson style forceps, pinch the posterior globe to express the lens and anterior hyloid membranes out of the corneal incision. Repeat the pinching maneuver with forceps to express the neural retina.

Then, use the forceps like a spoon to lift the retina away from the corneal incision. Place the retina directly into the warm medium in a three centimeter dish, or a six well tissue culture cluster plate. Next, use two pairs of fine angled McPherson’s style forceps to gently dissect remaining vitreous from the retina.

This is is best done by grasping the vitreous at the the periphery of the retinal cup, and pulling towards the center, with a final disinsertion of the vitreous body from the center as a whole. Then, remove any residual retinal pigment epithelium from the photo receptor surface of the retina. Cut a P1000 pipette tip with a razor blade to create an approximate four millimeter opening to prevent undue trauma to the delicate retinal tissue during transfer maneuvers.

Then, use the cut pipette tip to transfer the isolated retinal tissue into fresh medium. Finally, use a one millimeter biopsy punch equipped with a plunger, to cut punches of retina around the optic nerve. Set the retinal punches aside in clean medium, kept on a 37 degree Celsius block, or heating pad.

Once the desired number of punches have been collected, use cute P1000 tip to aspirate an individual punch in 450 microliters of medium, and transfer it to a 24 well eyelet capture micro plate on a 37 degree Celsius heat pad or heat block. Use fine, straight forceps to gently manipulate punches into the center of the micro plate. Keep punches oriented in the same direction.

For example, with the ganglion cell side upwards. For each experiment set aside three to four blank wells with 450 microliters of base medium to serve as negative controls. While avoiding air bubbles or excessive movement, gently position the eyelet capture mesh inserts into each well, and secure the inserts with a metal plunger or with a cut P1000 pipette tip.

Although the micro chamber has adequate depth to accommodate typical mouse retina, occasionally machine defects in the mesh inserts make cause tissues to become overly compressed during screen insertion. If this happens, simply make a note of this crushed tissue, and exclude that sample from the final analysis. Incubate the tissue plate in a 37 degree Celsius carbon dioxide free incubator for at least 60 minutes.

Program the extra cellular flux analyzer using mix, weight, measure, and repeat commands. As an example, a typical experiment with retinal tissue may include the following. Mix two minutes, wait two minutes, and measure five minutes.

Repeat the experiment with these three steps between five to eight times for the base line recording, and after injection of each compound being tested. Press the program start button on the extracellular flux analyzer, and follow the instructions on the screen to insert the sensor cartridge for calibration. At the end of the calibration, follow instructions on the screen to replace the calibrant plate with the plate containing the retinal samples, then allow the program to run as programmed.

At the completion of the run, follow instructions on the screen to eject the tissue plate. View the results of the run and store the data file. With a bent 20 gauge needle and forceps, remove all mesh inserts from the wells, leaving the retinal punch behind.

Carefully aspirate the medium from the tissue, and wash the tissue twice with 0.5 milliliters of cold phosphate buffered saline. After aspirating the second PBS wash, add 100 microliters of lysis buffer to each wash, and pipette up and down to homogenize tissue. Finally, quantitate input levels based on total double stranded DNA content by diluting lysed samples one to one, with 100 microliters of Tris-EDTA buffer, and transferring 100 microliters of the diluted sample to a 96 well plate.

Then, add 100 microliters of detection buffer to each well and mix for two to five minutes. Lastly, quantitate florescence after excitation at 480 nanometers and emission at 520 nanometers by comparing to a standard curve. Normalize the raw extracellular flux tracing to DNA content within the well.

At base line, in the presence of five milli muller glucose and in the presence of 10 milli muller pyruvate, there is no significant difference in rates of acid eflux, measured by the extracellular acidification rate. Between retinal tissues from wile type animals, or knock out mice that lack the machinery to close cyclic nucleotide gated ion channels in response to light stimuli. Similar patterns are seen after addition of 20 milli muller glucose and the glycolytic inhibitor 2DG.

These data may be mathematically transformed to represent fractional changes from baseline, a format which may allow for better comparison between different experimental interventions. Absolute oxygen consumption rate at baseline also is equivalent between control and mutant retinal tissue. The addition of 20 milli muller glucose increases mitochondrial respiration, but no change in observed between groups in terms of absolute quantification or in terms of change from baseline.

The addition of one milli muller FCCP, an uncoupling agent, to demonstrate maximal mitochondrial respiratory rates does not significantly increase OCR above the level seen with high glucose in control and mutant retinal tissue. However, signals from both mice drastically drop after the addition of RAA cocktail. In conclusion, we demonstrate a technique for quantitating rates of oxidated phosphorylation and glycolysis in organotypic explants of mouse retina.

With efficient isolation of retinal tissue, the technique yields reliable and reproudcable measurements. Real time effects of exposures, such as uncoupling agents, can be assessed. In addition to this procedure, other methods, like quantitative RTPCR and western blotting, can be performed on the surplus tissue isolated in the dissection step to obtain parallel information relevant to studies of retinal metabolism.