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.

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	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;"><a name="RANGE!A1:D30">Name of the Material/Equipment</a></td>
			<td style="width:168px;">Company</td>
			<td style="width:165px;">Catalog Number</td>
			<td style="width:1504px;">Comments/ Description (optional)</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Faraday cage</td>
			<td>80/20 Inc</td>
			<td>custom</td>
			<td>Custom designed aluminum &#34;Industrial Erector Set&#34; for Cage framework</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">PVA sponge</td>
			<td>Amazon</td>
			<td>B000ZOWG1C</td>
			<td>Provides a soft, moist platform for placement of zebrafish larvae</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;">Micromanipulator</td>
			<td>Drummond</td>
			<td>3-000-024-R</td>
			<td>Holding and positioning the recording microelectrode</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;width:333px;">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 height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;">Steriflip 50 ml filters</td>
			<td>Millipore</td>
			<td>SCGP00525</td>
			<td>Filtering solutions to prevent small articulates from blocking micropipettes</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">BNC adaptor</td>
			<td>Monoprice</td>
			<td>4127</td>
			<td>Connecting camera to BNC cable</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">BNC cable</td>
			<td>Monoprice</td>
			<td>626</td>
			<td>Connecting camera to video adaptor</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Camera lens</td>
			<td>Navitar</td>
			<td>1582232</td>
			<td>Visualizing the positioning of the recording microelectrode onto the larval cornea</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Camera coupler</td>
			<td>Navitar</td>
			<td>1501149</td>
			<td>Visualizing the positioning of the recording microelectrode onto the larval cornea</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;">Borosilicate glass</td>
			<td>Sutter</td>
			<td>BF-150-86-10</td>
			<td>Fire- polished borosilicate glass (metling temperature = 821&#176;C) with filament and dimensions of 1.5mm x 0.86 mm (outer diameter by inner diameter)&#160;</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">P97 Flaming/Brown puller</td>
			<td>Sutter</td>
			<td>P97</td>
			<td>For pulling glass micropipettes</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;">Breadboard</td>
			<td>Thorlabs</td>
			<td>B2436F</td>
			<td>Vibration isolation platfrom for ERG stimulator and zebrafish specimen</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Passive Isolation Mounts</td>
			<td>Thorlabs</td>
			<td>PWA074</td>
			<td>Provides vibration isolation to breadboard</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Copper mesh</td>
			<td>TWP</td>
			<td>022X022C0150W36T</td>
			<td>To line Faraday Cage</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Pipette pump</td>
			<td>VWR</td>
			<td>53502-233</td>
			<td>Used with Pasteur pipettes to carefully transfer zebrafish larvae</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Pasteur pipettes</td>
			<td>VWR</td>
			<td>14672-608</td>
			<td>Used with Pipette pump to carefully transfer zebrafish larvae</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Camera</td>
			<td>Watec</td>
			<td>WAT-902B</td>
			<td>Visualizing the positioning of the recording microelectrode onto the larval cornea</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Tricaine (MS-222)</td>
			<td>Western Chemical</td>
			<td>Tricaine-S</td>
			<td>Pharmaceutical-grade anesthetic,</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Micro-fil</td>
			<td>WPI</td>
			<td>MF28G-5</td>
			<td>Filling microelectrode holder and microelectrode glass</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Microelectrode holder</td>
			<td>WPI</td>
			<td>MEH2SW15</td>
			<td>Holds glass microelectrode, connects to ERG equipment</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;">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 vistas.  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

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

Drosophila larval locomotion is a splendid model system in developmental and physiological neuroscience, by virtue of the genetic accessibility of the underlying neuronal components in the circuits1-6. Application of optogenetics7,8 in the larval neural circuit allows us to manipulate neuronal activity in spatially and temporally patterned ways9-13. Typically, specimens are broadly illuminated with a mercury lamp or LED, so specificity of the target neurons is controlled by binary gene expression systems such as the Gal4-UAS system14,15. In this work, to improve the spatial resolution to "sub-genetic resolution", we locally illuminated a subset of neurons in the ventral nerve cord using lasers implemented in a conventional confocal microscope. While monitoring the motion of the body wall of the semi-intact larvae, we interactively activated or inhibited neural activity with channelrhodopsin16,17 or halorhodopsin18-20, respectively. By spatially and temporally restricted illumination of the neural tissue, we can manipulate the activity of specific neurons in the circuit at a specific phase of behavior. This method is useful for studying the relationship between the activities of a local neural assembly in the ventral nerve cord and the spatiotemporal pattern of motor output.

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

Here we describe a protocol for optogenetic manipulation of motoneuronal activity while monitoring changes in motor output (muscle contraction) in semi-intact Drosophila larvae using lasers within a conventional confocal microscope. This technique enables researchers to achieve local perturbation of neural activity in a few neuromeres to elucidate the dynamics of motor circuits.

Drosophila larval locomotion is a splendid model system in developmental and physiological neuroscience, by virtue of the genetic accessibility of the ...

Automated High-throughput Behavioral Analyses in Zebrafish Larvae

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system measures spontaneous behaviors and the response to an aversive stimulus, which is shown to the larvae via a PowerPoint presentation. The recorded images are analyzed with an ImageJ macro, which automatically splits the color channels, subtracts the background, and applies a threshold to identify individual larvae placement in the lanes. We can then import the coordinates into an Excel sheet to quantify swim speed, preference for edge or side of the lane, resting behavior, thigmotaxis, distance between larvae, and avoidance behavior. Subtle changes in behavior are easily detected using our system, making it useful for behavioral analyses after exposure to environmental toxicants or pharmaceuticals.

Our laboratory developed a novel high-throughput automated imaging system that is useful for the detection of several different behaviors in 7-day-old zebrafish larvae. The system can be used for detecting subtle changes in behavior after the larvae have been exposed to environmental toxicants or pharmaceuticals.

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system ...

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  ...

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 ...

Ablation of a Neuronal Population Using a Two-photon Laser and Its Assessment Using Calcium Imaging and Behavioral Recording in Zebrafish Larvae

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ablation of neurons is an effective method for this purpose when neurons are selectively labeled with fluorescent probes. In the present study, protocols for laser ablating a subpopulation of neurons using a two-photon microscope and testing of its functional and behavioral consequences are described. In this study, prey capture behavior in zebrafish larvae is used as a study model. The pretecto-hypothalamic circuit is known to underlie this visually-driven prey catching behavior. Zebrafish pretectum were laser-ablated, and neuronal activity in the inferior lobe of the hypothalamus (ILH; the target of the pretectal projection) was examined. Prey capture behavior after pretectal ablation was also tested.

Here, we present a protocol to ablate a genetically labeled subpopulation of neurons by a two-photon laser from Zebrafish larvae.

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ...

Using Optical Coherence Tomography and Optokinetic Response As Structural and Functional Visual System Readouts in Mice and Rats

Optical coherence tomography (OCT) is a fast, non-invasive, interferometric technique allowing high-resolution retinal imaging. It is an ideal tool for the investigation of processes of neurodegeneration, neuroprotection and neuro-repair involving the visual system, as these often correlate well with retinal changes. As a functional readout, visually evoked compensatory eye and head movements are commonly used in experimental models involving the visual function. Combining both techniques allows a quantitative in vivo investigation of structure and function, which can be used to investigate the pathological conditions or to evaluate the potential of novel therapeutics. A great benefit of the presented techniques is the possibility to perform longitudinal analyses allowing the investigation of dynamic processes, reducing variability and cuts down the number of animals needed for the experiments. The protocol described aims to provide a manual for acquisition and analysis of high quality retinal scans of mice and rats using a low cost customized holder with an option to deliver inhalational anesthesia. Additionally, the proposed guide is intended as an instructional manual for researchers using optokinetic response (OKR) analysis in rodents, which can be adapted to their specific needs and interests.

A detailed protocol for the assessment of structural and visual readouts in rodents by optical coherence tomography and optokinetic response is presented. The results provide valuable insights for ophthalmologic as well as neurologic research.

Optical coherence tomography (OCT) is a fast, non-invasive, interferometric technique allowing high-resolution retinal imaging. It is an ideal tool for ...

Using Zebrafish Larvae to Study the Pathological Consequences of Hemorrhagic Stroke

Despite being the most severe subtype of stroke with high global mortality, there is no specific treatment for patients with intracerebral hemorrhage (ICH). Modelling ICH pre-clinically has proven difficult, and current rodent models poorly recapitulate the spontaneous nature of human ICH. Therefore, there is an urgent requirement for alternative pre-clinical methodologies for study of disease mechanisms in ICH and for potential drug discovery.
The use of zebrafish represents an increasingly popular approach for translational research, primarily due to a number of advantages they possess over mammalian models of disease, including prolific reproduction rates and larval transparency allowing for live imaging. Other groups have established that zebrafish larvae can exhibit spontaneous ICH following genetic or chemical disruption of cerebrovascular development. The aim of this methodology is to utilize such models to study the pathological consequences of brain hemorrhage, in the context of pre-clinical ICH research. By using live imaging and motility assays, brain damage, neuroinflammation and locomotor function following ICH can be assessed and quantified.
This study shows that key pathological consequences of brain hemorrhage in humans are conserved in zebrafish larvae highlighting the model organism as a valuable in vivo&#160;system for pre-clinical investigation of ICH. The aim of this methodology is to enable the pre-clinical stroke community to employ the zebrafish larval model as an alternative complementary model system to rodents.

Here we present a protocol to quantify brain injury, locomotor deficits and neuroinflammation following bleeding in the brain in zebrafish larvae, in the context of human intracerebral hemorrhage (ICH).

Despite being the most severe subtype of stroke with high global mortality, there is no specific treatment for patients with intracerebral hemorrhage ...

In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at cellular and subcellular resolution. Advances in molecular and imaging technologies have allowed us to gain numerous detailed insights into cellular and molecular mechanisms of brain development in the transparent zebrafish embryo. Recently, processes of refinement of neuronal connectivity that occur at later larval stages several weeks after fertilization, which are for example control of social behavior, decision making or motivation-driven behavior, have moved into focus of research. At these stages, pigmentation of the zebrafish skin interferes with light penetration into brain tissue, and solutions for embryonic stages, e.g., pharmacological inhibition of pigmentation, are not feasible anymore.
Therefore, a minimally invasive surgical solution for microscopy access to the brain of awake zebrafish is provided that is derived from electrophysiological&#160;approaches. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, exposing underlying neurons and axonal tracts without damage. This allows for recording neuronal morphology, including synaptic structures and their molecular contents, and the observation of physiological changes such as Ca2+ transients or intracellular transport events. In addition, interrogation of these processes by means of pharmacological inhibition or optogenetic manipulation is feasible. This brain exposure approach provides information about structural and physiological changes in neurons as well as the correlation and interdependence of these events in live brain tissue in the range of minutes or hours. The technique is suitable for in vivo brain imaging of zebrafish larvae up to 30 days post fertilization, the latest developmental stage tested so far. It, thus, provides access to such important questions as synaptic refinement and scaling, axonal and dendritic transport, synaptic targeting of cytoskeletal cargo or local activity-dependent expression. Therefore, a broad use for this mounting and imaging approach can be anticipated.

Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at ...