Name: Electroretinogram Analysis of the Visual Response in Zebrafish Larvae (Video) | JoVE Journal
Uploaded: 2015-03-16T15:00:00.000+00:00
Duration: 9 min 44 s
Description: Learn electroretinogram analysis for visual function assessment in zebrafish larvae using micromanipulation techniques. This method enables in vivo investigation of cone visual function in the vertebrate retina.

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5 minutes

We thank members of the UNC Zebrafish Aquaculture facility for maintenance of the zebrafish. We would also like to thank Diagnosys, LLC for assistance with the setup of the ERG apparatus. Additional thanks go to Dr. Portia McCoy and the laboratory of Dr. Ben Philpot for assistance with electrophysiological methods. We also wish to thank Lizzy Griffiths for her illustration of a larval zebrafish. This work was supported by National Institutes of Health awards F32 EY022279 (to J.D.C) and R21 EY019758 (to E.R.W).

Animal upkeep and experimental protocols were approved by the Institutional Animal Care and Use Committees of the University of North Carolina at Chapel Hill, and meet all requirements of the NIH Office of Laboratory Animal Welfare and the Association for Assessment and Accreditation of Laboratory Animal Care International. 
NOTE: To obtain larvae for ERG analysis, published protocols for standard zebrafish husbandry and maintenance were employed18. Larvae are obtained through natural breeding and housed under a 14 hr light/10 hr dark cycle. This protocol has been optimized for larvae at 5-7 days post-fertilization (dpf), but could ideally be performed on older fish with small modifications to the procedure. Here, use the TL strain of wild-type zebrafish larvae at 5 dpf.
1. Micropipette Production
<ol>
	<li>Pull several micropipettes using 1.5 x 0.86 mm (outer diameter by inner diameter) fire-polished borosilicate glass capillaries with filament (melting temperature, 821 &#176;C) and a P-97 Flaming/Brown Micropipette Puller fitted with box heat filament. Use the program for fashioning micropipettes described in Table 1.</li>
	<li>Check each micropipette under a microscope with an appropriate graticule ruler to ensure that tips are 10-15 &#181;m in diameter and has a smooth tip opening (i.e., no jagged edges).</li>
	<li>Carefully store micropipettes to prevent tip damage and exposure to dust. Storage options include Petri dishes with lab tape, foam-lined boxes, or commercially available micropipette storage containers. 
	NOTE: Other micropipette pullers and glass capillaries can be used as long as the correct diameter of the micropipette and a high-quality tip is achieved.</li>
</ol>
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Pressure</td>
			<td>Heat</td>
			<td>Pull</td>
			<td>Velocity</td>
			<td>Time</td>
		</tr>
		<tr>
			<td>500</td>
			<td>560</td>
			<td>-</td>
			<td>30</td>
			<td>200</td>
		</tr>
		<tr>
			<td>500</td>
			<td>450</td>
			<td>-</td>
			<td>30</td>
			<td>200</td>
		</tr>
		<tr>
			<td>500</td>
			<td>410</td>
			<td>55</td>
			<td>40</td>
			<td>200</td>
		</tr>
	</tbody>
</table>
Table 1: Program for the production of micropipettes using a P-97 Flaming/brown Micropipette Puller fitted with a box heat filament. Micropipettes are made using 1.5 x 1.0 mm2 (outer diameter by inner diameter) fire-polished borosilicate glass capillaries with filament (melting temperature, 821 &#176;C).
2. Buffer Preparation
<ol>
	<li>Use filtered, oxygenated goldfish Ringer&#39;s buffer19 in the microelectrode capillary and to saturate the polyvinyl alcohol (PVA) sponge onto which the larvae are placed for experiments. Alternatively, use E3 embryo media or Hank&#39;s Balanced Salt Solution.</li>
	<li>Prepare 10x goldfish Ringer&#39;s solution as described in Table 2. Adjust to pH 7.8, and sterilize using a 0.22 &#181;m filter and store the 10x stock at 4 &#176;C.</li>
	<li>Create a working solution on the day of the experiment by diluting the 10x Ringer&#39;s solution to 1x with deionized, distilled water. Filter using a 0.22 &#181;m filter system. Oxygenate by bubbling with 95% O2/5% CO2 gas for 10 minutes. Cap tightly afterwards to ensure that the solution remains oxygenated.</li>
</ol>
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>NaCl</td>
			<td>1.25 M</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>26 mM</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>25 mM</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>glucose</td>
			<td>100 mM</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>100 mM</td>
		</tr>
	</tbody>
</table>
Table 2: Preparation of 10x goldfish Ringer&#8217;s solution.
3. Electroretinogram Platform
<ol>
	<li>Perform ERG experiments on an anti-vibration table inside a Faraday cage to improve the signal to noise ratio. Attach a custom steel platform to the anti-vibration table using hex nuts. Place a movable plastic platform with a viscoelastic urethane polymer shock-absorbing bottom on the table under the light source.</li>
	<li>Position the camera with a magnetized stand, aimed down at the movable plastic platform. Position the micromanipulator (which will hold the recording microelectrode) with a second magnetized stand to the right of the movable plastic platform. Ensure that the camera and micromanipulator will not be disturbed by the movement of other equipment and that they do not block illumination from the light source.</li>
	<li>Connect the camera to a video monitor and position it to view the eye of the larva for placing the electrode in the proper position.</li>
	<li>Ensure that the setup is properly grounded with copper wire. To check the noise, place the reference electrode and tip of the recording microelectrode in a 35 mm Petri dish filled with Ringer&#39;s solution. Check the electrical noise levels of the setup with an oscilloscope or a built-in feature of the ERG apparatus. Noise levels should be no more than &#177;10 &#181;V from baseline.</li>
</ol>
4. Sponge Preparation
<ol>
	<li>Cut a small rectangle of dry PVA sponge that will fit snugly in a 35 mm Petri dish. The thickness of the sponge should not be greater than the depth of the dish. Use a utility knife with a clean razor blade for cutting.</li>
	<li>Make an additional cut into the sponge to accommodate the reference electrode (either a shallow cut lengthwise on the bottom of the sponge or a butterfly cut vertically through one of the smaller ends).</li>
	<li>Use a chemically resistant marker to mark a small dot on the sponge (where the larva will be placed) that can be used for positioning the camera.</li>
	<li>Soak the PVA sponge in Ringer&#39;s solution until saturated. Remove and blot quickly on a paper towel 2-3 times. Place the sponge in a clean 35 mm Petri dish.</li>
	<li>Position the Petri dish containing the sponge on the plastic platform such that the mark can be visualized by the camera.</li>
</ol>
5. Electrode Preparation
NOTE: The zebrafish setup consists of a reference electrode in contact with the Ringer&#39;s solution-saturated PVA sponge and a recording electrode in contact with the cornea. The reference electrode consists of an Ag/AgCl pellet. The recording electrode is a pulled glass micropipette filled with Ringer&#39;s solution and held by a microelectrode holder containing an Ag wire.
<ol>
	<li>Chloride the electrodes by soaking them in 6-9% sodium hypochlorite (bleach) for 5 min (the recording microelectrode wire) or 15 min (the reference electrode). Air dry on a Kimwipe for 5 min.
	<ol>
		<li>Depending on the style of cut made in Step 4.2, place the Ag/AgCl pellet of the reference electrode into (for the vertical butterfly cut) or under (for the shallow cut lengthwise on the bottom) the sponge. Attach the reference electrode lead to the recording system.</li>
		<li>Alternatively, if the ERG setup has space constraints or there are particularly strong photovoltaic artifacts from the Ag/AgCl electrode, connect the reference electrode to the sponge via an agar salt bridge to move the electrode out of the light path.</li>
	</ol>
	</li>
	<li>Attach ~40 cm of appropriately sized tubing to a 5 ml non-Luer lock syringe. Fill the syringe with Ringer&#39;s solution. Microelectrode holders possessing pressure ports typically ship with adaptors to accommodate tubing with inner diameters of 1/16&#8221;, 3/32&#8221;, 1/8&#8221; or 5/32&#8221;.</li>
	<li>Fill a 1 ml non-Luer lock syringe with Ringer&#39;s solution and, using a Micro-fil, carefully fill the microelectrode holder. Prevent the formation of bubbles.</li>
	<li>Attach the 5 ml syringe to the pressure port of the microelectrode holder with tubing and use it to ensure that the microelectrode holder is full of Ringer&#39;s solution. Using the Micro-fil and 1-ml syringe filled with Ringer&#39;s solution, fill the micropipette glass from the tip and ensure that no bubbles are present.</li>
	<li>Attach the glass micropipette to the microelectrode holder, being careful to keep the electrode wire straight. Once secured, use the 5 ml syringe to carefully force Ringer&#39;s solution through the microelectrode until a tiny amount of solution is visible on the tip. Occasional application of pressure to the syringe (when not applied to cornea) will prevent formation of air bubbles, as well as occlusions due to dust or salt accumulation, in the micropipette tip.
	<ol>
		<li>If the solution comes out as a stream, replace the glass micropipette, as the tip opening is too large or is damaged.</li>
	</ol>
	</li>
	<li>Carefully place the recording microelectrode in the micromanipulator and attach the lead to the recording system.</li>
</ol>
6. Electroretinogram Analysis
NOTE: Due to the cone dominance of the larval retina, high quality ERG results can be obtained when preparations for recording are performed under low levels of indirect white light (&#60;1 lux) or with short periods (&#60;1 min) of higher intensity (&#8804;250 lux) working light. A short period of dark adaptation is still required prior to recording (see step 6.7). However, experiments can be performed under dim red or infrared light using an infrared-sensitive camera. All experiments were performed in filter-sterilized (0.22 &#181;m) system water from the UNC Zebrafish AquaCulture Facility but alternative embryo media can be used.
<ol>
	<li>Cut paper towel squares measuring approximately 1 cm2.</li>
	<li>If measuring isolated cone mass receptor potential, incubate 3-5 larvae in system water with 500 &#181;M (&#177;)-2-Amino-4-phosphonobutyric acid (APB) for 5 min. 
	NOTE: While APB is a racemic mixture of the active (L) and inactive (R) forms of AP4, it is as effective as L-AP4 and is less expensive.</li>
	<li>Anesthetize 3-5 larvae in system water with 0.02% (w/v) Tricaine until unresponsive, about 1-2 min.</li>
	<li>Use a Pasteur pipette and pipette pump to carefully transfer individual larvae onto paper towel squares under a dissecting stereoscope using minimal illumination (&#8804;250 lux for &#60;1 min). Check the position of each larva and choose a candidate that is dorsal side up with an unoccluded eye.
	<ol>
		<li>For extended recordings (&#62;30 min), keep the larva moist by glazing the body up to but not including the head with 3% methylcellulose using a fine camel-hair brush.</li>
	</ol>
	</li>
	<li>Using forceps, transfer the paper towel square with the larva to the damp PVA sponge.
	<ol>
		<li>For extended recordings (&#62;30 min), apply a continuous stream of water-saturated 100% O2 gas over the larva by bubbling the gas through an airstone in a side-arm flask containing distilled water. Position the tubing exiting the flask side-arm that conveys the humidified oxygen near the larva&#39;s head. 
		NOTE: Step 6.4.1 and step 6.5.1 will prolong the life of the fish16.</li>
	</ol>
	</li>
	<li>Under minimal illumination, use the micromanipulator and camera to position the microelectrode tip at the midpoint between the nasal and caudal ends of the eye and gently press onto the dorsal limit of the cornea. 
	NOTE: Misplacement of the electrode tip to the far distal areas of the cornea can result in ERG waveforms of reversed polarity.</li>
	<li>Allow larva to dark-adapt for 5-10 min.</li>
	<li>Record test flash responses to light provided from an LED light source or optical stimulator using the available stimulation and recording equipment. Adjust protocol parameters such as flash intensity, flash length, flash color, background intensity and color and filter settings to fit the experiment.</li>
	<li>When finished with the experiment, euthanize larvae according to AVMA/IACUC guidelines.</li>
</ol>

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</ol>

No conflicts of interest declared.

In this protocol a simple procedure for ERG recordings of larval zebrafish is detailed. This procedure allows for a quick and comprehensive assay of visual function.There are several critical steps throughout the procedure that should be kept in mind. The zebrafish larvae should be healthy before the experiment to prevent death during potential drug treatments and ensure prolonged livelihood during the ERG recordings. In addition, it is important that the larvae utilized in experiments are closely age-matched. This is du...

The electroretinogram (ERG) is a noninvasive electrophysiological method that has been used extensively in the clinic for determining the function of the retina in humans. The electrical activity in response to a light stimulus is measured by placing recording electrodes on the outer surface of the cornea. The characteristics of the stimulus paradigm and the response waveform define the retinal neurons contributing to the response. This method has been adapted for use with a number of animal models including mice and zebrafish. The typical vertebrate ERG response has four principal components: the a-wave, which is a cornea-negative potential derived from photoreceptor cell activity; the b-wave, a cornea-positive potential derived from the ON bipolar cells; the d-wave, a cornea-positive potential interpreted as the activity of the OFF bipolar cells; and the c-wave, which occurs several seconds after the b-wave and reflects activity in M&#252;ller glia and the retinal pigment epithelium1-4. Additional references for understanding the history and principles of ERG analysis in humans and model animals are the online textbook, Webvision, from the University of Utah and texts such as the Principles and Practice of Clinical Electrophysiology of Vision4,5.
Daniorerio (zebrafish) has long been favored as a model for vertebrate development, due to its rapid maturation and transparency, which allows for noninvasive morphological analysis of organ systems, behavioral assays and both forward and reverse genetic screens (for review, see Fadool and Dowling6). Zebrafish larvae are highly amenable to genetic and pharmacological manipulation, which, when coupled with their high fecundity, make them an excellent animal model for high-throughput biological analyses. The higher ratio of cones to rods in larval zebrafish &#8211; roughly 1:1 compared to mice (~3% cones) &#8211; make them particularly useful for the study of cone function7-9.
In the vertebrate retina, cones develop before rods10. Interestingly, zebrafish cones are operative as early as 4 dpf, allowing for selective electrophysiological analysis of cones at that stage6,11,12. In contrast, ERG responses in rods appear between 11 and 21 dpf13. Therefore, zebrafish larvae at 4-7 dpf serve functionally as an all-cone retina. However, the native photopic ERG response of 4-7 dpf larvae is dominated by the b-wave. Application of pharmacological agents, such as L-(+)-2-amino-4-phosphono-butyric acid (L-AP4), an agonist for the metabotropic glutamate (mGluR6) receptor expressed by the ON bipolar cells, effectively blocks the generation of the b-wave and reveals the isolated cone mass receptor potential, (the &#8220;a-wave&#8221;)14-17.
Here we describe a simple and reliable method for ERG analysis using commercially available ERG equipment designed for use with mice that have been adapted for use with zebrafish larvae. This system can be utilized on zebrafish larvae of varying genetic backgrounds, as well as those treated with pharmacological agents, to aid researchers in the identification of signaling pathways that contribute to visual sensitivity and light adaptation16. The experimental procedures outlined in this protocol will guide investigators in the use of ERG analysis to answer a variety of biological questions pertaining to vision, and demonstrate the construction of a flexible ERG setup.

<table><tbody><tr><td>Faraday cage</td><td>80/20 Inc</td><td>custom</td><td>Custom designed aluminum &quot;Industrial Erector Set&quot; for Cage framework</td></tr><tr><td>PVA sponge</td><td>Amazon</td><td>B000ZOWG1C</td><td>Provides a soft, moist platform for placement of zebrafish larvae</td></tr><tr><td>150 ml Sterile Filter systems</td><td>Corning</td><td>431154</td><td>Filtering solutions to prevent small articulates from blocking micropipettes</td></tr><tr><td>Espion E2</td><td>Diagnosys, LLC</td><td>contact</td><td>Modular electrophysiology system capable of generating visual stimuli for any stimulator and digital recording and analysis of responses using propietary software, more information at http://www.diagnosysllc.com</td></tr><tr><td>Colordome</td><td>Diagnosys, LLC</td><td>contact</td><td>Light stimulator with RGB LED and Xenon light sources for Ganzfeld ERG, more information at http://www.diagnosysllc.com</td></tr><tr><td>Micromanipulator</td><td>Drummond</td><td>3-000-024-R</td><td>Holding and positioning the recording microelectrode</td></tr><tr><td>Magnetic ring stand</td><td>Drummond</td><td>3-000-025-MB</td><td>Holding and positioning of the camera and refrence electrode</td></tr><tr><td>Lead extensions</td><td>Grass Technologies</td><td>F-LX</td><td>Spare female to male 1.5 mm lead cables for connecting electrodes</td></tr><tr><td>Male Pin to Female SAFELEAD Adaptor</td><td>Grass Technologies</td><td>DF-215/10</td><td>Connecting 2 mm pins to 1.5 headboard pins</td></tr><tr><td>Window screen frame (metal) and spline</td><td>Lowes or Home Depot</td><td>various</td><td>For attaching copper mesh to Faraday cage framework</td></tr><tr><td>Steriflip 50 ml filters</td><td>Millipore</td><td>SCGP00525</td><td>Filtering solutions to prevent small articulates from blocking micropipettes</td></tr><tr><td>BNC adaptor</td><td>Monoprice</td><td>4127</td><td>Connecting camera to BNC cable</td></tr><tr><td>BNC cable</td><td>Monoprice</td><td>626</td><td>Connecting camera to video adaptor</td></tr><tr><td>Camera lens</td><td>Navitar</td><td>1582232</td><td>Visualizing the positioning of the recording microelectrode onto the larval cornea</td></tr><tr><td>Camera coupler</td><td>Navitar</td><td>1501149</td><td>Visualizing the positioning of the recording microelectrode onto the larval cornea</td></tr><tr><td>Luna BNC to VGA + HDMI Converter</td><td>Sewell</td><td>SW-29297-PRO</td><td>BNC to VGA adaptor allowing camera image to project on computer monitor</td></tr><tr><td>APB</td><td>Sigma</td><td>A1910</td><td>mGluR6 agonist, blocks b-wave allowing analysis of the isolated cone mass receptor potential</td></tr><tr><td>Borosilicate glass</td><td>Sutter</td><td>BF-150-86-10</td><td>Fire- polished borosilicate glass (metling temperature = 821&amp;deg;C) with filament and dimensions of 1.5mm x 0.86 mm (outer diameter by inner diameter)&amp;nbsp;</td></tr><tr><td>P97 Flaming/Brown puller</td><td>Sutter</td><td>P97</td><td>For pulling glass micropipettes</td></tr><tr><td>Sorbothane sheet</td><td>Thorlabs</td><td>SB12A</td><td>Synthetic viscoelastic urethane polymer, placed under Passive Isolation Mounts and ERG platform to absorb shock and prevent slipping, can be cut to size</td></tr><tr><td>Breadboard</td><td>Thorlabs</td><td>B2436F</td><td>Vibration isolation platfrom for ERG stimulator and zebrafish specimen</td></tr><tr><td>Passive Isolation Mounts</td><td>Thorlabs</td><td>PWA074</td><td>Provides vibration isolation to breadboard</td></tr><tr><td>Copper mesh</td><td>TWP</td><td>022X022C0150W36T</td><td>To line Faraday Cage</td></tr><tr><td>Pipette pump</td><td>VWR</td><td>53502-233</td><td>Used with Pasteur pipettes to carefully transfer zebrafish larvae</td></tr><tr><td>Pasteur pipettes</td><td>VWR</td><td>14672-608</td><td>Used with Pipette pump to carefully transfer zebrafish larvae</td></tr><tr><td>Camera</td><td>Watec</td><td>WAT-902B</td><td>Visualizing the positioning of the recording microelectrode onto the larval cornea</td></tr><tr><td>Tricaine (MS-222)</td><td>Western Chemical</td><td>Tricaine-S</td><td>Pharmaceutical-grade anesthetic,</td></tr><tr><td>Micro-fil</td><td>WPI</td><td>MF28G-5</td><td>Filling microelectrode holder and microelectrode glass</td></tr><tr><td>Microelectrode holder</td><td>WPI</td><td>MEH2SW15</td><td>Holds glass microelectrode, connects to ERG equipment</td></tr><tr><td>Reference Electrode</td><td>WPI</td><td>DRIREF-5SH</td><td>Carefully break off last centimeter of casing to drain electrolyte and expose sintered Ag/AgCl pellet electrode</td></tr><tr><td>Reference Electrode (alternative)</td><td>WPI</td><td>EP1</td><td>Alternative to DRIREF-5SH. Ag/AgCl electrode that must be wired/soldered to connecting lead</td></tr><tr><td>Low-noise cable for Microelectrode holder</td><td>WPI</td><td>13620</td><td>Connecting recording microelctrode holder to adaptor/headboard</td></tr></tbody></table>

electroretinogram analysis of the visual response in zebrafish larvae

Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the surface of the cornea, electrical activity generated in response to light can be measured and used to assess the activity of retinal cells in vivo. This manuscript describes the use of the ERG to measure visual function in zebrafish. Zebrafish have long been utilized as a model for vertebrate development due to the ease of gene suppression by morpholino oligonucleotides and pharmacological manipulation. At 5-10 dpf, only cones are functional in the larval retina. Therefore, the zebrafish, unlike other animals, is a powerful model system for the study of cone visual function in vivo. This protocol uses standard anesthesia, micromanipulation and stereomicroscopy protocols that are common in laboratories that perform zebrafish research. The outlined methods make use of standard electrophysiology equipment and a low light camera to guide the placement of the recording microelectrode onto the larval cornea. Finally, we demonstrate how a commercially available ERG stimulator/recorder originally designed for use with mice can easily be adapted for use with zebrafish. ERG of larval zebrafish provides an excellent method of assaying cone visual function in animals that have been modified by morpholino oligonucleotide injection as well as newer genome engineering techniques such as Zinc Finger Nucleases (ZFNs), Transcription Activator-Like Effector Nucleases (TALENs), and Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9, all of which have greatly increased the efficiency and efficacy of gene targeting in zebrafish. In addition, we take advantage of the ability of pharmacological agents to penetrate zebrafish larvae to evaluate the molecular components that contribute to the photoresponse. This protocol outlines a setup that can be modified and used by researchers with various experimental goals.

Typically, ERGs are recorded from zebrafish larvae at 5 dpf, since a number of studies have published ERG recordings at this stage9,16,20. Larval responses were measured under dark-adapted conditions with no background illumination using a 20 msec stimulus of white LED light. We utilized a commercially available ERG system consisting of a Ganzfeld light stimulator and computer controller/recorder. The stimulator uses a tightly controlled proprietary pulse width modulation (PWM) system to control the...

We present a method for the electroretinographic (ERG) analysis of zebrafish larvae utilizing micromanipulation and electroretinography techniques. This is a simple and straightforward method for assaying visual function of zebrafish larvae in vivo.

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the ...

The overall goal of this procedure is to illustrate the steps necessary to perform electrographic analysis on larval zebra fish. This is accomplished by first generating the appropriately shaped and sized glass micro pipettes. The second step is to carefully set up equipment critical to the experiment, such as reference and recording electrodes.Next, the larval zebrafish is anesthetized and properly oriented relative to the stimulation and recording equipment. The final step is the proper placement of the glass micro electrode onto the cornea of the larval zebra fish. Ultimately larval zebra fish.ERG analysis is useful for investigating cone visual function in the all cone vertebrate retina in vivo. This method is very useful for answering key questions in the field of vision science, such as the identification of signaling pathways that contribute to visual sensitivity and light adaptation in cones. Visual demonstration of this method is critical as the proper positioning of the recording micro electrode onto the specimen is difficult to learn due to the small size of the larval zebrafish eye.To begin this procedure, pull several micro pipettes using fire polished boro silicate glass capillaries with filament. Check each micro pipette under a microscope with an appropriate grad ruler to ensure that the tips diameter is 10 to 15 micrometers and even in appearance. Create a working solution on the day of the experiment by diluting 10 x ringer solution to one x.With deionized distilled water filter, sterilize the solution to ensure that particulates will not block the micro pipette tip. Then oxygenated with 95%oxygen, 5%carbon dioxide for 10 minutes. Cap the solution tightly afterwards to ensure that the solution remains oxygenated.Now place a movable plastic platform with a viscoelastic urethane polymer shock absorbing bottom on the anti vibration table under the light source. Next, position the camera with a magnetized stand and aim it at the movable plastic platform. Then position the micro manipulator of the recording micro electrode on a second magnetized stand.Ensure that the camera and micro manipulator are not disturbed by the movement of other equipment, and do not block illumination from the light source. Afterward, connect the camera to a video monitor and position it towards the platform. Ensure that the setup is properly grounded with copper wire.In this step, cut a small rectangle of dry PVA sponge that will fit snugly in a 35 millimeter Petri dish with a clean razor blade. Make an additional cut into the sponge to accommodate the reference electrode. Then use a chemical resistant marker to Mark A.Small dot on the sponge where the larva will be placed.After that, soak the PVA sponge in ringer solution until saturated. Remove and blot it quickly on a paper towel two to three times before placing it in a clean 35 millimeter Petri dish. Subsequently, position the Petri dish containing the sponge on a plastic platform such that the mark can be visualized by the camera.Next, chloride the electrodes by soaking them in six to 9%sodium chloride. Then air dry them on a Kim wipe for five minutes. Place the silver, silver chloride pellet of the reference electrode into or under the sponge depending on the style of the cut made, and attach the reference electrode lead to the recording system.Next, attach approximately 40 centimeters of tubing to a five milliliter non lure lock syringe and fill the syringe with ringer solution. Afterward, fill a one milliliter non lure lock syringe with ringer solution. Using a microfill, carefully fill the micro electrode holder and prevent the formation of bubbles.Then attach the five milliliter syringe to the pressure port of the micro electrode holder with tubing using the microfill and one milliliter syringe filled with ringer solution. Fill the glass micro pipette from the tip and ensure that no bubbles are present. Next, attach the glass micro pipette to the micro electrode holder and be careful to keep the electrode wire straight.Once secured, use the five milliliter syringe to carefully force the ringer solution through the micro electrode until a tiny amount of solution is visible on the tip. Now carefully place the recording micro electrode in the micro manipulator and attach the lead to the recording system to check for system noise. Place the reference electrode and the tip of the recording micro electrode in a 35 millimeter Petri dish filled with ringer solution.Check the electrical noise levels of the setup with an oscilloscope or a built-in feature of the ERG apparatus. Noise levels should be no more than plus or minus 10 microvolts from the baseline. In this procedure, cut several one square centimeter paper towel squares, anesthetize three to five larvae in a 35 millimeter Petri dish by adding trica to a final concentration of 0.02%and waiting one to two minutes.After that, carefully transfer an individual larva onto a paper towel square under a dissecting stereoscope with minimal illumination. Check the position of the larvae choosing a candidate that is dorsal side up with an occluded eye for recording sessions that last longer than 30 minutes. Keep the larva moist by glazing the body with 3%methylcellulose using a fine camel hair brush.Then transfer the paper towel square with the larva to the damp PVA sponge. Next, apply a continuous stream of water saturated 100%oxygen over the larvae by bubbling the gas through an airstone in a sidearm flask containing distilled water. Position the humidified oxygen from the flask sidearm near the larvas head with tubing under minimal illumination.Use the micro manipulator and camera to position the micro electrode tip at the midpoint between the nasal and coddle ends of the eye and gently press onto the dorsal limit of the cornea. Allow the larvae to adapt to the dark for five to 10 minutes. Then record the test flash responses to light provided from an LED light source or optical stimulator.This figure shows a typical ERG recording in a five DPF larval zebra fish. The dark adapted fish were exposed to white LED light for 20 milliseconds with intensities ranging from one to 5, 000 canula per square meter with the onset of the light stimulus at zero seconds. The negative potential a wave is difficult to distinguish, whereas the positive potential B wave is the dominant peak of the wave form.This inset shows the averaged B wave response amplitudes that have been fit using the NACA rushed in equation. This figure shows a typical ERG recording in a dark adapted five DPF larval zebra fish treated with a PB.The fish are exposed to LED white light for 20 milliseconds with intensities ranging from one to 5, 000 canula per square meter, and the isolated cone mass receptor potential is the dominant element of the waveform. This inset shows the averaged response amplitudes of a PB treated larval zebra fish that had been fit using the NACA rushed in equation.This figure shows a typical ERG recording in a five D-P-F-A-P-B treated larval zebra fish. Using a dual flash paradigm. The dark adapted fish are exposed to successive 20 millisecond white light.LED flashes with an intensity of 1000 canula per square meter, and the ISI between the flashes is two seconds. The onset of each light stimulus is indicated here. This figure shows the ratio of the maximum isolated cone mass receptor potential response of the second stimulus to that of the initial stimulus with increasing ISI, which indicates the progressive recovery of photoreceptor sensitivity.While attempting this procedure is important to remember that the quality of the recordings is dictated by the integrity of the electrode corneal connection and the health of the larvae. The development of this technique has greatly enhanced the ability of researchers in the vision sciences to explore cone adaptation and recovery in the vertebrate retina in vivo.

electroretinogram-analysis-of-the-visual-response-in-zebrafish-larvae

Research

JoVE Journal

Neuroscience

We developed the Visual-Motor Response to quantitate the motor output of larval zebrafish in response to light increments and decrements.  We also examined zebrafish vision mutants, including the no optokinetic response (nrc) mutants, which were thought to be completely blind when tested by another vision assay, the optokinetic reflex.

A Behavioral Assay to Measure Responsiveness of Zebrafish to Changes in Light Intensities 

Optokinetic response has been widely used to assess the visual functions of larval zebrafish. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adults1-5. Here, we introduce how to measure the OKR of adult zebrafish using a new protocol which is established in our lab.

Using the optokinetic response to study visual function of zebrafish

The VisioTracker is an automated system for the quantitative analysis of visual performance of larval and small adult fish based on the recording of eye movements. It features full control over visual stimulus properties and real-time analysis, enabling high-throughput research in fields such as visual system development and function, pharmacology, neural circuit studies and sensorimotor integration.

VisioTracker, an Innovative Automated Approach to Oculomotor Analysis

This video describes the photomotor response assay of zebrafish and fathead minnow larvae using an automated tracking software. The protocol measures the behavioral response of the larval fish following a sudden change in lighting condition.

Photomotor Response Assay: A Method to Measure the Behavioral Response of Larval Zebrafish to a Sudden Change in Lighting Condition

This video describes the optokinetic response assay in zebrafish. The video measures the reflexive eye movements based on the visual moving stimulus and determines the visual acuity in zebrafish.

Optokinetic Response Assay: A Method to Measure Visual Acuity in Zebrafish

This video demonstrates the method for studying visual prey detection in transgenic zebrafish larvae using calcium indicator-based fluoresce imaging with their neurons expressing fluorescent calcium indicator complexes.

Analyzing the Prey-Evoked Neuronal Activity in a Transgenic Zebrafish Larva

Zebrafish have been a valuable tool for many research areas. Here we demonstrate a method to elicit a visual response and calculate a functional visual acuity in the adult zebrafish.

The Optokinetic Response as a Quantitative Measure of Visual Acuity in Zebrafish

We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

Here, we present a protocol for the development and the characterization of a zebrafish model of epilepsy resulting from the transient inhibition of the DEPDC5 gene.

Behavioral And Physiological Analysis In A Zebrafish Model Of Epilepsy

Here, we present a protocol that simplifies the measurement of light evoked electroretinogram responses from larval zebrafish. A novel cone-shaped sponge-tip electrode can help to make the study of visual development in larval zebrafish using the electroretinogram ERG easier to achieve with reliable outcomes and lower cost.

Electroretinogram Recording in Larval Zebrafish using A Novel Cone-Shaped Sponge-tip Electrode

The article explains how to surgically remove eyes from living zebrafish larvae as the first step toward investigating how retinal input influences optic tectum growth and development. In addition, the article provides information about larval anesthetization, fixation, and brain dissection, followed by immunohistochemistry and confocal imaging.

Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System

Learn electroretinogram analysis for visual function assessment in zebrafish larvae using micromanipulation techniques. This method enables in vivo investigation of cone visual function in the vertebrate retina.

Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

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