This work was supported by NIH/NINDS Grant R01NS070159 (to T.M.D, J.D.L and A.T.S).

All experimental procedures were conducted in strict accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and followed protocol #A2011 09-003, which was reviewed, approved, and overseen by the University of Georgia Institutional Animal Care and Use Committee.
1. Equipment Setup
<ol>
	<li>Perfusion system for craniotomy 
	Immobilization of the adult necessitates an intubation system to deliver dissolved oxygen to the fish.&#160; A variety of systems can be utilized, but a simple gravity system consisting of a 60 ml syringe is used. Elevate the pressure head to a height that consistently yields a flow rate of ~1 ml/min. This syringe can be placed in series with the syringes that are then used for the electrophysiological perfusion system.
	<ol>
		<li>Obtain a single 60 ml Luer lock syringe tube and remove the plunger.</li>
		<li>Suspend the syringe approximately 8 in&#160;above the base of a ring-stand using a clamp.</li>
		<li>Connect a one-way stopcock to the end of the syringe. To the opposite end of the stopcock, connect a piece of tubing, 2 mm in diameter, which is long enough to extend to the intubation base.</li>
	</ol>
	</li>
	<li>Perfusion system for electrophysiology 
	For electrophysiological experiments, it is often necessary to introduce a variety of compounds during the course of an experiment. A variety of systems are available, but a gravity system consisting of 60 ml syringes in series can be used. This setup allows for simple, serial introduction of solutions to be carried out by opening the corresponding stopcock.
	<ol>
		<li>Obtain two or more 60 ml Luer lock syringes and remove the plungers from each. One syringe is required to administer habitat water to the fish while each subsequent tube can be used to administer a variety of pharmacological compounds to the fish.</li>
		<li>Connect each syringe to a 3-way stopcock with a Luer connection.</li>
		<li>Use &#8539; in outer diameter (O.D.) tubing to connect the syringes in series with a male Luer lock on one end.</li>
		<li>Secure the device such that the pressure head is elevated to a height of 27 in above the base, or a height that consistently yields a flow rate of ~1 ml/min.</li>
	</ol>
	</li>
	<li>Intubation cannula 
	The intubation cannula facilitates the introduction of fluid to the fish. This setup provides both flexibility and firmness to best position the cannula within the animal&#39;s mouth.
	<ol>
		<li>Using any general pair of scissors (e.g. Fiskars) or a razor blade, remove a 1.5 cm section from the wide end of a P-200 pipette tip.</li>
		<li>Insert a 6 cm x 1 mm piece of tubing into a reducing valve with a female Luer lock cap and silicone ferrule, i.e. a Tuohy Borst adapter, and insert this cannula into the modified pipette tip. Hold the pipette tip in position with the Luer&#160;lock using a short portion of &#8539; in diameter tubing. <img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/51065/51065fig1highres.jpg" src="/files/ftp_upload/51065/51065fig1.jpg" /> 
		Figure 1
		</li>
	</ol>
	</li>
	<li>Intubation base 
	This dish is used to hold the immobilized animal in a stable, upright position and to facilitate the removal of the fluid entering the animal through intubation. If this is not established correctly, pooling can occur, leading to intrusive vibrational signals in the electrophysiological recording.
	<ol>
		<li>Obtain the base dish of a 100 mm x 15 mm plastic Petri dish.</li>
		<li>Using a soldering tool, melt a hole, 6 mm in diameter and 7.5 mm below the lip of the rim, into the side of the dish. Using the same tool, melt a hole, 10 mm in diameter and 11 mm below the lip of the dish ~74&#176; counter-clockwise from the smaller hole. This hole will serve as the drainage hole.</li>
		<li>Insert a tube,1 cm in diameter, into the hole with the end lifted such that the end cannot be blocked. Then insert a P-200 Pipetteman tip into the small hole; this will serve as a dummy cannula.</li>
		<li>With the Petri dish slightly elevated on the side of the larger hole, pour melted canning wax into the dish such that the cannula hole is just covered. The region will serve as a stop, allowing the cannula to be inserted ~3 mm into the mouth of the fish. This set up should be allowed to solidify.</li>
		<li>Using a flame-heated metal edge, carve a channel to hold the fish so that the channel begins from the tip of the cannula and extends approximately 1-2 in.
		<ol>
			<li>Caution must be taken to ensure that a continuous downward angle from the cannula area to the drainage port is formed. <img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/51065/51065fig2highres.jpg" src="/files/ftp_upload/51065/51065fig2.jpg" width="400px" /> 
			Figure 2</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Perfusion setup
	<ol>
		<li>Before beginning, flush the perfusion set-ups with habitat water, ensuring that all air bubbles are removed.</li>
		<li>Add ~30 ml of 0.016% (630 &#956;M) tricaine solution to the craniotomy perfusion setup and allow ~2 ml to run through tubing to ensure that tricaine will be delivered to the animal upon perfusion initiation. See step 2.2 for dilution.</li>
		<li>To the main perfusion setup, add ~50 ml of habitat water to the first tube. Add any desired experimental compounds to each of the remaining tubes.</li>
		<li>Place the cannula setup into the small hole of the intubation base.</li>
		<li>Twist a 1 in&#160;wide strip of 42 cm Kimwipe into a tight spiral and insert it into the drainage tube such that the Kimwipe extends on both ends. This tissue will serve as a wick to remove the perfused liquid and to prevent fluid buildup within the intubation base, which could interfere with electrophysiological recording.</li>
		<li>Insert one end of the tubing into the large hole of the Petri dish, allowing the lower end to extend into an inert dish that is large enough to collect all perfusion waste. This dish will serve as the outflow reservoir for the setup. Position the tissue so that one end will lie near the animal&#39;s abdomen region and the lower extension can drip into the dish.</li>
		<li>Place this completed intubation base near the dissecting microscope. Connect the Luer lock of the cannula to the tricaine perfusion tubing.</li>
	</ol>
	</li>
	<li>Capillary needle and electrodes
	<ol>
		<li>Obtain a primary electrode consisting of 2.5 in&#160;of 0.010 in&#160;silver wire and a secondary electrode which should be made up of 15 in&#160;of the same wire to ground the setup. For the secondary electrode, use a 15 in&#160;section of 0.010 in&#160;silver wire with a soldered tip, which allows it to fit in back of a head-stage.
		<ol>
			<li>Electroplate the tips of both the primary and secondary silver wires with chloride ions.</li>
			<li>Using a micropipette puller, pull a thin-walled, borosilicate capillary needle with a resistance of &#60;15 m&#937;.
			<ol>
				<li>Fill the capillary needle with 2-3 &#956;l of 2 M potassium chloride, or enough to partially cover the chloride-coated tip of the silver wire.</li>
				<li>Insert the primary electrode into the capillary head and mount it in the electrode holder.</li>
				<li>Mount the head-stage into a micromanipulator and then mount the secondary electrode into a second micromanipulator. With the setup, first position each electrode and then fit the soldered tip of the secondary electrode into the back of the head-stage.</li>
			</ol>
			</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Preparation of Solutions, Perfusion System, and Electrophysiological Recording Equipment
<ol>
	<li>Obtain 1 L of habitat water from aquarium of fish.</li>
	<li>0.016% T]tricaine methanesulfonate (tricaine) (50 ml of 630 &#956;M)24.
	<ol>
		<li>Thaw an aliquot of 0.4% Tris-buffered tricaine, pH 7.2.</li>
		<li>Add 2.1 ml of 0.4% Tris-buffered tricaine to 47.9 ml of habitat water and mix.</li>
	</ol>
	</li>
	<li>Thaw stock concentration of desired water-soluble experimental compound (e.g. 300 mM pentylenetetrazol, a common chemoconvulsant).</li>
	<li>Thaw out an aliquot of 1 &#956;g/&#956;l pancuronium bromide in Ringer&#39;s solution.</li>
	<li>Fill both anesthetizing and experimental intubation systems with habitat water and drain at least enough to remove all bubbles from tubes. This must be done because bubbles obstruct fluid flow, leading to asphyxiation of the animal.
	<ol>
		<li>Completely drain the perfusion tubes that will hold drugs without allowing air into the connecting tubing.</li>
		<li>Fill the anesthetizing tube with 0.016% (630 &#956;M) tricaine solution and place any experimental compound(s) in additional tube(s) on experimental perfusion system.
		<ol>
			<li>The first tube of experimental perfusion should contain only habitat water.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Moisten the small pore sponge with habitat water and place in an empty 60 mm x 15 mm Petri dish. This dish will be used to hold the fish while the anesthetic is injected.</li>
</ol>
3. Craniotomy
<ol>
	<li>Add enough of the 630 &#956;M tricaine solution to a 15 x 60 mm petri dish to fill it about three-quarters of the way. Weigh the filled dish. This will be used to immobilize the animal so that further anesthesia can be injected intraperitoneally.</li>
	<li>Immerse the animal into the dish containing tricaine and weigh the dish again.
	<ol>
		<li>Subtract the difference between the weights from steps 1 and 2 to obtain the weight of the fish.</li>
		<li>Allow the animal to remain in the tricaine solution until calm and most movement has ceased.</li>
	</ol>
	</li>
	<li>Using a pair of broad forceps, transfer the fish to the premoistened sponge and position it laterally. Transferring the fish by the tail works well for this process.
	<ol>
		<li>Place the sponge and fish under the dissecting microscope.</li>
	</ol>
	</li>
	<li>With a Nanofil syringe containing a 34 G needle, measure out enough pancuronium bromide such that there is 1 &#956;g/g of fish weight.</li>
	<li>Using a popsicle stick along the dorsal side of the fish to hold it steady, administer the pancuronium bromide intraperitoneally using the Nanofil syringe.</li>
	<li>Using the fine forceps, grab the fish by the lower jaw and quickly transfer the animal to the intubation base.</li>
	<li>Position the animal so that it is dorsal-up on the wax form and such that the 1 mm cannula can be inserted into the mouth. Use the fine forceps to maneuver the fish and open the mouth around the cannula. Due to the makeup of the intubation tray, a stop was engineered into the base, allowing the cannula to be inserted 3 mm into the mouth of the fish.
	<ol>
		<li>Once in position, turn on the gravity-fed perfusion tube containing tricaine.</li>
	</ol>
	</li>
	<li>Cut a 3 cm2 section of Kimqipe tissue and wet it with habitat water.
	<ol>
		<li>Place the tissue over the animal to prevent desiccation. The Kimwipe section can also be positioned to aid in holding the animal dorsal-up.</li>
	</ol>
	</li>
	<li>Under the dissection microscope, use vanna spring scissors to remove ~2 mm2 section of the cranium covering the optic tectum. This area looks like a dark, bony plate that sits behind the eye.
	 <img alt="Figure 3" fo:content-width="5in" fo:src="/files/ftp_upload/51065/51065fig3highres.jpg" src="/files/ftp_upload/51065/51065fig3.jpg" width="400px" /> 
	Figure 3
	<ol>
		<li>Insert one blade of the vanna scissors at the edge of the plate and push with enough force to penetrate the bone, without piercing the brain. Close the scissors to snip the bone.</li>
		<li>Remove the piece of bone.</li>
		<li>If any blood is present, remove using the edge of a Kimwipe. Bleeding generally stops shortly after carrying out the craniotomy.</li>
	</ol>
	</li>
</ol>
4. Electrophysiology
<ol>
	<li>Shut off the intubation stop-cock and quickly disconnect the Luer&#160;lock connecting the intubation base to the tricaine drip.
	<ol>
		<li>Move the intact intubation setup to the electrophysiology microscope and connect to the perfusion system.</li>
		<li>Turn on the habitat water and perfuse at a rate of 1 ml/min for ~1 hr, in order to wash out the tricaine.</li>
	</ol>
	</li>
	<li>Using a micromanipulator under visual control, position the secondary electrode so that the electrode tip can be inserted into the animal&#39;s nostril or into the dip behind the upper jaw.</li>
	<li>Insert the primary electrode needle into the craniotomy opening. Insert the needle into the tissue, such that the tip is positioned fairly superficial within the optic tectum.
	<ol>
		<li>If the electrode is too deep, the electrical signal may be small.</li>
	</ol>
	</li>
	<li>Collect and analyze the electrical difference recorded between the primary and secondary reference electrodes. This process will allow for extracellular recordings to be obtained.
	<ol>
		<li>Collect the data in Gap-free mode at a 5&#160;kHz sample rate, with a low pass filter of 0.1 kHz and a high pass filter of 1 Hz.</li>
		<li>Do not begin recording electrophysiological activity until habitat water has perfused for a minimum of 45 min.</li>
		<li>Record a baseline of native activity for at least 15 min prior to the addition of experimental drugs. Begin perfusion with experimental drugs of choice and record for the desired amount of time. In healthy preparations, it is possible to record the neurological activity for 2-3 hr.</li>
	</ol>
	</li>
</ol>
5. Clean Up
<ol>
	<li>At the end of the experiment, remove the electrodes from the cranium and nostril and remove the intubation setup.
	<ol>
		<li>Euthanize the animal by drug overdose using tricaine in accordance with the accepted practices, as outlined by the American Veterinary Medical Association (AVMA) Guidelines for the Euthanasia of Animals (2013).</li>
		<li>Zebrafish to be euthanized should be placed in a tricaine solution at 200-500 mg/L. Fish are to be left in the solution for a minimum of 10 min after cessation of rhythmic opercular activity, after which they are subjected to a physical means of euthanasia (cervical transection) to ensure death.</li>
	</ol>
	</li>
	<li>Disconnect the Luer&#160;lock connecting the intubation base to the perfusion setup. Collect any remaining liquid in the perfusion system for appropriate disposal.
	<ol>
		<li>Run DI water through all tubes and collect for appropriate disposal, if hazardous.</li>
	</ol>
	</li>
	<li>To sanitize the perfusion system, run 70% ethanol through all tubes and collect for appropriate disposal.
	<ol>
		<li>Leave the stopcocks for each tube in the open position to facilitate air&#160;drying.</li>
		<li>Soak the electrodes in 70% ethanol and allow to air dry.</li>
	</ol>
	</li>
	<li>Dispose of the capillary needle used for the primary electrode in a sharps container.</li>
	<li>Dismantle the intubation setup and rinse all pieces with water.
	<ol>
		<li>Clean each piece with 70% ethanol and allow to air dry.</li>
	</ol>
	</li>
</ol>

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Neurosci. 27, 1606-1615 (2007).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The zebrafish book: a guide for the laboratory use of zebrafish (Brachydanio rerio). , (1993).</li><li>Lazarova, M., Samanin, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Potentiation+by+yohimbine+of+pentylenetetrazol-induced+seizures+in+rats:+role+of+alpha+2+adrenergic+receptors.">Potentiation by yohimbine of pentylenetetrazol-induced seizures in rats: role of alpha 2 adrenergic receptors.</a> Pharmacol. Res. Commun. 15, 419-425 (1983).</li><li>Loscher, W., Honack, D., Fassbender, C. P., Nolting, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+technical,+biological+and+pharmacological+factors+in+the+laboratory+evaluation+of+anticonvulsant+drugs.+III.+Pentylenetetrazole+seizure+models.">The role of technical, biological and pharmacological factors in the laboratory evaluation of anticonvulsant drugs. III. Pentylenetetrazole seizure models.</a> Epilepsy res. 8, 171-189 (1991).</li><li>DeMicco, A., Cooper, K. R., Richardson, J. R., White, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+neurotoxicity+of+pyrethroid+insecticides+in+zebrafish+embryos.">Developmental neurotoxicity of pyrethroid insecticides in zebrafish embryos.</a> Toxicol Sci. 113, 177-186 (2010).</li><li>Arnolds, D. E., 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=Physiological+effects+of+tricaine+on+the+supramedullary/dorsal+neurons+of+the+cunner,+Tautogolabrus+adspersus.">Physiological effects of tricaine on the supramedullary/dorsal neurons of the cunner, Tautogolabrus adspersus.</a> Biol. Bull. 203, 188-189 (2002).</li></ol>

The authors declare that they have no competing financial interests.

This protocol has been used to measure the neural activity of adult zebrafish in vivo. With practice, neural activity can be observed consistently, though the characteristics (amplitude and shape of events) of the recorded activity can vary between individual fish. Utilization of the extracellular recording technique can explain this observation. The method provides simultaneous monitoring of a large number of cells within a region1, so variations in the positioning of the primary electrode can play a...

In this article, a protocol is described for obtaining in vivo recordings of neural activity in adult zebrafish. Extracellular recording methods are used, providing voltage measurements of electrical activity within a small region of neural tissue. This method of investigation involves monitoring a large number of cells in a behaving animal1. Previously, slice recordings have been performed in both adults and larvae, as have eye cup preparations and electroretinogram recordings. These experiments have largely been performed to detail physiological responses of various sensory systems2-5. Until recently, intact brain preparations have only been available for performing electrophysiology with zebrafish larva3,6,7, where respiration and oxygen diffusion can occur through the skin. Our preparation allows the native neurological activity of an adult zebrafish to be measured while the animal remains fully conscious and aware of its surroundings.
Zebrafish (Danio rerio) currently play a fundamental role as a model for genetic, toxicological, pharmacological, and physiopathological studies3.&#160;Zebrafish have gained visibility within the field of neuroscience because they share extensive homology with mammals at the genetic, neural and endocrine levels8. Over the past decade, standard neuroanatomic and immunohistochemical techniques have been used to determine the detailed characteristic organization of the zebrafish nervous system9-12 and of the distribution of different neurotransmitters3,8,13. More recently, researchers have shifted their focus to functional studies14,15, many of which center on behavioral processes16-19 and electrophysiological characteristics of sensory systems2,13,20. A small number of these studies have concentrated on the electrical activity of specific areas of the adult zebrafish brain21-23, but were not carried out using an in vivo approach.
This protocol can be adapted for electrophysiological studies of both spontaneous and evoked activity within the zebrafish nervous system to describe the patterns of activity in specific brain regions. The use of this technique allows comparisons to be made between the neurological activity of young larval stages and adults. Further, our protocol permits comparisons between genetic or pharmacological alterations. Together with other approaches, such as genetic engineering or pharmacological tests, this method offers a new possibility for the functional analysis of neuronal communication and plasticity in the intact adult animal as well as for potential applications, such as studying late onset epilepsy or neurodegenerative processes.

<table border="0" cellpadding="0" cellspacing="0" width="539">
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	<tbody>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">70% Ethanol</td>
			<td style="width:91px;">Decon Laboratories</td>
			<td style="width:111px;">2750HC</td>
			<td style="width:132px;">Dilute 100% to 70% with DI water</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:207px;">2 M Potassium Chloride</td>
			<td style="width:91px;">J.T. Baker</td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:207px;">2 M Sodium Chloride</td>
			<td style="width:91px;">J.T. Baker</td>
			<td style="width:111px;">3624-05</td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="47">
			<td height="47" style="height:47px;width:207px;">0.4% Tris-Buffered Tricaine</td>
			<td style="width:91px;">Sigma-Aldrich</td>
			<td style="width:111px;">E10521</td>
			<td style="width:132px;">pH 7.2-7.4; stored at -20 oC</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:207px;">Pancuronium Bromide</td>
			<td style="width:91px;">Sigma-Aldrich</td>
			<td style="width:111px;">P1918</td>
			<td style="width:132px;">Diluted to 1 &#956;g/&#956;l in 1x phosphate buffered saline</td>
		</tr>
		<tr height="127">
			<td height="127" style="height:127px;width:207px;">Habitat water</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;">pH 7.0-7.4, conductivity of 400-450 &#956;S; maintained by Instant Ocean and Sodium Bicarbonate</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:207px;">Pentylenetetrazol</td>
			<td style="width:91px;">Sigma-Aldrich</td>
			<td style="width:111px;">P6500</td>
			<td style="width:132px;">Diluted to 300 mM in 1x phosphate buffered saline</td>
		</tr>
		<tr height="85">
			<td height="85" style="height:85px;width:207px;">Nanofil syringe</td>
			<td style="width:91px;">World Precision Instruments, Inc.</td>
			<td style="width:111px;">06A</td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="85">
			<td height="85" style="height:85px;width:207px;">34 G Beveled needle</td>
			<td style="width:91px;">World Precision Instruments, Inc.</td>
			<td style="width:111px;">NF34BV</td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">Sponge</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;">Small pore and chemical-free</td>
		</tr>
		<tr height="47">
			<td height="47" style="height:47px;width:207px;">Foam-backed fine sand paper</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;">5 x 5 cm2 is large enough</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:207px;">9 V Battery</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:207px;">Wires with alligator clips</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;">Need 2</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:207px;">37 cm x 42 cm Kimwipe</td>
			<td style="width:91px;">Kimberly-Clark Professional</td>
			<td style="width:111px;">TW31KEM</td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:207px;">11 cm x 21 cm Kimwipe</td>
			<td style="width:91px;">Kimberly-Clark Professional</td>
			<td style="width:111px;">TW31KWP</td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:207px;">1/8 in diameter tube</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:207px;">1 cm diameter tube</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:207px;">1 mm diameter tube</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">Reducing valve with female Luer lock cap and silicone ferrule</td>
			<td style="width:91px;">Qosina</td>
			<td style="width:111px;">51505</td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">Microscope (Leica MZ APO)</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;">Another microscope can be used</td>
		</tr>
		<tr height="85">
			<td height="85" style="height:85px;width:207px;">Vanna scissors</td>
			<td style="width:91px;">Roboz Surgical Instruments Co., Inc.</td>
			<td style="width:111px;">15018-10</td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="85">
			<td height="85" style="height:85px;width:207px;">60 ml Luer lock syringe tubes</td>
			<td style="width:91px;">Becton, Dickinson and Company</td>
			<td style="width:111px;">309653</td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">3-way Stopcocks with Luer connections</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">1-way Stopcock with Luer connection</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">Fisherbrand 100 mm x 15 mm Petri dish</td>
			<td style="width:91px;">Fisher Scientific</td>
			<td style="width:111px;">NC9299146</td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">Fisherbrand 60 mm x 15 mm Petri dish</td>
			<td style="width:91px;">Fisher Scientific</td>
			<td style="width:111px;">S67961</td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:207px;">4 in Borosilicate capillary tube</td>
			<td style="width:91px;">World Precision Instruments</td>
			<td style="width:111px;">TW100F-4</td>
			<td style="width:132px;">Can contain a filament to aid in filling with solution</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:207px;">P-97 Flaming/Brown Micropipette Puller</td>
			<td style="width:91px;">Sutter Instrument Co.</td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">Digidata 1440</td>
			<td style="width:91px;">Molecular Devices</td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">Axon Aloclamp 900A</td>
			<td style="width:91px;">Molecular Devices</td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">Axoclamp software</td>
			<td style="width:91px;">Molecular Devices</td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">HS-9Ax 1U headstage</td>
			<td style="width:91px;">Molecular Devices</td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">0.010 in Silver wire</td>
			<td style="width:91px;">A-M Systems, Inc.</td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">Q-series electrode holder</td>
			<td style="width:91px;">Warner Instruments</td>
			<td style="width:111px;">QSW-A10P</td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:207px;">10 ml Luer lock syringe</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:207px;">1 mm x 15 in Tubing</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;">Connect Luer lock syringe to Q-series electrode holder</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:207px;">Micromanipulator</td>
			<td style="width:91px;">Warner Instruments</td>
			<td style="width:111px;"></td>
			<td style="width:132px;">Need 2</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:207px;">Microsoft-based PC</td>
			<td style="width:91px;">Dell</td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:207px;">Faraday Cage</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:207px;">Air Table</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:207px;">Dissecting Microscope</td>
			<td style="width:91px;"></td>
			<td style="width:111px;"></td>
			<td style="width:132px;"></td>
		</tr>
	</tbody>
</table>

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 protocol has been used to measure the neural activity of adult zebrafish in vivo. These electrophysiological recordings are consistently and reproducibly obtained. Figure 5 shows a representative example of native and induced alterations of the neural activity of an adult zebrafish when pentylenetetrazol (PTZ), a common chemoconvulsant6,7,25,26, is introduced in the intubation setup.
The native neurological activity of the adult zebrafish was monitored...

Watch this Scientific Journal Video about Electrophysiological Recording in the Brain of Intact Adult Zebrafish at JoVE.com

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

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 ...

electrophysiological-recording-in-the-brain-of-intact-adult-zebrafish

Research

JoVE Journal

Neuroscience

21.3K Views.  University of Georgia. 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.

Video: Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Electrophysiological Characterization of GFP-Expressing Cell Populations in the Intact Retina

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack conspicuous morphological features or show a low population density. This applies particularly to retinal amacrine cells, an exceptionally multiform class of interneurons that comprise roughly 30 subtypes in mammals1. Though being a crucial part of the visual processing by shaping the retinal output2, most of these subtypes have not been studied up to now in a functional context because encountering these cells with a recording electrode is a rare event.
Recently, a multitude of transgenic mouse lines is available that express fluorescent markers like green fluorescent protein (GFP) under the control of promoters for membrane receptors or enzymes that are specific to only a subset of neurons in a given tissue3,4. These pre-labeled cells are therefore accessible to directed microelectrode targeting under microscopic control, permitting the systematic study of their physiological properties in situ. However, excitation of fluorescent markers is accompanied by the risk of phototoxicity for the living tissue. In the retina, this approach is additionally hampered by the problem that excitation light causes appropriate stimulation of the photoreceptors, thus inflicting photopigment bleaching and transferring the retinal circuits into a light-adapted condition. These drawbacks are overcome by using infrared excitation delivered by a mode-locked laser in short pulses of the femtosecond range. Two-photon excitation provides energy sufficient for fluorophore excitation and at the same time restricts the excitation to a small tissue volume minimizing the hazards of photodamage5. Also, it leaves the retina responsive to visual stimuli since infrared light (&gt;850 nm) is only poorly absorbed by photopigments6.
In this article we demonstrate the use of a transgenic mouse retina to attain electrophysiological in situ recordings from GFP-expressing cells that are visually targeted by two-photon excitation. The retina is prepared and maintained in darkness and can be subjected to optical stimuli which are projected through the condenser of the microscope (Figure 1). Patch-clamp recording of light responses can be combined with dye filling to reveal the morphology and to check for gap junction-mediated dye coupling to neighboring cells, so that the target cell can by studied on different experimental levels.

This article depicts the recording of individual cells from fluorescently tagged neuronal populations in the intact mouse retina. By using two-photon infrared excitation transgenetically labeled cells were targeted for patch-clamp recording to study their light responses, receptive field properties, and morphology.

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack ...

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 ...

Recording Electrical Activity from Identified Neurons in the Intact Brain of Transgenic Fish

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be performed in an intact brain preparation where the neural circuitry of the CNS remains intact. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with green fluorescent protein for identification in the intact brain. Fish have multiple populations of GnRH neurons, and their functions are dependent on their location in the brain and the GnRH gene that they express1 . We have focused our demonstration on GnRH3 neurons located in the terminal nerves (TN) associated with the olfactory bulbs using the intact brain of transgenic medaka fish (Figure 1B and C). Studies suggest that medaka TN-GnRH3 neurons are neuromodulatory, acting as a transmitter of information from the external environment to the central nervous system; they do not play a direct role in regulating pituitary-gonadal functions, as do the well-known hypothalamic GnRH1 neurons2, 3 .The tonic pattern of spontaneous action potential firing of TN-GnRH3 neurons is an intrinsic property4-6, the frequency of which is modulated by visual cues from conspecifics2 and the neuropeptide kisspeptin 15. In this video, we use a stable line of transgenic medaka in which TN-GnRH3 neurons express a transgene containing the promoter region of Gnrh3 linked to enhanced green fluorescent protein7 to show you how to identify neurons and monitor their electrical activity in the whole brain preparation6.

In this video, we will demonstrate how to record electrical activity from identified single neurons in a whole brain preparation, which preserves complex neural circuits. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with a fluorescent protein for identification in the intact brain preparation.

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be ...

Micromanipulation of Gene Expression in the Adult Zebrafish Brain Using Cerebroventricular Microinjection of Morpholino Oligonucleotides

Manipulation of gene expression in tissues is required to perform functional studies. In this paper, we demonstrate the cerebroventricular microinjection (CVMI) technique as a means to modulate gene expression in the adult zebrafish brain. By using CVMI, substances can be administered into the cerebroventricular fluid and be thoroughly distributed along the rostrocaudal axis of the brain. We particularly focus on the use of antisense morpholino oligonucleotides, which are potent tools for knocking down gene expression in vivo. In our method, when applied, morpholino molecules are taken up by the cells lining the ventricular surface. These cells include the radial glial cells, which act as neurogenic progenitors. Therefore, knocking down gene expression in the radial glial cells is of utmost importance to analyze the widespread neurogenesis response in zebrafish, and also would provide insight into how vertebrates could sustain adult neurogenesis response. Such an understanding would also help the efforts for clinical applications in human neurodegenerative disorders and central nervous system regeneration. Thus, we present the cerebroventricular microinjection method as a quick and efficient way to alter gene expression and neurogenesis response in the adult zebrafish forebrain. We also provide troubleshooting tips and other useful information on how to carry out the CVMI procedure.

In this article, we demonstrate a method for manipulation of gene expression in the ventricular cells of the adult zebrafish telencephalon using antisense morpholino oligonucleotides. We present this method as an efficient and quick protocol that can be used for functional studies in the adult vertebrate brain.

Manipulation of gene expression in tissues is  required to perform functional studies. In this paper, we demonstrate the  cerebroventricular ...

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...

Modeling Amyloid-&#946;42 Toxicity and Neurodegeneration in Adult Zebrafish Brain

Alzheimer's disease (AD) is a debilitating neurodegenerative disease in which accumulation of toxic amyloid-&#946;42 (A&#946;42) peptides leads to synaptic degeneration, inflammation, neuronal death, and learning deficits. Humans cannot regenerate lost neurons in the case of AD in part due to impaired proliferative capacity of the neural stem/progenitor cells (NSPCs) and reduced neurogenesis. Therefore, efficient regenerative therapies should also enhance the proliferation and neurogenic capacity of NSPCs. Zebrafish (Danio rerio) is a regenerative organism, and we can learn the basic molecular programs with which we could design therapeutic approaches to tackle AD. For this reason, the generation of an AD-like model in zebrafish was necessary. Using our methodology, we can introduce synthetic derivatives of A&#946;42 peptide with tissue penetrating capability into the adult zebrafish brain, and analyze the disease pathology and the regenerative response. The advantage over the existing methods or animal models is that zebrafish can teach us how a vertebrate brain can naturally regenerate, and thus help us to treat human neurodegenerative diseases better by targeting endogenous NSPCs. Therefore, the amyloid-toxicity model established in the adult zebrafish brain may open new avenues for research in the field of neuroscience and clinical medicine. Additionally, the simple execution of this method allows for cost-effective and efficient experimental assessment. This manuscript describes the synthesis and injection of A&#946;42 peptides into zebrafish brain.

Modeling Amyloid-&amp;#946;42 Toxicity and Neurodegeneration in Adult Zebrafish Brain

This protocol describes the synthesis, characterization, and injection of monomeric amyloid-&#946;42 peptides for generating amyloid toxicity in adult zebrafish to establish an Alzheimer&#39;s disease model, followed by histological analyses and detection of aggregations.

Alzheimer's disease (AD) is a debilitating neurodegenerative disease in which accumulation of toxic amyloid-&#946;42 (A&#946;42) peptides leads to ...

Wireless Electrophysiological Recording of Neurons by Movable Tetrodes in Freely Swimming Fish

The neural mechanisms governing fish behavior remain mostly unknown, although fish constitute the majority of all vertebrates. The ability to record brain activity from freely moving fish would advance research on the neural basis of fish behavior considerably. Moreover, precise control of the recording location in the brain is critical to studying coordinated neural activity across regions in fish brain. Here, we present a technique that records wirelessly from the brain of freely swimming fish while controlling for the depth of the recording location. The system is based on a neural logger associated with a novel water-compatible implant that can adjust the recording location by microdrive-controlled tetrodes. The capabilities of the system are illustrated through recordings from the telencephalon of goldfish.

A novel wireless technique for recording extracellular neural signals from the brain of freely swimming goldfish is presented. The recording device is composed of two tetrodes, a microdrive, a neural data logger, and a waterproof case. All parts are custom-made except for the data logger and its connector.

The neural mechanisms governing fish behavior remain mostly unknown, although fish constitute the majority of all vertebrates. The ability to record brain ...

A Scalable Model to Study the Effects of Blunt-Force Injury in Adult Zebrafish

Blunt-force traumatic brain injuries (TBI) are the most common form of head trauma, which spans a range of severities and results in complex and heterogenous secondary effects. While there is no mechanism to replace or regenerate the lost neurons following a TBI in humans, zebrafish possess the ability to regenerate neurons throughout their body, including the brain. To examine the breadth of pathologies exhibited in zebrafish following a blunt-force TBI and to study the mechanisms underlying the subsequent neuronal regenerative response, we modified the commonly used rodent Marmarou weight drop for the use in adult zebrafish. Our simple blunt-force TBI model is scalable, inducing a mild, moderate, or severe TBI, and recapitulates many of the phenotypes observed following human TBI, such as contact- and post-traumatic seizures, edema, subdural and intracerebral hematomas, and cognitive impairments, each displayed in an injury severity-dependent manner. TBI sequelae, which begin to appear within minutes of the injury, subside and return to near undamaged control levels within 7 days post-injury. The regenerative process begins as early as 48 hours post-injury (hpi), with the peak cell proliferation observed by 60 hpi. Thus, our zebrafish blunt-force TBI model produces characteristic primary and secondary injury TBI pathologies similar to human TBI, which allows for investigating disease onset and progression, along with the mechanisms of neuronal regeneration that is unique to zebrafish.

We modified the Marmarou weight drop model for adult zebrafish to examine a breadth of pathologies following blunt-force traumatic brain injury (TBI) and the mechanisms underlying subsequent neuronal regeneration. This blunt-force TBI model is scalable, induces a mild, moderate, or severe TBI, and recapitulates injury heterogeneity observed in human TBI.

Blunt-force traumatic brain injuries (TBI) are the most common form of head trauma, which spans a range of severities and results in complex and ...

Deep-Tissue Three-Photon Fluorescence Microscopy in Intact Mouse and Zebrafish Brain

Multiphoton microscopy techniques, such as two-photon microscopy (2PM) and three-photon microscopy (3PM), are powerful tools for deep-tissue in vivo imaging with subcellular resolution. 3PM has two major advantages for deep-tissue imaging over 2PM that has been widely used in biology laboratories: (i) longer attenuation length in scattering tissues by employing ~1,300 nm or ~1,700 nm excitation laser; (ii) less background fluorescence generation due to higher-order nonlinear excitation. As a result, 3PM allows high-contrast structural and functional imaging deep within scattering tissues such as intact mouse brain from the cortical layers to the hippocampus and the entire forebrain of adult zebrafish.
Today, laser sources suitable for 3PM are commercially available, enabling the conversion of an existing two-photon (2P) imaging system to a three-photon (3P) system. Additionally, multiple commercial 3P microscopes are available, which makes this technique readily available to biology research laboratories. This paper shows the optimization of a typical 3PM setup, particularly targeting biology groups that already have a 2P setup, and demonstrates intravital 3D imaging in intact mouse and adult zebrafish brains. This protocol covers the full experimental procedure of 3P imaging, including microscope alignment, prechirping of ~1,300 and ~1,700 nm laser pulses, animal preparation, and intravital 3P fluorescence imaging deep in adult zebrafish and mouse brains.

Three-photon microscopy enables high-contrast fluorescence imaging deep in living biological tissues, such as mouse and zebrafish brains, with high spatiotemporal resolution.

Multiphoton microscopy techniques, such as two-photon microscopy (2PM) and three-photon microscopy (3PM), are powerful tools for deep-tissue in vivo ...