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

Podsumowanie

This technique describes real time recording of oxygen consumption and extracellular acidification rates in explanted mouse retinal tissues using an extracellular flux analyzer.

Streszczenie

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.

Wprowadzenie

The retina is among the most energy-demanding tissues in the central nervous system¹. Like most tissues, it generates adenosine triphosphate (ATP) via glycolysis in the cytosol or via oxidative phosphorylation in mitochondria. The energetic advantage of oxidative phosphorylation over glycolysis to produce ATP from one molecule of glucose is clear: 36 molecules of ATP generated from the former vs. 2 molecules of ATP generated from the latter. Accordingly, retinal neurons primarily depend on mitochondrial respiration for energy supply and this is reflected by their high density of mitochondria². Yet, the retina also relies heavily on glycolytic machinery even when oxygen is abundant. This process of aerobic glycolysis was originally described in cancer cells by Otto Warburg³, who once noted that the retina was the only post-mitotic tissue capable of this form of metabolism⁴. Since those initial observations, many post-mitotic tissues have been described to engage in varying degrees of glycolysis in addition to oxidative phosphorylation to meet their ATP demands.

Phototransduction, visual pigment recycling, biosynthesis of photoreceptor outer segments, and synaptic activity are all energy demanding processes in photoreceptors, the predominant neuronal subclass in the retina. But the need to actively transport ions against their electrical and concentration gradients is by far the most energetically consuming process in neurons¹. Photoreceptors are peculiar neurons in the sense that they are depolarized in the absence of stimulation (i.e., in the dark), whereas a light stimulus triggers channel closure and subsequent hyperpolarization. Therefore, in the dark, the retina consumes large quantities of ATP to maintain its depolarization or "dark current" as it is commonly called. From an adaptive standpoint, a major challenge in supplying these vast quantities of ATP is the need for organisms to maintain visual clarity through the optical axis. The inverted retinal architecture seen in modern creatures is the dominant solution, as it keeps the dense capillary network supplying photoreceptors away from the path of light. But this marvel of natural bioengineering places the retina at a precipice in terms of metabolic reserve. Even small insults to retina can potentially disrupt the delicate balance of energy supply to demand, and visual dysfunction or frank blindness may ensue quickly.

Given the unique energetic demands of the neural retina, coupled with its tight restriction of vascular supply, accurate measurement of ATP consumption in the retina and its changes during disease could have profound implications in understanding and treating blinding conditions such as retinitis pigmentosa and diabetic retinopathy. Traditionally, these measurements require costly, custom-designed equipment with most studies emerging from a handful of laboratories entirely dedicated to measurements of metabolic activity^2,5,6,7,8. Techniques include individual assays for specific metabolites, tracer studies using radio-labeled precursors, oxygen consumption recording using Clark electrodes, and metabolomic profiling⁹.

With advances in high-throughput technology and increased availability of commercial devices, techniques to record retinal metabolism are increasingly accessible and affordable. The method described here measures both oxidative phosphorylation and glycolysis in retina using explanted tissue and a commercially-available extracellular flux analyzer^9,10,11,12. This analyzer separately records oxygen consumption rate (OCR) and the extracellular acidification rate (ECAR), serving as indirect indicators of oxidative phosphorylation and glycolysis, respectively¹³. These measurements are done by a probe submersed within a microchamber created over the tissue of interest. This adaptation of previously published methods uses a capture plate originally designed for pancreatic Islets of Langerhans to record metabolic activity in small, circular sections of mouse retina. Multiple pharmacologic exposures can be delivered to the tissue during the course of a single recording because the system contains 4 injection ports for each sample well. Using this system with separate protocols optimized for ECAR and OCR recordings, the responses of wild-type retinas can be compared to retinas lacking transducin (Gnat1^-/-), a cause of congenital stationary night blindness in humans¹⁴.

Protokół

Protocols followed the Association for Research in Vision and Ophthalmology Statement for the Use of Animals and were approved by Washington University.

1. Animal Preparation

1. Keep animals in standard housing with a 12 hours dark to 12 hours light cycle. Begin experiments at standardized times to avoid circadian effects, typically in the morning shortly after lights are turned on.

2. Solution Preparation

1. Prepare base media by dissolving 8.4 mg of powdered media in double-distilled H₂O (ddH₂O) and adjust pH to 7.4 with either HCl or NaOH, to a final volume of 1 L. Filter sterilize this solution with a 0.22 µm tissue culture filter.
   1. Add 1 M glucose and 100 mM sodium pyruvate to the media to achieve final concentrations of 5 mM glucose and 1 mM pyruvate.
   2. Place a 50 mL aliquot of the base media in a 37 °C water bath.
2. Prepare lysis solution, for tissue quantitation at the end of the flux analysis, by adding tris base to 10 mM, polyethylene glycol tert-octylphenyl ether to 0.2% (v/v), and EDTA to 1 mM, all in ddH₂O. Mix until all components completely dissolved and adjust pH to 8.0.
3. For the mitochondrial stress protocol, prepare an 11 µM stock of oligomycin, an ATP Synthase inhibitor, dissolved in base media to target a final concentration of 1 µM in the well. Prepare an 11 µM stock of carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone (FCCP), an uncoupling agent, dissolved in base media to target a final concentration of 1 µM in the well. Prepare a cocktail of the electron transport chain inhibitors, rotenone and antimycin A (RAA) by adding rotenone to 11 µM and antimycin A to 22 µM, dissolved in base media to target a final concentration of 1 µM rotenone and 2 µM antimycin A in the well.
4. For the glycolysis protocol, prepare a 220 mM stock of glucose dissolved in base media, to target a final concentration of 20 mM in the well. Also prepare a 1.1 M stock of 2-deoxyglucose (2-DG), a competitive inhibitor of glucose and glycolytic antagonist, by dissolving it in base media. This will target a final concentration of 100 mM 2-DG in the well.

3. Instrument Calibration

1. To calibrate the instrumentation for an extracellular flux assay, add 1 mL of calibration solution to each well of the sensor cartridge (24 wells total) and incubate at 37 °C in a CO₂-free incubator overnight (or at least 8 h).
2. Load additives for the mitochondrial stress protocol or the glycolysis protocol into each injector port, A through D, on the sensor cartridge, adjusting for volume changes in the well.
   Note: As an example, a typical assay would include 45 µL of an additive into the first injector port, 49.5 µL into the second, 54.5 µL into the third, and 60 µL into the last, assuming an initial well volume of 450 µL.
3. During the first several experiments, load base media into port A to gauge how much tissue movement/artefact occurs after a port injection.
4. Allow the loaded sensor plate to incubate at 37 °C in a non-CO₂ incubator for at least 60 min.

4. Isolation of Fresh Mouse Retinal Tissue

Note: This step is adapted from the technique of Dr. Barry Winkler¹⁵.

1. After administering deep anesthesia using a standard ketamine/xylazine cocktail, euthanize mice by cervical dislocation.
2. 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 (Figure 1A).
   1. With a clean razor blade, create a limbus to limbus incision across the cornea using a single, deliberate pass of the blade.
   2. Using fine, angled McPherson-style forceps, pinch the posterior globe to express the lens and anterior hyaloid membranes out of the corneal incision. Discard these tissues.
   3. Repeat the pinching maneuver with forceps to express the neural retina.
   4. Using the forceps like a spoon to lift the retina, rather than to grasp the tissue, transfer the retina away from the corneal incision and place directly into warm media in a 3-cm dish or a 6-well tissue culture cluster plate.
3. Use two pairs of fine, angled McPherson-style forceps to gently dissect remaining vitreous from the retina. This is best done by grasping the vitreous at 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. Using the forceps, remove any residual retinal pigment epithelium from the photoreceptor surface of the retina.
   1. Cut a P1000 pipet tip with a razor blade to create a ~4 mm opening to prevent undue trauma to the delicate retinal tissue during transfer maneuvers.
   2. Transfer the isolated retinal tissue into fresh media using a cut P1000 pipet tip.
   3. Cut 1 mm punches of retina around the optic nerve with a 1 mm biopsy punch equipped with a plunger to dislodge the tissue in case it becomes lodged into the bore (Figure 1B). Set the retinal punches aside in clean media kept on a 37 °C block or heating pad.

5. Assay Protocol

1. Transfer individual punches to a 24-well islet capture microplate using a cut P1000 tip, aspirating 450 µL of media along with the tissue.
2. Use fine, straight forceps to gently manipulate punches into the center of the microplate. Keep punches orientated in the same direction (e.g., ganglion cell side upwards).
   1. Rest the capture plate on a heating pad or heating block set to 37 °C as these steps are repeated for the remaining samples.
      Note: For each experiment, set aside 3-4 blank wells with 450 µL of base media to serve as negative controls. In the remaining 20-21 wells, test each biologic replicate in triplicate. Therefore, each 24-well plate will allow for recording from 6-7 different animals or treatment conditions.
3. Avoiding air bubbles, gently position the islet capture mesh inserts into each well using forceps and secure the inserts with a metal plunger, or with a cut P1000 pipet tip (Figure 1C).
   Note: Avoid excessive tissue movement during this step to maintain retinal punch position within the center of the sample microchamber. Air bubbles severely distort OCR readings. Although the microchamber has adequate depth to accommodate typical mouse retina (Figure 1C), occasionally machining-defects in the mesh inserts may cause tissues to become overly compressed during screen insertion. If this happens, simply make a note of the crushed tissue and exclude that sample from the final analysis.
4. Incubate the tissue plate in a 37 °C CO₂-free incubator for at least 60 min.
5. Program the extracellular flux analyzer using Mix, Wait, Measure, and Repeat commands. As an example, a typical experiment with retinal tissue may include the following: Mix 2 minutes, Wait 2 minutes, and Measure 5 minutes. Repeat the experiment with these three steps between 5-8 times (cycles) for a baseline recording, and after injection of each compound being tested during the run.
6. Press the program START button on the extracellular flux analyzer and follow the instructions on the screen to insert the sensor cartridge for calibration.
7. At the end of the calibration, follow instructions on the screen to replace the calibrant plate with the plate containing the retinal samples.
8. 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.
9. With a bent 20 gauge needle and forceps, remove all mesh inserts from the wells, leaving the retinal punch behind.
10. Carefully aspirate the media from the tissue and wash the tissue twice with 0.5 mL of cold phosphate-buffered saline (PBS). Aspirate the PBS after washing.
11. Add 100 µL of lysis buffer to each well and pipet up and down to homogenize tissue.
12. Quantitate input levels based on total double-stranded DNA (dsDNA) or total protein content.
    Note: The following is based on a commercially available dsDNA assay.
13. Dilute lysed samples 1:1 by adding 100 µL of Tris-EDTA (TE) buffer and transfer 100 µL of the diluted sample to a 96 well plate.
    1. Add 100 µL of detection buffer into each well. After mixing for 2-5 minutes, quantitate fluorescence after excitation at 480 nm and emission at 520 nm, comparing to a standard curve.
    2. Normalize the raw extracellular flux tracing to DNA content within the well.
       Note: In the results presented here, samples are normalized to 50 ng dsDNA - a typical amount in a 1 mm retinal punch. Therefore, absolute values presented here may be compared to studies presenting data "per retinal punch". Normalize tracings to an internal standard, such as the baseline run at a glucose concentration of 5 mM.

Wyniki

Using the described techniques (summarized in Figure 1), retinal explants from 8 week-old wild type (WT) mice were compared to age- and background-matched transducin null mice (Gnat1^-/-). Because Gnat1^-/- animals lack the machinery to close cyclic-nucleotide gated ion channels in response to light stimuli, their rod photoreceptors remain depolarized even in light¹⁴. The subsequent n...

Dyskusje

OCR and ECAR are readily measured in explanted retinal tissue using a bioanalyzer using the described techniques. This method departs from those of other groups in several critical steps. Retinal tissues are isolated through a large corneal incision without enucleating the globe, as originally described by Winkler¹⁵. This method of retinal isolation allows for a rapid transfer from the living eye into the tissue capture plate (often within 5 minutes). Tissues are kept at 37 °C throughout the ...

Materiały

Name	Company	Catalog Number	Comments
Seahorse XF24 Extracellular Flux Analyzer	Agilent, Santa Clara, CA
Seahorse XF24 Islet Capture FluxPak (includes: Islet Capture Microplate, Sensor Cartridge and Calibrant Solution)	Agilent, Santa Clara, CA	101174-100	Includes islet capture microplate, sensor cartridge and calibrant solution
RPMI 1640 Media (Powdered medium)	Millipore-Sigma	R1383	RPMI 1640 Media with L-Glutamine and without glucose or sodium bicarbonate
D-Glucose	Millipore-Sigma	G8270	1M D-Glucose filtered, for media preparation
Sodium pyruvate	Corning	25000CI	100 mM sodium pyruvate
Antimycin-A	Millipore-Sigma	A8674	Mitochondrial stress protocol component
FCCP	Millipore-Sigma	C2920	Mitochondrial stress protocol component
Rotenone	Millipore-Sigma	R8875	Mitochondrial stress protocol component
2-deoxyglucose	Millipore-Sigma	D6134	Glycolysis protocol component
1 mm skin biopsy punches with plunger	Integra-Miltex	33-31AA-P/25	Explanting retinal tissue tool
Dumont Mini-Forceps Straight	Fine Science Tools	11200-10	Explanting retinal tissue tool
Dumont Medical #5/45 Forceps- Angled 45 degrees	Fine Science Tools	11253-25	Explanting retinal tissue tool
Dumont #7 Forceps - Curved	Fine Science Tools	11271-30	Explanting retinal tissue tool
Quant-iT Picogreen dsDNA Assay Kit	Fisher Scientific	P7589	Loading normalization assay
Trizma base (Tris base)	Millipore-Sigma	T6066	Component of lysis buffer
Triton X-100 (polyethylene glycol tert-octylphenyl ether)	Millipore-Sigma	X100	Component of lysis buffer
0.5M EDTA pH 8.0	Ambion	AM9262	Component of lysis buffer
C57BL/6J mice	Jackson Laboratories	Strain 000664	Animals
Gnat1-/- and background-matched Gnat1+/+	Vladimir Kefalov, PhD; Washington University School of Medicine	Animals