This work is supported by the Intramural Research Program of the National Eye Institute (ZIAEY000450 and ZIAEY000546).

All mouse protocols were approved by the Animal Care and Use Committee of the National Eye Institute (NEI ASP# 650). Mice were housed in 12 h light-dark conditions and cared for by following the recommendations of the Guide for the Care and Use of Laboratory Animals, the Institute of Laboratory Animal Resources, and the Public Health Service Policy on Humane Care and Use of Laboratory Animals.
1. Hydrating sensor cartridge and preparation of the assay medium
<ol>
	<li>The day before the experiment, add 1 mL of the calibration medium to each well of the utility plate. Place the Hydro-Booster cover on the top and lower the sensor cartridge through the opening on the cover. Check to ensure that the sensor is submerged in the calibration medium. Incubate the sensor cartridge overnight in a CO2-free incubator at 37 &#730;C to activate the fluorophores. 
	NOTE: To prevent evaporation, the incubator is humidified by keeping a tray of water inside, and the sensor cartridge cassette is wrapped with clear plastic wrap.</li>
	<li>Prepare the assay medium by reconstituting the Seahorse DMEM medium with the addition of glucose, pyruvate, and glutamine to the desired concentrations. In the assays reported in this article, the final concentration of substrates in the assay medium are: 6 mM of glucose, 0.12 mM of pyruvate, and 0.5 mM of glutamine. For each assay plate, 40 mL of the assay medium is prepared fresh on the day of the experiment.</li>
	<li>Set up the assay program in the analyzer following the manufacturer&#39;s instruction26. In the assay demonstrated here, the protocol is set as follows: 5 cycles of measurements for baseline, then inject port A, followed by 4 cycles of measurements, then inject port B and followed by 4 cycles of measurements. Each cycle is composed of mix (3 min), wait (2 min) and measure (3 min).</li>
</ol>
2. Coating mesh inserts of islet capture microplate
<ol>
	<li>Prepare the coating mix by combining 20 &#181;L of the cell attachment medium (e.g., Cell-Tak) with 171 &#181;L of 0.1 M sodium bicarbonate and 9 &#181;L of 1 M NaOH.</li>
	<li>Open the lid of the cassette containing mesh inserts. Pipette 8 &#181;L of the coating mix to each mesh inserts. Use a pipette tip to gently smear/spread the droplet around to distribute the coating mix equally throughout the mesh insert.</li>
	<li>Close the cassette and allow the mesh inserts to incubate at room temperature for at least 25 min for adsorption.</li>
	<li>Wash the mesh insert by pipetting 4 mL of the assay medium directly onto the mesh inserts. Gently shake the cassette to ensure all mesh inserts are washed with the assay medium.</li>
	<li>Keep the mesh insert aside. It is ready to use.</li>
</ol>
3. Preparing injection compounds
<ol>
	<li>Take out stock aliquots of Bam15 (10 mM), Rotenone (10 mM), Antimycin A (10 mM) and 2-DG (500 mM) from -80 &#176;C freezer and thaw at room temperature. 
	NOTE: The 2-DG stock is ready to use. The other drugs need to be diluted to working stock.</li>
	<li>Warm up 10 mL of the assay medium in a 37 &#176;C water bath.</li>
	<li>Dilute 10 mM Bam15 stock to 50 &#181;M working stock using a two-step dilution procedure: mix 20 &#181;L of 10 mM stock with 20 &#181;L of DMSO to get 5 mM intermediate stock. Then mix 10 &#181;L of the above 5 mM intermediate stock with 990 &#181;L of pre-warmed assay medium to get the final 50 &#181;M working stock.</li>
	<li>Dilute and combine 10 mM Rotenone and 10 mM Antimycin A stock to 10 &#181;M Rotenone/Antimycin A (Rot/AA) working stock by two steps of dilutions: mix 10 &#181;L each of 10 mM Rotenone and 10 mM Antimycin A stock with 80 &#181;L of DMSO to get 1 mM Rot/AA intermediate stock. Then mix 10 &#181;L of the above 1 mM intermediate stock with 990 &#181;L of pre-warmed assay medium to get the final 10 &#181;M Rot/AA working stock.</li>
	<li>Freshly prepare the above-mentioned working stocks of injection compounds on the day of the experiment and set them aside at room temperature until loading into injection ports of the sensor cartridge.</li>
</ol>
4. Retinal dissection and retinal punch preparation
<ol>
	<li>Euthanize a mouse by CO2 asphyxiation following AVMA Guidelines on Euthanasia27. 
	NOTE: Do not leave the animal in a CO2 chamber longer than the time needed for euthanasia.</li>
	<li>Enucleate eyes and place into ice-old 1x PBS buffer in a Petri-dish and then place it under a dissection microscope.</li>
	<li>Carefully remove, by cutting with microscissors, the extra rectus muscles attached outside the eyeball and cut off the optic nerve.</li>
	<li>Use a 30 G needle to punch a hole at the edge of the cornea (limbus); this serves as the insertion site for the microscissors. Then, use a fine dissection microscissors to make a circular cut along the edge of the cornea, separating it from the posterior eye cup.</li>
	<li>Use sharp dissection forceps to remove the cornea, lens, and the vitreous humor away from the eye cup.</li>
	<li>Use fine dissection microscissors to make several small cuts on the scleral layer at the rim of eye cup. Avoid cutting the retina layer. Use two sharp dissection forceps to hold on to the scleral tissue at each side of the cut and very carefully pull on the scleral layer to remove it from the neural retina. Repeat this around the eye cup until all sclera is removed and an intact retinal cup is obtained.</li>
	<li>Use dissection microscissors and make radial cuts on the retinal cup to flatten it and generate several distinct sections. 
	NOTE: Depending on the person&#39;s dissection skills and experience in handling fresh retinal tissue, the retinal cup can be cut to generate 3 to 5 distinct sections.</li>
	<li>Use 1 mm diameter biopsy puncher to cut one retinal disk from each section of the flattened retinal cup. 
	NOTE: Care should be taken to get the retinal disks punched at equal distance from the optic nerve head.</li>
	<li>Use forceps to transfer the pre-coated mesh inserts into the dissecting petri-dish. With the help of two superfine eyelash brushes, place the retinal punch disk onto the mesh insert. The retinal punch disk is placed at the center of the mesh insert with ganglion cell layer side down touching the mesh and photoreceptor layer facing up. 
	NOTE: Frequently, some RPE cells remain attached to the photoreceptors, and the pigmentation of these cells can be used as an indicator of the retinal punch disk orientation.</li>
</ol>
5. Loading the sensor cartridge injection ports and calibration
<ol>
	<li>Take the hydrated sensor cartridge plate cassette out of the 37 &#176;C incubator. Remove the Hydro-Booster cover and place the sensor cartridge back on the utility plate.</li>
	<li>Load the desired volume of injection compound working stock solutions into appropriate ports. Hold the pipette tip at 45 &#176;&#160;angle. Insert the pipette tip halfway into an injection port with the bevel of the tip against the opposite wall of the injection port and gently load the compound into each port. Avoid introducing air bubbles.</li>
	<li>Refer to the instrument user guide for the volume of the compound loaded in each injection port for a specific assay. In the experiments presented in this paper, 68 &#181;L of 50 &#181;M Bam15 working stock (for mitochondrial stress assay) or 68 &#181;L of 10 &#181;M Rot/AA working stock (for glycolytic rate assay) is loaded into port A; 75 &#181;L of 10 &#181;M Rot/AA working stock (for mitochondrial stress assay) or 75 &#181;L of 500 mM 2-DG working stock (for glycolytic rate assay) is loaded into port B.</li>
	<li>Load injection ports of all wells of the plate including background correction wells and blank wells to ensure proper injection. Load the respective compound solution in each port for the background correction wells. Assay medium can be substituted, instead of the compound solution, in each of the ports of the bank wells.</li>
	<li>Place the loaded sensor cartridge plate, with lid off, into the analyzer machine to start calibration prior to the assay run. After the calibration is over, the program will automatically pause, waiting for the replacement of the utility plate with the islet capture plate containing retinal punches.</li>
</ol>
6. Loading the islet capture plate and start assay run
<ol>
	<li>Add 607 &#181;L of the assay medium to each well of the islet capture plate</li>
	<li>Use forceps to grab the rim of the mesh insert containing retinal punch disks on top and take it out from the Petri-dish. Lightly tap the bottom of the mesh insert on an absorbing wipe tissue to remove extra liquid and put it into the well of the islet capture plate. Repeat this step until all mesh inserts with retinal punches are placed into the islet capture plate. Fill background correction wells and blank wells with empty mesh inserts.</li>
	<li>Use two Graefe forceps to carefully and gently press the rim of each mesh insert and make sure that these are securely inserted at the bottom of the islet capture plate.</li>
	<li>Place the loaded islet capture plate into a 37 &#176;C incubator for 5 min to warm up.</li>
	<li>Eject the utility plate after the calibration is complete and replace it with an and replace it with the islet capture plate, with lid off, containing&#160;retinal punches.</li>
	<li>Resume the assay run.</li>
</ol>
7. Run termination and data storage
<ol>
	<li>After the run is complete, eject the sensor cartridge and islet capture plate containing retinal punches. The data is automatically saved as .asyr file.</li>
	<li>Use the associated data analysis software to view and analyze the data following the manufacturer&#39;s user guide26.</li>
	<li>Use the Export function to export .xslx file of the data, which can be viewed and analyzed using spreadsheet software.</li>
</ol>
8. Saving the retinal punch sample
<ol>
	<li>After the assay, take out the plate from the machine, remove the sensor cartridge and gently remove the assay medium from each well using a pipette.</li>
	<li>Apply the cover back on and seal the sides of the plate with the parafilm strip.</li>
	<li>Store at -80 &#176;C.</li>
	<li>For normalization, quantify the total DNA or protein content of the punch in each well.</li>
</ol>
9. Data analysis
<ol>
	<li>Mitochondrial stress assay 
	NOTE: The measured OCR value (totalOCR) represents total oxygen consumption by the tissue. After Bam15 (uncoupler) injection, OCR increases from the basal level (totalOCRbasal) to the maximum level (totalOCRmax) and goes down following the Rot/AA injection. The residual OCR value after Rot/AA injection (totalOCRRot/AA) represents non-mitochondrial oxygen consumption.
	<ol>
		<li>Calculate mitochondria-related oxygen consumption as: 
		<img alt="Equation 1" src="/files/ftp_upload/62914/62914eq01.jpg" /> (Eq. 1)28</li>
		<li>Calculate the mitochondrial reserve capacity (MRC) as: 
		<img alt="Equation 2" src="/files/ftp_upload/62914/62914eq2v4.png" /> (Eq. 2)29 
		NOTE: The last reading among the 5 measurements before Bam15 injection is taken as the &#34;basal&#34; value (for totalOCRbasal and mitoOCRbasal). The highest reading among the 4 measurements following Bam15 injection is used as &#34;max&#34; value (for totalOCRmax and mitoOCRmax). The lowest reading among the 4 measurements following the Rot/AA injection is used as totalOCRRot/AA.</li>
	</ol>
	</li>
	<li>Glycolytic rate assay 
	NOTE: The measured ECAR value (totalECAR) represents the total acidification of the medium by the tissue&#39;s metabolic activity. In general, acidification of the extracellular micro-environment results mainly by extrusion of the glycolytic product, lactate. Catabolism of substrates in mitochondrial TCA cycle results in the production of CO2, which also acidifies the extracellular medium through hydration to bicarbonate.
	<ol>
		<li>Substract mitochondrial contributed medium acidification (mitoECAR) from totalECAR to obtain the glycoECAR. 
		<img alt="Equation 3" src="/files/ftp_upload/62914/62914eq03.jpg" /> (Eq. 3)28 
		NOTE: Mitochondrial respiration and TCA cycle are strongly coupled processes. Production of CO2 from mitochondria is a function of the rate of OXPHOS, which is measurable by mitoOCR.</li>
		<li>Calculate the mitoECAR as: 
		<img alt="Equation 4" src="/files/ftp_upload/62914/62914eq04.jpg" /> (Eq. 4)28 
		where, the CCF (CO2 Contribution Factor) is an empirically calculated ratio value, representing the amount of H+ contribution from CO2-mediated acidification vs each O2 consumption from OXPHOS. CCF for this system is pre-determined to be 0.6028. Accurate measurement of medium acidification is determined by the buffer capacity of the medium, the sensitivity of instrument pH sensor, and the effective measurement chamber capacity. Here, the BF (Buffer Factor) is a parameter of the in situ experimental buffer capacity, representing the amount of H+ or OH- added to the effective measurement chamber to change the pH level by 1 unit. When customized assay medium is used, the BF can be determined by titrating known amounts of acid into the assay medium following the Buffer Factor protocol30. The Seahorse DMEM medium pH 7.4 used in this protocol has a pre-determined BF of 2.60 mmol H+/L/pH. The islet capture plate used in this protocol has a Volmicrochamber = 16.6 &#181;L31. The volume scaling factor, Kvol, is an empirically determined constant. Kvol value is not available for the islet capture plate but can be calculated from the value of the microplate28, accounting for the volume difference in their microchambers, to be 0.41. 
		NOTE: Injection of the Rot/AA shuts down mitochondrial respiration and forces the tissue to switch to glycolysis for ATP production, leading to higher lactate extrusion and an increase in ECAR measurement. Glycolysis is ceased with 2-DG injection, and the residual ECAR measurement reveals non-glycolytic and non-mitochondrial acidification of medium.</li>
		<li>Calculate the glycolytic reserve capacity (GRC) as: 
		<img alt="Equation 5" src="/files/ftp_upload/62914/62914eq05.jpg" /> (Eq. 5)32 
		where, the last reading among the 5 measurements before Rot/AA injection is taken as the &#34;basal&#34; value (glycoECARbasal). The highest reading among the 4 measurements following Rot/AA injection is used as &#34;max&#34; value (glycoECARmax). The lowest reading among the 4 measurements following 2-DG injection is used as glycoECAR2-DG.</li>
	</ol>
	</li>
	<li>Normalization 
	NOTE: Normalization is essential when comparing the readings from retinal tissues of different age groups or between wild-type and pathological/degenerative samples, which might differ in cell numbers.
	<ol>
		<li>Use commerically available kits to assess the DNA content in each retinal punch disk33,34.</li>
		<li>Alternatively, use radioimmunoprecipitation assay buffer (RIPA buffer) to extract total protein from the retinal punch and use the protein content for normalization. 
		NOTE: The surface area of an adult mouse retina has been previously determined to be around 20 mm2, and each retina contains ~6.5 million cells35. Hence, each 1 mm diameter retinal punch is ~1/25 of a single retina and contains ~260K cells. One can refer to these numbers when comparing the data from a retinal punch to those from other tissue samples or cultured cells,</li>
	</ol>
	</li>
</ol>

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Report Generator User Guide Agilent Seahorse XF Cell Mito Stress Test Available from: <a target="_blank" href="https://www.agilent.com/cs/library/usermanuals/public/Report_Generator_User_Guide_Seahorse_XF_Cell_Mito_Stress_Test_Single_File.pdf">https://www.agilent.com/cs/library/usermanuals/public/Report_Generator_User_Guide_Seahorse_XF_Cell_Mito_Stress_Test_Single_File.pdf</a> (2021)</li><li>. Agilent Seahorse XF Buffer Factor Protocol Available from: <a target="_blank" href="https://www.agilent.com/cs/library/usermanuals/public/usermanual-xf-buffer-factor-protocol-cell-analysis-S7888-10010en-agilent.pdf">https://www.agilent.com/cs/library/usermanuals/public/usermanual-xf-buffer-factor-protocol-cell-analysis-S7888-10010en-agilent.pdf</a> (2021)</li><li>. Agilent sensor cartridges and cell culture microplates Available from: <a target="_blank" href="https://www.agilent.com/cs/library/brochures/5991-8657EN_seahorse_plastics_brochure.pdf">https://www.agilent.com/cs/library/brochures/5991-8657EN_seahorse_plastics_brochure.pdf</a> (2021)</li><li>. Agilent Seahorse XF Glycolysis Stress Test Kit User Guide Available from: <a target="_blank" href="https://www.agilent.com/cs/library/usermanuals/public/XF_Glycolysis_Stress_Test_Kit_User_Guide.pdf">https://www.agilent.com/cs/library/usermanuals/public/XF_Glycolysis_Stress_Test_Kit_User_Guide.pdf</a> (2021)</li><li>Fan, Y. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+bioassay+to+measure+energy+metabolism+in+mouse+colonic+crypts,+organoids,+and+sorted+stem+cells.">A bioassay to measure energy metabolism in mouse colonic crypts, organoids, and sorted stem cells.</a> American Journal of Physiology-Gastrointestinal and Liver Physiology. 309, 1-9 (2015).</li><li>Huang, L., 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=Ductal+pancreatic+cancer+modeling+and+drug+screening+using+human+pluripotent+stem+cell-+and+patient-derived+tumor+organoids.">Ductal pancreatic cancer modeling and drug screening using human pluripotent stem cell- and patient-derived tumor organoids.</a> Nature Medicine. 21 (11), 1364-1371 (2015).</li><li>Jeon, C. J., Strettoi, E., Masland, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+major+cell+populations+of+the+mouse+retina.">The major cell populations of the mouse retina.</a> Journal of Neuroscience. 18 (21), 8936-8946 (1998).</li><li>Akimoto, M., 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=Targeting+of+GFP+to+newborn+rods+by+Nrl+promoter+and+temporal+expression+profiling+of+flow-sorted+photoreceptors.">Targeting of GFP to newborn rods by Nrl promoter and temporal expression profiling of flow-sorted photoreceptors.</a> Proceedings of the National Academy of Science U. S. A. 103 (10), 3890-3895 (2006).</li><li>Kenwood, B. M., 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=Identification+of+a+novel+mitochondrial+uncoupler+that+does+not+depolarize+the+plasma+membrane.">Identification of a novel mitochondrial uncoupler that does not depolarize the plasma membrane.</a> Molecular Metabolism. 3 (2), 114-123 (2014).</li><li>Corso-Diaz, X., 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=Genome-wide+profiling+identifies+DNA+methylation+signatures+of+aging+in+rod+photoreceptors+associated+with+alterations+in+energy+metabolism.">Genome-wide profiling identifies DNA methylation signatures of aging in rod photoreceptors associated with alterations in energy metabolism.</a> Cell Reports. 31 (3), 107525 (2020).</li><li>Berkowitz, B. 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=Mitochondrial+respiration+in+outer+retina+contributes+to+light-evoked+increase+in+hydration+in+vivo.">Mitochondrial respiration in outer retina contributes to light-evoked increase in hydration in vivo.</a> Investigative Ophthalmology and Visual Science. 59 (15), 5957-5964 (2018).</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></ol>

The authors have nothing to disclose.

Provided here are detailed instructions for performing microplate-based assays of mitochondrial respiration and glycolysis activity using ex vivo, freshly dissected retinal punch disks. The protocol has been optimized to: 1) ensure the use of a suitable assay medium for ex vivo retinal tissue; 2) employ proper size of retinal punch disks to obtain OCR and ECAR readings that fall within the machine&#39;s optimal detecting range; 3) coating mesh inserts to enhance the adhesiveness of retinal punch for sta...

Mitochondria are essential organelle that regulates cellular metabolism, signaling, homeostasis, and apoptosis by coordinating multiple crucial physiological processes1. Mitochondria serve as the powerhouse in the cell to generate adenosine triphosphate (ATP) through oxidative phosphorylation (OXPHOS) and provide energy that supports almost all cellular events. The majority of cellular oxygen is metabolized in mitochondria, where it serves as the final electron acceptor in the electron transport chain (ETC) during aerobic respiration. Low amounts of ATP can also be produced from glycolysis in the cytosol, where glucose is converted to pyruvate, which can be further converted to lactate or be transported into mitochondria and oxidized to acetyl-CoA, a substrate in the tricarboxylic acid cycle (TCA cycle).
The retina is one of the most metabolically active tissues in mammals2, displaying high levels of mitochondrial respiration and extremely high oxygen consumption3. The rod and cone photoreceptors contain a high density of mitochondria4, and OXPHOS generates most ATP in the retina5. In addition, the retina also relies heavily on aerobic glycolysis6,7 by converting glucose to lactate5. Mitochondrial defects are associated with various neurodegenerative diseases8,9; and with its unique high energy demands, the retina is especially vulnerable to metabolic defects, including those affecting mitochondrial OXPHOS4 and glycolysis10. Mitochondrial dysfunction and defects in glycolysis are implicated in retinal11,12 and macular13 degenerative diseases, age-related macular degeneration10,14,15,16, and diabetic retinopathy17,18. Therefore, accurate measurements of mitochondrial respiration and glycolysis can provide important parameters for assessing the integrity and health of the retina.
Mitochondrial respiration can be measured through the determination of oxygen consumption rate (OCR). Given that the conversion of glucose to pyruvate and subsequently to lactate results in extrusion of protons into and acidification of the extracellular environment, measurements of the extracellular acidification rate (ECAR) provide an indication of glycolysis flux. As the retina is composed of multiple cell types with intimate relationships and active synergy, including the exchange of substrates6, it is imperative to analyze mitochondrial function and metabolism in the context of whole retinal tissue with intact lamination and circuitry. For the past several decades, the Clark type O2 electrodes and other oxygen microelectrodes have been used to measure oxygen consumption in the retina19,20,21. These oxygen electrodes have major limitations in sensitivity, requirement of a large sample volume, and the need for continuous stirring of suspending sample, which usually leads to the disruption of cellular and tissue context. The protocol described here was developed using a microplate-based, fluorescence technique to measure mitochondrial energy metabolism in freshly dissected ex vivo mouse retina tissue. It allows mid-throughput real-time measurements of both OCR and ECAR simultaneously using a small sample (1 mm punch) of ex vivo retinal tissue while avoiding the need for suspension and continuous stirring.
Demonstrated here is the experimental procedure for mitochondrial stress assay and glycolytic rate assay on freshly dissected retinal punch disks. This protocol allows the measurement of mitochondria-related metabolic activities in an ex vivo tissue context. Different from the assays performed using cultured cells, the readings obtained here reflect combined energy metabolism at the tissue level and are influenced by interactions between the different cell types within the tissue. The protocol is modified from a previously published version22,23 to adapt to the new generation of the Agilent Seahorse extracellular flux 24-wells (XFe24) analyzer with Islet Capture plate. The assay medium, injection compound concentrations, and number/duration of assay cycles have also been optimized for retinal tissue. A detailed step-by-step protocol is given for the preparation of retinal punch disks. More information on the program setup and data analysis can be obtained from the manufacturer&#39;s user guide24,25,26.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1X PBS</td>
			<td>Thermo Fisher</td>
			<td>14190-144</td>
			<td></td>
		</tr>
		<tr>
			<td>2-Deoxy glucose (2-DG), 500 mM stock solution</td>
			<td>Sigma</td>
			<td>D6134</td>
			<td>Dissolve in Seahorse XF DMEM medium, prepare ahead of time</td>
		</tr>
		<tr>
			<td>30-gauge needle</td>
			<td>BD Precision Glide</td>
			<td>305106</td>
			<td></td>
		</tr>
		<tr>
			<td>Antimycin A, 10 mM stock solution</td>
			<td>Sigma</td>
			<td>A8674</td>
			<td>Dissolve in DMSO, prepare ahead of time</td>
		</tr>
		<tr>
			<td>Bam15, 10 mM stock solution</td>
			<td>TimTec</td>
			<td>ST056388</td>
			<td>Dissolve in DMSO, prepare ahead of time</td>
		</tr>
		<tr>
			<td>Biopsy puncher, 1 mm</td>
			<td>Integra Miltex</td>
			<td>33-31AA</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell-Tak</td>
			<td>Corning Life Sciences</td>
			<td>CB40240</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2 asphyxiation chamber</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection forceps-Dumont #5</td>
			<td>Fine Science Tools</td>
			<td>11251-10</td>
			<td>Stright tip</td>
		</tr>
		<tr>
			<td>Dissection forceps-Dumont #7</td>
			<td>Fine Science Tools</td>
			<td>11274-20</td>
			<td>Curved tip</td>
		</tr>
		<tr>
			<td>Dissection microscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma</td>
			<td>D2438</td>
			<td></td>
		</tr>
		<tr>
			<td>Graefe forceps</td>
			<td>Fine Science Tools</td>
			<td>11051-10</td>
			<td>Curved, Serrated tip</td>
		</tr>
		<tr>
			<td>Microscissors</td>
			<td>Fine Science Tools</td>
			<td>15004-08</td>
			<td>Curved tip</td>
		</tr>
		<tr>
			<td>NaOH solution, 1 M</td>
			<td>Sigma-Aldrich</td>
			<td>S8263</td>
			<td>Aqueous solution, prepare ahead of time</td>
		</tr>
		<tr>
			<td>Rotenone, 10 mM stock solution</td>
			<td>Sigma</td>
			<td>R8875</td>
			<td>Dissolve in DMSO, prepare ahead of time</td>
		</tr>
		<tr>
			<td>Seahorse calibration medium</td>
			<td>Agilent</td>
			<td>100840-000</td>
			<td></td>
		</tr>
		<tr>
			<td>Seahorse XF 1.0 M glucose</td>
			<td>Agilent</td>
			<td>103577-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Seahorse XF 100 mM pyruvate</td>
			<td>Agilent</td>
			<td>103578-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Seahorse XF 200 mM glutamine</td>
			<td>Agilent</td>
			<td>103579-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Seahorse XF DMEM medium</td>
			<td>Agilent</td>
			<td>103575-100</td>
			<td>pH 7.4, with 5 mM HEPES</td>
		</tr>
		<tr>
			<td>Seahorse XFe24 Islet Capture FluxPak</td>
			<td>Agilent</td>
			<td>103518-100</td>
			<td>Containing Sensor Cartridge and Islet Capture microplate</td>
		</tr>
		<tr>
			<td>Seahorse XFe24, Extra Cellular Flux Analyzer</td>
			<td>Agilent</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium bicarbonate solution, 0.1 M</td>
			<td>Sigma-Aldrich</td>
			<td>S5761</td>
			<td>Aqueous solution, prepare ahead of time</td>
		</tr>
		<tr>
			<td>Superfine eyelash brush</td>
			<td>Ted Pella</td>
			<td>113</td>
			<td></td>
		</tr>
	</tbody>
</table>

determination of mitochondrial respiration and glycolysis in ex vivo retinal tissue samples

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. In addition, photoreceptors also exhibit high aerobic glycolysis like cancer cells. Precise measurements of these metabolic activities can provide valuable insights into cellular homeostasis under physiological conditions and in disease states. High throughput microplate-based assays have been developed to measure mitochondrial respiration and various metabolic activities in live cells. However, a vast majority of these are developed for cultured cells and have not been optimized for intact tissue samples and for application ex vivo. Described here is a detailed step-by-step protocol, using microplate-based fluorescence technology, to directly measure oxygen consumption rate (OCR) as an indicator of mitochondrial respiration, as well as extracellular acidification rate (ECAR) as an indicator of glycolysis, in intact ex vivo retinal tissue. This method has been used to successfully assess metabolic activities in adult mouse retina and demonstrate its application in investigating cellular mechanisms of aging and disease.

The data reported here are representative mitochondrial stress assay showing OCR trace (Figure 1) and glycolytic rate assay showing OCR trace and ECAR trace (Figure 2), which were performed using freshly dissected 1 mm retinal punch disks from 4 months old transgenic Nrl-L-EGFP mice36 (C57B/L6 background). These mice express GFP specifically in rod photoreceptors without altering normal retinal development, histology, and physiol...

Watch this Scientific Journal Video about Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples at JoVE.com

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Described here is a detailed protocol for performing mitochondrial stress assay and glycolytic rate assay in ex vivo retinal tissue samples using a commercial bioanalyzer.

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. ...

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Neuroscience

4.1K Views.  National Institutes of Health. Described here is a detailed protocol for performing mitochondrial stress assay and glycolytic rate assay in ex vivo retinal tissue samples using a commercial bioanalyzer.

Video: Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP. A significant loss of &Delta;&Psi;m renders cells depleted of energy with subsequent death. Reactive oxygen species (ROS) are important signaling molecules, but their accumulation in pathological conditions leads to oxidative stress. The two major sources of ROS in cells are environmental toxins and the process of oxidative phosphorylation. Mitochondrial dysfunction and oxidative stress have been implicated in the pathophysiology of many diseases; therefore, the ability to determine &Delta;&Psi;m and ROS can provide important clues about the physiological status of the cell and the function of the mitochondria. 
Several fluorescent probes (Rhodamine 123, TMRM, TMRE, JC-1) can be used to determine &Delta;&psi;m in a variety of cell types, and many fluorescence indicators (Dihydroethidium, Dihydrorhodamine 123, H2DCF-DA) can be used to determine ROS. Nearly all of the available fluorescence probes used to assess &Delta;&Psi;m or ROS are single-wavelength indicators, which increase or decrease their fluorescence intensity proportional to a stimulus that increases or decreases the levels of &Delta;&Psi;m or ROS. Thus, it is imperative to measure the fluorescence intensity of these probes at the baseline level and after the application of a specific stimulus. This allows one to determine the percentage of change in fluorescence intensity between the baseline level and a stimulus. This change in fluorescence intensity reflects the change in relative levels of &Delta;&Psi;m or ROS. In this video, we demonstrate how to apply the fluorescence indicator, TMRM, in rat cortical neurons to determine the percentage change in TMRM fluorescence intensity between the baseline level and after applying FCCP, a mitochondrial uncoupler. The lower levels of TMRM fluorescence resulting from FCCP treatment reflect the depolarization of mitochondrial membrane potential. We also show how to apply the fluorescence probe H2DCF-DA to assess the level of ROS in cortical neurons, first at baseline and then after application of H2O2. This protocol (with minor modifications) can be also used to determine changes in &#x2206;&#x03A8;m and ROS in different cell types and in neurons isolated from other brain regions.

We demonstrate application of the fluorescence indicator, TMRM, in cortical neurons to determine the relative changes in TMRM fluorescence intensity before and after application of a specific stimulus. We also show application of the fluorescence probe H2DCF-DA to assess the relative level of reactive oxygen species in cortical neurons.

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP.  A ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

Ex vivo Culturing of Whole, Developing Drosophila Brains

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for investigating the cell biology and development of central brain structures by allowing acute pharmacological interventions and live imaging of cellular processes. As an example of the technique, prior work from our lab1 has shown that a previously unrecognized subcellular compartment lies between the axonal and somatodendritic compartments of axons of the Drosophila central brain. The development of this compartment, referred to as the axon initial segment (AIS)2, was shown genetically to depend on the neuron-specific cyclin-dependent kinase, Cdk5. We show here that ex vivo treatment of wild-type Drosophila larval brains with the Cdk5-specific pharmacological inhibitors roscovitine and olomoucine3 causes acute changes in actin organization, and in localization of the cell-surface protein Fasciclin 2, that mimic the changes seen in mutants that lack Cdk5 activity genetically.
A second example of the ex vivo culture technique is provided for remodeling of the connections of embryonic mushroom body (MB) gamma neurons during metamorphosis from larva to adult. The mushroom body is the center of olfactory learning and memory in the fly4, and these gamma neurons prune their axonal and dendritic branches during pupal development and then re-extend branches at a later timepoint to establish the adult innervation pattern5. Pruning of these neurons of the MB has been shown to occur via local degeneration of neurite branches6, by a mechanism that is triggered by ecdysone, a steroid hormone, acting at the ecdysone receptor B17, and that is dependent on the activity of the ubiquitin-proteasome system6. Our method of ex vivo culturing can be used to interrogate further the mechanism of developmental remodeling. We found that in the ex vivo culture setting, gamma neurons of the MB recapitulated the process of developmental pruning with a time course similar to that in vivo. It was essential, however, to wait until 1.5 hours after puparium formation before explanting the tissue in order for the cells to commit irreversibly to metamorphosis; dissection of animals at the onset of pupariation led to little or no metamorphosis in culture. Thus, with appropriate modification, the ex vivo culture approach can be applied to study dynamic as well as steady state aspects of central brain biology.

Ex vivo Culturing of Whole, Developing Drosophila Brains

This article describes a method by which one can mimic in vivo development of the Drosophila mushroom body in an ex vivo culture system.

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for ...

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

The process by which the anterior region of the neural plate gives rise to the vertebrate retina continues to be a major focus of both clinical and basic research. In addition to the obvious medical relevance for understanding and treating retinal disease, the development of the vertebrate retina continues to serve as an important and elegant model system for understanding neuronal cell type determination and differentiation1-16. The neural retina consists of six discrete cell types (ganglion, amacrine, horizontal, photoreceptors, bipolar cells, and M&uuml;ller glial cells) arranged in stereotypical layers, a pattern that is largely conserved among all vertebrates 12,14-18.
While studying the retina in the intact developing embryo is clearly required for understanding how this complex organ develops from a protrusion of the forebrain into a layered structure, there are many questions that benefit from employing approaches using primary cell culture of presumptive retinal cells 7,19-23. For example, analyzing cells from tissues removed and dissociated at different stages allows one to discern the state of specification of individual cells at different developmental stages, that is, the fate of the cells in the absence of interactions with neighboring tissues 8,19-22,24-33. Primary cell culture also allows the investigator to treat the culture with specific reagents and analyze the results on a single cell level 5,8,21,24,27-30,33-39. Xenopus laevis, a classic model system for the study of early neural development 19,27,29,31-32,40-42, serves as a particularly suitable system for retinal primary cell culture 10,38,43-45.
Presumptive retinal tissue is accessible from the earliest stages of development, immediately following neural induction 25,38,43. In addition, given that each cell in the embryo contains a supply of yolk, retinal cells can be cultured in a very simple defined media consisting of a buffered salt solution, thus removing the confounding effects of incubation or other sera-based products 10,24,44-45.
However, the isolation of the retinal tissue from surrounding tissues and the subsequent processing is challenging. Here, we present a method for the dissection and dissociation of retinal cells in Xenopus laevis that will be used to prepare primary cell cultures that will, in turn, be analyzed for calcium activity and gene expression at the resolution of single cells. While the topic presented in this paper is the analysis of spontaneous calcium transients, the technique is broadly applicable to a wide array of research questions and approaches (Figure 1).

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

Xenopus laevis provides an ideal model system for studying cell fate specification and physiological function of individual retinal cells in primary cell culture. Here we present a technique for dissecting retinal tissues and generating primary cell cultures that are imaged for calcium activity and analyzed by in situ hybridization.

The process by which  the anterior region of the neural plate gives rise to the vertebrate retina  continues to be a major focus of both clinical and ...

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of existing mouse lines for genetic cell lineage tracing: a tamoxifen-inducible Cre line and a Cre reporter line expressing dsRed upon Cre-mediated recombination. By using a relatively low level of tamoxifen, we were able to induce recombination in a small number of cells, permitting us to follow individual cell divisions. Additionally, we observed the transcriptional response to Sonic Hedgehog (Shh) signaling using an Olig2-eGFP transgenic line 1-3 and we monitored formation of cilia by infecting the cultured slice with virus expressing the cilia marker, Sstr3-GFP 4. In order to image the neuroepithelium, we harvested embryos at E8.5, isolated the neural tube, mounted the neural slice in proper culturing conditions into the imaging chamber and performed time-lapse confocal imaging. Our ex vivo live imaging method enables us to trace single cell divisions to assess the relative timing of primary cilia formation and Shh response in a physiologically relevant manner. This method can be easily adapted using distinct fluorescent markers and provides the field the tools with which to monitor cell behavior in situ and in real time.

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

Here we develop the tools necessary for ex vivo live imaging to trace single cell divisions in the mouse E8.5 neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular recording has historically been a common technique used to study sensitivities of individual photoreceptor cells to different light stimuli that is still being used today. However, there remains a dearth of detailed methodology in the literature for researchers wishing to replicate intracellular recording experiments in the eye. Here we present the insect as a model for examining eye physiology more generally. Insect photoreceptor cells are located near the surface of the eye and are therefore easy to reach, and many of the mechanisms involved in vision are conserved across animal phyla. We describe the basic procedure for in vivo intracellular recording of photoreceptor cells in the eye of a butterfly, with the goal of making this technique more accessible to researchers with little prior experience in electrophysiology. We introduce the basic equipment needed, how to prepare a live butterfly for recording, how to insert a glass microelectrode into a single cell, and finally the recording procedure itself. We also explain the basic analysis of raw response data for determining spectral sensitivity of individual cell types. Although our protocol focuses on determining spectral sensitivity, other stimuli (e.g., polarized light) and variations of the method are applicable to this setup.

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

The electrophysiological technique of intracellular recording is demonstrated and used to determine spectral sensitivities of single photoreceptor cells in the compound eye of a butterfly.

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular ...