Funding for this project was provided by a grant from the Melbourne Neuroscience Institute (to PTG, PRJ &#38; BVB).

All electroretinogram (ERG) procedures were performed according to the provisions of the Australian National Health and Medical Research Council code of practice for the care and use of animals and were approved by the institutional animal ethics committee at the University of Melbourne.
1. Buffer Preparation
<ol>
	<li>Prepare the 10x goldfish Ringer&#8217;s buffer (1.25 M NaCl, 26 mM KCl, 25 mm CaCl2, 10 mM MgCl2, 100 mM glucose, 100 mM HEPES) using reverse osmosis (RO) water. Adjust the buffer to pH 7.8 and sterilize the buffer using 0.22 &#956;m filter. Store the 10x buffer at 4 &#176;C as the solution stock3. 
	NOTE: The 10x Ringer&#8217;s buffer should be used within 3 months.</li>
	<li>On the day of the experiment, make 1x goldfish Ringer&#8217;s buffer by diluting the 10x goldfish Ringer&#8217;s buffer using reverse osmosis water. 
	NOTE: The 1x goldfish Ringer&#8217;s buffer is used to saturate the PVA sponge, including the sponge used for the placement of larval zebrafish, and the sponge on the recording electrode tip.</li>
</ol>
2. Electrode Preparation
<ol>
	<li>Prepare the cone-shaped sponge recording electrode.
	<ol>
		<li>Cut the male end from the platinum electrode lead extension and remove from the end 10 mm of the outer polytetrafluoroethylene insulation coating using a scalpel blade. Take care not to damage the inner wire of the electrode lead.</li>
		<li>Cut a 40 mm length of silver wire (0.3 mm diameter) and securely attach this to the electrode lead by entwining the silver wire with the exposed inner wire. Encase the joint using insulating tape, leaving a ~15 mm length of silver wire exposed (Figure 1A).</li>
		<li>Electroplate the exposed silver wire with chloride using a 9 V DC source for 60 s to improve signal conduction. Immerse the exposed silver tip in normal saline and connect the other end to the positive terminal of the battery. Connect another wire to the negative terminal of the battery and immerse the other end of the wire into the saline2. 
		NOTE: Alternatively, chlorinate the silver wire by soaking it for 1 hour in a bleach solution (active ingredient 42 g/L sodium hypochlorite).</li>
		<li>Cut a ~20 mm x 20 mm square of PVA sponge using scissors to make a cone (Figure 1A). Saturate the sponge using 1x Ringer&#8217;s buffer. Under a microscope with a scale bar on the eyepiece, use a scalpel blade to shape the apex of the cone to ~40 &#181;m diameter. Air dry the cone-shaped sponge on absorbent paper tissue until it is solid. 
		NOTE: The PVA sponge expands significantly when saturated, thus it is important that the sponge is first saturated with saline before shaping the apex of the cone.</li>
		<li>After chloriding, air dry the silver wire on an absorbent tissue for 5 min. Insert the silver wire into the dried, solid, cone-shaped PVA sponge through the base of the cone. Insulate any excess exposed metal using mask tape to reduce photovoltaic artifacts (Figure 1B-C). 
		NOTE: After each experimental session, remove the sponge from the silver wire. Wash the sponge using reverse osmosis water and air dry for reuse. To ensure optimal signal collection, single use of silver wire is recommended. PVA sponges should not be reused more than 5 times.</li>
		<li>On the day of the experiment, immerse the sponge-tip of the recording electrode into 1x Ringer&#8217;s buffer for at least 15 min to fully saturate the sponge.</li>
	</ol>
	</li>
	<li>Prepare reference electrodes as described above, but without attaching the sponge tip.</li>
	<li>Obtain the ground electrode commercially.</li>
</ol>
3. Zebrafish Preparation
<ol>
	<li>Dark adapt zebrafish larvae overnight (&#62;8 h) prior to recordings by placing zebrafish in a 15 mL tube (&#60;20 larvae per tube) wrapped in aluminum foil in a dark incubator. Remove the lid to ensure adequate oxygen supply.</li>
	<li>On the day of recording, tighten the lid to the foil-wrapped falcon tube containing larvae and ensure that the tube is light-proof. Transport larvae to the ERG lab.</li>
	<li>Pour the fish into Petri dishes in the dark with the assistance of dim red illumination from a light-emitting diode (LED; 17.4 cd.m-2, &#955;max 600 nm). Cover Petri dishes using light-proof towels to minimize light exposure.</li>
</ol>
4. Sponge Platform Preparation
<ol>
	<li>Cut a rectangle of dry PVA sponge to fit snugly in a 35 mm Petri dish. Ensure that the thickness of the sponge should be roughly equal to the depth of the petri dish.</li>
	<li>Make a small cut vertically through one end of the sponge to accommodate the silver wire of the reference electrode.</li>
	<li>Soak the PVA sponge in 1x goldfish Ringer&#8217;s buffer until saturated. Then, place the sponge in a clean 35-mm petri dish. Use a paper towel to absorb extra liquid until no solution exudes from the sponge in response to a light finger press.</li>
</ol>
5. Animal and Electrode Positioning
<ol>
	<li>Anesthetize the larvae using 0.02 % tricaine diluted in 1x goldfish Ringer&#8217;s buffer.</li>
	<li>Use a 3 mL Pasteur pipette to transfer an anesthetized larva onto a square of paper towel (~3 cm2).</li>
	<li>Place the paper towel containing the larva on the moist sponge platform using forceps. Use a fine brush soaked in Ringer&#8217;s buffer to adjust the position of the larva. Ensure that one eye faces upwards, isolated from any nearby liquid on the square of paper towel underneath the larva.</li>
	<li>Glaze the larval body, excluding the head, with moisturizing eye gel to keep the larva moist throughout the ERG recording.</li>
	<li>Position the Petri dish with sponge platform on a small water-heated platform in front of the Ganzfeld bowl light stimulus situated inside a Faraday cage (Figure 1D). 
	NOTE: Maintenance of the temperature of the sponge and the larval body ensures stable ERG signals.</li>
	<li>Insert the reference electrode into the cut made in the platform sponge (Figure 1D).</li>
	<li>Connect the commercially obtained ground electrode to the Faraday cage.</li>
	<li>Attach the recording electrode to an electrode holder and secure the holder to the stereotaxic arm of a micromanipulator (Figure 1D). Use a 3 mL Pasteur pipette to drip one drop of 1x Ringer&#8217;s solution on the sponge tip of the electrode for re-saturation.</li>
	<li>Position the microscope in the Faraday cage over the ERG platform for placement of the electrode. 
	NOTE: Illumination should be provided by a dim red LED (17.4 cd.m-2, &#955;max 600 nm) to allow observation of the larva and placement of the active electrode, whilst maintaining dark-adaptation.</li>
	<li>Adjust the position of the sponge platform to allow observation of the larva under the microscope. Then, use absorbent tissue to remove excess liquid from the electrode sponge tip.</li>
	<li>Position the active electrode so that it gently touches the central corneal surface of the larval zebrafish eye (Figure 1E).</li>
	<li>Move the Ganzfeld bowl towards the sponge platform and ensure that the larva is covered by the bowl.</li>
	<li>Close the Faraday cage to reduce extraneous electromagnetic noise.</li>
</ol>
6. Electroretinogram Recording
<ol>
	<li>Use the computer software of the particular ERG system (see Table of Materials for details) to trigger the stimulus and acquire data based on the settings recommended below2.
	<ol>
		<li>Set the sampling rate of the system to 4 KHz over a 650 ms recording window (2,560 points) in the acquisition software.</li>
		<li>Set the gain of the system to 1,000&#215;.</li>
		<li>Set band-pass filtering of the system to 1&#8211;300 Hz.</li>
		<li>Use a notch filter to reduce 60 Hz (or 50 Hz, depending on local utility frequency) noise. 
		NOTE: The ideal noise level should be no more than &#177;10 &#181;V.</li>
	</ol>
	</li>
	<li>Commence data collection using the procedure described below.
	<ol>
		<li>Use a single test-flash (0.06 log cd.s/m2) to measure a test response from the eye to assess the positioning of electrodes. 
		NOTE: This intensity of test flash should result in a b-wave amplitude greater than 25 &#181;V in 4-dpf larvae. If a robust response cannot be measured, then reposition the electrodes and do another test flash to confirm that electrodes are well positioned.</li>
		<li>Following the test-flash, allow the animal to dark adapt for 3 min in complete darkness before recordings.</li>
		<li>Present flashes from dimmer to brighter light intensities.</li>
		<li>Average signals across repeats according to the signal-to-noise level. 
		NOTE: Generally, average more signals at the dimmer light levels (no fewer than 3 repeats) and fewer at the brighter light levels (usually 1 repeat). Gradually lengthen the inter-stimulus interval from 10 to 60 s from the dimmest to brightest light level. A sample protocol is shown in Table 1.</li>
		<li>After the recordings, humanely kill larvae using 0.1% tricaine.</li>
	</ol>
	</li>
</ol>
7. Analysis
<ol>
	<li>Measure the a-wave amplitude from baseline to the negative a-wave trough and the b-wave amplitude from the negative a-wave trough to the positive b-wave peak.</li>
	<li>Measure the a- and b-wave implicit times from stimulus onset to the trough of the a-wave and the peak of the b-wave, respectively.</li>
</ol>

<ol>
	<li>Roper, C., Tanguay, R. L., Slikker, W., Paule, M. G., Wang, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Handbook of Developmental Neurotoxicology (Second Edition). , 143-151 (2018).</li><li>Nguyen, C. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+Recording+of+Electroretinography+and+Visual+Evoked+Potentials+in+Anesthetized+Rats.">Simultaneous Recording of Electroretinography and Visual Evoked Potentials in Anesthetized Rats.</a> Journal of visualized experiments: JoVE. , e54158 (2016).</li><li>Chrispell, J. D., Rebrik, T. I., Weiss, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroretinogram+analysis+of+the+visual+response+in+zebrafish+larvae.">Electroretinogram analysis of the visual response in zebrafish larvae.</a> Journal of visualized expriment: JoVE. (97), (2015).</li><li>Seeliger, M. W., Rilk, A., Neuhauss, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ganzfeld+ERG+in+zebrafish+larvae.">Ganzfeld ERG in zebrafish larvae.</a> Documenta Ophthalmologica. 104 (1), 57-68 (2002).</li><li>Fleisch, V. C., Jametti, T., Neuhauss, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electroretinogram+(ERG)+Measurements+in+Larval+Zebrafish.">Electroretinogram (ERG) Measurements in Larval Zebrafish.</a> Cold Spring Harbor Protocols. 2008, (2008).</li><li>Biehlmaier, O., Neuhauss, S. C., Kohler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+plasticity+and+functionality+at+the+cone+terminal+of+the+developing+zebrafish+retina.">Synaptic plasticity and functionality at the cone terminal of the developing zebrafish retina.</a> Developmental Neurobiololgy. 56 (3), 222-236 (2003).</li><li>Gestri, G., Link, B. A., Neuhauss, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+visual+system+of+zebrafish+and+its+use+to+model+human+ocular+diseases.">The visual system of zebrafish and its use to model human ocular diseases.</a> Developmental Neurobiololgy. 72 (3), 302-327 (2012).</li><li>Saszik, S., Bilotta, J., Givin, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ERG+assessment+of+zebrafish+retinal+development.">ERG assessment of zebrafish retinal development.</a> Visual Neuroscience. 16 (5), 881-888 (1999).</li><li>Niklaus, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cocaine+accumulation+in+zebrafish+eyes+leads+to+augmented+amplitudes+in+the+electroretinogram.">Cocaine accumulation in zebrafish eyes leads to augmented amplitudes in the electroretinogram.</a> Matters. 3 (6), e201703000003 (2017).</li><li>Tanvir, Z., Nelson, R. F., DeCicco-Skinner, K., Connaughton, V. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=One+month+of+hyperglycemia+alters+spectral+responses+of+the+zebrafish+photopic+ERG.">One month of hyperglycemia alters spectral responses of the zebrafish photopic ERG.</a> Disease models &#38; mechanisms. , (2018).</li><li>Kakiuchi, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oscillatory+potentials+in+electroretinogram+as+an+early+marker+of+visual+abnormalities+in+vitamin+A+deficiency.">Oscillatory potentials in electroretinogram as an early marker of visual abnormalities in vitamin A deficiency.</a> Molecular medicine reports. 11 (2), 995-1003 (2015).</li><li>Emran, F., Rihel, J., Adolph, A. R., Dowling, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+larvae+lose+vision+at+night.">Zebrafish larvae lose vision at night.</a> Proceedings of the National Academy of Sciences. , (2010).</li><li>Makhankov, Y. V., Rinner, O., Neuhauss, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+inexpensive+device+for+non-invasive+electroretinography+in+small+aquatic+vertebrates.">An inexpensive device for non-invasive electroretinography in small aquatic vertebrates.</a> Journal of Neuroscience Methods. 135 (1-2), 205-210 (2004).</li><li>Bilotta, J., Saszik, S., Sutherland, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rod+contributions+to+the+electroretinogram+of+the+dark-adapted+developing+zebrafish.">Rod contributions to the electroretinogram of the dark-adapted developing zebrafish.</a> Developmental Dynamics. 222 (4), 564-570 (2001).</li><li>Cameron, M. A., Barnard, A. R., Lucas, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+electroretinogram+as+a+method+for+studying+circadian+rhythms+in+the+mammalian+retina.">The electroretinogram as a method for studying circadian rhythms in the mammalian retina.</a> Journal of genetics. 87 (5), 459-466 (2008).</li><li>Bui, B. V., Armitage, J. A., Vingrys, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extraction+and+modelling+of+oscillatory+potentials.">Extraction and modelling of oscillatory potentials.</a> Documenta Ophthalmologica. 104 (1), 17-36 (2002).</li><li>Bui, B. V., Fortune, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ganglion+cell+contributions+to+the+rat+full-field+electroretinogram.">Ganglion cell contributions to the rat full-field electroretinogram.</a> The Journal of Physiology. 555 (1), 153-173 (2004).</li></ol>

The authors have no disclosures relevant to this work.

Functional readouts such as the ERG have become increasingly important in the suite of tools used to study larval zebrafish8,9,12,14. Due to the small size of the larval zebrafish eye, glass micropipettes have been adapted as recording electrodes in most published protocols3,4,5,<sup class="...

The zebrafish (Danio rerio) has become a widely used genetic vertebrate model, including studies of the visual neurosciences. The increasing popularity of this species can be attributed to advantages including ease of genetic manipulation, the highly conserved vertebrate visual system (neuron types, anatomical morphology and organization, and underlying genetics), high fecundity and lower cost of husbandry compared to mammalian models1. The non-invasive electroretinogram (ERG) has long been used clinically to assess human visual function, and in the laboratory setting to quantify vision in a range of large and small species including rodents and larval zebrafish2,3,4,5. The most commonly analyzed ERG components are the a-wave and b-wave, originating from the light-sensing photoreceptors and bipolar interneurons, respectively. In larval zebrafish, distinct layers in the retina are established by 3 days post-fertilization (dpf) and the morphology of the photoreceptor cone terminal synapses mature before 4 dpf6,7. Outer retinal function of larval zebrafish is thus established before 4 dpf, meaning that the ERG is measurable from this early age onwards. Because of the short experimental cycle and the high-throughput properties of the model, the ERG has been applied to larval zebrafish for functional assessment of disease models, analyzing color vision and retinal development, studying visual circadian rhythms and testing drugs8,9,10,11,12.
However, current approaches for larval zebrafish ERG has some complexities that may make it harder to adopt. Published larval zebrafish ERG protocols commonly use a glass micropipette filled with conductive liquid as the recording electrode3,4,5,13, which requires a high quality micropipette tip3. Specialized equipment, such as a micropipette puller and in some cases a microforge, are required for their manufacture. This can be a challenge for laboratories with limited resources and leads to extra costs even when adapting available small animal ERG systems for measurement of larval zebrafish visual function. Even when smoothed, the sharp micropipette tip can damage the surface of the larval eye. Additionally, commercial micropipette holders for electrophysiology are constructed with a fixed silver wire. These fixed wires become passivated after repetitive use, requiring the purchase of new holders leading to increased maintenance costs.
Here we describe an ERG method using a cone-shaped sponge-tip recording electrode, that is particularly useful for adapting established small-animal ERG setups for larval zebrafish ERG measurements. The electrode is easily made using common polyvinyl acetate (PVA) sponge and fine silver wire without any other specialized equipment. Our data show that this novel electrode is sensitive and reliable enough to demonstrate the functional development of retinal neural circuits in larval zebrafish between 4 and 7 dpf. This economical and practical sponge-tip electrode may be useful to researchers establishing new ERG systems or modifying existing small-animal systems, for zebrafish studies.

<table>
	<tbody>
		<tr>
			<td>0.22 &#181;m filter</td>
			<td>Millex GP</td>
			<td>SLGP033RS</td>
			<td>Filters the 10&#215; goldfish ringer&#39;s buffer for sterilizatio</td>
		</tr>
		<tr>
			<td>1-mL syringe</td>
			<td>Terumo</td>
			<td>DVR-5175</td>
			<td>With a 30G &#215; &#189;&#34; needle to add drops of saline to the electrode sponge tip to prevent drying and increased noisein the ERG signals.</td>
		</tr>
		<tr>
			<td>30G &#215; &#189;&#34; needle</td>
			<td>Terumo</td>
			<td>NN*3013R</td>
			<td>For adding saline toteh sopnge tip electrode.</td>
		</tr>
		<tr>
			<td>Bioamplifier</td>
			<td>ADInstruments</td>
			<td>ML135</td>
			<td>For amplifying ERG signals.</td>
		</tr>
		<tr>
			<td>Bleach solution&#160;</td>
			<td>King White</td>
			<td>9333441000973</td>
			<td>For an alternative method of sliver electrode chlorination. Active ingredient: 42 g/L sodium hypochlorite.</td>
		</tr>
		<tr>
			<td>Circulation water bath</td>
			<td>Lauda-Ko&#776;nigshoffen</td>
			<td>MGW Lauda</td>
			<td>Used to make the water-heated platfrom.</td>
		</tr>
		<tr>
			<td>Electrode lead</td>
			<td>Grass Telefactor</td>
			<td>F-E2-30</td>
			<td>Platinum cables for connecting silver wire electrodes to the amplifier.</td>
		</tr>
		<tr>
			<td>Faraday Cage</td>
			<td>Photometric Solution International&#160;</td>
			<td></td>
			<td>For maintianing dark adaptation and enclosing the Ganzfeld setup to improve signal-to-noise ratio.</td>
		</tr>
		<tr>
			<td>Ganzfeld Bowl</td>
			<td>Photometric Solution International&#160;</td>
			<td></td>
			<td>Custom designed light stimulator: 36 mm diameter, 13 cm aperture size.</td>
		</tr>
		<tr>
			<td>Luxeon LEDs</td>
			<td>Phillips Light Co.</td>
			<td></td>
			<td>For light stimulation twenty 5W and one 1W LEDs.</td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>Harvard Apparatus</td>
			<td>BS4 50-2625</td>
			<td>Holds the recording electrode during experiments.</td>
		</tr>
		<tr>
			<td>Microsoft Office Excel</td>
			<td>Microsoft</td>
			<td>version 2010</td>
			<td>Spreadsheet software for data analysis.</td>
		</tr>
		<tr>
			<td>Moisturizing eye gel</td>
			<td>GenTeal Gel</td>
			<td>9319099315560</td>
			<td>Used to cover zebrafish larvae during recordings to avoiding dehydration. Active ingredient: 0.3 % Hypromellose and 0.22 % carbomer 980.</td>
		</tr>
		<tr>
			<td>Pasteur pipette</td>
			<td>Copan</td>
			<td>200C</td>
			<td>Used to caredully transfer larval zebrafish.</td>
		</tr>
		<tr>
			<td>Powerlab data acquisition system</td>
			<td>ADInstruments</td>
			<td>ML785</td>
			<td>Controls the LEDs to generate stimuli.</td>
		</tr>
		<tr>
			<td>PVA sponge</td>
			<td>MeiCheLe</td>
			<td>R-1675</td>
			<td>For the placement of larval zebrafish and making the cone-shaped electrode ti</td>
		</tr>
		<tr>
			<td>Saline solution</td>
			<td>Aaxis Pacific</td>
			<td>13317002</td>
			<td>For electroplating silver wire electrode.</td>
		</tr>
		<tr>
			<td>Scope Software</td>
			<td>ADInstruments</td>
			<td>version 3.7.6</td>
			<td>Simultaneously triggers the stimulus through the Powerlab system and collects data</td>
		</tr>
		<tr>
			<td>Silver (fine round wire)</td>
			<td>A&#38;E metal</td>
			<td>0.3 mm</td>
			<td>Used to make recording and reference ERG electrodes.</td>
		</tr>
		<tr>
			<td>Stereo microscope&#160;</td>
			<td>Leica</td>
			<td>M80</td>
			<td>Used to shape and measure the cone-shaped sponge apex (with scale bar on eyepiece). Positioned in the Faraday cage for electrode placement.</td>
		</tr>
		<tr>
			<td>Tricaine&#160;</td>
			<td>Sigma-aldrich</td>
			<td>E10521-50G</td>
			<td>For anaethetizing larval zebrafish.</td>
		</tr>
		<tr>
			<td>Water-heated platform</td>
			<td>custom-made</td>
			<td></td>
			<td>For maintianing the temperature of the sponge platform and the larval body during ERG recordings</td>
		</tr>
	</tbody>
</table>

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.

This section provides representative results for ERG measurements taken daily from 4 to 7 dpf. From 4 dpf, ERG responses show robust a- and b-wave components, which arise from photoreceptors and bipolar cells, respectively. At each age tested, the amplitude of the b-wave increased with light intensity (Figure 2; Figure 3). Notably, the sensitivity of the larval zebrafish retina to dimmer flashes increased with age. The a- and b-wave were not ...

Watch this Scientific Journal Video about Electroretinogram Recording in Larval Zebrafish using A Novel Cone-Shaped Sponge-tip Electrode at JoVE.com

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

Électrorétinogramme enregistrement chez le poisson zèbre larvaire, à l’aide d’électrode de pointe éponge en forme de cône de roman A

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 ...

electroretinogram-recording-larval-zebrafish-using-novel-cone-shaped

Research

JoVE Journal

Developmental Biology

6.3K vues.  University of Melbourne. Ce protocole permet de simplifier les mesures des électrorétogrammes ou des ERG chez le poisson zèbre larvaire qui fournit une lecture fonctionnelle importante des modèles de développement visuel et de maladies génétiques. Les extrémités douces d’éponge de l’enregistrement des électrodes utilisées dans ce protocole est facile à fabriquer en utilisant des matériaux économiques et facilement disponibles et évite les dommages potentiels aux yeux larvaire. L’électrode à po...

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

Live Imaging of Innate Immune and Preneoplastic Cell Interactions Using an Inducible Gal4/UAS Expression System in Larval Zebrafish Skin

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS expression system1 in zebrafish larvae, and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells. The KalTA4-ERT2/UAS system is both inducible and reversible which allows us to induce cell transformation with precise temporal/spatial resolution in vivo. This provides us with a unique opportunity to live image how individual preneoplastic cells interact with host tissues as soon as they emerge, then follow their progression as well as regression. Recent studies in zebrafish larvae have shown a trophic function of innate immunity in the earliest stages of tumorigenesis2,3. Our inducible system would allow us to live image the onset of cellular transformation and the subsequent host response, which may lead to important insights on the underlying mechanisms for the regulation of oncogenic trophic inflammatory responses. We also discuss how one might adapt our protocol to achieve temporal and spatial control of ectopic gene expression in any tissue of interest.

Studying the earliest events of preneoplastic cell progression and innate immune cell interaction is pivotal to understand and treat cancer. Here we describe a method to conditionally induce epithelial cell transformations and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells in zebrafish larvae.

Imagerie en direct d&#39;Innate immunitaire et prénéoplasiques Interactions cellulaires L&#39;utilisation d&#39;un système d&#39;expression / UAS inductible Gal4 dans des larves de poisson zèbre peau

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS ...

Recherche

Biologie du développement

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

Microbead implantation dans l&#39;embryon de poisson zèbre

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Tracking Cells in GFP-transgenic Zebrafish Using the Photoconvertible PSmOrange System

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing developmental processes as they happen in the living animal. Cells of interest can be labeled by using a tissue specific promoter to drive the expression of a fluorescent protein (FP) for the generation of transgenic lines. Using fluorescent photoconvertible proteins for this purpose additionally allows to precisely follow defined structures within the expression domain. Illuminating the protein in the region of interest, changes its emission spectrum and highlights a particular cell or cell cluster leaving other transgenic cells in their original color. A major limitation is the lack of known promoters for a large number of tissues in the zebrafish. Conversely, gene- and enhancer trap screens have generated enormous transgenic resources discretely labeling literally all embryonic structures mostly with GFP or to a lesser extend red or yellow FPs. An approach to follow defined structures in such transgenic backgrounds would be to additionally introduce a ubiquitous photoconvertible protein, which could be converted in the cell(s) of interest. However, the photoconvertible proteins available involve a green and/or less frequently a red emission state1 and can therefore often not be used to track cells in the FP-background of existing transgenic lines. To circumvent this problem, we have established the PSmOrange system for the zebrafish2,3. Simple microinjection of synthetic mRNA encoding a nuclear form of this protein labels all cell nuclei with orange/red fluorescence. Upon targeted photoconversion of the protein, it switches its emission spectrum to far red. The quantum efficiency and stability of the protein makes PSmOrange a superb cell-tracking tool for zebrafish and possibly other teleost species.

We established the photoconvertible PSmOrange system as a powerful, straight-forward and cost inexpensive tool for in vivo cell tracking in GFP transgenic backgrounds. This protocol describes its application in the zebrafish model system.

Suivi des cellules dans le poisson zèbre transgénique de la GFP-Utilisation du système photoconvertible PSmOrange

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing ...

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration have been thoroughly analyzed in vitro. However, in vivo cell migration somehow differs from in vitro migration, and has proven more difficult to analyze, being less accessible to direct observation and manipulation. This protocol uses the migration of the prospective prechordal plate in the early zebrafish embryo as a model system to study the function of candidate genes in cell migration. Prechordal plate progenitors form a group of cells which, during gastrulation, undergoes a directed migration from the embryonic organizer to the animal pole of the embryo. The proposed protocol uses cell transplantation to create mosaic embryos. This offers the combined advantages of labeling isolated cells, which is key to good imaging, and of limiting gain/loss of function effects to the observed cells, hence ensuring cell-autonomous effects. We describe here how we assessed the function of the TORC2 component Sin1 in cell migration, but the protocol can be used to analyze the function of any candidate gene in controlling cell migration in vivo.

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Combining cell transplantation, cytoskeletal labeling and loss/gain of function approaches, this protocol describes how the migrating zebrafish prospective prechordal plate can be used to analyze the function of a candidate gene in in vivo cell migration.

en cours d&#39;analyse<em&gt; In Vivo</em&gt; Migration cellulaire en utilisant Transplantations cellulaires et imagerie time-lapse en Zebrafish Embryons

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration ...

Using Touch-evoked Response and Locomotion Assays to Assess Muscle Performance and Function in Zebrafish

Zebrafish muscle development is highly conserved with mammalian systems making them an excellent model to study muscle function and disease. Many myopathies affecting skeletal muscle function can be quickly and easily assessed in zebrafish over the first few days of embryogenesis. By 24 hr post-fertilization (hpf), wildtype zebrafish spontaneously contract their tail muscles and by 48 hpf, zebrafish exhibit controlled swimming behaviors. Reduction in the frequency of, or other alterations in, these movements may indicate a skeletal muscle dysfunction. To analyze swimming behavior and assess muscle performance in early zebrafish development, we utilize both touch-evoked escape response and locomotion assays.
Touch-evoked escape response assays can be used to assess muscle performance during short burst movements resulting from contraction of fast-twitch muscle fibers. In response to an external stimulus, which in this case is a tap on the head, wildtype zebrafish at 2 days post-fertilization (dpf) typically exhibit a powerful burst swim, accompanied by sharp turns. Our method quantifies skeletal muscle function by measuring the maximum acceleration during a burst swimming motion, the acceleration being directly proportional to the force produced by muscle contraction.
In contrast, locomotion assays during early zebrafish larval development are used to assess muscle performance during sustained periods of muscle activity. Using a tracking system to monitor swimming behavior, we obtain an automated calculation of the frequency of activity and distance in 6-day old zebrafish, reflective of their skeletal muscle function. Measurements of swimming performance are valuable for phenotypic assessment of disease models and high-throughput screening of mutations or chemical treatments affecting skeletal muscle function.

Zebrafish are an excellent model to study muscle function and disease. During early embryogenesis zebrafish begin regular muscle contractions producing rhythmic swimming behavior, which is altered when the muscle is disrupted. Here we describe a touch-evoked response and locomotion assay to examine swimming performance as a measure of muscle function.

Utilisation de la réponse tactile évoqué et Locomotion essais pour évaluer la performance de la fonction musculaire et en Zebrafish

Zebrafish muscle development is highly conserved with mammalian systems making them an excellent model to study muscle function and disease. Many ...

Dissection of Larval Zebrafish Gonadal Tissue

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex determination system, where several genes distributed throughout the genome collectively determine the sex identities of individual fish. Currently, the genes involved in regulating gonad development and how they work remain elusive. Normally, isolating gonadal tissue is the first step to examine the sex developmental processes. Here, we present a procedure to isolate gonadal tissue from 17 dpf (days post fertilization) and 25 dpf zebrafish larvae. The isolated gonadal tissue may be subsequently examined by morphology and gene expression profiling.

Here, we present a protocol for isolating gonadal tissue of larval zebrafish, which will facilitate investigations of zebrafish sex differentiation and maintenance.

Dissection de tissu gonadique Larval Zebrafish

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Induction de l&#39;hypoxie dans les écrevisses vivantes de grenouille et de poisson zèbre

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

Chagas disease is a parasitic infection caused by Trypanosoma cruzi, whose motility is not only important for localization, but also for cellular binding and invasion. Current animal models for the study of T. cruzi allow limited observation of parasites in vivo, representing a challenge for understanding parasite behavior during the initial stages of infection in humans. This protozoan has a flagellar stage in both vector and mammalian hosts, but there are no studies describing its motility in vivo.The objective of this project was to establish a live vertebrate zebrafish model to evaluate T. cruzi motility in the vascular system. Transparent zebrafish larvae were injected with fluorescently labeled trypomastigotes and observed using light sheet fluorescence microscopy (LSFM), a noninvasive method to visualize live organisms with high optical resolution. The parasites could be visualized for extended periods of time due to this technique&#39;s relatively low risk of photodamage compared to confocal or epifluorescence microscopy. T. cruzi parasites were observed traveling in the circulatory system of live zebrafish in different-sized blood vessels and the yolk. They could also be seen attached to the yolk sac wall and to the atrioventricular valve despite the strong forces associated with heart contractions. LSFM of T. cruzi-inoculated zebrafish larvae is a valuable method that can be used to visualize circulating parasites and evaluate their tropism, migration patterns, and motility in the dynamic environment of the cardiovascular system of a live animal.

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

In this protocol, fluorescently labeled T. cruzi&#160;were injected into transparent zebrafish larvae, and parasite motility was observed in vivo using light sheet fluorescence microscopy.

Mise en place du poisson-zèbre larvaire comme modèle Animal pour enquêter sur Trypanosoma cruzi motilité In Vivo

Chagas disease is a parasitic infection caused by Trypanosoma cruzi, whose motility is not only important for localization, but also for cellular binding ...

Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in vivo imaging techniques can provide a clear window into these processes in the living organism. For example, dividing cells and their progeny can be followed in real time as the nervous system forms. In recent years, technical advances in multicolor techniques have expanded the types of questions that can be investigated. The multicolor Brainbow approach can be used to not only distinguish among like cells, but also to color-code multiple different clones of related cells that each derive from one progenitor cell. This allows for a multiplex lineage analysis of many different clones and their behaviors simultaneously during development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells over real time within the developing zebrafish nervous system. This is particularly useful for following cellular interactions among like cells, which are difficult to label differentially using traditional promoter-driven colors. Our approach can be used for tracking lineage relationships among multiple different clones simultaneously. The large datasets generated using this technique provide rich information that can be compared quantitatively across genetic or pharmacological manipulations. Ultimately the results generated can help to answer systematic questions about how the nervous system develops.

In vivo imaging is a powerful tool that can be used to investigate the cellular mechanisms underlying nervous system development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells in real time within the developing zebrafish nervous system.

Visualiser le cerveau en développement dans le poisson zèbre vivant à l’aide de Brainbow et Time-lapse Confocal Imaging

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in ...

Quantifying Liver Size in Larval Zebrafish Using Brightfield Microscopy

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval liver size in zebrafish HCC models provides a means to rapidly assess the effects of drugs and other manipulations on an oncogene-related phenotype. Here we show how to fix zebrafish larvae, dissect the tissues surrounding the liver, photograph livers using bright-field microscopy, measure liver area, and analyze results. This protocol enables rapid, precise quantification of liver size. As this method involves measuring liver area, it may underestimate differences in liver volume, and complementary methodologies are required to differentiate between changes in cell size and changes in cell number. The dissection technique described herein is an excellent tool to visualize the liver, gut, and pancreas in their natural positions for myriad downstream applications including immunofluorescence staining and in situ hybridization. The described strategy for quantifying larval liver size is applicable to many aspects of liver development and regeneration.

Here we demonstrate a method for quantifying liver size in larval zebrafish, providing a way to assess the effects of genetic and pharmacologic manipulations on liver growth and development.

Quantifier la taille du foie chez le poisson zèbre larvaire à l'aide de la microscopie Brightfield

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval ...