Name: Electrophysiological Recording in the Brain of Intact Adult Zebrafish (Video) | JoVE Journal
Uploaded: 2013-11-19T17:00:00.000+00:00
Duration: 9 min 42 s
Description: Learn electrophysiological recording in intact adult zebrafish using immobilization, intubation, and craniotomy techniques. This method enables in vivo neural activity studies for neurobiology research.

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5 minutes

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.  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 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 ⅛ 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'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 lock using a short portion of ⅛ in diameter tubing. <img alt="Hypodermic needle anatomy diagram with measurements alongside a ruler for size reference." fo:content-width="5in" fo:src="/files/ftp_upload/51065/51065fig1highres.jpg" src="/files/ftp_upload/51065/51065fig1.jpg" data-altupdatedat="1787630408000" data-alt="Figure 1"> 
		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° 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="Static equilibrium diagram; setup with pipes A, B; ice formation experiment; ruler for scale." fo:content-width="5in" fo:src="/files/ftp_upload/51065/51065fig2highres.jpg" src="/files/ftp_upload/51065/51065fig2.jpg" width="400px" data-altupdatedat="1787630408000" data-alt="Figure 2"> 
			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 μM) 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 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'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 of 0.010 in silver wire and a secondary electrode which should be made up of 15 in of the same wire to ground the setup. For the secondary electrode, use a 15 in section of 0.010 in 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 &lt;15 mΩ.
			<ol>
				<li>Fill the capillary needle with 2-3 μl 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 μM)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 μg/μl pancuronium bromide in Ringer'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 μM) 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 μM 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 μg/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="Fish cranial nerve stimulation, black and white close-up, scientific study of neural responses." fo:content-width="5in" fo:src="/files/ftp_upload/51065/51065fig3highres.jpg" src="/files/ftp_upload/51065/51065fig3.jpg" width="400px" data-altupdatedat="1787630408000" data-alt="Figure 3"> 
	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 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'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 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 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 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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<li>Lazarova, M., Samanin, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Potentiation+by+yohimbine+of+pentylenetetrazol-induced+seizures+in+rats%3A+role+of+alpha+2+adrenergic+receptors%5BTitle%5D)+AND+%22Pharmacol.+Res.+Commun%22%5BJournal%5D)">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/pubmed?term=((The+role+of+technical%2C+biological+and+pharmacological+factors+in+the+laboratory+evaluation+of+anticonvulsant+drugs.+III.+Pentylenetetrazole+seizure+models%5BTitle%5D)+AND+%22Epilepsy+res%22%5BJournal%5D)">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/pubmed?term=((Developmental+neurotoxicity+of+pyrethroid+insecticides+in+zebrafish+embryos%5BTitle%5D)+AND+%22Toxicol+Sci%22%5BJournal%5D)">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/pubmed?term=((Physiological+effects+of+tricaine+on+the+supramedullary%2Fdorsal+neurons+of+the+cunner%2C+Tautogolabrus+adspersus%5BTitle%5D)+AND+%22Biol.+Bull%22%5BJournal%5D)">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><tbody><tr><td>70% Ethanol</td><td>Decon Laboratories</td><td>2750HC</td><td>Dilute 100% to 70% with DI water</td></tr><tr><td>2 M Potassium Chloride</td><td>J.T. Baker</td><td></td><td></td></tr><tr><td>2 M Sodium Chloride</td><td>J.T. Baker</td><td>3624-05</td><td></td></tr><tr><td>0.4% Tris-Buffered Tricaine</td><td>Sigma-Aldrich</td><td>E10521</td><td>pH 7.2-7.4; stored at -20 &lt;sup&gt;o&lt;/sup&gt;C</td></tr><tr><td>Pancuronium Bromide</td><td>Sigma-Aldrich</td><td>P1918</td><td>Diluted to 1 &amp;mu;g/&amp;mu;l in 1x phosphate buffered saline</td></tr><tr><td>Habitat water</td><td></td><td></td><td>pH 7.0-7.4, conductivity of 400-450 &amp;mu;S; maintained by Instant Ocean and Sodium Bicarbonate</td></tr><tr><td>Pentylenetetrazol</td><td>Sigma-Aldrich</td><td>P6500</td><td>Diluted to 300 mM in 1x phosphate buffered saline</td></tr><tr><td>Nanofil syringe</td><td>World Precision Instruments, Inc.</td><td>06A</td><td></td></tr><tr><td>34 G Beveled needle</td><td>World Precision Instruments, Inc.</td><td>NF34BV</td><td></td></tr><tr><td>Sponge</td><td></td><td></td><td>Small pore and chemical-free</td></tr><tr><td>Foam-backed fine sand paper</td><td></td><td></td><td>5 x 5 cm&lt;sup&gt;2&lt;/sup&gt; is large enough</td></tr><tr><td>9 V Battery</td><td></td><td></td><td></td></tr><tr><td>Wires with alligator clips</td><td></td><td></td><td>Need 2</td></tr><tr><td>37 cm x 42 cm Kimwipe</td><td>Kimberly-Clark Professional</td><td>TW31KEM</td><td></td></tr><tr><td>11 cm x 21 cm Kimwipe</td><td>Kimberly-Clark Professional</td><td>TW31KWP</td><td></td></tr><tr><td>1/8 in diameter tube</td><td></td><td></td><td></td></tr><tr><td>1 cm diameter tube</td><td></td><td></td><td></td></tr><tr><td>1 mm diameter tube</td><td></td><td></td><td></td></tr><tr><td>Reducing valve with female Luer lock cap and silicone ferrule</td><td>Qosina</td><td>51505</td><td></td></tr><tr><td>Microscope (Leica MZ APO)</td><td></td><td></td><td>Another microscope can be used</td></tr><tr><td>Vanna scissors</td><td>Roboz Surgical Instruments Co., Inc.</td><td>15018-10</td><td></td></tr><tr><td>60 ml Luer lock syringe tubes</td><td>Becton, Dickinson and Company</td><td>309653</td><td></td></tr><tr><td>3-way Stopcocks with Luer connections</td><td></td><td></td><td></td></tr><tr><td>1-way Stopcock with Luer connection</td><td></td><td></td><td></td></tr><tr><td>Fisherbrand 100 mm x 15 mm Petri dish</td><td>Fisher Scientific</td><td>NC9299146</td><td></td></tr><tr><td>Fisherbrand 60 mm x 15 mm Petri dish</td><td>Fisher Scientific</td><td>S67961</td><td></td></tr><tr><td>4 in Borosilicate capillary tube</td><td>World Precision Instruments</td><td>TW100F-4</td><td>Can contain a filament to aid in filling with solution</td></tr><tr><td>P-97 Flaming/Brown Micropipette Puller</td><td>Sutter Instrument Co.</td><td></td><td></td></tr><tr><td>Digidata 1440</td><td>Molecular Devices</td><td></td><td></td></tr><tr><td>Axon Aloclamp 900A</td><td>Molecular Devices</td><td></td><td></td></tr><tr><td>Axoclamp software</td><td>Molecular Devices</td><td></td><td></td></tr><tr><td>HS-9Ax 1U headstage</td><td>Molecular Devices</td><td></td><td></td></tr><tr><td>0.010 in Silver wire</td><td>A-M Systems, Inc.</td><td></td><td></td></tr><tr><td>Q-series electrode holder</td><td>Warner Instruments</td><td>QSW-A10P</td><td></td></tr><tr><td>10 ml Luer lock syringe</td><td></td><td></td><td></td></tr><tr><td>1 mm x 15 in Tubing</td><td></td><td></td><td>Connect Luer lock syringe to Q-series electrode holder</td></tr><tr><td>Micromanipulator</td><td>Warner Instruments</td><td></td><td>Need 2</td></tr><tr><td>Microsoft-based PC</td><td>Dell</td><td></td><td></td></tr><tr><td>Faraday Cage</td><td></td><td></td><td></td></tr><tr><td>Air Table</td><td></td><td></td><td></td></tr><tr><td>Dissecting Microscope</td><td></td><td></td><td></td></tr></tbody></table>

electrophysiological recording in the brain of intact adult zebrafish

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...

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 ...

The overall goal of this procedure is to obtain in vivo recordings of neural activity in the brain of intact adult zebrafish. This is accomplished by first obtaining an immobilizing, an adult zebra fish through the bath application of Trica Methyls sulfonate. The second step is to intubate the immobilized zebra fish.Next, a craniotomy is performed under trica anesthesia to provide access to the brain. The final step is to position a primary electrode within the craniotomy window, which allows extracellular brain activity to be recorded. Ultimately in vivo.Electrophysiology in the adult zebrafish can be used to show how different pharmacological compounds or genetic mutations can lead to alterations in the native neurological activity of the zebrafish. This method can help answer key questions in both developmental biology and neurobiology about how changes in brain structure can affect changes in neural activity, both in normal development and in the case of genetic mutation. Generally, individuals new to this technique will struggle because extra care must be taken to remove only a small portion of the bony plate covering the head without damaging the brain tissue.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's mouth. For this setup, remove a 1.5 centimeter section from the wide end of a P 200 pipette tip with a pair of scissors or a razor blade.Next, insert a six centimeter by one millimeter piece of tubing into a reducing valve. Then insert the modified pipette tip into a short portion of one eighth of an inch diameter tubing. The cannula can then be inserted into the pipette tip using the tubing to hold the pipette tip in position with the lure lock.Before the experiment flushed a profusion setup with habitat water, ensuring that all the air bubbles are removed. Next, add 30 milliliters of 630 micromolar trica solution to the craniotomy perfusion setup, and allow two milliliters of it to run through the tubing to ensure the delivery of trica to the animal upon profusion initiation. For the main profusion setup, add 50 milliliters of habitat water to the first tube.Then add any desired experimental compounds to each of the remaining tubes. After that, place a cannula setup into the small hole of the intubation base twist a one inch wide strip of Kim wipe into a tight spiral and insert it into the drainage tube such that the Kim wipe 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 that could interfere with electrophysiological recording.Next, 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 profusion waste. This dish will serve as the outflow reservoir for the setup. Then position the tissue so that one end will lie near the animal's abdominal region, and the lower extension can drip into the dish.Place this completed intubation base near the dissection microscope. Subsequently connect the lu lock of the cannula to the trican profusion tubing. In this procedure, add 630 micromolar trica solution to a 15 by 60 millimeter Petri dish up to three quarters.Next, weigh the fields dish. This will be used to immobilize the animal so that further anesthesia can be injected into peritoneal. Now immerse the animal into the dish containing trica and weigh the dish again.Then calculate the weight of the fish by subtracting the weight of the filled dish from the total weight. Allow the animal to remain in the trica solution until it is calm and most movement has ceased. While the fish is immersed in the trica solution, use a NFI syringe containing a 34 gauge needle to withdraw enough pantonium bromide such that there is one microgram per gram of fish weight.Once the fish is calmed, use a pair of broad forceps to transfer the fish by the tail to a pre-moistened sponge and position it laterally. Next place a sponge and the fish under the dissecting microscope. Hold the dorsal side of the fish with a popsicle stick.Then administer the pan keon bromide interperitoneal. Now use fine forceps. Grab the fish by the lower jaw and quickly transfer the animal to the intubation base.Position the animal dorsal side up on the wax form such that the one millimeter cannula can be inserted into the mouth. Next, use the fine forceps to maneuver the fish and open the mouth around the cannula. In our setup.A stop was engineered into the base, allowing the cannula to be inserted three millimeters into the mouth of the fish. Once it is in position, turn on the gravity fed perfusion tube containing the trica. Then cut a three square centimeter section of Kim wipe tissue and wed it with habitat water.Place the tissue over the animal to prevent desiccation. Under the dissection microscope, the fish is prepared and waits to have the craniotomy carried out. This area looks like a dark bony plate that sits behind the eye.Next, insert one blade of the van of scissors at the edge of the plate and push it with enough force to penetrate the bone without piercing the brain. Then close the scissors to snip the bone. After that, remove the piece of bone for electrophysiology recording.Shut off the intubation. Stop cock and quickly disconnect the lure lock connecting the intubation base to the trica drip. Move the intact intubation setup to the electrophysiology microscope and connect it to the perfusion system.Turn on the habitat, water and perfuse at a rate of one milliliter per minute for about one hour. In order to wash out the trica using a micro manipulator, position the secondary electrode so that the electrode tip can be inserted into the animal's nostril or into the dip under the upper jaw. Subsequently, insert the primary electrode needle into the craniotomy opening.Then insert the needle into the tissue such that the tip is positioned fairly superficial within the optic tum. Deeper positions of the electrode may result in a small electrical signal for data collection gap free mode at 5, 000 hertz sampling rate, together with a low pass filter of 0.1 kilohertz and a high pass filter of one hertz are used. Begin the recording of the electrophysiological activity when the habitat water has perfused for a minimum of 45 minutes.Record a baseline of native activity for at least 15 minutes prior to the addition of experimental drugs. The native neurological activity of the adult zebrafish was monitored in this experiment, this figure shows the spontaneous baseline activity of the optic tum with a small amplitude around two to 10 millivolts throughout the duration of the recording. Following the introduction of 15 millimolar PTZ to the system, spontaneous epileptic form like discharges began to develop within five minutes.Continuous exposure to PTZ led to a consistent pattern of up to 15 millivolt, large amplitude discharges, followed by smaller, more frequent bursts of activity. While carrying out this procedure, it is important to remember to be patient and to take your time while handling the animal and carrying out the craniotomy. After watching this video, you should have a good idea of how to obtain in vivo recordings of neural activity in the brain of an intact adult zebra fish.This is accomplished by using a profusion system and performing a craniotomy on the immobilized animal.

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

Research

JoVE Journal

Neuroscience

This video shows how to perform electrocardiography (ECG or EKG) measurements, a test which measures the electrical activity of the heart, on a zebrafish.

Zebrafish Electrocardiography (ECG): A Minimally Invasive Assay to Evaluate Cardiac Function

The protocol describes the procedure to perform electrophysiology in adult zebrafish. This method can be used for measuring neural activity in vivo.

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

This video demonstrates the recording of electrical signals from the afferent neurons in a paralyzed zebrafish larva by positioning a micropipette near the lateral line ganglion and forming a loose seal for signal capture. The extracellular solution maintains neuronal viability during this process.

Monitoring Electrical Signals from the Afferent Neurons in an Immobilized Zebrafish Larva

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.

Forebrain Electrophysiological Recording in Larval Zebrafish

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.

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

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

Here, we present a reliable, minimally invasive, and cost-effective method to record and interpret electrocardiograms in live anesthetized adult zebrafish.

In Vivo Surface Electrocardiography for Adult Zebrafish

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

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

The present protocol describes the acute isolation of viable cardiac and vascular smooth muscle cells from adult, juvenile, larval, and embryonic zebrafish (Danio rerio), suitable for electrophysiological studies.

Isolation of Cardiac and Vascular Smooth Muscle Cells from Adult, Juvenile, Larval and Embryonic Zebrafish for Electrophysiological Studies

Here, we present a protocol to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish.

Two-Photon Calcium Imaging of Forebrain Activity in Behaving Adult Zebrafish

The study introduces Zebra II, an advanced system for prolonged ECG acquisition and analysis in zebrafish. This system features an independent perfusion system and multiple-point electrodes for handling multiple fish in controlled environments. Acute effects of Amiodarone were assessed and analyzed with wild-type zebrafish.

Zebra II as A Novel System to Record Electrophysiological Signals in Zebrafish

Learn electrophysiological recording in intact adult zebrafish using immobilization, intubation, and craniotomy techniques. This method enables in vivo neural activity studies for neurobiology research.

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

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