We thank Dr. Alexander Kolesnikov and Dr. Vladimir Kefalov for providing Gnat1-/- mice, for helpful feedback and advice, and for reading the manuscript.
This work was supported by NIH EY025269 (RR), the Diabetes Research Center at Washington University - NIH DK020579 (JRM and RR), a Career Development Award from Research to Prevent Blindness (RR), the Horncrest Foundation (RR), a Career Development Award from JDRF (JRM), NIH DK101392 (CFS), DK020579 (CFS), DK056341 (CFS), and DK114233 (JRM).

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
<ol>
	<li>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.</li>
</ol>
2. Solution Preparation
<ol>
	<li>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 &#181;m tissue culture filter.
	<ol>
		<li>Add 1 M glucose and 100 mM sodium pyruvate to the media to achieve final concentrations of 5 mM glucose and 1 mM pyruvate.</li>
		<li>Place a 50 mL aliquot of the base media in a 37 &#176;C water bath.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>For the mitochondrial stress protocol, prepare an 11 &#181;M stock of oligomycin, an ATP Synthase inhibitor, dissolved in base media to target a final concentration of 1 &#181;M in the well. Prepare an 11 &#181;M stock of carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone (FCCP), an uncoupling agent, dissolved in base media to target a final concentration of 1 &#181;M in the well. Prepare a cocktail of the electron transport chain inhibitors, rotenone and antimycin A (RAA) by adding rotenone to 11 &#181;M and antimycin A to 22 &#181;M, dissolved in base media to target a final concentration of 1 &#181;M rotenone and 2 &#181;M antimycin A in the well.</li>
	<li>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.</li>
</ol>
3. Instrument Calibration
<ol>
	<li>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 &#176;C in a CO2-free incubator overnight (or at least 8 h).</li>
	<li>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 &#181;L of an additive into the first injector port, 49.5 &#181;L into the second, 54.5 &#181;L into the third, and 60 &#181;L into the last, assuming an initial well volume of 450 &#181;L.</li>
	<li>During the first several experiments, load base media into port A to gauge how much tissue movement/artefact occurs after a port injection.</li>
	<li>Allow the loaded sensor plate to incubate at 37 &#176;C in a non-CO2 incubator for at least 60 min.</li>
</ol>
4. Isolation of Fresh Mouse Retinal Tissue
Note: This step is adapted from the technique of Dr. Barry Winkler15.
<ol>
	<li>After administering deep anesthesia using a standard ketamine/xylazine cocktail, euthanize mice by cervical dislocation.</li>
	<li>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).
	<ol>
		<li>With a clean razor blade, create a limbus to limbus incision across the cornea using a single, deliberate pass of the blade.</li>
		<li>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.</li>
		<li>Repeat the pinching maneuver with forceps to express the neural retina.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>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.</li>
		<li>Transfer the isolated retinal tissue into fresh media using a cut P1000 pipet tip.</li>
		<li>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 &#176;C block or heating pad.</li>
	</ol>
	</li>
</ol>
5. Assay Protocol
<ol>
	<li>Transfer individual punches to a 24-well islet capture microplate using a cut P1000 tip, aspirating 450 &#181;L of media along with the tissue.</li>
	<li>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).
	<ol>
		<li>Rest the capture plate on a heating pad or heating block set to 37 &#176;C as these steps are repeated for the remaining samples. 
		Note: For each experiment, set aside 3-4 blank wells with 450 &#181;L 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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Incubate the tissue plate in a 37 &#176;C CO2-free incubator for at least 60 min.</li>
	<li>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.</li>
	<li>Press the program START button on the extracellular flux analyzer and follow the instructions on the screen to insert the sensor cartridge for calibration.</li>
	<li>At the end of the calibration, follow instructions on the screen to replace the calibrant plate with the plate containing the retinal samples.</li>
	<li>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.</li>
	<li>With a bent 20 gauge needle and forceps, remove all mesh inserts from the wells, leaving the retinal punch behind.</li>
	<li>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.</li>
	<li>Add 100 &#181;L of lysis buffer to each well and pipet up and down to homogenize tissue.</li>
	<li>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.</li>
	<li>Dilute lysed samples 1:1 by adding 100 &#181;L of Tris-EDTA (TE) buffer and transfer 100 &#181;L of the diluted sample to a 96 well plate.
	<ol>
		<li>Add 100 &#181;L 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.</li>
		<li>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 &#34;per retinal punch&#34;. Normalize tracings to an internal standard, such as the baseline run at a glucose concentration of 5 mM.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Wong-Riley, M. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Energy+metabolism+of+the+visual+system.">Energy metabolism of the visual system.</a> Eye and brain. 2, 99-116 (2010).</li><li>Ames, A., Li, Y. Y., Heher, E. C., Kimble, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Energy+metabolism+of+rabbit+retina+as+related+to+function:+high+cost+of+Na++transport.">Energy metabolism of rabbit retina as related to function: high cost of Na+ transport.</a> The Journal of neuroscience: the official journal of the Society for Neuroscience. 12 (3), 840-853 (1992).</li><li>Warburg, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=On+the+origin+of+cancer+cells.">On the origin of cancer cells.</a> Science. 123 (3191), 309-314 (1956).</li><li>Wubben, T. J., 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=Photoreceptor+metabolic+reprogramming+provides+survival+advantage+in+acute+stress+while+causing+chronic+degeneration.">Photoreceptor metabolic reprogramming provides survival advantage in acute stress while causing chronic degeneration.</a> Scientific reports. 7 (1), 17863 (2017).</li><li>Du, J., Linton, J. D., Hurley, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Probing+Metabolism+in+the+Intact+Retina+Using+Stable+Isotope+Tracers.">Probing Metabolism in the Intact Retina Using Stable Isotope Tracers.</a> Methods in enzymology. 561, 149-170 (2015).</li><li>Felder, A. E., Wanek, J., Tan, M. R., Blair, N. P., Shahidi, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Method+for+Combined+Retinal+Vascular+and+Tissue+Oxygen+Tension+Imaging.">A Method for Combined Retinal Vascular and Tissue Oxygen Tension Imaging.</a> Scientific reports. 7 (1), 10622 (2017).</li><li>Hurley, J. B., Lindsay, K. J., Du, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glucose,+lactate,+and+shuttling+of+metabolites+in+vertebrate+retinas.">Glucose, lactate, and shuttling of metabolites in vertebrate retinas.</a> Journal of neuroscience research. 93 (7), 1079-1092 (2015).</li><li>Winkler, B. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glycolytic+and+oxidative+metabolism+in+relation+to+retinal+function.">Glycolytic and oxidative metabolism in relation to retinal function.</a> The Journal of general physiology. 77 (6), 667-692 (1981).</li><li>Pelletier, M., Billingham, L. K., Ramaswamy, M., Siegel, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracellular+flux+analysis+to+monitor+glycolytic+rates+and+mitochondrial+oxygen+consumption.">Extracellular flux analysis to monitor glycolytic rates and mitochondrial oxygen consumption.</a> Methods in enzymology. 542, 125-149 (2014).</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><li>Kooragayala, K., 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=Quantification+of+Oxygen+Consumption+in+Retina+Ex+Vivo+Demonstrates+Limited+Reserve+Capacity+of+Photoreceptor+Mitochondria.">Quantification of Oxygen Consumption in Retina Ex Vivo Demonstrates Limited Reserve Capacity of Photoreceptor Mitochondria.</a> Investigative ophthalmology &amp; visual science. 56 (13), 8428-8436 (2015).</li><li>Pearsall, E. 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=PPARalpha+is+essential+for+retinal+lipid+metabolism+and+neuronal+survival.">PPARalpha is essential for retinal lipid metabolism and neuronal survival.</a> BMC biology. 15 (1), 113 (2017).</li><li>Nicholls, D. G., 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=Bioenergetic+profile+experiment+using+C2C12+myoblast+cells.">Bioenergetic profile experiment using C2C12 myoblast cells.</a> Journal of visualized experiments. (46), (2010).</li><li>Lobanova, E. 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=Transducin+translocation+in+rods+is+triggered+by+saturation+of+the+GTPase-activating+complex.">Transducin translocation in rods is triggered by saturation of the GTPase-activating complex.</a> The Journal of neuroscience: the official journal of the Society for Neuroscience. 27 (5), 1151-1160 (2007).</li><li>Winkler, B. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+electroretinogram+of+the+isolated+rat+retina.">The electroretinogram of the isolated rat retina.</a> Vision research. 12 (6), 1183-1198 (1972).</li><li>Jastroch, M., Divakaruni, A. S., Mookerjee, S., Treberg, J. R., Brand, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+proton+and+electron+leaks.">Mitochondrial proton and electron leaks.</a> Essays in biochemistry. 47, 53-67 (2010).</li><li>Divakaruni, A. S., Paradyse, A., Ferrick, D. A., Murphy, A. N., Jastroch, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+and+interpretation+of+microplate-based+oxygen+consumption+and+pH+data.">Analysis and interpretation of microplate-based oxygen consumption and pH data.</a> Methods in enzymology. 547, 309-354 (2014).</li><li>Du, J., 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=Phototransduction+Influences+Metabolic+Flux+and+Nucleotide+Metabolism+in+Mouse+Retina.">Phototransduction Influences Metabolic Flux and Nucleotide Metabolism in Mouse Retina.</a> The Journal of biological chemistry. 291 (9), 4698-4710 (2016).</li></ol>

The authors have nothing to disclose.

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 &#176;C throughout the ...

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 &#34;dark current&#34; 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 activity2,5,6,7,8. 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 analyzer9,10,11,12. 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.

<table border="0" cellpadding="0" cellspacing="0" style="width:777px;" width="778">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:337px;">Seahorse XF24 Extracellular Flux Analyzer</td>
			<td style="width:155px;">Agilent, Santa Clara, CA</td>
			<td style="width:119px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="80">
			<td height="80" style="height:80px;width:337px;">Seahorse XF24 Islet Capture FluxPak (includes: Islet Capture Microplate, Sensor Cartridge and Calibrant Solution)</td>
			<td>Agilent, Santa Clara, CA</td>
			<td>101174-100</td>
			<td style="width:167px;">Includes islet capture microplate, sensor cartridge and calibrant solution</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">RPMI 1640 Media (Powdered medium)</td>
			<td>Millipore-Sigma</td>
			<td>R1383</td>
			<td style="width:167px;">RPMI 1640 Media with L-Glutamine and without glucose or sodium bicarbonate</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">D-Glucose</td>
			<td>Millipore-Sigma</td>
			<td>G8270</td>
			<td style="width:167px;">1M D-Glucose filtered, for media preparation</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:337px;">Sodium pyruvate</td>
			<td>Corning</td>
			<td>25000CI</td>
			<td style="width:167px;">100 mM sodium pyruvate</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:337px;">Antimycin-A</td>
			<td>Millipore-Sigma</td>
			<td>A8674</td>
			<td style="width:167px;">Mitochondrial stress protocol component</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:337px;">FCCP</td>
			<td>Millipore-Sigma</td>
			<td>C2920</td>
			<td style="width:167px;">Mitochondrial stress protocol component</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:337px;">Rotenone</td>
			<td>Millipore-Sigma</td>
			<td>R8875</td>
			<td style="width:167px;">Mitochondrial stress protocol component</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:337px;">2-deoxyglucose</td>
			<td>Millipore-Sigma</td>
			<td>D6134</td>
			<td style="width:167px;">Glycolysis protocol component</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:337px;">1 mm skin biopsy punches with plunger</td>
			<td>Integra-Miltex</td>
			<td>33-31AA-P/25</td>
			<td style="width:167px;">Explanting retinal tissue tool</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:337px;">Dumont Mini-Forceps Straight</td>
			<td>Fine Science Tools</td>
			<td>11200-10</td>
			<td style="width:167px;">Explanting retinal tissue tool</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:337px;">Dumont Medical #5/45 Forceps- Angled 45 degrees</td>
			<td>Fine Science Tools</td>
			<td>11253-25</td>
			<td style="width:167px;">Explanting retinal tissue tool</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:337px;">Dumont #7 Forceps - Curved</td>
			<td>Fine Science Tools</td>
			<td>11271-30</td>
			<td style="width:167px;">Explanting retinal tissue tool</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:337px;">Quant-iT Picogreen dsDNA Assay Kit</td>
			<td>Fisher Scientific</td>
			<td>P7589</td>
			<td style="width:167px;">Loading normalization assay</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Trizma base (Tris base)</td>
			<td>Millipore-Sigma</td>
			<td>T6066</td>
			<td style="width:167px;">Component of lysis buffer</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Triton X-100 (polyethylene glycol tert-octylphenyl ether)</td>
			<td>Millipore-Sigma</td>
			<td>X100</td>
			<td style="width:167px;">Component of lysis buffer</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">0.5M EDTA pH 8.0</td>
			<td>Ambion</td>
			<td>AM9262</td>
			<td style="width:167px;">Component of lysis buffer</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">C57BL/6J mice&#160;</td>
			<td>Jackson Laboratories</td>
			<td>&#160;Strain 000664</td>
			<td style="width:167px;">Animals</td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">Gnat1-/- and background-matched Gnat1+/+&#160;</td>
			<td colspan="2">Vladimir Kefalov, PhD; Washington University School of Medicine</td>
			<td style="width:167px;">Animals</td>
		</tr>
	</tbody>
</table>

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.

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-/-&#160;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...

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

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 ...

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9.4K Views.  Washington University School of Medicine. This technique describes real time recording of oxygen consumption and extracellular acidification rates in explanted mouse retinal tissues using an extracellular flux analyzer. 

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

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called photoreceptors to sense and adapt to light. The red/far-red light-absorbing phytochrome photoreceptors have been studied extensively. Phytochromes exist as a family of proteins with distinct and overlapping functions in all higher plant systems in which they have been studied1. Phytochrome-mediated light responses, which range from seed germination through flowering and senescence, are often localized to specific plant tissues or organs2. Despite the discovery and elucidation of individual and redundant phytochrome functions through mutational analyses, conclusive reports on distinct sites of photoperception and the molecular mechanisms of localized pools of phytochromes that mediate spatial-specific phytochrome responses are limited. We designed experiments based on the hypotheses that specific sites of phytochrome photoperception regulate tissue- and organ-specific aspects of photomorphogenesis, and that localized phytochrome pools engage distinct subsets of downstream target genes in cell-to-cell signaling. We developed a biochemical approach to selectively reduce functional phytochromes in an organ- or tissue-specific manner within transgenic plants. Our studies are based on a bipartite enhancer-trap approach that results in transactivation of the expression of a gene under control of the Upstream Activation Sequence (UAS) element by the transcriptional activator GAL43. The biliverdin reductase (BVR) gene under the control of the UAS is silently maintained in the absence of GAL4 transactivation in the UAS-BVR parent4. Genetic crosses between a UAS-BVR transgenic line and a GAL4-GFP enhancer trap line result in specific expression of the BVR gene in cells marked by GFP expression4. BVR accumulation in Arabidopsis plants results in phytochrome chromophore deficiency in planta5-7. Thus, transgenic plants that we have produced exhibit GAL4-dependent activation of the BVR gene, resulting in the biochemical inactivation of phytochrome, as well as GAL4-dependent GFP expression. Photobiological and molecular genetic analyses of BVR transgenic lines are yielding insight into tissue- and organ-specific phytochrome-mediated responses that have been associated with corresponding sites of photoperception4, 7, 8. Fluorescence Activated Cell Sorting (FACS) of GFP-positive, enhancer-trap-induced BVR-expressing plant protoplasts coupled with cell-type-specific gene expression profiling through microarray analysis is being used to identify putative downstream target genes involved in mediating spatial-specific phytochrome responses. This research is expanding our understanding of sites of light perception, the mechanisms through which various tissues or organs cooperate in light-regulated plant growth and development, and advancing the molecular dissection of complex phytochrome-mediated cell-to-cell signaling cascades.

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

The molecular basis of spatial-specific phytochrome responses is being investigated using transgenic plants that exhibit tissue- and organ-specific phytochrome deficiencies. The isolation of specific cells exhibiting induced phytochrome chromophore depletion by Fluorescence-Activated Cell Sorting followed by microarray analyses is being utilized to identify genes involved in spatial-specific phytochrome responses.

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called ...

OLIgo Mass Profiling (OLIMP) of Extracellular Polysaccharides

The direct contact of cells to the environment is mediated in many organisms by an extracellular matrix. One common aspect of extracellular matrices is that they contain complex sugar moieties in form of glycoproteins, proteoglycans, and/or polysaccharides. Examples include the extracellular matrix of humans and animal cells consisting mainly of fibrillar proteins and proteoglycans or the polysaccharide based cell walls of plants and fungi, and the proteoglycan/glycolipid based cell walls of bacteria. All these glycostructures play vital roles in cell-to-cell and cell-to-environment communication and signalling.
An extraordinary complex example of an extracellular matrix is present in the walls of higher plant cells. Their wall is made almost entirely of sugars, up to 75% dry weight, and consists of the most abundant biopolymers present on this planet. Therefore, research is conducted how to utilize these materials best as a carbon-neutral renewable resource to replace petrochemicals derived from fossil fuel. The main challenge for fuel conversion remains the recalcitrance of walls to enzymatic or chemical degradation due to the unique glycostructures present in this unique biocomposite.
Here, we present a method for the rapid and sensitive analysis of plant cell wall glycostructures. This method OLIgo Mass Profiling (OLIMP) is based the enzymatic release of oligosaccharides from wall materials facilitating specific glycosylhydrolases and subsequent analysis of the solubilized oligosaccharide mixtures using matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF/MS)1 (Figure 1). OLIMP requires walls of only 5000 cells for a complete analysis, can be performed on the tissue itself2, and is amenable to high-throughput analyses3. While the absolute amount of the solubilized oligosaccharides cannot be determined by OLIMP the relative abundance of the various oligosaccharide ions can be delineated from the mass spectra giving insights about the substitution-pattern of the native polysaccharide present in the wall.
OLIMP can be used to analyze a wide variety of wall polymers, limited only by the availability of specific enzymes4. For example, for the analysis of polymers present in the plant cell wall enzymes are available to analyse the hemicelluloses xyloglucan using a xyloglucanase5, 11, 12, 13, xylan using an endo-&beta;-(1-4)-xylanase 6,7, or for pectic polysaccharides using a combination of a polygalacturonase and a methylesterase 8. Furthermore, using the same principles of OLIMP glycosylhydrolase and even glycosyltransferase activities can be monitored and determined 9.

OLIgo Mass Profiling OLIMP of Extracellular Polysaccharides

A rapid way is described to gain insights into the structure of polysaccharides in an extracellular matrix. The method takes advantage of the specificity of glycosylhydrolases and the sensitivity of mass spectrometry allowing minute amounts of materials to be analyzed. This technique is adaptable to be used directly on tissue itself.

The direct contact of cells to the environment is mediated in many organisms by an extracellular matrix. One common aspect of extracellular matrices is ...

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts

Cellular invasion into local tissues is a process important in development and homeostasis. Malregulated invasion and subsequent cell movement is characteristic of multiple pathological processes, including inflammation, cardiovascular disease and tumor cell metastasis1. Focalized proteolytic degradation of extracellular matrix (ECM) components in the epithelial or endothelial basement membrane is a critical step in initiating cellular invasion. In tumor cells, extensive in vitro analysis has determined that ECM degradation is accomplished by ventral actin-rich membrane protrusive structures termed invadopodia2,3. Invadopodia form in close apposition to the ECM, where they moderate ECM breakdown through the action of matrix metalloproteinases (MMPs). The ability of tumor cells to form invadopodia directly correlates with the ability to invade into local stroma and associated vascular components3. 
Visualization of invadopodia-mediated ECM degradation of cells by fluorescent microscopy using dye-labeled matrix proteins coated onto glass coverslips has emerged as the most prevalent technique for evaluating the degree of matrix proteolysis and cellular invasive potential4,5. Here we describe a version of the standard method for generating fluorescently-labeled glass coverslips utilizing a commercially available Oregon Green-488 gelatin conjugate. This method is easily scaled to rapidly produce large numbers of coated coverslips. We show some of the common microscopic artifacts that are often encountered during this procedure and how these can be avoided. Finally, we describe standardized methods using readily available computer software to allow quantification of labeled gelatin matrix degradation mediated by individual cells and by entire cellular populations. The described procedures provide the ability to accurately and reproducibly monitor invadopodia activity, and can also serve as a platform for evaluating the efficacy of modulating protein expression or testing of anti-invasive compounds on extracellular matrix degradation in single and multicellular settings.

We describe the prototypical method for producing microscope coverslips coated with fluorescent gelatin for visualizing invadopodia-mediated matrix degradation. Computational techniques using available software are presented for quantifying the resultant levels of matrix proteolysis by single cells within a mixed population and for multicellular groups encompassing entire microscopic fields.

Cellular invasion into local tissues is a process important in development and homeostasis.  Malregulated invasion and subsequent cell movement is ...

Multi-parameter Measurement of the Permeability Transition Pore Opening in Isolated Mouse Heart Mitochondria

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with a molecular mass smaller than 1.5 kDa. Although the definitive molecular identity of the pore is still under debate, proteins such as cyclophilin D, VDAC and ANT contribute to mtPTP formation. While the involvement of mtPTP opening in cell death is well established1, accumulating evidence indicates that the mtPTP serves a physiologic role during mitochondrial Ca2+ homeostasis2, bioenergetics and redox signaling 3.
mtPTP opening is triggered by matrix Ca2+ but its activity can be modulated by several other factors such as oxidative stress, adenine nucleotide depletion, high concentrations of Pi, mitochondrial membrane depolarization or uncoupling, and long chain fatty acids4. In vitro, mtPTP opening can be achieved by increasing Ca2+ concentration inside the mitochondrial matrix through exogenous additions of Ca2+ (calcium retention capacity). When Ca2+ levels inside mitochondria reach a certain threshold, the mtPTP opens and facilitates Ca2+ release, dissipation of the proton motive force, membrane potential collapse and an increase in mitochondrial matrix volume (swelling) that ultimately leads to the rupture of the outer mitochondrial membrane and irreversible loss of organelle function.
Here we describe a fluorometric assay that allows for a comprehensive characterization of mtPTP opening in isolated mouse heart mitochondria. The assay involves the simultaneous measurement of 3 mitochondrial parameters that are altered when mtPTP opening occurs: mitochondrial Ca2+ handling (uptake and release, as measured by Ca2+ concentration in the assay medium), mitochondrial membrane potential, and mitochondrial volume. The dyes employed for Ca2+ measurement in the assay medium and mitochondrial membrane potential are Fura FF, a membrane impermeant, ratiometric indicator which undergoes a shift in the excitation wavelength in the presence of Ca2+, and JC-1, a cationic, ratiometric indicator which forms green monomers or red aggregates at low and high membrane potential, respectively. Changes in mitochondrial volume are measured by recording light scattering by the mitochondrial suspension. Since high-quality, functional mitochondria are required for the mtPTP opening assay, we also describe the steps necessary to obtain intact, highly coupled and functional isolated heart mitochondria.

A spectrofluorometric protocol for the measurement of the mitochondrial permeability transition pore opening in isolated mouse heart mitochondria is presented here. The assay involves the simultaneous measurement of mitochondria Ca2+ handling, mitochondrial membrane potential and mitochondrial volume. The procedure for obtaining high-quality and functional heart mitochondria is also described.

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with ...

Techniques for the Analysis of Extracellular Vesicles Using Flow Cytometry

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help regulate a diverse range of biological processes. Analysis of EVs using flow cytometry (FCM) has been notoriously difficult due to their small size and lack of discrete populations positive for markers of interest. Methods for EV analysis, while considerably improved over the last decade, are still a work in progress. Unfortunately, there is no one-size-fits-all protocol, and several aspects must be considered when determining the most appropriate method to use. Presented here are several different techniques for processing EVs and two protocols for analyzing EVs using either individual detection or a bead-based approach. The methods described here will assist with eliminating the antibody aggregates commonly found in commercial preparations, increasing signal&#8211;to-noise ratio, and setting gates in a rational fashion that minimizes detection of background fluorescence. The first protocol uses an individual detection method that is especially well suited for analyzing a high volume of clinical samples, while the second protocol uses a bead-based approach to capture and detect smaller EVs and exosomes.

Many different methods exist for the measurement of extracellular vesicles (EVs) using flow cytometry (FCM). Several aspects should be considered when determining the most appropriate method to use. Two protocols for measuring EVs are presented, using either individual detection or a bead-based approach.

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help ...

Measurement of Extracellular Ion Fluxes Using the Ion-selective Self-referencing Microelectrode Technique

Cells from animals, plants and single cells are enclosed by a barrier called the cell membrane that separates the cytoplasm from the outside. Cell layers such as epithelia also form a barrier that separates the inside from the outside or different compartments of multicellular organisms. A key feature of these barriers is the differential distribution of ions across cell membranes or cell layers. Two properties allow this distribution: 1) membranes and epithelia display selective permeability to specific ions; 2) ions are transported through pumps across cell membranes and cell layers. These properties play crucial roles in maintaining tissue physiology and act as signaling cues after damage, during repair, or under pathological condition. The ion-selective self-referencing microelectrode allows measurements of specific fluxes of ions such as calcium, potassium or sodium at single cell and tissue levels. The microelectrode contains an ionophore cocktail which is selectively permeable to a specific ion. The internal filling solution contains a set concentration of the ion of interest. The electric potential of the microelectrode is determined by the outside concentration of the ion. As the ion concentration varies, the potential of the microelectrode changes as a function of the log of the ion activity. When moved back and forth near a source or sink of the ion (i.e. in a concentration gradient due to ion flux) the microelectrode potential fluctuates at an amplitude proportional to the ion flux/gradient. The amplifier amplifies the microelectrode signal and the output is recorded on computer. The ion flux can then be calculated by Fick&#8217;s law of diffusion using the electrode potential fluctuation, the excursion of microelectrode, and other parameters such as the specific ion mobility. In this paper, we describe in detail the methodology to measure extracellular ion fluxes using the ion-selective self-referencing microelectrode and present some representative results.

Transporters in cell membranes allow differential segregation of ions across cell membranes or cell layers and play crucial roles during tissue physiology, repair and pathology. We describe the ion-selective self-referencing microelectrode that allows the measurement of specific ion fluxes at single cells and tissues in vivo.

Cells from animals, plants and single cells are enclosed by a barrier called the cell membrane that separates the cytoplasm from the outside. Cell layers ...

Measurement and Analysis of Extracellular Acid Production to Determine Glycolytic Rate

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both mitochondrial (respiration) and non-mitochondrial (other redox) reactions consume oxygen, but these reactions can be easily distinguished by chemical inhibition of mitochondrial respiration. However, while mitochondrial oxygen consumption is an unambiguous and direct measurement of respiration rate2, the same is not true for extracellular acid production and its relationship to glycolytic rate 3-6. Extracellular acid produced by cells is derived from both lactate, produced by anaerobic glycolysis, and CO2, produced in the citric acid cycle during respiration. For glycolysis, the conversion of glucose to lactate- + H+ and the export of products into the assay medium is the source of glycolytic acidification. For respiration, the export of CO2, hydration to H2CO3 and dissociation to HCO3- + H+ is the source of respiratory acidification. The proportions of glycolytic and respiratory acidification depend on the experimental conditions, including cell type and substrate(s) provided, and can range from nearly 100% glycolytic acidification to nearly 100% respiratory acidification 6. Here, we demonstrate the data collection and calculation methods needed to determine respiratory and glycolytic contributions to total extracellular acidification by whole cells in culture using C2C12 myoblast cells as a model.

Glycolysis is a defining metabolic marker in multiple biological systems. Monitoring glycolysis by measuring the extracellular flux of H+ is common, but requires correction to be quantitative and unambiguous. Here, we demonstrate how to gather and correct extracellular flux data to distinguish between respiratory and glycolytic sources of extracellular acidification.

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both ...

Isolation, Characterization, And High Throughput Extracellular Flux Analysis of Mouse Primary Renal Tubular Epithelial Cells

Mitochondrial dysfunction in the renal tubular epithelial cells (TECs) can lead to renal fibrosis, a major cause of&#160;chronic kidney disease (CKD). Therefore, assessing mitochondrial function in primary TECs may provide valuable insight into the bioenergetic status of the cells, providing insight into the pathophysiology of CKD. While there are a number of complex protocols available for the isolation and purification of proximal tubules in different species, the field lacks a cost-effective method optimized for&#160;tubular cell isolation without the need for purification. Here, we provide an isolation protocol that allows for studies focusing on both primary mouse proximal and distal renal TECs. In addition to cost-effective reagents and minimal animal procedures required in this protocol, the isolated cells maintain high energy levels after isolation and can be sub-cultured up to four&#160;passages, allowing for continuous studies. Furthermore, using a high throughput extracellular flux analyzer, we assess the mitochondrial respiration directly in the isolated TECs in a 96-well plate for which we provide recommendations for the optimization of cell density and compound concentration. These observations suggest that this protocol can be used for renal tubular ex vivo studies with a consistent, well-standardized production of renal TECs. This protocol may have broader future applications to study mitochondrial dysfunction associated with renal disorders for drug discovery or drug characterization purposes.

This protocol provides a cost-effective approach to isolate and characterize mouse primary renal tubular cells that can subsequently be sub-cultured to assess renal biological functions ex vivo, including mitochondrial bioenergetics.

Mitochondrial dysfunction in the renal tubular epithelial cells (TECs) can lead to renal fibrosis, a major cause of&#160;chronic kidney disease (CKD). ...

Adipo-Clear: A Tissue Clearing Method for Three-Dimensional Imaging of Adipose Tissue

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte precursors, immune cells, fibroblasts, blood vessels, and nerve projections. Although the molecular control of cell type specification and how these cells interact have been increasingly delineated, a more comprehensive understanding of these adipose-resident cells can be achieved by visualizing their distribution and architecture throughout the whole tissue. Existing immunohistochemistry and immunofluorescence approaches to analyze adipose histology rely on thin paraffin-embedded sections. However, thin sections capture only a small portion of tissue; as a result, the conclusions can be biased by what portion of tissue is analyzed. We have therefore developed an adipose tissue clearing technique, Adipo-Clear, to permit comprehensive three-dimensional visualization of molecular and cellular patterns in whole adipose tissues. Adipo-Clear was adapted from iDISCO/iDISCO+, with specific modifications made to completely remove the lipid stored in the tissue while preserving native tissue morphology. In combination with light-sheet fluorescence microscopy, we demonstrate here the use of the Adipo-Clear method to obtain high-resolution volumetric images&#160;of an entire adipose tissue.

Due to the high lipid content, adipose tissue has been challenging to visualize using traditional histological methods. Adipo-Clear is a tissue clearing technique that allows robust labeling and high-resolution volumetric fluorescent imaging of adipose tissue. Here, we describe the methods for sample preparation, pretreatment, staining, clearing, and mounting for imaging.

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte ...

Laser-Capture Microdissection RNA-Sequencing for Spatial and Temporal Tissue-Specific Gene Expression Analysis in Plants

The development of a complex multicellular organism is governed by distinct cell types that have different transcriptional profiles. To identify transcriptional regulatory networks that govern developmental processes it is necessary to measure the spatial and temporal gene expression profiles of these individual cell types. Therefore, insight into the spatio-temporal control of gene expression is essential to gain understanding of how biological and developmental processes are regulated. Here, we describe a laser-capture microdissection (LCM) method to isolate small number of cells from three barley embryo organs over a time-course during germination followed by transcript profiling. The method consists of tissue fixation, tissue processing, paraffin embedding, sectioning, LCM and RNA extraction followed by real-time PCR or RNA-seq. This method has enabled us to obtain spatial and temporal profiles of seed organ transcriptomes from varying numbers of cells (tens to hundreds), providing much greater tissue-specificity than typical bulk-tissue analyses. From these data we were able to define and compare transcriptional regulatory networks as well as predict candidate regulatory transcription factors for individual tissues. The method should be applicable to other plant tissues with minimal optimization.

Presented here is a protocol for laser-capture microdissection (LCM) of plant tissues. LCM is a microscopic technique for isolating areas of tissue in a contamination-free manner. The procedure includes tissue fixation, paraffin embedding, sectioning, LCM and RNA extraction. RNA is used in the downstream tissue-specific, temporally resolved analysis of transcriptomes.

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

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Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

OLIgo Mass Profiling (OLIMP) of Extracellular Polysaccharides

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts

Multi-parameter Measurement of the Permeability Transition Pore Opening in Isolated Mouse Heart Mitochondria

Techniques for the Analysis of Extracellular Vesicles Using Flow Cytometry

Measurement of Extracellular Ion Fluxes Using the Ion-selective Self-referencing Microelectrode Technique

Measurement and Analysis of Extracellular Acid Production to Determine Glycolytic Rate

Isolation, Characterization, And High Throughput Extracellular Flux Analysis of Mouse Primary Renal Tubular Epithelial Cells

Adipo-Clear: A Tissue Clearing Method for Three-Dimensional Imaging of Adipose Tissue

Laser-Capture Microdissection RNA-Sequencing for Spatial and Temporal Tissue-Specific Gene Expression Analysis in Plants