This work was supported by NEI intramural funds. The authors sincerely acknowledge Dr. Sheldon Miller for his scientific guidance, technical advice, and expertise in RPE physiology and disease. The authors thank Megan Kopera and the animal care staff for managing the mouse colonies, and Dr. Tarun Bansal, Raymond Zhou, and Yuan Wang for technical support.

This protocol follows the animal care guidelines outlined in the animal study protocol approved by the Animal Care and Use Committee of the National Eye Institute.
1. Importing light stimulation protocols for DC-ERG
NOTE: Follow the directions below to import the light stimulation protocols for the DC-ERG into the ERG system software (Table of Materials). The protocol consists of a 0.5 min pre-stimulus interval, followed by a step of light (10 cd/m2) for 7 min, and ending with a 1.5 min post-stimulus interval. The light intensity of 10 cd/m2 (1 log10 cd/m2) was selected since it evokes approximately half the maximal response for all the components of the DC-ERG in WT mice18,21. The c-wave and fast oscillation are of particular interest as the origins of these electrical responses are well characterized and can be isolated and studied further in vitro RPE models (e.g., iPSC-RPE). The application of other light intensities can extract additional information, for instance, the off response undergoes a reversal of polarity at brighter light stimuli and may show differences at the intensity at which this reversal takes place. The user is free to change the light intensity settings at their discretion.
<ol>
	<li>Open the ERG system software.</li>
	<li>Click on Database Center.</li>
	<li>Click on New (provide a new database file name). Click on Save. The popup box will display: &#8220;Database created, do you want to connect to the new database file.&#8221; Click on Yes. The current database name should now reflect the new file name.</li>
	<li>Click on Transfer In in the Database Control Center Window.</li>
	<li>Select Supplementary File 1: LightProtocols - TRANSFER.EXP. Click on Open.</li>
	<li>Click on Close (Database Control Center) upon completion of the progress bar.</li>
	<li>Click on the green Start button.</li>
	<li>Click on New on the Select Patient Window. Create a new patient family name describing the mouse model, enter the date of birth (DOB, mm/dd/yyyy), toggle the gender (M/F) button to the appropriate description. Click on Close to save the experimental details.</li>
	<li>Click on Protocols. The dark-adapted ERG and DC-ERG protocols should now be visible.</li>
	<li>Lastly, place the Long Flash.col file into the following folder C:\ERG User Files\Long Flash.col.</li>
</ol>
2. Capillary electrode preparation
<ol>
	<li>Cut the 1.5 mm glass capillaries in half by using a ceramic tile (Table of Materials) to score the glass and break them cleanly using a table to provide physical counterforce and to stabilize the glass. 
	NOTE: Blunt the cut ends as necessary.</li>
	<li>Using a Bunsen burner (Table of Materials) allow the heat to make a small bend in the capillary while holding it over the flame with forceps.</li>
</ol>
3. Filling capillary electrodes
<ol>
	<li>Connect the vacuum desiccator to the laboratory&#8217;s vacuum line through an in-line filter (Table of Materials).</li>
	<li>Pour 30 mL of Hanks&#8217; Balanced Salt Solution (HBSS) (Table of Materials) into an open 50 mL conical tube and place it (with the cap removed) into the vacuum chamber.</li>
	<li>Turn on the vacuum and degas the HBSS while turning on the computer and the recording equipment.</li>
	<li>After 5&#8210;10 min turn off the vacuum and use the degassed HBSS to fill a 12 mL syringe through an attached 25 G needle. Use it to fill the bases of the electrode holders taking extra precaution not to introduce bubbles.
	<ol>
		<li>To do this, remove the threaded screw cap and carefully slide the syringe needle through the silicone rubber gasket to reach the back wall (silver/silver chloride pellet) of the holder. 
		NOTE: The silver/silver chloride pellets are advantageous over holders that use silver wire as they provide more surface area resulting in a stable low-noise baseline. However, silver/silver chloride pellets require a liquid interface free from air bubbles to achieve a good connection. Therefore, take great care not to introduce air bubbles during this process.</li>
		<li>Gradually inject HBSS to fill the entire microelectrode while slowly retracting the syringe needle. Reattach the threaded cap but do not tighten. Reinsert the syringe needle to fill the empty space within the cap with HBSS. Then fill the glass capillary while holding it horizontal to prevent solution from escaping from the other end.</li>
		<li>Hold the glass capillary from the bent end and slowly insert the opposite end through the loosened cap and then tighten the screw cap into place. 
		NOTE: Glass electrodes filled with HBSS maintain lubrication of the mouse&#8217;s eye and prevent corneal dehydration that would occur with the use of standard gold loop electrodes.</li>
	</ol>
	</li>
	<li>Position the electrode holders with the capillary electrodes tilted upwards to allow bubbles to flow out. Run the vacuum for 5&#8210;10 min to degas. Gas escaping from the glass and plastic surfaces will push HBSS out from the electrode holders.</li>
	<li>Stop and then slowly release the vacuum. Refill the electrode holders and glass capillaries as described previously. 
	NOTE: Bubbles tend to collect on or near the silicone rubber gasket and can also hide in the grooves of the threaded cap, therefore special attention must be paid to keep these areas bubble-free.</li>
	<li>Install and secure the microelectrode holder into the custom-made T-clip/Magnetic ball joint stand (Figure 1A, inset). To make the custom stand for the microelectrode holder, modify a T-clip (5/16&#8221;-11/32&#8221; OD Tubing) #8 (Table of Materials) by removing the black polyacetal clips on one side. Have the cylinder base of the magnetic ball joints machined in half to adjust the height (Table of Materials). Secure the modified T-clips to the magnetic ball mounting screws with M3 sized nuts.</li>
	<li>Place the microelectrode holder into the modified T-clip and hold it tightly in place by sliding in approximately a 1-inch tapered wooden handle made from breaking a cotton tipped cleaning stick (Table of Materials) at an angle. Use the rare earth magnet cylinder base to securely position the customized electrode holder stand onto the metal plate of the stage enabling 360&#176; rotation on a 180&#176; axis.</li>
</ol>
4. Test electrodes
<ol>
	<li>Pour HBSS into a small container (e.g., the cap of the 50 mL conical tube).</li>
	<li>Gently lower the fully assembled, HBSS-filled capillary microelectrodes into the cap containing HBSS to pre-equilibrate the electrodes and place the needle ground electrode (tail/rear leg) and the Ag/AgCl sintered pellet reference electrode (mouth) in the same HBSS to complete the circuit (Figure 1A). 
	NOTE: Perform all subsequent steps under dim red light. Use a red-light flashlight to position the mouse and capillary electrodes. Remember to completely turn off all light sources prior to beginning the recording.</li>
	<li>Select or create an appropriate identifier (family name) to describe the mouse to be tested and select the DC-ERG protocol to be performed by completing the registration according to the following order.
	<ol>
		<li>Click on Protocols. Select DC-ERG. Click on Run. A dialog box will pop-up: &#8220;Current patient is XXX [DOB: XX/XX/XX) Is this correct for the test being performed?&#8221; Click on Yes. Then proceed to &#8220;Step 1/6.&#8221;</li>
	</ol>
	</li>
	<li>Close the doors to the faraday cage.</li>
	<li>Display impedance mode by clicking on Impedance and verify that the values for the mouth reference, tail ground, and recording electrodes are acceptable (see Figure 1B).</li>
	<li>Test the baseline stability (noise and drift) by clicking on Step (forward arrow) to select Step 4/6&#160;&#8220;Long Flash No Light.&#8221; 
	NOTE: The amount of drift observed when the electrodes are placed in the HBSS bath is generally less than 500 &#181;V per 80 s once they have stabilized and is equivalent to the drift observed when the electrodes are connected to the mouse. Thus, the electrical readout of the electrodes in the HBSS bath are an important indicator of the status of the electrodes. The noise, measured as peak-to-peak, is generally ~ 10&#8210;15% greater in the mouse than in the HBSS bath. This is probably due to the addition of motion artifacts from breathing.</li>
	<li>Begin viewing the traces by clicking on Preview. The traces should be low noise with a peak-to-peak amplitude &#60;200 &#181;V. A slight drift (&#60;500 &#181;V/80 s) that gradually fades to baseline is acceptable (Figure 1C).</li>
</ol>
5. Mouse and electrode positioning
<ol>
	<li>Keep the mice overnight in a well-ventilated light-tight box for dark adaptation.</li>
	<li>Anesthetize the animals by intraperitoneal injection of ketamine (80 mg/kg) and xylazine (8 mg/kg).</li>
	<li>Apply a drop of 0.5% proparacaine HCl topically to anaesthetize the cornea as well as a drop of 2.5% phenylephrine HCl and 0.5% tropicamide to dilate the pupils.</li>
	<li>Trim the mouse whiskers with scissors to prevent inadvertent twitching from disturbing the glass capillary electrodes during the recording. 
	NOTE: The DC-ERG stimulus protocol within the ERG system has several built-in stimulus routines that can be selected by clicking on the Step (forward arrow) or Step (backward arrow). Only Steps 1, 4, and 5 in the software are required to prepare and perform the DC-ERG recording.</li>
	<li>In the ERG system software verify that the correct patient is selected. Click on the green Protocols button. Under Protocol Description select DC-ERG. Then click on Run. Verify that this is the correct test being performed by clicking on Yes.</li>
	<li>Use Step 1/6 designated Red Light stimulus to turn on the red light inside the dome to help position the mouse and electrodes while observing the changes in impedance.</li>
	<li>Place the mouse on a heated recording table and carefully tent the skin of the rear leg using forceps. Hold the needle electrode firmly in one hand and insert it subcutaneously into the rear leg to secure it into place.</li>
	<li>Place the reference Ag/AgCl electrode (Table of Materials) inside the mouth so that the sintered pellet rests along the back cheek and is held in place behind the teeth. 
	NOTE: Gold wire electrodes should not be used as the mouth reference electrode as they have different impedance characteristics and increase the peak-to-peak noise.</li>
	<li>Prior to placing the capillary electrodes to the mouse&#8217;s eye, hold the electrode holder with the glass capillaries vertical, flick the electrode holder with the index finger to remove any bubbles that may have been introduced. Fill the tip with HBSS using a 25 G needle attached to a syringe and inspect to ensure that there is no air bubble trapped at the tip. Position the electrode holder stand so that the open tip of the HBSS-filled capillaries are in gentle contact with the cornea.</li>
	<li>Use special precaution to avoid introducing air bubbles by holding the lubricant eye gel dispenser inverted and discard the initial drops. Place a drop of lubricant eye gel on each eye to maintain conductivity and prevent desiccation during the recording.</li>
</ol>
6. DC-ERG recording
<ol>
	<li>Click on Step (forward arrow) to select Step 4/6 &#8220;Long Flash No Li.&#8221;</li>
	<li>Click on Impedance. Use the Impedance Checking screen to examine the resistances of the left and right eyes. The impedance values for the recording electrodes at each eye are expected to be similar (~39 k&#8486;). The impedance values for both the ground and reference electrodes are expected to be less than 10 k&#8486;).</li>
	<li>Click on Preview to view the traces for the left and right eye. Wait for a stable baseline to be achieved (&#60;10 min). Click on Stop to exit the trace preview. 
	NOTE: Abrupt changes in drift direction or aberrant noise in the recording will not improve with time and will require identifying the capillary electrode that requires attention. The most likely cause is a bubble introduced to the tip of the electrode. Refill the tip with HBSS. Click on Preview again to check the baseline.</li>
	<li>Click on Step (forward arrow) to select Step 5/6 &#8220;Long Flash 10 cd 7 min.&#34;</li>
	<li>Click Run to start the recording (Figure 1D).</li>
</ol>
7. Data export
<ol>
	<li>Select the &#8220;Patient&#8221; (Family Name) describing the mouse recording to be exported.</li>
	<li>Click on Old Tests.</li>
	<li>Under Protocol Description select DC-ERG. Click on the green Load button to load the previously acquired data.</li>
	<li>Click on Step (forward arrow) to advance to Step 5/6 &#8220;Long Flash 10 cd 7 min.&#8221;</li>
	<li>Click on Export.</li>
	<li>Provide the filename (e.g., filename.csv). A valid filename must begin with a letter, followed by letters, numbers, or underscores. Do not use special characters or hyphens. The data analysis program (DCERG_Analysis.exe) requires that table entries meet the requirements for variable names.</li>
	<li>Place a checkmark next to Data Table. Next to separator, select Tab. Place checkmarks next to Options (Titles, Vertical), Include All (Steps, Chans, Results), Data Columns (Contents, Results, Sweeps), and Format (File).</li>
	<li>Then click on Export (Figure 1E). This saves the *.csv file to the C:\Multifocal folder.</li>
</ol>
8. Data analysis
<ol>
	<li>Download and install the appropriate runtime installer (Table of Materials)</li>
	<li>Download and install the DCERG_Analysis.exe installer. 
	NOTE: This installs the script that will perform the analysis of the DC-ERG components and creates a shortcut to run the program in the Start Menu folder.</li>
	<li>Click on the shortcut created in the Start Menu &#62; Programs folder.</li>
	<li>Select the exported data file or files (*.csv) for analysis. Use Ctrl + left click on the mouse to select more than one file. 
	NOTE: The executable file generates two types of plots: 1) the raw data is plotted with a best fit line indicating the measured drift; 2) the drift corrected response is plotted after being smoothed with a moving average (with a span of ~5 s). From this plot the amplitudes and time-to-peak of the DC-ERG components are identified: c-wave, fast oscillation, light peak, and off response. The data is then exported in table format to excel where each sheet corresponds to a different mouse recording. These sheets are followed by two summary sheets: (i) compiled DC-ERG amplitudes (mV); (ii) compiled DC-ERG time-to-peaks (light onset, t = 0 min).</li>
</ol>

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The authors have nothing to disclose.

Critical Steps
A good DC-ERG recording requires stable electrodes that are free from bubbles that create artifacts and unwanted drift as they are extremely sensitive to outgassing and temperature changes. It is essential that a stable baseline is achieved when the electrodes are placed in the HBSS bath solution before proceeding forward with the mouse recording. Small bubbles tend to collect at the base of the capillary electrode or around the silicone gasket and are difficult...

The retinal pigment epithelium (RPE) is a monolayer of specialized cells that line the posterior segment of the eye and exert critical functions to maintain retinal homeostasis1. The RPE supports photoreceptors by regenerating their photon-capturing visual pigment in a process called the visual cycle2, by participating in the diurnal phagocytosis of shed outer segment tips3, and in the transport of nutrients and metabolic products between photoreceptors and the choriocapillaris4,5. Abnormalities in RPE function underlie numerous human retinal diseases, such as age-related macular degeneration6, Leber&#8217;s congenital amaurosis7,8 and Best vitelliform macular dystrophy9. As donor eye tissues are often difficult to obtain solely for research purposes, animal models with genetic modifications can provide an alternative way to study the development of retinal diseases10,11. Additionally, the emergence and application of CRISPR cas9 technology now permits genomic introductions (knock-in) or deletions (knock-out) in a simple, one-step process surpassing limitations of prior gene targeting technologies12. The boom in the availability of new mouse models13 necessitates a more efficient recording protocol to non-invasively evaluate RPE function.
Measurement of the light-evoked electrical responses of the RPE can be achieved using a direct-coupled electroretinogram (DC-ERG) technique. When used in combination with conventional ERG recordings that measure the photoreceptor (a-wave) and bipolar (b-wave) cell responses14, the DC-ERG can define how the response properties of the RPE change with retinal degeneration15,16,17 or whether RPE dysfunction precedes photoreceptor loss. This protocol describes a method adapted from the work of Marmorstein, Peachey, and colleagues who first developed the DC-ERG technique16,18,19,20 and improves upon the reproducibility and ease of use.
The DC-ERG recording is difficult to perform because of the long acquisition time (9 min) during which any interruption or introduction of noise can complicate the interpretation of the data. The advantage of this new method is that the baselines reach steady state within a shorter amount of time reducing the likelihood that the animal will awaken prematurely from anesthesia and is less prone to bubble formation in the capillary electrodes.

<table><tbody><tr><td>Ag/AgCl (mouth) Electrode</td><td>WPI Inc</td><td>EP1</td><td>Mouth reference electrode for mouse</td></tr><tr><td>Ceramic Tile</td><td>Sutter Instrument</td><td>CTS</td><td>Used to cut the glass capillary tube to an appropriate size</td></tr><tr><td>Cotton Tipped Cleaning Stick</td><td>Puritan Medical Products</td><td>867-WC No Glue</td><td>To be used as a spacer to improve the fit of the electrode holder assembly</td></tr><tr><td>Electroretinogram (ERG) System</td><td>Diagnosys LLC</td><td>E3 System</td><td>Visual electrophysiology system to diagnose ophthalmic conditions in vision research and drug trials</td></tr><tr><td>Bunsen Burner</td><td>Argos Technologies</td><td>BW20002460</td><td>Or equivlaent to shape glass under flame</td></tr><tr><td>Glass Capillary Tube (1.5 mm)</td><td>Sutter Instruments</td><td>BF150-75</td><td>For filling with HBSS and making contact to the cornea</td></tr><tr><td>Hank&amp;rsquo;s Buffered Salt Solution (HBSS)</td><td>Thermo Fisher Scientific Inc</td><td>14175-095</td><td>Commercially available. Maintain at RT</td></tr><tr><td>In-Line Filter</td><td>Whatman</td><td>6722-5001</td><td>To protect vacuum pump from aerosols</td></tr><tr><td>Low Noise Cable for Microelectrode Holders</td><td>WPI Inc</td><td>5372</td><td>Suggested for improving the length and placement of the cables and electrode holder assemblies</td></tr><tr><td>Magnetic Ball Joint</td><td>WPI Inc</td><td>500871</td><td>For magnetically positioning the electrode holder assembly on the stage</td></tr><tr><td>MatLab</td><td>Mathworks</td><td></td><td>MatLab: For editing the analysis software</td></tr><tr><td>MatLab Curvefit Toolbox</td><td>Mathworks</td><td></td><td>Toolbox for MatLab (only required for editing the analysis software)</td></tr><tr><td>MatLab Compiler</td><td>Mathworks</td><td></td><td>Toolbox for MatLab (only required for editing and re-releasing the analysis software)</td></tr><tr><td>MatLab Runtime version 9.5</td><td>Mathworks</td><td>R2018b (9.5)</td><td>Required to run the analysis software: https://www.mathworks.com/products/compiler/matlab-runtime.html</td></tr><tr><td>Microelectrode Holders (45 degrees)</td><td>WPI Inc</td><td>MEH345-15</td><td>For holding the capillaries</td></tr><tr><td>Needle (25 ga)</td><td>Covidien</td><td>8881250313</td><td>For filling the capillary tubes with HBSS</td></tr><tr><td>Needle (ground) Electrode</td><td>Rhythmlink</td><td>13mm - one elctrode</td><td>Subdermal needle electrode (ground) for mouse (13mm long, 0.4mm diameter needle, 1.5m leadwire)</td></tr><tr><td>Regulator/Power Conditioner</td><td>Furman</td><td>P-1800</td><td>Or equivalent to remove DC-offset from noise introduced through power line</td></tr><tr><td>Syringe (12 mL)</td><td>Monoject</td><td>1181200777</td><td>For filling the capillary tubes with HBSS</td></tr><tr><td>T-clip</td><td>Cole-Parmer</td><td>06852-20</td><td>For electrode holder assembly</td></tr><tr><td>Vacuum Desiccator</td><td>Bel-Art</td><td>420120000</td><td>Clear polycarbonate bottom &amp;amp; cover</td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Pharmacological treatment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Lubricant eye gel</td><td>Alcon</td><td>0078-0429-47</td><td>Helps lubricate corneal surface and maintain electrical contact with capillary electrodes</td></tr><tr><td>Phenylephrine Hydrochloride 2.5%</td><td>Akorn</td><td>17478-201-15</td><td>Short acting mydriatic eye drops (for pupil dilation)</td></tr><tr><td>Proparacaine Hydrochloride 0.5%</td><td>Akorn</td><td>17478-263-12</td><td>Local anesthetic for ophthalmic instillation</td></tr><tr><td>Tropicamide 0.5%</td><td>Akorn</td><td>17478-101-12</td><td>Short acting mydriatic eye drops (for pupil dilation)</td></tr><tr><td>Xylazine</td><td>AnaSed</td><td>sc-362949Rx</td><td>Analgesic and muscle relaxant</td></tr><tr><td>Zetamine (Ketamine HCl)</td><td>VetOne</td><td>501072</td><td>Anesthetic for intramuscular injections</td></tr></tbody></table>

direct-coupled electroretinogram (dc-erg) for recording the light-evoked electrical responses of the mouse retinal pigment epithelium

The retinal pigment epithelium (RPE) is a specialized monolayer of cells strategically located between the retina and the choriocapillaris that maintain the overall health and structural integrity of the photoreceptors. The RPE is polarized, exhibiting apically and basally located receptors or channels, and performs vectoral transport of water, ions, metabolites, and secretes several cytokines.
In vivo noninvasive measurements of RPE function can be made using direct-coupled ERGs (DC-ERGs). The methodology behind the DC-ERG was pioneered by Marmorstein, Peachey, and colleagues using a custom-built stimulation recording system and later demonstrated using a commercially available system. The DC-ERG technique uses glass capillaries filled with Hank&#8217;s buffered salt solution (HBSS) to measure the slower electrical responses of the RPE elicited from light-evoked concentration changes in the subretinal space due to photoreceptor activity. The prolonged light stimulus and length of the DC-ERG recording make it vulnerable to drift and noise resulting in a low yield of useable recordings. Here, we present a fast, reliable method for improving the stability of the recordings while reducing noise by using vacuum pressure to reduce/eliminate bubbles that result from outgassing of the HBSS and electrode holder. Additionally, power line artifacts are attenuated using a voltage regulator/power conditioner. We include the necessary light stimulation protocols for a commercially available ERG system as well as scripts for analysis of the DC-ERG components: c-wave, fast oscillation, light peak, and off response. Due to the improved ease of recordings and rapid analysis workflow, this simplified protocol is particularly useful in measuring age-related changes in RPE function, disease progression, and in the assessment of pharmacological intervention.

Figure 2 is a sample dataset from miR-204 ko/ko cre/+ (conditional KO) and wild type (WT) mice. MiR-204 ko/ko cre/+ are mice with a conditional knockout of microRNA 204 in the retinal pigment epithelium. These mice are generated by crossing floxed miR-204 mice (produced by NEIGEF)22 with VMD2-CRE mice23. MiR-204 is highly expressed in the RPE where it regulates the expression of proteins critical for epithelial function that maintain tight junc...

Watch this Scientific Journal Video about Direct-Coupled Electroretinogram DC-ERG for Recording the Light-Evoked Electrical Responses of the Mouse Retinal Pigment Epithelium at JoVE.com

Direct-Coupled Electroretinogram DC-ERG for Recording the Light-Evoked Electrical Responses of the Mouse Retinal Pigment Epithelium

Here, we present a method for recording light-evoked electrical responses of the retinal pigment epithelium (RPE) in mice using a technique known as DC-ERGs first described by Marmorstein, Peachey, and colleagues in the early 2000s.

Direct-Coupled Electroretinogram (DC-ERG) for Recording the Light-Evoked Electrical Responses of the Mouse Retinal Pigment Epithelium

The retinal pigment epithelium (RPE) is a specialized monolayer of cells strategically located between the retina and the choriocapillaris that maintain ...

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Research

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Chemistry

5.1K Views.  National Institutes of Health. Here, we present a method for recording light-evoked electrical responses of the retinal pigment epithelium (RPE) in mice using a technique known as DC-ERGs first described by Marmorstein, Peachey, and colleagues in the early 2000s.

Video: Direct-Coupled Electroretinogram DC-ERG for Recording the Light-Evoked Electrical Responses of the Mouse Retinal Pigment Epithelium

An Isolated Retinal Preparation to Record Light Response from Genetically Labeled Retinal Ganglion Cells

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1. This information is then segregated into what are known as the ON and OFF pathways. The photoreceptors signal light information to the bipolar cells (BCs), which depolarize in response to increases (On BCs) or decreases (Off BCs) in light intensity. This segregation of light information is maintained at the level of the retinal ganglion cells (RGCs), which have dendrites stratifying in either the Off sublamina of the inner plexiform layer (IPL), where they receive direct excitatory input from Off BCs, or stratifying in the On sublamina of the IPL, where they receive direct excitatory input from On BCs. This segregation of information regarding increases or decreases in illumination (the On and Off pathways) is conserved and signaled to the brain in parallel.
The RGCs are the output cells of the retina, and are thus an important cell to study in order to understand how light information is signaled to visual nuclei in the brain. Advances in mouse genetics over recent decades have resulted in a variety of fluorescent reporter mouse lines where specific RGC populations are labeled with a fluorescent protein to allow for identification of RGC subtypes 2 3 4 and specific targeting for electrophysiological recording. Here, we present a method for recording light responses from fluorescently labeled ganglion cells in an intact, isolated retinal preparation. This isolated retinal preparation allows for recordings from RGCs where the dendritic arbor is intact and the inputs across the entire RGC dendritic arbor are preserved. This method is applicable across a variety of ganglion cell subtypes and is amenable to a wide variety of single-cell physiological techniques.

This article provides a description of how to dissect and record from the isolated retinal preparation in mouse. In particular, we describe how to record light responses from a fluorescently labeled ganglion cell population and subsequently identify and analyze its morphology.

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1.  This information is then ...

Neuroscience

Transretinal ERG Recordings from Mouse Retina: Rod and Cone Photoresponses

There are two distinct classes of image-forming photoreceptors in the vertebrate retina: rods and cones. Rods are able to detect single photons of light whereas cones operate continuously under rapidly changing bright light conditions. Absorption of light by rod- and cone-specific visual pigments in the outer segments of photoreceptors triggers a phototransduction cascade that eventually leads to closure of cyclic nucleotide-gated channels on the plasma membrane and cell hyperpolarization. This light-induced change in membrane current and potential can be registered as a photoresponse, by either classical suction electrode recording technique1,2 or by transretinal electroretinogram recordings (ERG) from isolated retinas with pharmacologically blocked postsynaptic response components3-5. The latter method allows drug-accessible long-lasting recordings from mouse photoreceptors and is particularly useful for obtaining stable photoresponses from the scarce and fragile mouse cones. In the case of cones, such experiments can be performed both in dark-adapted conditions and following intense illumination that bleaches essentially all visual pigment, to monitor the process of cone photosensitivity recovery during dark adaptation6,7. In this video, we will show how to perform rod- and M/L-cone-driven transretinal recordings from dark-adapted mouse retina. Rod recordings will be carried out using retina of wild type (C57Bl/6) mice. For simplicity, cone recordings will be obtained from genetically modified rod transducin &alpha;-subunit knockout (T&alpha;-/-) mice which lack rod signaling8.

We describe a relatively simple method of transretinal electroretinogram (ERG) recordings for obtaining rod and cone photoresponses from intact mouse retina. This approach takes advantage of the block of synaptic transmission from photoreceptors to isolate their light responses and record them using field electrodes placed across the isolated flat-mounted retina.

There are two distinct classes of image-forming photoreceptors in the vertebrate retina: rods and cones. Rods are able to detect single photons of light ...

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment are open and allow cations to flow steadily inwards across the membrane, depolarizing the cell. Light exposure triggers the closure of the CNG channels, blocks the inward cation current flow, and thus results in cell hyperpolarization. Based on the polarity of photoreceptors, a suction recording method was developed in 1970s that, unlike the classic patch-clamp technique, does not require penetrating the plasma membrane 1. Drawing the outer segment into a tightly-fitting glass pipette filled with extracellular solution allows recording the current changes in individual cells upon test-flash exposure. However, this well-established "outer-segment-in (OS-in)" suction recording is not suitable for mouse cone recordings, because of the low percentage of cones in the mouse retina (3%) and the difficulties in identifying the cone outer segments. Recently, an inner-segment-in (IS-in) recording configuration was developed to draw the inner segment/nuclear region of the photoreceptor into the recording pipette 2,3. In this video, we will show how to record from individual mouse cone photoresponses using single-cell suction electrode.

We will show how to record flash responses from single mouse cones using a suction electrode.

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment ...

Biology

Using the Electroretinogram to Assess Function in the Rodent Retina and the Protective Effects of Remote Limb Ischemic Preconditioning

The ERG is the sum of all retinal activity. The ERG is usually recorded from the cornea, which acts as an antenna that collects and sums signals from the retina. The ERG is a sensitive measure of changes in retinal function that are pan-retinal, but is less effective for detecting damage confined to a small area of retina. In the present work we describe how to record the &#8216;flash&#8217; ERG, which is the potential generated when the retina is exposed to a brief light flash. We describe methods of anaesthesia, mydriasis and corneal management during recording; how to keep the retina dark adapted; electrode materials and placement; the range and calibration of stimulus energy; recording parameters and the extraction of data. We also describe a method of inducing ischemia in one limb, and how to use the ERG to assess the effects of this remote-from-the-retina ischemia on retinal function after light damage. A two-flash protocol is described which allows isolation of the cone-driven component of the dark-adapted ERG, and thereby the separation of the rod and cone components. Because it can be recorded with techniques that are minimally invasive, the ERG has been widely used in studies of the physiology, pharmacology and toxicology of the retina. We describe one example of this usefulness, in which the ERG is used to assess the function of the light-damaged retina, with and without a neuroprotective intervention; preconditioning by remote ischemia.

The electroretinogram (ERG) is an electrical potential generated by the retina in response to light. This paper describes how to use the ERG to assess retinal function, in dark-adapted rats, and how it can be can be used to assess a neuroprotective intervention, in the present case remote ischemic preconditioning.

The ERG is the sum of all retinal activity. The ERG is usually recorded from the cornea, which acts as an antenna that collects and sums signals from the ...

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 ...

Simultaneous Recording of Electroretinography and Visual Evoked Potentials in Anesthetized Rats

The electroretinogram (ERG) and visual evoked potential (VEP) are commonly used to assess the integrity of the visual pathway. The ERG measures the electrical responses of the retina to light stimulation, while the VEP measures the corresponding functional integrity of the visual pathways from the retina to the primary visual cortex following the same light event. The ERG waveform can be broken down into components that reflect responses from different retinal neuronal and glial cell classes. The early components of the VEP waveform represent the integrity of the optic nerve and higher cortical centers. These recordings can be conducted in isolation or together, depending on the application. The methodology described in this paper allows simultaneous assessment of retinal and cortical visual evoked electrophysiology from both eyes and both hemispheres. This is a useful way to more comprehensively assess retinal function and the upstream effects that changes in retinal function can have on visual evoked cortical function.

This protocol describes simultaneous measurement of electroretinogram and visual evoked potentials in anesthetized rats.

The electroretinogram (ERG) and visual evoked potential (VEP) are commonly used to assess the integrity of the visual pathway. The ERG measures the ...

Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors

The Drosophila G-protein-coupled photopigment rhodopsin (R) is composed of a protein (opsin) and a chromophore. The activation process of rhodopsin is initiated by photon absorption-inducing isomerization of the chromophore, promoting conformational changes of the opsin and resulting in a second dark-stable photopigment state (metarhodopsin, M). Investigation of this bi-stable photopigment using random mutagenesis requires simple and robust methods for screening mutant flies. Therefore, several methods for measuring reductions in functional photopigment levels have been designed. One such method exploits the charge displacements within the photopigment following photon absorption and the huge amounts of photopigment molecules expressed in the photoreceptors. This electrical signal, named the early receptor potential (or early receptor current), is measured by a variety of electrophysiological methods (e.g., electroretinogram and whole-cell recordings) and is linearly proportional to functional photopigment levels. The advantages of this method are the high signal-to-noise ratio, direct linear measurement of photopigment levels, and independence of phototransduction mechanisms downstream to rhodopsin or metarhodopsin activation. An additional electrophysiological method called prolonged depolarizing afterpotential (PDA) exploits the bi-stability of Drosophila photopigment and the absorption-spectral differences of fly R and M pigment states. The PDA is induced by intense blue light, converting saturating amounts of rhodopsin to metarhodopsin, resulting in the failure of light-response termination for an extended time in darkness, but it can be terminated by metarhodopsin to rhodopsin conversion using intense orange light. Since the PDA is a robust signal that requires massive photopigment conversion, even small defects in the biogenesis of the photopigment lead to readily detected abnormal PDA. Indeed, defective PDA mutants led to the identification of novel signaling proteins important for phototransduction.

Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors

We present a protocol to electrophysiologically characterize bi-stable photopigments: (i) exploiting the charge displacements within the photopigment molecules following photon-absorption and their huge amount in the photoreceptors, and (ii) exploiting the absorption-spectra differences of rhodopsin and metarhodopsin photopigment states. These protocols are useful to screen for mutations affecting bi-stable photopigment systems.&#160;

The Drosophila G-protein-coupled photopigment rhodopsin (R) is composed of a protein (opsin) and a chromophore. The activation process of rhodopsin is ...

Light-Evoked Postsynaptic Responses in Neurons of Mouse Retinal Slices

Source: Hellmer, C. B., et al. <a href="https://app.jove.com/v/52422">Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques.</a> J. Vis. Exp. (2015).
This video demonstrates the recording of light-evoked post-synaptic responses in a dark-adapted mouse retinal tissue slice preparation. The retinal slice is placed in a recording chamber, and a recording pipette is used to establish electrical continuity with a retinal ganglion cell. Light pulses are applied to generate excitatory post-synaptic potentials in the ganglion cells, which are recorded by the pipette.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Preparation of Experimental ...

JoVE Encyclopedia of Experiments

Neurophysiology

Imaging and Optical Techniques

Direct-Coupled Electroretinogram Recording of the Retinal Pigment Epithelium in a Mouse

Source: Miyagishima, K. J., et al. <a href="https://app.jove.com/v/61491">Direct-Coupled Electroretinogram (DC-ERG) for Recording the Light-Evoked Electrical Responses of the Mouse Retinal Pigment Epithelium.</a> J. Vis. Exp. (2020).
This video demonstrates the recording of a direct-coupled electroretinogram (DC-ERG) on a dark-adapted, anesthetized mouse retinal pigment epithelium (RPE). After applying dilating drops, the animal is placed in a recording setup, and the capillary electrode filled with a salt buffer is placed on the cornea. After establishing the baseline, DC-ERG is recorded.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Mouse ...

Electrophysiological Recording of Voltage Responses of Drosophila Retinal Photoreceptors to Light Stimuli

Source: Juusola, M., et al. <a href="https://app.jove.com/v/54142">Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo.</a> J. Vis. Exp. (2016).
This video outlines the protocol for electrophysiological recording of photoreceptor activity in the eye of an immobilized Drosophila, enabling precise measurement of voltage changes in response to light stimuli.

1. Recording from R1-R6 Photoreceptors 

	Always be grounded when operating the microelectrode amplifier (for example by touching the metal surface of the ...

Direct-Coupled Electroretinogram (DC-ERG) for Recording the Light-Evoked Electrical Responses of the Mouse Retinal Pigment Epithelium

Summary

Explore More Videos

Chapters in this video

An Isolated Retinal Preparation to Record Light Response from Genetically Labeled Retinal Ganglion Cells

Transretinal ERG Recordings from Mouse Retina: Rod and Cone Photoresponses

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Using the Electroretinogram to Assess Function in the Rodent Retina and the Protective Effects of Remote Limb Ischemic Preconditioning

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

Simultaneous Recording of Electroretinography and Visual Evoked Potentials in Anesthetized Rats

Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors

Light-Evoked Postsynaptic Responses in Neurons of Mouse Retinal Slices

Direct-Coupled Electroretinogram Recording of the Retinal Pigment Epithelium in a Mouse

Electrophysiological Recording of Voltage Responses of Drosophila Retinal Photoreceptors to Light Stimuli