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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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

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...

Watch this Scientific Journal Video about Electroretinogram Analysis of the Visual Response in Zebrafish Larvae at JoVE.com

Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

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 ...

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

Research

JoVE Journal

Neuroscience

15.5K Views.  University of North Carolina at Chapel Hill. 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.

Video: Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

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

The optokinetic reflex (OKR) is a basic visual reflex exhibited by most vertebrates and plays an important role in stabilizing the eye relative to the visual scene. However, the OKR requires that an animal detect moving stripes and it is possible that fish that fail to exhibit an OKR may not be completely blind. One zebrafish mutant, the no optokinetic response c (nrc) has no OKR under any light conditions tested and was reported to be completely blind. Previously, we have shown that OFF-ganglion cell activity can be recorded in these mutants. To determine whether mutant fish with no OKR such as the nrc mutant can detect simple light increments and decrements we developed the visual motor behavioral assay (VMR). In this assay, single zebrafish larvae are placed in each well of a 96-well plate allowing the simultaneous monitoring of larvae using an automated video-tracking system. The locomotor responses of each larva to 30 minutes light ON and 30 minutes light OFF were recorded and quantified. WT fish have a brief spike of motor activity upon lights ON, known as the startle response, followed by return to lower-than baseline activity, called a freeze. WT fish also sharply increase their locomotor activity immediately following lights OFF and only gradually (over several minutes) return to baseline locomotor activity. The nrc mutants respond similarly to light OFF as WT fish, but exhibit a slight reduction in their average activity as compared to WT fish. Motor activity in response to light ON in nrc mutants is delayed and sluggish. There is a slow rise time of the nrc mutant response to light ON as compared to WT light ON response. The results indicate that nrc fish are not completely blind. Because teleosts can detect light through non-retinal tissues, we confirmed that the immediate behavioral responses to light-intensity changes require intact eyes by using the chokh (chk) mutants, which completely lack eyes from the earliest stages of development. In our VMR assay, the chk mutants exhibit no startle response to either light ON or OFF, showing that the lateral eyes mediate this behavior. The VMR assay described here complements the well-established OKR assay, which does not test the ability of zebrafish larvae to respond to changes in light intensities. Additionally, the automation of the VMR assay lends itself to high-throughput screening for defects in light-intensity driven visual responses.

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

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.

The optokinetic reflex (OKR) is a basic visual reflex exhibited by most vertebrates and plays an important role in stabilizing the eye relative to the ...

Biology

VisioTracker, an Innovative Automated Approach to Oculomotor Analysis

Investigations into the visual system development and function necessitate quantifiable behavioral models of visual performance that are easy to elicit, robust, and simple to manipulate. A suitable model has been found in the optokinetic response (OKR), a reflexive behavior present in all vertebrates due to its high selection value. The OKR involves slow stimulus-following movements of eyes alternated with rapid resetting saccades. The measurement of this behavior is easily carried out in zebrafish larvae, due to its early and stable onset (fully developed after 96 hours post fertilization (hpf)), and benefitting from the thorough knowledge about zebrafish genetics, for decades one of the favored model organisms in this field. Meanwhile the analysis of similar mechanisms in adult fish has gained importance, particularly for pharmacological and toxicological applications.
Here we describe VisioTracker, a fully automated, high-throughput system for quantitative analysis of visual performance. The system is based on research carried out in the group of Prof. Stephan Neuhauss and was re-designed by TSE Systems. It consists of an immobilizing device for small fish monitored by a high-quality video camera equipped with a high-resolution zoom lens. The fish container is surrounded by a drum screen, upon which computer-generated stimulus patterns can be projected. Eye movements are recorded and automatically analyzed by the VisioTracker software package in real time.
Data analysis enables immediate recognition of parameters such as slow and fast phase duration, movement cycle frequency, slow-phase gain, visual acuity, and contrast sensitivity.
Typical results allow for example the rapid identification of visual system mutants that show no apparent alteration in wild type morphology, or the determination of quantitative effects of pharmacological or toxic and mutagenic agents on visual system performance.

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.

Investigations into the visual system development and function necessitate quantifiable behavioral models of visual performance that are easy to elicit, ...

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

Zebrafish are a proven model for vision research, however many of the earlier methods generally focused on larval fish or demonstrated a simple response. More recently adult visual behavior in zebrafish has become of interest, but methods to measure specific responses are new coming. To address this gap, we set out to develop a methodology to repeatedly and accurately utilize the optokinetic response (OKR) to measure visual acuity in adult zebrafish. Here we show that the adult zebrafish's visual acuity can be measured, including both binocular and monocular acuities. Because the fish is not harmed during the procedure, the visual acuity can be measured and compared over short or long periods of time. The visual acuity measurements described here can also be done quickly allowing for high throughput and for additional visual procedures if desired. This type of analysis is conducive to drug intervention studies or investigations of disease progression.

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.

Zebrafish are a proven model for vision research, however many of the  earlier methods generally focused on larval fish or demonstrated a simple  ...

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

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs survival and regeneration experiments. Many studies have been performed in mammalian animals to research RGCs survival after optic nerve injury. However, retrograde labeling of RGCs in adult zebrafish has not yet been reported, though some alternative methods can count cell numbers in retinal ganglion cell layers (RGCL). Considering the small size of the adult zebrafish skull and the high risk of death after drilling on the skull, we open the skull with the help of acid-etching and seal the hole with a light curing bond, which could significantly improve the survival rate. After absorbing the dyes for 5 days, almost all the RGCs are labeled. As this method does not need to transect the optic nerve, it is irreplaceable in the research of RGCs survival after optic nerve crush in adult zebrafish. Here, we introduce this method step by step and provide representative results.

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

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs ...

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

The zebrafish (Danio rerio) is commonly used as a vertebrate model in developmental studies and is particularly suitable for visual neuroscience. For functional measurements of visual performance, electroretinography (ERG) is an ideal non-invasive method, which has been well established in higher vertebrate species. This approach is increasingly being used for examining the visual function in zebrafish, including during the early developmental larval stages. However, the most commonly used recording electrode for larval zebrafish ERG to date is the glass micropipette electrode, which requires specialized equipment for its manufacture, presenting a challenge for laboratories with limited resources. Here, we present a larval zebrafish ERG protocol using a cone-shaped sponge-tip electrode. The novel electrode is easier to manufacture and handle, more economical, and less likely to damage the larval eye than the glass micropipette. Like previously published ERG methods, the current protocol can assess outer retinal function through photoreceptor and bipolar cell responses, the a- and b-wave, respectively. The protocol can clearly illustrate the refinement of visual function throughout the early development of zebrafish larvae, supporting the utility, sensitivity, and reliability of the novel electrode. The simplified electrode is particularly useful when establishing a new ERG system or modifying existing small-animal ERG apparatus for zebrafish measurement, aiding researchers in the visual neurosciences to use the zebrafish model organism.

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.

The zebrafish (Danio rerio) is commonly used as a vertebrate model in developmental studies and is particularly suitable for visual neuroscience. For ...

Developmental Biology

Using the optokinetic response to study visual function of zebrafish

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the visual functions of larval zebrafish1-5. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adult zabrafish. Here, we introduce how to measure the OKR of adult zebrafish with our simple custom-built apparatus using a new protocol which is established in our lab. Both our apparatus and step-by-step procedure of OKR in adult zebrafish are illustrated in this video. In addition, the measurements of the larval OKR, as well as the optomotor response (OMR) test of adult zebrafish, are also demonstrated in this video. This OKR assay of adult zebrafish in our experiment may last for up to 4 hours. Such OKR test applied in adult fish will benefit to visual function investigation more efficiently when the adult fish vision system is manipulated.
Su-Qi Zou and Wu Yin contributed equally to this paper.

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.

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the ...

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

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis support continuous growth of the entire visual system, both in the eye and the brain. Coordinated growth between the retinae and the optic tectum ensures accurate retinotopic mapping as new neurons are added in the eyes and brain. To address whether retinal axons provide crucial information for regulating tectal stem and progenitor cell behaviors such as survival, proliferation, and/or differentiation, it is necessary to be able to compare innervated and denervated tectal lobes within the same animal and across animals. 
Surgical removal of one eye from living larval zebrafish followed by observation of the optic tectum achieves this goal. The accompanying video demonstrates how to anesthetize larvae, electrolytically sharpen tungsten needles, and use them to remove one eye. It next shows how to dissect brains from fixed zebrafish larvae. Finally, the video provides an overview of the protocol for immunohistochemistry and a demonstration of how to mount stained embryos in low-melting-point agarose for microscopy.

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.

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis ...

Analysis of Skeletal Muscle Defects in Larval Zebrafish by Birefringence and Touch-evoke Escape Response Assays

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular dystrophies1-3, congenital myopathies4,5, and disruptions in sarcomeric assembly6,7, due to high genomic and structural conservation with mammals8. Muscular disorganization and locomotive impairment can be quickly assessed in the zebrafish over the first few days post-fertilization. Two assays to help characterize skeletal muscle defects in zebrafish are birefringence (structural) and touch-evoked escape response (behavioral).
Birefringence is a physical property in which light is rotated as it passes through ordered matter, such as the pseudo-crystalline array of muscle sarcomeres9. It is a simple, noninvasive approach to assess muscle integrity in translucent zebrafish larvae early in development. Wild-type zebrafish with highly organized skeletal muscle appear very bright amidst a dark background when visualized between two polarized light filters, whereas muscle mutants have birefringence patterns specific to the primary muscular disorder they model. Zebrafish modeling muscular dystrophies, diseases characterized by myofiber degeneration followed by repeated rounds of regeneration, exhibit degenerative dark patches in skeletal muscle under polarized light. Nondystrophic myopathies are not associated with necrosis or regenerative changes, but result in disorganized myofibers and skeletal muscle weakness. Myopathic zebrafish typically show an overall reduction in birefringence, reflecting the disorganization of sarcomeres.
The touch-evoked escape assay involves observing an embryo&#39;s swimming behavior in response to tactile stimulation10-12. In comparison to wild-type larvae, mutant larvae frequently display a weak escape contraction, followed by slow swimming or other type of impaired motion that fails to propel the larvae more than a short distance12. The advantage of these assays is that disease progression in the same fish type can be monitored in vivo for several days, and that large numbers of fish can be analyzed in a short time relative to higher vertebrates.

The zebrafish is now an established and powerful tool for modeling muscular dystrophies, congenital myopathies, and related neuromuscular diseases. Birefringence and touch-evoked escape behavior are two common noninvasive assays used to determine the degree of muscular disorganization and locomotive impairment of zebrafish embryos during early development.

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular ...

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

Source: Steele, W. B., et al. <a href="https://www.jove.com/v/57938/" target="_blank">Experimental Protocol for Examining Behavioral Response Profiles in Larval Fish: Application to the Neuro-stimulant Caffeine</a>.&#160;J. Vis. Exp.&#160;(2018).
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.

Source: Steele, W. B., et al. Experimental Protocol for Examining Behavioral Response Profiles in Larval Fish: Application to the Neuro-stimulant ...

JoVE Encyclopedia of Experiments

Danio rerio (zebrafish)

Larva

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

Source: Cameron, D. J., et al. <a href="https://www.jove.com/v/50832/" target="_blank">The Optokinetic Response as a Quantitative Measure of Visual Acuity in Zebrafish</a>.&#160;J. Vis. Exp.&#160;(2013)
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.

Source: Cameron, D. J., et al. The Optokinetic Response as a Quantitative Measure of Visual Acuity in Zebrafish.&#160;J. Vis. Exp.&#160;(2013)
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Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

Summary

Explore More Videos

Chapters in this video

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

VisioTracker, an Innovative Automated Approach to Oculomotor Analysis

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

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

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

Using the optokinetic response to study visual function of zebrafish

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

Analysis of Skeletal Muscle Defects in Larval Zebrafish by Birefringence and Touch-evoke Escape Response Assays

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

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