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 &amp;micro;m filter</td><td>Millex GP</td><td>SLGP033RS</td><td>Filters the 10&amp;times; goldfish ringer&#039;s buffer for sterilizatio</td></tr><tr><td>1 mL syringe</td><td>Terumo</td><td>DVR-5175</td><td>With a 30G &amp;times; &amp;frac12;&quot; needle to add drops of saline to the electrode sponge tip to prevent drying and increased noisein the ERG signals.</td></tr><tr><td>30 G &amp;times; &amp;frac12;&quot; 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&amp;nbsp;</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?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&amp;nbsp;</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&amp;nbsp;</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&amp;amp;E metal</td><td>0.3 mm</td><td>Used to make recording and reference ERG electrodes.</td></tr><tr><td>Stereo microscope&amp;nbsp;</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&amp;nbsp;</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

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

Microsoft Bing

Research

JoVE Journal

Developmental Biology

6.4K Views.  University of Melbourne. 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.

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

Transretinal ERG Recordings from Mouse Retina: Rod and Cone Photoresponses

There are two distinct classes of image-forming photoreceptors in the vertebrate retina: rods and cones. Rods are able to detect single photons of light whereas cones operate continuously under rapidly changing bright light conditions. Absorption of light by rod- and cone-specific visual pigments in the outer segments of photoreceptors triggers a phototransduction cascade that eventually leads to closure of cyclic nucleotide-gated channels on the plasma membrane and cell hyperpolarization. This light-induced change in membrane current and potential can be registered as a photoresponse, by either classical suction electrode recording technique1,2 or by transretinal electroretinogram recordings (ERG) from isolated retinas with pharmacologically blocked postsynaptic response components3-5. The latter method allows drug-accessible long-lasting recordings from mouse photoreceptors and is particularly useful for obtaining stable photoresponses from the scarce and fragile mouse cones. In the case of cones, such experiments can be performed both in dark-adapted conditions and following intense illumination that bleaches essentially all visual pigment, to monitor the process of cone photosensitivity recovery during dark adaptation6,7. In this video, we will show how to perform rod- and M/L-cone-driven transretinal recordings from dark-adapted mouse retina. Rod recordings will be carried out using retina of wild type (C57Bl/6) mice. For simplicity, cone recordings will be obtained from genetically modified rod transducin &alpha;-subunit knockout (T&alpha;-/-) mice which lack rod signaling8.

We describe a relatively simple method of transretinal electroretinogram (ERG) recordings for obtaining rod and cone photoresponses from intact mouse retina. This approach takes advantage of the block of synaptic transmission from photoreceptors to isolate their light responses and record them using field electrodes placed across the isolated flat-mounted retina.

There are two distinct classes of image-forming photoreceptors in the vertebrate retina: rods and cones. Rods are able to detect single photons of light ...

Neuroscience

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment are open and allow cations to flow steadily inwards across the membrane, depolarizing the cell. Light exposure triggers the closure of the CNG channels, blocks the inward cation current flow, and thus results in cell hyperpolarization. Based on the polarity of photoreceptors, a suction recording method was developed in 1970s that, unlike the classic patch-clamp technique, does not require penetrating the plasma membrane 1. Drawing the outer segment into a tightly-fitting glass pipette filled with extracellular solution allows recording the current changes in individual cells upon test-flash exposure. However, this well-established "outer-segment-in (OS-in)" suction recording is not suitable for mouse cone recordings, because of the low percentage of cones in the mouse retina (3%) and the difficulties in identifying the cone outer segments. Recently, an inner-segment-in (IS-in) recording configuration was developed to draw the inner segment/nuclear region of the photoreceptor into the recording pipette 2,3. In this video, we will show how to record from individual mouse cone photoresponses using single-cell suction electrode.

We will show how to record flash responses from single mouse cones using a suction electrode.

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment ...

Biology

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.

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 ...

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing organisms, combined with the embryonic optical clarity, served as initial compelling attributes of this model. Over the past two decades, the success of this model has been further propelled by its amenability to large-scale mutagenesis screens and by the ease of transgenesis. More recently, gene-editing approaches have extended the power of the model.
For neurodevelopmental studies, the zebrafish embryo and larva provide a model to which multiple methods can be applied. Here, we focus on methods that allow the study of an essential property of neurons, electrical excitability. Our preparation for the electrophysiological study of zebrafish spinal neurons involves the use of veterinarian suture glue to secure the preparation to a recording chamber. Alternative methods for recording from zebrafish embryos and larvae involve the attachment of the preparation to the chamber using a fine tungsten pin1,2,3,4,5. A tungsten pin is most often used to mount the preparation in a lateral orientation, although it has been used to mount larvae dorsal-side up4. The suture glue has been used to mount embryos and larvae in both orientations. Using the glue, a minimal dissection can be performed, allowing access to spinal neurons without the use of an enzymatic treatment, thereby avoiding any resultant damage. However, for larvae, it is necessary to apply a brief enzyme treatment to remove the muscle tissue surrounding the spinal cord. The methods described here have been used to study the intrinsic electrical properties of motor neurons, interneurons, and sensory neurons at several developmental stages6,7,8,9.

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

This manuscript describes methods for electrophysiological recordings from spinal neurons of zebrafish embryos and larvae. The preparation maintains neurons in situ and often involves minimum dissection. These methods allow for the electrophysiological study of a variety of spinal neurons, from the initial electrical excitability acquisition through the early larval stages.

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing ...

Behavioral And Physiological Analysis In A Zebrafish Model Of Epilepsy

Epilepsy represents one of the most common neurological disorders, affecting an estimated 50 million people worldwide. Recent advances in genetic research have uncovered a large spectrum of genes implicated in various forms of epilepsy, highlighting the heterogeneous nature of this disorder. Appropriate animal models are essential for investigating the pathological mechanisms triggered by genetic mutations implicated in epilepsy and for developing specialized, targeted therapies. In recent years, zebrafish has emerged as a valuable vertebrate organism for modeling epilepsies, with the use of both genetic manipulation and exposure to known epileptogenic drugs, such as pentylenetetrazole (PTZ), to identify novel anti-epileptic therapeutics. Deleterious mutations in the mTOR regulator DEPDC5 have been associated with various forms of focal epilepsies and knock-down of the zebrafish orthologue causes hyperactivity associated with spontaneous seizure-like episodes, as well as enhanced electrographic activity and characteristic turn wheel swimming. Here, we described the method involved in generating the DEPDC5 loss-of-function model and illustrate the protocol for assessing motor activity at 28 and 48 h post fertilization (hpf), as well as a method for recording field activity in the zebrafish optic tectum. An illustration of the effect of the epileptogenic drug PTZ on neuronal activity over time is also provided.

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

Epilepsy represents one of the most common neurological disorders, affecting an estimated 50 million people worldwide. Recent advances in genetic research ...

Activity of Posterior Lateral Line Afferent Neurons during Swimming in Zebrafish

Sensory systems gather cues essential for directing behavior, but animals must decipher what information is biologically relevant. Locomotion generates reafferent cues that animals must disentangle from relevant sensory cues of the surrounding environment. For example, when a fish swims, flow generated from body undulations is detected by the mechanoreceptive neuromasts, comprising hair cells, that compose the lateral line system. The hair cells then transmit fluid motion information from the sensor to the brain via the sensory afferent neurons. Concurrently, corollary discharge of the motor command is relayed to hair cells to prevent sensory overload. Accounting for the inhibitory effect of predictive motor signals during locomotion is, therefore, critical when evaluating the sensitivity of the lateral line system. We have developed an in vivo electrophysiological approach to simultaneously monitor posterior lateral line afferent neuron and ventral motor root activity in zebrafish larvae (4-7 days post fertilization) that can last for several hours. Extracellular recordings of afferent neurons are achieved using the loose patch clamp technique, which can detect activity from single or multiple neurons. Ventral root recordings are performed through the skin with glass electrodes to detect motor neuron activity. Our experimental protocol provides the potential to monitor endogenous or evoked changes in sensory input across motor behaviors in an intact, behaving vertebrate.

We describe a protocol to monitor changes in the afferent neuron activity during motor commands in a model vertebrate hair cell system.

Sensory systems gather cues essential for directing behavior, but animals must decipher what information is biologically relevant. Locomotion generates ...

Adult Zebrafish Electrophysiology: A Method to Measure the Extracellular Electrical Activity of Neurons In Vivo

Source: Johnston, L., et al. <a href="https://www.jove.com/v/51065/" target="_blank">Electrophysiological Recording in the Brain of Intact Adult Zebrafish</a>. &#160;J. Vis. Exp. (2013)
The protocol describes the procedure to perform electrophysiology in adult zebrafish. This method can be used for measuring neural activity in vivo.

Elektrofizjologia dorosłego danio pręgowanego: metoda pomiaru zewnątrzkomórkowej aktywności elektrycznej neuronów in vivo

Source: Johnston, L., et al. Electrophysiological Recording in the Brain of Intact Adult Zebrafish. &#160;J. Vis. Exp. (2013)
The protocol describes the ...

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

Podsumowanie

Przeglądaj więcej filmów

Rozdziały w tym wideo

Transretinal ERG Recordings from Mouse Retina: Rod and Cone Photoresponses

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Forebrain Electrophysiological Recording in Larval Zebrafish

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

Behavioral And Physiological Analysis In A Zebrafish Model Of Epilepsy

Activity of Posterior Lateral Line Afferent Neurons during Swimming in Zebrafish

Elektrofizjologia dorosłego danio pręgowanego: metoda pomiaru zewnątrzkomórkowej aktywności elektrycznej neuronów in vivo