This work was supported by National Eye Institute grants EY02665 and EY031706 and International Retinal Research Foundation to Dr. Vinberg, National Institutes of Health Core Grant (EY014800), and an Unrestricted Grant from Research to Prevent Blindness, New York, NY, to the Department of Ophthalmology &#38; Visual Sciences, University of Utah. Dr. Frans Vinberg is also a recipient of a Research to Prevent Blindness/Dr. H. James and Carole Free Career Development Award, and Dr. Silke Becker of an ARVO EyeFind grant. We thank Dr. Anne Hanneken from The Scripps Research Institute for providing the donor eye used for recordings shown in Figure 2E.

All experiments using mice were conducted in accordance with the NIH Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Studies Committee at the University of Utah. Pig eyes for demonstration of this video were obtained postmortem from the slaughterhouse (Sustainable Swine Resources, Johnsonville). Eyes were obtained from human donors after brain or cardiac death with consent for research use through the Utah Lions Eye Bank, the San Diego Eye Bank or Lifesharing, which are fully accredited by the FDA, the Association of Organ Procurement Organizations (AOPO) and the Eye Banks of America Association. The use of human donor eyes had exempt status at the University of Utah (IRB no. 00106658) and ScrippsHealth IRB (IRB no. 16-6781).
1. Setting up the ex vivo ERG
<ol>
	<li>To convert a multielectrode array setup, connect the ex vivo ERG specimen holder via a head stage to a differential amplifier, which is plugged into the analog input of the interface board of the multielectrode array system. Use the recording software for the multielectrode array to record and store the input data from the ex vivo ERG. Set the gain of the differential amplifier to 100 and add an additional 10x voltage amplification depending on the digitizer specifications. Set the low pass filter to 100 Hz.</li>
	<li>To convert a patch clamp setup, connect the ex vivo ERG specimen holder via a head stage to a differential amplifier, which is connected to the head stage of the patch clamp amplifier. Use the patch clamp system software and digitizer to record and store the input data from the ex vivo ERG. Set the gain of the differential amplifier to 100 and apply an additional 10x voltage amplification via the patch clamp head stage. Set the low pass filter to 100 Hz.</li>
	<li>Connect LEDs with the appropriate wavelengths (e.g., approximately 530 nm to elicit rod photoreceptor and ON-bipolar cell responses) to the microscope. Control the LEDs with recording software that enables the triggering of light stimuli. To control light stimuli, use an LED driver controlled by analog outputs from a digitizer.</li>
	<li>Calibrate the light output of the LEDs at the position of the retina in the specimen holder using a photodiode. If necessary, insert neutral density filters&#160;into the light path to dim the light intensity.</li>
	<li>Use a commercially available or custom built ex vivo ERG specimen holder. 
	NOTE: Drawings for CNC machining from polycarbonate can be obtained from the authors upon request.</li>
	<li>To prepare the electrode, insert an Ag/AgCl pellet electrode into a threaded luer connector. Fill the inside of the luer connector with hot glue and insert a 2 mm socket into the hot glue from the non-threaded side. Solder a silver wire of the EP1 electrode to the 2 mm socket. Screw the finished electrode, with an o-ring on the thread, into the electrode ports of the ex vivo ERG specimen holder. 
	&#8203;NOTE: Electrodes can be used for a long time. If the pellet surface accumulates dirt and/or gets dark (this can cause high offset voltage and/or electrical drift), it can be &#34;polished&#34; using fine sandpaper.</li>
	<li>At least 1 day prior to experimentation, glue filter papers onto the domes of the ex vivo ERG specimen holder using epoxy glue, ensuring that the glue does not obstruct the electrode channel (see 3 for video).</li>
</ol>
2.&#160; Animal preparation
<ol>
	<li>Dark adapt animals for at least 6 h or overnight.</li>
</ol>
3. Equipment preparation
<ol>
	<li>Fill the perfusion line of the&#160;ex vivo&#160;ERG with Ames&#39; medium bubbled with 5% carbon dioxide and 95% oxygen. Connect the perfusion line to a heating element with a DC current source or heat controller near the specimen holder to warm the Ames&#39; medium, so that the retina is kept at approximately 35-38 &#176;C.</li>
	<li>Fill both halves of the&#160;ex vivo&#160;ERG specimen holder with electrode solution, seal the perfusion line with luer stoppers, connect the electrodes, and assemble the specimen holder with four screws (see&#160;3&#160;for video).</li>
	<li>Check the resistance and offset voltage between the electrodes in the assembled specimen holder by inserting the probes of the multimeter into the electrodes. 
	NOTE: If there are no blockages, and electrodes are in good condition, the resistance should be below 100 k&#937; and offset voltage below 5 mV.</li>
	<li>Connect the assembled specimen holder to the perfusion line and place onto the stage of a microscope set up to deliver light flashes. Ensure that the specimen holder and perfusion line do not contain any air bubbles.</li>
</ol>
4. Tissue preparation
<ol>
	<li>Sacrifice the animal with CO2 and immediately enucleate the eyes, or obtain large animal or human donor eyes.</li>
	<li>Clean the globe of the remaining connective tissue and extraocular muscles. Trim off the optic nerve.</li>
	<li>On filter paper, carefully place a small incision with a razor blade along the ora serrata, approximately 1 mm from the limbus in the mouse eye. Insert fine vannas scissors into the incision and cut along the limbus to remove the anterior portion of the eye with the lens.</li>
	<li>Move the eye cup into a dish containing Ames&#39; medium. Grasp the sclera with fine forceps, taking care to not touch the retina. Insert vannas scissors between the retina and the sclera and cut the sclera from the peripheral toward the central part. Take care not to touch or damage the retina.</li>
	<li>Immobilize the sclera by holding one side of the incision with vannas scissors against the bottom of the dissection dish. Grasp the sclera with forceps on the other side of the incision. Remove the sclera without touching or damaging the retina by pulling apart the sclera on either side of the incision to allow isolation of the retina with minimal damage to the tissue.</li>
	<li>Remove the anterior segment with the lens from the fellow eye and store the eye cup at room temperature in Ames&#39; solution bubbled with 5% carbon dioxide and 95% oxygen. 
	NOTE: Functional light responses from eyes stored in this way can be obtained for several hours.</li>
	<li>In large eyes, including human donor eyes, clean the globe of the remaining connective tissue and remove the anterior segment and lens, similarly to the procedure described for the mouse eye. Use a scalpel to make a cut approximately 3 mm from the limbus. Insert curved dissection scissors into the incision and cut along the limbus to remove the anterior portion of the eye with the lens. Obtain retinal specimens for ex vivo electroretinography with a 3-6 mm disposable biopsy punch.</li>
</ol>
5. Mounting the tissue on the specimen holder
<ol>
	<li>Place the lower half of the specimen holder into a large Petri dish and fill with oxygenated Ames&#39; medium so the dome of the specimen holder is just submerged.</li>
	<li>Carefully grasp the edge of the retina with fine forceps and transfer the retina onto the dome of the ex vivo specimen holder, photoreceptor side facing up.</li>
	<li>Lift the specimen holder from the Ames&#39; solution, taking care that the retina stays in place.</li>
	<li>Completely dry the plate of the specimen holder to minimize noise, electrical crosstalk, and signal shunting.</li>
	<li>Assemble both halves of the specimen holder with four screws and connect the perfusion line. Dry the electrode in the lower half of the specimen holder and connect the anode cable to the inner retinal side and the cathode cable to the photoreceptor side.</li>
	<li>Perfuse the specimen holder with at least 1 mL/min of oxygenated Ames&#39; medium for 10-20 min to allow time for light responses to stabilize.</li>
</ol>
6. Recording retinal neuronal cell function
<ol>
	<li>Record response families by exposing the retina to light flashes of increasing intensity. For example, record mouse rod photoreceptor response families with light intensities ranging from approximately 10 to 1,000 photons/&#181;m2 and those from mouse ON-bipolar cells with light intensities ranging from approximately 0.6 to 20 photons/&#181;m2.</li>
	<li>Measure photoreceptor light responses (a-wave) in the presence of 100 &#181;M barium chloride, which blocks potassium channels in M&#252;ller glial cells, and 40 &#181;M DL-AP4, which blocks glutamatergic signal transmission to ON-bipolar cells (Figure 2B).</li>
	<li>To isolate the b-wave, which originates from the function of the ON-bipolar cells, first record combined light responses from both photoreceptor and ON-bipolar cells in the presence of barium chloride alone (Figure 2A). Then, perfuse for 5-10 min with Ames&#39; medium containing barium chloride, as well as DL-AP4, and record photoreceptor responses to the same light stimuli as before (Figure 2B). Subtract photoreceptor responses from combined photoreceptor and ON-bipolar cell responses, thus calculating ON-bipolar cell light responses alone (Figure 2C).</li>
</ol>
7. Optimizing ON-bipolar cell function
NOTE: The b-wave, which originates from the ON-bipolar cells, is highly sensitive to the temperature in the specimen holder and the perfusion rate.
<ol>
	<li>Maintain a perfusion rate of at least 0.5 mL/min to obtain the b-wave. 
	NOTE: A higher perfusion rate of 1-2 mL/min is preferable to maintain a large and stable response from ON-bipolar cells.</li>
	<li>Ensure that for a given perfusion rate, the temperature in the specimen holder near the retina is close to body temperature (i.e., approximately 35-38 &#176;C). 
	NOTE: It is important adjust the temperature of the perfusate, which is heated before it reaches the ex vivo ERG specimen holder, so that it is within the optimal temperature range at the retina.</li>
	<li>Store eye cups for later experimentation protected from light and in oxygenated Ames&#39; medium at room temperature to maintain normal a- and b-waves for several hours.</li>
</ol>

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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Principles and Practice of Clinical Electrophysiology of Vision. , (2006).</li><li>Vinberg, F., Kolesnikov, A. V., Kefalov, V. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ex+vivo+ERG+analysis+of+photoreceptors+using+an+in+vivo+ERG+system.">Ex vivo ERG analysis of photoreceptors using an in vivo ERG system.</a> Vision Research. 101, 108-117 (2014).</li><li>Winkler, B. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+and+the+fast+and+slow+components+of+P3+of+the+electroretinogram+of+the+isolated+rat+retina.">Calcium and the fast and slow components of P3 of the electroretinogram of the isolated rat retina.</a> Vision Research. 14 (1), 9-15 (1974).</li><li>Green, D. G., Kapousta-Bruneau, N. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+dissection+of+the+electroretinogram+from+the+isolated+rat+retina+with+microelectrodes+and+drugs.">A dissection of the electroretinogram from the isolated rat retina with microelectrodes and drugs.</a> Visual Neuroscience. 16 (4), 727-741 (1999).</li><li>Abbas, F., 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=Revival+of+light+signalling+in+the+postmortem+mouse+and+human+retina.">Revival of light signalling in the postmortem mouse and human retina.</a> Nature. , (2022).</li><li>Becker, S., Carroll, L. S., Vinberg, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diabetic+photoreceptors:+Mechanisms+underlying+changes+in+structure+and+function.">Diabetic photoreceptors: Mechanisms underlying changes in structure and function.</a> Visual Neuroscience. 37, (2020).</li><li>Kantola, L., Piccolino, M., Wade, N. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+action+of+light+on+the+retina:+Translation+and+commentary+of+Holmgren+(1866).">The action of light on the retina: Translation and commentary of Holmgren (1866).</a> Journal of the History of the Neurosciences. 28 (4), 399-415 (2019).</li><li>Frank, R. N., Dowling, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rhodopsin+photoproducts:+effects+on+electroretinogram+sensitivity+in+isolated+perfused+rat+retina.">Rhodopsin photoproducts: effects on electroretinogram sensitivity in isolated perfused rat retina.</a> Science. 161 (3840), 487-489 (1968).</li><li>Hamasaki, D. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+sodium+ion+concentration+on+the+electroretinogram+of+the+isolated+retina+of+the+frog.">The effect of sodium ion concentration on the electroretinogram of the isolated retina of the frog.</a> Journal of Physiology. 167 (1), 156-168 (1963).</li><li>Granit, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+components+of+the+retinal+action+potential+in+mammals+and+their+relation+to+the+discharge+in+the+optic+nerve.">The components of the retinal action potential in mammals and their relation to the discharge in the optic nerve.</a> Journal of Physiology. 77 (3), 207-239 (1933).</li><li>Donner, K., Hemila, S., Koskelainen, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temperature-dependence+of+rod+photoresponses+from+the+aspartate-treated+retina+of+the+frog+(Rana+temporaria).">Temperature-dependence of rod photoresponses from the aspartate-treated retina of the frog (Rana temporaria).</a> Acta Physiologica Scandinavica. 134 (4), 535-541 (1988).</li><li>Green, D. G., Kapousta-Bruneau, N. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+properties+of+a+new+isolated+rat+retina+preparation.">Electrophysiological properties of a new isolated rat retina preparation.</a> Vision Research. 39 (13), 2165-2177 (1999).</li><li>Luke, 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=The+isolated+perfused+bovine+retina--a+sensitive+tool+for+pharmacological+research+on+retinal+function.">The isolated perfused bovine retina--a sensitive tool for pharmacological research on retinal function.</a> Brain Research Protocols. 16 (1-3), 27-36 (2005).</li><li>Becker, S., Carroll, L. S., Vinberg, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rod+phototransduction+and+light+signal+transmission+during+type+2+diabetes.">Rod phototransduction and light signal transmission during type 2 diabetes.</a> BMJ Open Diabetes Research and Care. 8 (1), 001571 (2020).</li><li>Nymark, S., Haldin, C., Tenhu, H., Koskelainen, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+method+for+measuring+free+drug+concentration:+retinal+tissue+as+a+biosensor.">A new method for measuring free drug concentration: retinal tissue as a biosensor.</a> Investigative Ophthalmology &amp; Visual Science. 47 (6), 2583-2588 (2006).</li><li>Winkler, B. S., Kapousta-Bruneau, N., Arnold, M. J., Green, D. 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None of the authors has any conflicts of interest to disclose.

Originally developed in 1865 by Holmgren to measure retinal light responses from the amphibian retina10, technical constraints initially prevented the ERG from being widely used. Nevertheless, seminal studies by Ragnar Granit and others identified the cellular origins of the ERG and measured photoreceptor and ON-bipolar cell responses ex vivo11,12,13. Since then, refined methods have allowed more widespread use of ex vivo ERG recordings14,15, although response amplitudes, particularly those from ON-bipolar cells, remained comparatively small16. To overcome these limitations, retinal function is today more commonly measured with in vivo ERG, despite experimental constraints and more complicated interpretation of the recorded waveforms. Although the ex vivo ERG attempts to replicate physiological conditions as closely as possible, it nevertheless measures retinal function in an artificial environment and in the absence of systemic factors and pathological changes. However, when combined with the in vivo ERG, this limitation can be used to answer important questions, such as whether changes in retinal function in disease are intrinsic to retinal cells or caused by systemic changes17.
Recently, newly designed ex vivo ERG specimen holders have simplified the methodology, making the ex vivo ERG accessible to a broader research community2,3,5,18. Modifications to existing patch clamp or multielectrode array equipment described in this protocol will enable more laboratories to perform ex vivo ERG with minimal financial investments and space requirements. In particular, recent developments in ex vivo ERG technology have amplified the responsiveness of mammalian isolated retinas and resulted in a superior signal-to-noise ratio, which remains good despite additional amplification steps described in this protocol. Nevertheless, the decline of the light responses, mainly from ON-bipolar cells19, has perhaps hampered the more widespread use of the ex vivo ERG. The use of Ames&#39; or Locke&#39;s medium results in larger photoreceptor and ON-bipolar cell responses, making them preferable to, for example, HEPES-buffered Ringer solution for ex vivo ERG3. Other laboratories stabilized ex vivo ON-bipolar cell function by supplementing with glutamine or glutamate19. This report demonstrates how to obtain large and stable light responses from isolated retinas, including from human donor eyes, as previously reported8.
Important experimental parameters include the temperature of the retina, which should be kept within the physiological range, and a rapid perfusion rate. The most noticeable effects of lowering the temperature in the ex vivo ERG specimen holder are slower response kinetics in both photoreceptors and ON-bipolar cells and an attenuated ON-bipolar cell amplitude. Although lowering the temperature appears to have little effect on the photoreceptor amplitude in the combined response from photoreceptors and ON-bipolar cells, this may be an artifact due to differential changes in the response kinetics from both cell types.
A sufficient perfusion rate appears to be especially critical for retinal function, most likely to supply oxygen and nutrients and remove waste products. While both photoreceptor and ON-bipolar cell amplitudes are somewhat diminished by a moderate reduction in the perfusion rate, even a short cessation of perfusion abolished ON-bipolar but not photoreceptor function. This implies that photoreceptor responses are more robust and may be more easily preserved in eyes that undergo experimentation with a significant delay, such as human donor eyes8. In this context, it is notable that ON-bipolar cell function is not diminished by storing eye cups in oxygenated Ames&#39; medium for several hours. We therefore hypothesize that the loss of ON-bipolar cell function when the perfusion in the ex vivo specimen holder is stopped may be due to rapid depletion of oxygen and other nutrients in the small volume surrounding the retina in the specimen holder. This is supported by reports that a short delay from death to enucleation is critical to record photoreceptor and, particularly, ON-bipolar cell responses from human retinas, and that postmortem hypoxia is the most likely candidate for irreversible damage to retinal neuronal function8. While experimental parameters for ex vivo ERG were optimized in the mouse retina, they nevertheless successfully improved recording conditions for retinal specimens from large animals and human donor eyes.

Electroretinography measures retinal function in response to light1. It is integral to studying retinal physiology and pathophysiology, and measuring the success of therapies for retinal diseases. The in vivo ERG is widely used to assess retinal function in intact organisms, but it has significant limitations2,3. Amongst these, the quantitative analysis of individual retinal cell types in the in vivo ERG is hampered, since it records the sum of potential changes, and therefore overlaying responses, from all retinal cells to light stimuli4. Furthermore, it does not readily allow addition of drugs to the retina, is vulnerable to systemic influences, and has a relatively low signal-to-noise ratio. These disadvantages are eliminated in the ex vivo ERG that investigates the function of the isolated retina2,3,5,6. The ex vivo ERG allows the recording of large and stable responses from specific retinal cell types by addition of pharmacological inhibitors and easy evaluation of therapeutic agents, which can be added to the superfusate. At the same time, it removes influences of systemic effects and eliminates physiological noise (e.g., heartbeat or breathing).
In the ex vivo ERG, retinas or retinal samples are isolated and mounted photoreceptor-side up on the dome of the specimen holder3,5. The specimen holder is assembled, connected to a perfusion system that supplies the retina with heated, oxygenated media, and placed onto the stage of a microscope, which has been modified to deliver computer-controlled light stimuli. To record the responses elicited by light, the specimen holder is connected to an amplifier, digitizer, and recording system (Figure 1). This technique allows isolation of responses from rod and cone photoreceptors, ON-bipolar cells, and M&#252;ller glia by changing the parameters of the light stimuli and adding pharmacological agents.
An existing patch clamp or multi-electrode array (MEA) setup can be converted to record ex vivo ERG, either in conjunction with a commercially available ex vivo ERG adapter or a custom polycarbonate computer numerical control (CNC)-machined specimen holder, to measure light responses in retinas from small animal models, such as mice. This modification increases the accessibility of ex vivo ERG while minimizing the need for specialized equipment. The design of the specimen holder simplifies the mounting technique and integrates electrodes, eliminating the need for manipulation of microelectrodes compared to previously reported transretinal ex vivo ERG methods7. The perfusion rate and temperature inside the specimen holder are important factors that affect the response properties from photoreceptors and ON-bipolar cells. By adjusting these conditions, the ex vivo ERG can be reliably recorded from the isolated mouse retina over prolonged periods of time. Optimized experimental conditions allow ex vivo ERG recordings in retinal punches from larger retinas, including large animal eyes and human donor eyes8.

<table><tbody><tr><td>2 mm socket</td><td>WPI</td><td>2026-10</td><td>materials to prepare electrode</td></tr><tr><td>Ag/AgCl Electrode</td><td>World Precision Instruments</td><td>EP1</td><td>materials to prepare electrode</td></tr><tr><td>Ames&#039; medium</td><td>Sigma Aldrich</td><td>A1420</td><td>perfusion media</td></tr><tr><td>barium chloride</td><td>Sigma Aldrich</td><td>B0750</td><td>potassium channel blocker</td></tr><tr><td>DL-AP4</td><td>Tocris</td><td>0101</td><td>broad spectrum glutamatergic antagonist</td></tr><tr><td>OcuScience Ex Vivo ERG Adapter</td><td>OcuScience</td><td>n/a</td><td>ex vivo ERG specimen holder</td></tr><tr><td>Threaded luer connector</td><td>McMaster-Carr</td><td>51525K222 or 51525K223</td><td>materials to prepare electrode</td></tr></tbody></table>

optimizing the setup and conditions for ex vivo electroretinogram to study retina function in small and large eyes

Measurements of retinal neuronal light responses are critical to investigating the physiology of the healthy retina, determining pathological changes in retinal diseases, and testing therapeutic interventions. The ex vivo electroretinogram (ERG) allows the quantification of contributions from individual cell types in the isolated retina by addition of specific pharmacological agents and evaluation of tissue-intrinsic changes independently of systemic influences. Retinal light responses can be measured using a specialized ex vivo ERG specimen holder and recording setup, modified from existing patch clamp or microelectrode array equipment. Particularly, the study of ON-bipolar cells, but also of photoreceptors, has been hampered by the slow but progressive decline of light responses in the ex vivo ERG over time. Increased perfusion speed and adjustment of the perfusate temperature improve ex vivo retinal function and maximize response amplitude and stability. The ex vivo ERG uniquely allows the study of individual retinal neuronal cell types. In addition, improvements to maximize response amplitudes and stability allow the investigation of light responses in retina samples from large animals, as well as human donor eyes, making the ex vivo ERG a valuable addition to the repertoire of techniques used to investigate retinal function.

Ex vivo ERG enables recording of reproducible and stable photoreceptor and ON-bipolar cell light responses, for example, from the mouse retina (Figure 2A-C). Recording of photoreceptor responses from human donor retinas is possible with up to 5 h postmortem delay of enucleation (Figure 2D) and of ON-bipolar cell responses with a &#60;20 min enucleation delay (Figure 2E). Important parameters to obtain large responses include a careful dissection technique, high perfusion rate, and perfusion temperature close to physiological values (35-38 &#176;C in the mammalian retina). Under these conditions, response amplitudes and kinetics in both cell types were relatively stable over time but slowly declined approximately 40-45 min after retinas were mounted on the specimen holder (Figure 3).
Compared to photoreceptors, ON-bipolar cell function is more easily disrupted, for example, by damage to the retina during dissection and mounting or by a drop in the temperature and/or perfusion speed. While reduced temperature in the specimen holder greatly slowed the kinetics of both photoreceptors and ON-bipolar cells, it decreased the amplitude of the b-wave but not the a-wave (Figure 4A). Conversely, slowing the perfusion rate from 2.1 mL/min to 0.6 mL/min reduced the amplitudes of both photoreceptor and ON-bipolar cell responses but did not affect the implicit time (time from stimulus onset to response peak) of either the a- or the b-wave (Figure 4B). Cessation of perfusion for 10 min followed by reperfusion resulted in a complete loss of ON-bipolar cell function with preserved photoreceptor responses (Figure 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62763/62763fig01.jpg" /> 
Figure 1: Ex vivo electroretinogram specimen holder and recording setup. (A,B) The ex vivo ERG specimen holder comprises a dome to mount the isolated retina, which is connected to a perfusion line to continuously deliver Ames&#39; medium. Electrodes are connected through narrow channels to both the photoreceptor side of the retina via the perfusion line and the inner retina through the filter paper glued to the dome. These electrodes are connected to a differential amplifier, which enables measurement of potential differences in the retina in response to light stimuli. (C) The specimen holder is placed onto the stage of a microscope, which has been modified to deliver light flashes and connected to the perfusion line, which delivers heated, oxygenated Ames&#39; medium by gravity. The entire recording setup is shielded by a Faraday cage to minimize electrical noise. This figure has been modified from 9. Abbreviation: ERG = electroretinogram. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62763/62763fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62763/62763fig02.jpg" /> 
Figure 2: Example traces of ex vivo photoreceptor and ON-bipolar cell responses. Addition of pharmacological agents to the perfusate allows quantification of contributions from individual retinal cell types to the ex vivo electroretinogram. Photoreceptor (PR) light responses are isolated in the presence of 100 &#181;M barium chloride (BaCl2), a blocker of K+ channels expressed by M&#252;ller glial cells, as well as 40 &#181;M DL-AP4, a glutamate receptor blocker, which inhibits signal transmission from photoreceptors to ON-bipolar cells (B). ON-bipolar cell (ON-BPC) function (C) is determined by subtracting the photoreceptor component (B) from the combined photoreceptor and ON-bipolar cell response in the presence of barium chloride alone (A). Photoreceptor light responses can be obtained from human donor retinas with a death to enucleation delay of &#60;5 h (D), whereas retinas enucleated within 20 min of death frequently also give ON-bipolar cell responses (E) (see 8 for further information). Figure 2A-C is modified from 9. Abbreviations: PR = photoreceptor; ON-BPC = ON-bipolar cell. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62763/62763fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62763/62763fig03.jpg" /> 
Figure 3: Stability of photoreceptor and ON-bipolar cell function in the ex vivo electroretinogram over time. (A) Light responses were recorded every minute from photoreceptors alone in the presence of both 100 &#181;M barium chloride and 40 &#181;M DL-AP4. (B) Dim light flashes in the presence of 100 &#181;M barium chloride alone are heavily dominated by ON-bipolar cell function, although they contain a small photoreceptor component. Light responses from isolated retinas typically stabilize after perfusing the ex vivo specimen holder for 15-20 min, and were stable for at least 20-25 min before starting to decline. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62763/62763fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62763/62763fig04.jpg" /> 
Figure 4: Combined photoreceptor and ON-bipolar cell responses at different temperatures and perfusion speeds. (A) Lowering of the temperature inside the specimen holder from 37 &#176;C to room temperature greatly slowed photoreceptor and ON-bipolar cell kinetics but only reduced the ON-bipolar cell amplitude in the mixed photoreceptor and ON-bipolar cell response. (B) Reduction of the perfusion rate from 2.1 mL/min to 0.6 mL/min resulted in decreased photoreceptor and ON-bipolar cell amplitudes but no change in the response kinetics. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62763/62763fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62763/62763fig05.jpg" /> 
Figure 5: ON-bipolar cell responses are more sensitive to cessation of perfusion. (A) Large photoreceptor and ON-bipolar cell responses were recorded following perfusion with 2.1 mL/min for 20 min. (B) After cessation of perfusion for 10 min followed by reperfusion for 10 min at 2.1 mL/min, photoreceptor responses were present, whereas ON-bipolar cells responses were completely lost. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62763/62763fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes at JoVE.com

Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

Modification of existing multielectrode array or patch clamp equipment makes the ex vivo electroretinogram more widely accessible. Improved methods to record and maintain ex vivo light responses facilitate the study of photoreceptor and ON-bipolar cell function in the healthy retina, animal models of eye diseases, and human donor retinas.

Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

Measurements of retinal neuronal light responses are critical to investigating the physiology of the healthy retina, determining pathological changes in ...

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Neuroscience

2.2K Views.  University of Utah. Modification of existing multielectrode array or patch clamp equipment makes the ex vivo electroretinogram more widely accessible. Improved methods to record and maintain ex vivo light responses facilitate the study of photoreceptor and ON-bipolar cell function in the healthy retina, animal models of eye diseases, and human donor retinas.

Video: Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

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 ...

Electrophysiological Characterization of GFP-Expressing Cell Populations in the Intact Retina

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack conspicuous morphological features or show a low population density. This applies particularly to retinal amacrine cells, an exceptionally multiform class of interneurons that comprise roughly 30 subtypes in mammals1. Though being a crucial part of the visual processing by shaping the retinal output2, most of these subtypes have not been studied up to now in a functional context because encountering these cells with a recording electrode is a rare event.
Recently, a multitude of transgenic mouse lines is available that express fluorescent markers like green fluorescent protein (GFP) under the control of promoters for membrane receptors or enzymes that are specific to only a subset of neurons in a given tissue3,4. These pre-labeled cells are therefore accessible to directed microelectrode targeting under microscopic control, permitting the systematic study of their physiological properties in situ. However, excitation of fluorescent markers is accompanied by the risk of phototoxicity for the living tissue. In the retina, this approach is additionally hampered by the problem that excitation light causes appropriate stimulation of the photoreceptors, thus inflicting photopigment bleaching and transferring the retinal circuits into a light-adapted condition. These drawbacks are overcome by using infrared excitation delivered by a mode-locked laser in short pulses of the femtosecond range. Two-photon excitation provides energy sufficient for fluorophore excitation and at the same time restricts the excitation to a small tissue volume minimizing the hazards of photodamage5. Also, it leaves the retina responsive to visual stimuli since infrared light (&gt;850 nm) is only poorly absorbed by photopigments6.
In this article we demonstrate the use of a transgenic mouse retina to attain electrophysiological in situ recordings from GFP-expressing cells that are visually targeted by two-photon excitation. The retina is prepared and maintained in darkness and can be subjected to optical stimuli which are projected through the condenser of the microscope (Figure 1). Patch-clamp recording of light responses can be combined with dye filling to reveal the morphology and to check for gap junction-mediated dye coupling to neighboring cells, so that the target cell can by studied on different experimental levels.

This article depicts the recording of individual cells from fluorescently tagged neuronal populations in the intact mouse retina. By using two-photon infrared excitation transgenetically labeled cells were targeted for patch-clamp recording to study their light responses, receptive field properties, and morphology.

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack ...

Glutamate and Hypoxia as a Stress Model for the Isolated Perfused Vertebrate Retina

Neuroprotection has been a strong field of investigation in ophthalmological research in the past decades and affects diseases such as glaucoma, retinal vascular occlusion, retinal detachment, and diabetic retinopathy. It was the object of this study to introduce a standardized stress model for future preclinical therapeutic testing.
Bovine retinas were prepared and perfused with an oxygen saturated standard solution, and the ERG was recorded. After recording stable b-waves, hypoxia (pure N2) or glutamate stress (250 &#181;m glutamate) was exerted for 45 min. To investigate the effects on photoreceptor function alone, 1 mM aspartate was added to obtain a-waves. ERG-recovery was monitored for 75 min.
For hypoxia, a decrease in a-wave amplitude of 87.0% was noted (p &#60;0.01) after an exposition time of 45 min (decrease of 36.5% after the end of the washout p = 0.03). Additionally, an initial decrease in b-wave amplitudes of 87.23% was recorded, that reached statistical significance (p &#60;0.01, decrease of 25.5% at the end of the washout, p = 0.03).
For 250 &#181;m glutamate, an initial 7.8% reduction of a-wave amplitudes (p &#62;0.05) followed by a reduction of 1.9% (p &#62;0.05). A reduction of 83.7% of b-wave amplitudes (p &#60;0.01) was noted; after a washout of 75 min the reduction was 2.3% (p = 0.62). In this study, a standardized stress model is presented that may be useful to identify possible neuroprotective effects in the future.

With this study, we introduce a standardized stress model for the isolated superfused bovine retina for future preclinical therapeutic testing. The effect of either hypoxia (pure N2) or glutamate stress (250 &#181;M glutamate) on retinal function represented by a- and b-wave amplitudes was evaluated.

Neuroprotection has been a strong field of investigation in ophthalmological research in the past decades and affects diseases such as glaucoma, retinal ...

Medicine

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 ...

In Vivo Methods to Assess Retinal Ganglion Cell and Optic Nerve Function and Structure in Large Animals

The optic nerve collects axons signals from the retinal ganglion cells and transmits visual signal to the brain. Large animal models of optic nerve injury are essential for translating novel therapeutic strategies from rodent models to clinical application due to their closer similarities to humans in size and anatomy. Here we describe some in vivo methods to evaluate the function and structure of the retinal ganglion cells (RGCs) and optic nerve (ON) in large animals, including visual evoked potential (VEP), pattern electroretinogram (PERG) and optical coherence tomography (OCT). Both goat and non-human primate were employed in this study. By presenting these in vivo methods step by step, we hope to increase experimental reproducibility among different labs and facilitate the usage of large animal models of optic neuropathies.

In Vivo Methods to Assess Retinal Ganglion Cell and Optic Nerve Function and Structure in Large Animals

Here we demostrate several in vivo tests (flash visual evoked potential, pattern electroretinogram and optic coherence tomography) in goat and rhesus macaque to understand the structure and function of the optic nerve and its neurons.

The optic nerve collects axons signals from the retinal ganglion cells and transmits visual signal to the brain. Large animal models of optic nerve injury ...

Electroretinogram Recording for Infants and Children under Anesthesia to Achieve Optimal Dark Adaptation and International Standards

Electroretinogram (ERG) is the only clinical objective test available to assess retinal function. Full-field ERG (ffERG) measures the panretinal rod and cone photoreceptor function as well as inner retinal function and is an important measure in the diagnosis and management of inherited retinal diseases as well as inflammatory, toxic, and nutritional retinopathies. Adhering to international standards and maintaining retinal dark adaptation are critical to acquire valid and reliable dark-adapted (scotopic) and light-adapted (photopic) ffERG responses. Performing ffERG in infants and children is challenging and often requires general anesthesia in the operating room. However, maintaining retinal dark adaptation in the operating room is becoming increasingly difficult given the numerous light sources from anesthesiology monitoring systems and other equipment. A practical and widely applicable method for ffERG testing is described in the operating room that optimizes retinal dark adaptation. The method reduces operating room time by dark-adapting the patient before general anesthesiology is instituted. The operating room is modified for dark adaptation and any remaining light source in the darkened operating room is minimized with the use of a modified portable foldable darkroom that encloses the patient&#8217;s head and the ERG examiner during ffERG scotopic recordings. The simple method adheres to ffERG international standards and provides valid reliable scotopic and photopic ffERG recordings that are critical to assess objective retinal function in this young age group where subjective assessment of visual function such as visual acuity and visual fields are not possible. Furthermore, the ffERG is the gold standard clinical test in detecting early onset inherited retinal diseases including Leber congenital amaurosis where approved gene therapy has become available. In sedated conditions, very low amplitude ffERG signals can be detected due to minimal orbicularis muscle activity interference, which is particularly relevant in patients after gene therapy to detect improved amplitude responses.

Adhering to international standards and maintaining retinal dark adaptation are critical to acquire valid full-field electroretinogram responses in the diagnosis and management of inherited retinal diseases. A practical protocol using a portable darkroom is provided to obtain full-field&#160;electroretinogram for infants and children under sedation or general anesthesia in the operating room setting.

Electroretinogram (ERG) is the only clinical objective test available to assess retinal function. Full-field ERG (ffERG) measures the panretinal rod and ...

Full-Circle Cauterization of Limbal Vascular Plexus for Surgically Induced Glaucoma in Rodents

Glaucoma, the second leading cause of blindness worldwide, is a heterogeneous group of ocular disorders characterized by structural damage to the optic nerve and retinal ganglion cell (RGC) degeneration, resulting in visual dysfunction by interrupting the transmission of visual information from the eye to the brain. Elevated intraocular pressure is the most important risk factor; thus, several models of ocular hypertension have been developed in rodents by either genetic or experimental approaches to investigate the causes and effects of the disease. Among those, some limitations have been reported such as surgical invasiveness, inadequate functional assessment, requirement of extensive training, and highly variable extension of retinal damage. The present work characterizes a simple, low-cost, and efficient method to induce ocular hypertension in rodents, based on low-temperature, full-circle cauterization of the limbal vascular plexus, a major component of aqueous humor drainage. The new model provides a technically easy, noninvasive, and reproducible subacute ocular hypertension, associated with progressive RGC and optic nerve degeneration, and a unique post-operative clinical recovery rate that allows in vivo functional studies by both electrophysiological and behavioral methods.

The goal of this protocol is to characterize a novel model of glaucomatous neurodegeneration based on 360&#176; thermic cauterization of limbal vascular plexus, inducing subacute ocular hypertension.

Glaucoma, the second leading cause of blindness worldwide, is a heterogeneous group of ocular disorders characterized by structural damage to the optic ...

Retinal Explant of the Adult Mouse Retina as an Ex Vivo Model for Studying Retinal Neurovascular Diseases

One of the challenges in retina research is studying the cross-talk between different retinal cells such as retinal neurons, glial cells, and vascular cells. Isolating, culturing, and sustaining retinal neurons in vitro have technical and biological limitations. Culturing retinal explants may overcome these limitations and offer a unique ex vivo model to study the cross-talk between various retinal cells with well-controlled biochemical parameters and independent of the vascular system. Moreover, retinal explants are an effective screening tool for studying novel pharmacological interventions in various retinal vascular and neurodegenerative diseases such as diabetic retinopathy. Here, we describe a detailed protocol for retinal explants' isolation and culture for an extended period. The manuscript also presents some of the technical problems during this procedure that may affect the desired outcomes and reproducibility of the retinal explant culture. The immunostaining of the retinal vessels, glial cells, and neurons demonstrated intact retinal capillaries and neuroglial cells after 2 weeks from the beginning of the retinal explant culture. This establishes retinal explants as a reliable tool for studying changes in the retinal vasculature and neuroglial cells under conditions that mimic retinal diseases such as diabetic retinopathy.

Retinal Explant of the Adult Mouse Retina as an Ex Vivo Model for Studying Retinal Neurovascular Diseases

This protocol presents and describes steps for the isolation, dissection, culturing, and staining of retinal explants obtained from an adult mouse. This method is beneficial as an ex vivo model for studying different retinal neurovascular diseases such as diabetic retinopathy.

One of the challenges in retina research is studying the cross-talk between different retinal cells such as retinal neurons, glial cells, and vascular ...

Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

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Transretinal ERG Recordings from Mouse Retina: Rod and Cone Photoresponses

Electrophysiological Characterization of GFP-Expressing Cell Populations in the Intact Retina

Glutamate and Hypoxia as a Stress Model for the Isolated Perfused Vertebrate Retina

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

In Vivo Methods to Assess Retinal Ganglion Cell and Optic Nerve Function and Structure in Large Animals

Electroretinogram Recording for Infants and Children under Anesthesia to Achieve Optimal Dark Adaptation and International Standards

Full-Circle Cauterization of Limbal Vascular Plexus for Surgically Induced Glaucoma in Rodents

Retinal Explant of the Adult Mouse Retina as an Ex Vivo Model for Studying Retinal Neurovascular Diseases