We are grateful to Nachum Ulanovsky and the members of the Ulanovsky lab for all their help. In addition, we are grateful to Tal Novoplansky-Tzur for helpful technical assistance. We gratefully acknowledge financial support from THE ISRAEL SCIENCE FOUNDATION - FIRST Program (grant no. 281/15), and the Helmsley Charitable Trust through the Agricultural, Biological and Cognitive Robotics Initiative of Ben-Gurion University of the Negev.

All surgery procedures must be approved by the local ethics committees on animal welfare (e.g., IACUC).
1. Construction of the Microdrive Housing
<ol>
	<li>To construct the housing, cut a 1 mm wide brass plate into a 19 mm x 29 mm x 1 mm plate using a saw. Cut two 5.5 mm slits on each of the long sides perpendicular to the edge, such that each slit is 6.5 mm away from the narrow sides (Figure 2A).</li>
	<li>Using pliers, fold the area between the slits on the long sides inward, then fold the bottom part inward and the upper side outward to obtain the housing (Figure 2B,C).</li>
	<li>Using a 3 mm drill bit, make holes for screws in the microdrive housing. 
	NOTE: These holes will be used later to attach the housing to the logger box (Figure 2D).</li>
	<li>Solder the sides of the housing.</li>
	<li>Using a fine circular file, generate a small, 1.5 mm in radius, semicircular slit at the bottom of the housing (Figure 2E). 
	NOTE: This will be used later to insert the stainless steel tube to guide the electrodes.</li>
	<li>Use a 1 mm drill bit to make a hole in the back of the housing for the tetrodes (Figure 2F). 
	NOTE: A 3D model of the housing is found in the Supplementary housing.stl file.</li>
</ol>
2. Construction of the Microdrive
<ol>
	<li>Using a cutter, break off a three-pin piece from a single row male pin header strip (Figure 1H). Using pliers, pull out the middle pin.</li>
	<li>Using a cutter, cut the remaining pins to 10 mm in length (2 mm less than the screw length). Another possibility is to use a longer screw (see step 2.4).</li>
	<li>Drill a hole using a #65 drill bit through the middle pin hole. Drill a thread using a 00&#8722;99 tap.</li>
	<li>Assemble the microdrive and the brass plates (7.5 mm x 2.5 mm x 0.6 mm, see Supplementary Figure 4) such that the brass plates are touching the pins. Insert a screw (#00-90 round head, 12 mm, brass) through the first brass plate, then through the pin header thread and the second brass plate. Finally, place a nut on the screw and gently tighten the assembled microdrive.</li>
	<li>Solder the pins together with the brass plates, and the nut with the tip of the screw.</li>
	<li>Solder the microdrive into the microdrive housing at four points on the sides of the microdrive brass plates.</li>
	<li>Cut one stainless steel tube 6 mm long with an inner diameter of 1.5 mm and another stainless steel tube 3 mm long with an inner diameter of 1.2 mm. Polish the ends of the tubes to avoid sharp ends.</li>
	<li>Glue the 6 mm long tube to the small semicircular slit at the bottom of the microdrive housing using epoxy. Glue the 3 mm long stainless steel to the pin header, lined up with the 6 mm long tube on the housing.</li>
	<li>Cut two 5 cm long silicone tube segments with diameter of 0.64 mm and one 5 cm long polyimide tube with diameter of 0.250 mm.</li>
	<li>Insert the three tubes into the two stainless steel tubes. Glue the tubes to the stainless steel tube attached to the pin header using cyanoacrylate glue. Screw the microdrive all the way up and cut off the excess tubing from the top and bottom of the two steel tubes. 
	NOTE: The microdrive with the housing is now ready for use (Figure 1C).</li>
</ol>
3. Preparing the Tetrode Array
<ol>
	<li>To fabricate a two-tetrode implant with four electrodes on each tetrode, prepare eight wires, each 12 cm long, Formvar insulated, out of 25 &#181;m diameter tungsten wire. 
	NOTE: The same design can accommodate four tetrodes.</li>
	<li>Place a holder for a 16-channel electrode interface board (EIB-16) PCB (see Table of Materials) under the microscope.</li>
	<li>Using a soft tipped tweezer and a lighter, remove the coating off each of the eight wires on one side using the flame. 
	NOTE: This is to ensure that the wire will be properly connected to the PCB connector later on.</li>
	<li>Push a wire into one of the holes in the EIB-16 with the coated side in the hole. Place a pin and press it using pliers. Check the connectivity by measuring the resistance between the pin and the uncoated side of the wire. 
	NOTE: The resistance is on the order of tens of Ohms.</li>
	<li>Repeat step 3.4 with all eight wires.</li>
	<li>Tape two groups of four wires together using duct tape at the end of each wire. 
	NOTE: Each group will be glued together later to form a tetrode.</li>
	<li>Cut one piece of tungsten wire 12 cm long with a 50 &#181;m diameter. Connect it to one of the EIB-16 connections. 
	NOTE: This wire will serve as the reference electrode.</li>
	<li>Cut two bare silver wires 12 cm long with a 75 &#181;m diameter that will serve as grounds for the recording logger. Solder the two wires to the ground connection in the EIB-16.</li>
	<li>Hold the EIB-16 above a motorized turning device and place the duct tape end of one group of four wires on the motorized tuning device. Apply 130 rounds clockwise followed by 20 counter- clockwise rotations. Apply cyanoacrylate glue to cover the tetrode.</li>
	<li>Wait for the glue to cure. Cut the tetrode close to the duct tape.</li>
	<li>Repeat steps 3.9 and 3.10 with the second tetrode. 
	NOTE: This produces the finished two-tetrode array (Figure 1D).</li>
</ol>
4. Assembling the Implant
<ol>
	<li>Screw the microdrive all the way down.</li>
	<li>Using 1 x 3M Phillips round head screws, attach the EIB-16 to the PVC plate.</li>
	<li>Using soft end tweezers, pull all the tetrodes and wires through the hole in the front of the logger box cover.</li>
	<li>Using the 2 x 6M Phillips flat head screws, attach the PVC plate to the logger box cover. Keep the EIB-16 connector in the correct orientation so that the logger can be mounted on the EIB-16. Make sure the EIB-16 is fixed in place to avoid motion artifacts in the recorded signal.</li>
	<li>Seal the wires to the box using epoxy. Apply as little as possible because the primary sealing will be done by room temperature vulcanizing (RTV) later on.</li>
	<li>Attach the microdrive housing to the logger box cover using 2 mm screws.</li>
	<li>Thread the tetrodes and all wires through the hole at the back of the microdrive housing. Thread the tetrodes through the two silicone tubes in the microdrive. Thread the 50 &#181;m tungsten wire through the polyimide tube in the microdrive.</li>
	<li>Glue the tetrodes and wires to their tubes by applying cyanoacrylate glue to the top end of the tubes, to ensure the movement is consistent with the microdrive. Screw the microdrive all the way to the top.</li>
	<li>Apply soft petroleum (see Table of Materials) on the exposed tetrode and wires inside the microdrive housing to prevent motion.</li>
	<li>Cut a 12 mm x 14.5 mm Petri dish bottom window using a heated razor blade. Attach the window to the front of the microdrive housing with epoxy. Keep the ground wires outside the window.</li>
	<li>Apply RTV coating to the exposed tetrodes and wires between the logger box cover and the microdrive housing.</li>
	<li>After the RTV is cured, close the box with a small weight inside, and submerge in water overnight to ensure there is no water leakage into the box.</li>
	<li>Cut the tetrodes and reference wire to the desired length using sharp scissors.</li>
	<li>Attach the marked extruded polystyrene foam (see Table of Materials) to the box. Adjust its size so its buoyancy is balanced when submerged in a water bath.</li>
	<li>Dip the tetrode tips in platinum black solution and use a direct current (-0.2 &#181;A) to coat the electrodes and set the electrodes&#39; impedance as desired. Use a multielectrode impedance tester (see Table of Materials) for coating and impedance measurements. 
	NOTE: In the goldfish pallium, a value of 40 kOhm is best. Depending on the application, the electrode impedance can be adjusted by modifying the platinum black coating10,11.</li>
</ol>
5. Anesthesia Preparation &#8212; 1% MS-222 Stock Solution
CAUTION: Anesthesia preparation includes the use of powdered MS-222, a carcinogen. Hence, steps 5.2 and 5.3 must be done in a chemical hood using gloves.
<ol>
	<li>Add 100 mL of water to a tube that can contain more than 100 mL.</li>
	<li>In a chemical hood, place a disposable weighting plate on a scale. Add 1 g of MS-222 powder using a spatula, then add the powder to the tube.</li>
	<li>Shake the tube well. 
	NOTE: In liquid form, MS-222 can be used outside the chemical hood wearing gloves but does not require a mask.</li>
	<li>Place a disposable weighting plate on a scale. Add 2 g of sodium bicarbonate using a spatula, then add the powder to the tube. Shake the tube well.</li>
</ol>
6. Preparing the Fish Skull
NOTE: At this stage, the fish is ready for implant surgery. Prior to surgery, make sure that all components and supplies have been sterilized by the appropriate procedures. For this step, an out of water U-shaped fish holder is needed. In this protocol, an aluminum holder that fits a 15 cm head to tail long goldfish is used. This system holds the fish out of water while perfusing the gills with oxygenated water. For details see Vinepinsky et al.8.
<ol>
	<li>Place the fish in a 0.02% MS-222 water bath for 20 min until the fish is asleep.</li>
	<li>Using sterile gloves, take the fish out of the water and place it in the holder. 
	NOTE: The oxygenated water perfusing the fish contains MS-222 at a concentration of 0.02%, so that the fish remains anesthetized during surgery.</li>
	<li>Using a sterile spatula, apply lidocaine 5% paste on the skin above the designated place for surgery for 10 min, then remove the lidocaine. 
	NOTE: Consult an appropriate brain atlas to target the specific brain region.</li>
	<li>Using a sterile 15 blade scalpel, remove the skin above the skull in the region of implant.</li>
	<li>Using a dental drill with 0.7 mm drill bits, drill 4 holes in the skull. Insert a 1 mm screw (3 mm long) into each hole and apply cyanoacrylate glue on the holes right before inserting the screw.</li>
	<li>Using a dental burnisher, apply dental cement on the screws and on the periphery of the exposed skull.</li>
	<li>Using the dental drill, make a 5 mm diameter hole in the skull above the brain region of interest. Remove the fatty tissue between the skull and the brain and expose the brain region target using fine tweezers and soft tissue paper. Be careful not to damage the large blood vessels underneath the skull. 
	NOTE: By the end of this stage, the fish is prepared to implant the probe. Only the main steps specific to this protocol are described here. Several postoperative procedures (such as detailed documentation on the animal&#39;s health and sterilization of the surgery tools and area) are not presented or discussed because they are applicable to all surgeries with fish or small animals.</li>
</ol>
7. Implanting the Probe
NOTE: To complete the final step in the protocol, a manipulator that can hold the implant in place while it is inserted into the brain is needed.
<ol>
	<li>Use the manipulator to hold the logger box cover with the tetrodes pointing down toward the fish brain.</li>
	<li>Bend the reference electrode such that when the tetrodes are lowered into the brain, the reference stays outside the brain.</li>
	<li>Cut the grounds such that they fit into the skull. Optionally, connect one ground wire to one of the skull screws.</li>
	<li>Lower the implant such that the electrodes are inserted into the brain while the bottom part of the microdrive housing is near the skull.</li>
	<li>Start attaching the implant to the skull by applying a small quantity of dental cement between the housing and the nearest skull screw.</li>
	<li>After the first part of the dental cement is cured, apply dental cement and close the hole above the skull and the entire exposed skull. 
	NOTE: Usually several rounds of dental cement applications are needed in order to cover the entire exposed skull.</li>
	<li>Install the logger and the battery in the box and seal the box with all the screws.</li>
	<li>Apply antibiotics and local painkillers according to the type of fish used for the experiments.</li>
	<li>Flush the fish&#39;s gills with fresh water until the fish starts to wake up. Remove the fish from the holder and place it back in its home tank. 
	NOTE: The fish is fully recovered within 60 min after surgery.</li>
	<li>Make sure the fish is able to swim freely with the implant (Figure 3, Supplementary Video 1). If needed, readjust the size of the extruded polystyrene foam above the logger box such that the fish can balance easily.</li>
</ol>

<ol>
	<li>Jacobson, M., Gaze, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Types+of+visual+response+from+single+units+in+the+optic+tectum+and+optic+nerve+of+the+goldfish.">Types of visual response from single units in the optic tectum and optic nerve of the goldfish.</a> Quarterly Journal of Experimental Physiology and Cognate Medical Sciences. 49 (2), 199-209 (1964).</li><li>Ben-Tov, M., Donchin, O., Ben-Shahar, O., Segev, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pop-out+in+visual+search+of+moving+targets+in+the+archer+fish.">Pop-out in visual search of moving targets in the archer fish.</a> Nature Communications. 6, 6476 (2015).</li><li>Zottoli, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlation+of+the+startle+reflex+and+Mauthner+cell+auditory+responses+in+unrestrained+goldfish.">Correlation of the startle reflex and Mauthner cell auditory responses in unrestrained goldfish.</a> Journal of Experimental Biology. 66 (1), 243-254 (1977).</li><li>Canfield, J. G., Rose, G. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+Mauthner+neurons+during+prey+capture.">Activation of Mauthner neurons during prey capture.</a> Journal of Comparative Physiology A. 172 (5), 611-618 (1993).</li><li>Canfield, J. G., Mizumori, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+chronic+neural+recording+in+the+telencephalon+of+freely+behaving+fish.">Methods for chronic neural recording in the telencephalon of freely behaving fish.</a> Journal of Neuroscience Methods. 133 (1-2), 127-134 (2004).</li><li>Orger, M. B., de Polavieja, G. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+behavior:+opportunities+and+challenges.">Zebrafish behavior: opportunities and challenges.</a> Annual Review of Neuroscience. 40, 125-147 (2017).</li><li>Vanwalleghem, G. C., Ahrens, M. B., Scott, E. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Integrative+whole-brain+neuroscience+in+larval+zebrafish.">Integrative whole-brain neuroscience in larval zebrafish.</a> Current Opinion in Neurobiology. 50, 136-145 (2018).</li><li>Vinepinsky, E., Donchin, O., Segev, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wireless+electrophysiology+of+the+brain+of+freely+swimming+goldfish.">Wireless electrophysiology of the brain of freely swimming goldfish.</a> Journal of Neuroscience Methods. 278, 76-86 (2017).</li><li>Vandecasteele, M., 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=Large-scale+recording+of+neurons+by+movable+silicon+probes+in+behaving+rodents.">Large-scale recording of neurons by movable silicon probes in behaving rodents.</a> JoVE (Journal of Visualized Experiments). (61), e3568 (2012).</li><li>Ferguson, J. E., Boldt, C., Redish, A. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Creating+low-impedance+tetrodes+by+electroplating+with+additives.">Creating low-impedance tetrodes by electroplating with additives.</a> Sensors and Actuators A: Physical. 156 (2), 388-393 (2009).</li><li>Arcot Desai, S., Rolston, J. D., Guo, L., Potter, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improving+impedance+of+implantable+microwire+multi-electrode+arrays+by+ultrasonic+electroplating+of+durable+platinum+black.">Improving impedance of implantable microwire multi-electrode arrays by ultrasonic electroplating of durable platinum black.</a> Frontiers in Neuroengineering. 3, 5 (2010).</li><li>Lewicki, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+review+of+methods+for+spike+sorting:+the+detection+and+classification+of+neural+action+potentials.">A review of methods for spike sorting: the detection and classification of neural action potentials.</a> Network: Computation in Neural Systems. 9 (4), R53-R78 (1998).</li><li>Teixeira, F. B., Freitas, P., Pessoa, L. M., Campos, R. L., Ricardo, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+IEEE+802.11+underwater+networks+operating+at+700+MHz,+2.4+GHz+and+5+GHz.">Evaluation of IEEE 802.11 underwater networks operating at 700 MHz, 2.4 GHz and 5 GHz.</a> Proceedings of the 10th International Conference on Underwater Networks &amp; Systems. , (2015).</li><li>Sendra, S., Lloret, J., Rodrigues, J. J., Aguiar, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Underwater+wireless+communications+in+freshwater+at+2.4+GHz.">Underwater wireless communications in freshwater at 2.4 GHz.</a> IEEE Communications Letters. 17 (9), 1794-1797 (2013).</li><li>Lloret, J., Sendra, S., Ardid, M., Rodrigues, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Underwater+wireless+sensor+communications+in+the+2.4+GHz+ISM+frequency+band.">Underwater wireless sensor communications in the 2.4 GHz ISM frequency band.</a> Sensors. 12 (4), 4237-4264 (2012).</li><li>Hoogerwerf, A. C., Wise, K. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+three-dimensional+microelectrode+array+for+chronic+neural+recording.">A three-dimensional microelectrode array for chronic neural recording.</a> IEEE Transactions on Biomedical Engineering. 41 (12), 1136-1146 (1994).</li><li>Harris, K. D., Quiroga, R. Q., Freeman, J., Smith, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improving+data+quality+in+neuronal+population+recordings.">Improving data quality in neuronal population recordings.</a> Nature Neuroscience. 19 (9), 1165 (2016).</li></ol>

The authors have nothing to disclose.

This protocol details the steps involved in implanting a tetrode array into the telencephalon of freely swimming goldfish. This technique implements a neural logger that amplifies and records the signals acquired from up to 16 channels along with a microdrive that can adjust the tetrode position in the brain. The microdrive makes it possible to adjust the position in the brain to optimize the recording.
This protocol can easily be modified for recording from other brain regions (see Vinepinsky...

Fish are the largest and most diverse group of vertebrates, and like other vertebrates they exhibit complex cognitive abilities such as navigating, socializing, sleeping, hunting, etc. Nevertheless, the neural mechanisms governing fish behavior remain for the most part unknown.
In the past few decades, extracellular recordings from immobilized fish have primarily been implemented to investigate different aspects of the neural basis of behavior1,2. Although this technique is appropriate for some sensory systems, investigation of the full spectrum of the neural basis of behavior is difficult if not impossible in immobilized animals. The first advances involved recording from the Mauthner cells of tethered swimming fish3,4. However, Mauthner cells are disproportionately large and the recorded action potential amplitudes, which can go as high as a few mV, facilitate recording. Later, Canfield et al. described a proof of concept when using a tethered animal to record from the telencephalon of fish5. Another recent technique for recording neural activity from fish is calcium imaging (see reviews by Orger and de Polavieja6, and Vanwalleghem et al.7). This technique was developed for use with zebrafish larvae because the skin and skull are transparent during the larval stage. However, this technique cannot be used to study complex behaviors in later stages of development.
Here, we present a novel technique for recording extracellular neural activity from the brains of freely swimming fish. This is a modified version of the protocol described in Vinepinsky et al.8. The main innovation is the addition of a microdrive that makes it possible to control the position of the electrodes after surgery. The technique is designed for recording from the telencephalon of goldfish using a set of tetrodes that are connected to a neural data logger via a microdrive. The whole setup is wireless and anchored to the fish&#39;s skull. The specific weight of the system is equalized to the water-specific weight by adding a small float that allows the fish to swim freely.
The technique is based on the use of a neural data logger that amplifies, digitizes, and stores the signal in an onboard memory device. The logger telemetry system is used to start and stop the recordings, and for synchronization with the video camera. In this protocol, a 16-channel neural logger is used, embedded in a waterproof box together with the microdrive.
The microdrive assembly is fabricated from two main components: the microdrive itself and the microdrive housing (Figure 1A,B). The housing holds the microdrive and the tetrodes, and also acts as the anchor between the skull and the logger box (Figure 1C). The PVC logger box is fabricated using a machine process and is sealed using an O-ring (Figure 1E-G, see also Supplementary Figure 1, Supplementary Figure 2, and Supplementary Figure 3 for a three-dimensional [3D] diagram). At one end, a piece of polystyrene foam is attached to the logger box to compensate for the weight of the implant and provide the fish with a buoyancy-neutral implant. The construction of the microdrive described in the protocol follows the procedure presented by Vandecasteele et al.9 with a modification to attach the microdrive to the housing (Figure 1A). All major steps are presented.
The procedure described in the protocol to prepare the fish skull is similar to the one presented in Vinepinsky et al.8 and is described briefly in the protocol. One day after surgery, the fish are normally fully recovered from the effects of anesthesia and are ready for the behavioral experiments. Note that the tetrode location can be adjusted by turning the microdrive screw. The screw has a spacing of 300 &#181;m per full rotation and an advancement of 75 &#181;m is recommended until the target brain location is reached. An appropriate brain atlas should be consulted to target the specific brain region of interest. It is advisable to test the electrode impedance each time the fish is anesthetized for battery or memory card replacement.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.7 mm round drill bits</td>
			<td></td>
			<td></td>
			<td>Compatible with the drill.</td>
		</tr>
		<tr>
			<td>15-blade Scalpel</td>
			<td>Sigma-Aldrich</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>16 channel PCB board</td>
			<td>Neurlynx</td>
			<td>EIB-16</td>
			<td></td>
		</tr>
		<tr>
			<td>1X3M phillips flat head screws</td>
			<td></td>
			<td></td>
			<td>Stainless steel. Any type.</td>
		</tr>
		<tr>
			<td>1X3M phillips round head screws</td>
			<td></td>
			<td></td>
			<td>Stainless steel. Any type.</td>
		</tr>
		<tr>
			<td>27 cm X 19 cm X 1 mm brass plate</td>
			<td></td>
			<td></td>
			<td>See Figure 2</td>
		</tr>
		<tr>
			<td>2X6M phillips flat head screws</td>
			<td></td>
			<td></td>
			<td>Stainless steel. Any type.</td>
		</tr>
		<tr>
			<td>3140 RTV coating</td>
			<td>Dow Crowning</td>
			<td>2767996</td>
			<td></td>
		</tr>
		<tr>
			<td>75 &#181;m Silver wire</td>
			<td>A-M Systems</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Brass machine screws #00-90</td>
			<td></td>
			<td>947-1006</td>
			<td></td>
		</tr>
		<tr>
			<td>Brass plates 7.5mm X 2.5mm X 0.6mm</td>
			<td></td>
			<td></td>
			<td>A 3D drawing is provided. See supplementary 1</td>
		</tr>
		<tr>
			<td>Coated Tungsten wire 25&#181;m</td>
			<td>California Fine Wire Company</td>
			<td>5000160</td>
			<td>Depending on the appication the tetrodes can be fabricated from any type of wire. Popular wires are nicrome wires that can be found with lower diameters (eg. A-M systems, 762000)</td>
		</tr>
		<tr>
			<td>Coated Tungsten wire 50&#181;m</td>
			<td>A-M Systems</td>
			<td>795500</td>
			<td>Can be replaced with any other wire with low impedance</td>
		</tr>
		<tr>
			<td>Cyanoacrilic glue</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dental Burnisher</td>
			<td>ComDent UK</td>
			<td></td>
			<td>Any small sterille stainless-still tool will do.</td>
		</tr>
		<tr>
			<td>Dental cement - GCFujiPLUS</td>
			<td>GC</td>
			<td>431011</td>
			<td>Other dental cements would probably will work as well although we have never tried any other.</td>
		</tr>
		<tr>
			<td>Dental drill or nail polish drill</td>
			<td></td>
			<td></td>
			<td>Dental drills are expensive, a nail polish drill can be a cheap replacement.</td>
		</tr>
		<tr>
			<td>Drill bit #65</td>
			<td></td>
			<td>947-65</td>
			<td></td>
		</tr>
		<tr>
			<td>Fast curing epoxy</td>
			<td></td>
			<td></td>
			<td>Any 5 minutes curing epoxy can be used here.</td>
		</tr>
		<tr>
			<td>Logger box with O-ring sealing</td>
			<td></td>
			<td></td>
			<td>A 3D drawing is provided. See supplementary 1-3. The box should be machine fabricated (do not use 3D printers). Use transperant material, to be able to see the indicator LEDs on the logger.</td>
		</tr>
		<tr>
			<td>Motorized turning device</td>
			<td></td>
			<td></td>
			<td>Custom made as described in &#34;open ephys&#34; website. Can also be purchusaed from neurolynx (&#34;Tetrode Spinner 2.0&#34;) or bulit by other means.</td>
		</tr>
		<tr>
			<td>Mouselog-16 Neural logger</td>
			<td>Deuteron Technologies Ltd</td>
			<td></td>
			<td>There are several neural loggers available on the market, including: SpikeGadget (UH32 32channels) and Neurologger 2/2A/2B of Alexei Vyssotski. It should be noted that weight is not a major contraint since it can be counterbalanced with floating Styrofoam</td>
		</tr>
		<tr>
			<td>MS-222</td>
			<td>Sigma Aldrich</td>
			<td>E10521</td>
			<td>Ethtl 3-aminobenzoate methanesulfonate 98%</td>
		</tr>
		<tr>
			<td>Nano-Z plating</td>
			<td>White Matter LLC</td>
			<td></td>
			<td>The nano-Z can be bought from several supllieres. Any impedance meter can be used, e.g. IMP-1 / 6662 / 2788, BAK Electronics.</td>
		</tr>
		<tr>
			<td>PCB pins</td>
			<td>Neurlynx</td>
			<td>Neuralynx EIB Pins</td>
			<td></td>
		</tr>
		<tr>
			<td>Polymide tubing 250&#181;m</td>
			<td>A-M Systems</td>
			<td>822000</td>
			<td></td>
		</tr>
		<tr>
			<td>Rechargable battery</td>
			<td></td>
			<td></td>
			<td>3.7 Lipo battery, 370 mAh. Holds about 6 hours of recording. Smaller or larger battries can be used to reduce the weight or extend recording time.</td>
		</tr>
		<tr>
			<td>Silicone tubing 0.64 mm</td>
			<td>A-M Systems</td>
			<td>806100</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless steel 1.5 mm</td>
			<td>A-M Systems</td>
			<td>846000</td>
			<td></td>
		</tr>
		<tr>
			<td>Sudium Bicarbonate</td>
			<td>Sigma Aldrich</td>
			<td>S9625</td>
			<td></td>
		</tr>
		<tr>
			<td>Tap #00-90</td>
			<td></td>
			<td>947-1301</td>
			<td></td>
		</tr>
		<tr>
			<td>Vaseline</td>
			<td></td>
			<td></td>
			<td>Any type of soft petroleum skin protectant can be used here.</td>
		</tr>
	</tbody>
</table>

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.

During a recording session the goldfish swam freely in a square water tank while the neural activity in its telencephalon was recorded. The goal of these experiments was to study how the neural activity of single cells determines the fish&#39;s behavior. To do so, spiking activity needed to be identified in the recorded data. The brain activity, while being recorded, was digitized at 31,250 Hz and high-pass filtered at 300 Hz by the data logger. Then, offline, a band-pass filter (300&#872...

Watch this Scientific Journal Video about Wireless Electrophysiological Recording of Neurons by Movable Tetrodes in Freely Swimming Fish at JoVE.com

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

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 ...

wireless-electrophysiological-recording-neurons-movable-tetrodes

Research

JoVE Journal

Neuroscience

8.7K Views.  Ben-Gurion University of the Negev. 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.

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

Large-scale Recording of Neurons by Movable Silicon Probes in Behaving Rodents

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of neurons and circuits requires methods with the spatial selectivity and temporal resolution appropriate for mechanistic analysis of neural ensembles in the behaving animal, i.e. recording of representatively large samples of isolated single neurons. Ensemble monitoring of neuronal activity has progressed remarkably in the past decade in both small and large-brained animals, including human subjects1-11. Multiple-site recording with silicon-based devices are particularly effective because of their scalability, small volume and geometric design.
Here, we describe methods for recording multiple single neurons and local field potential in behaving rodents, using commercially available micro-machined silicon probes with custom-made accessory components. There are two basic options for interfacing silicon probes to preamplifiers: printed circuit boards and flexible cables. Probe supplying companies (<a href="http://www.neuronexustech.com/" >http://www.neuronexustech.com/</a>; <a href="http://www.sbmicrosystems.com/">http://www.sbmicrosystems.com/</a>; <a href="http://www.acreo.se/">http://www.acreo.se/</a>) usually provide the bonding service and deliver probes bonded to printed circuit boards or flexible cables. Here, we describe the implantation of a 4-shank, 32-site probe attached to flexible polyimide cable, and mounted on a movable microdrive. Each step of the probe preparation, microdrive construction and surgery is illustrated so that the end user can easily replicate the process.

We describe methods for large-scale recording of multiple single units and local field potential in behaving rodents with silicon probes. Drive fabrication, probe attachment to the drive and probe implantation processes are illustrated in sufficient details for easy replication.

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of ...

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 ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Long-term Behavioral Tracking of Freely Swimming Weakly Electric Fish

Long-term behavioral tracking can capture and quantify natural animal behaviors, including those occurring infrequently. Behaviors such as exploration and social interactions can be best studied by observing unrestrained, freely behaving animals. Weakly electric fish (WEF) display readily observable exploratory and social behaviors by emitting electric organ discharge (EOD). Here, we describe three effective techniques to synchronously measure the EOD, body position, and posture of a free-swimming WEF for an extended period of time. First, we describe the construction of an experimental tank inside of an isolation chamber designed to block external sources of sensory stimuli such as light, sound, and vibration. The aquarium was partitioned to accommodate four test specimens, and automated gates remotely control the animals&#39; access to the central arena. Second, we describe a precise and reliable real-time EOD timing measurement method from freely swimming WEF. Signal distortions caused by the animal&#39;s body movements are corrected by spatial averaging and temporal processing stages. Third, we describe an underwater near-infrared imaging setup to observe unperturbed nocturnal animal behaviors. Infrared light pulses were used to synchronize the timing between the video and the physiological signal over a long recording duration. Our automated tracking software measures the animal&#39;s body position and posture reliably in an aquatic scene. In combination, these techniques enable long term observation of spontaneous behavior of freely swimming weakly electric fish in a reliable and precise manner. We believe our method can be similarly applied to the study of other aquatic animals by relating their physiological signals with exploratory or social behaviors.

We describe a set of techniques for studying spontaneous behavior of freely swimming weakly electric fish over an extended period of time, by synchronously measuring the animal&#39;s electric organ discharge timing, body position and posture both accurately and reliably in a specially designed aquarium tank inside a sensory isolation chamber.

Long-term behavioral tracking can capture and quantify natural animal behaviors, including those occurring infrequently. Behaviors such as exploration and ...

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

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

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

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

Recording Single Neurons' Action Potentials from Freely Moving Pigeons Across Three Stages of Learning

While the subject of learning has attracted immense interest from both behavioral and neural scientists, only relatively few investigators have observed single-neuron activity while animals are acquiring an operantly conditioned response, or when that response is extinguished. But even in these cases, observation periods usually encompass only a single stage of learning, i.e. acquisition or extinction, but not both (exceptions include protocols employing reversal learning; see Bingman&#160;et al.1 for an example). However, acquisition and extinction entail different learning mechanisms and are therefore expected to be accompanied by different types and/or loci of neural plasticity.
Accordingly, we developed a behavioral paradigm which institutes three stages of learning in a single behavioral session and which is well suited for the simultaneous recording of single neurons&#39; action potentials. Animals are trained on a single-interval forced choice task which requires mapping each of two possible choice responses to the presentation of different novel visual stimuli (acquisition). After having reached a predefined performance criterion, one of the two choice responses is no longer reinforced (extinction). Following a certain decrement in performance level, correct responses are reinforced again (reacquisition). By using a new set of stimuli in every session, animals can undergo the acquisition-extinction-reacquisition process repeatedly. Because all three stages of learning occur in a single behavioral session, the paradigm is ideal for the simultaneous observation of the spiking output of multiple single neurons. We use pigeons as model systems, but the task can easily be adapted to any other species capable of conditioned discrimination learning.

Learning new stimulus-response associations engages a wide range of neural processes which are ultimately reflected in changing spike output of individual neurons. Here we describe a behavioral protocol allowing for the continuous registration of single-neuron activity while animals acquire, extinguish, and reacquire a conditioned response within a single experimental session.

While the subject of learning has attracted immense interest from both behavioral and neural scientists, only relatively few investigators have observed ...

Extracellular Wire Tetrode Recording in Brain of Freely Walking Insects

Increasing interest in the role of brain activity in insect motor control requires that we be able to monitor neural activity while insects perform natural behavior. We previously developed a technique for implanting tetrode wires into the central complex of cockroach brains that allowed us to record activity from multiple neurons simultaneously while a tethered cockroach turned or altered walking speed. While a major advance, tethered preparations provide access to limited behaviors and often lack feedback processes that occur in freely moving animals. We now present a modified version of that technique that allows us to record from the central complex of freely moving cockroaches as they walk in an arena and deal with barriers by turning, climbing or tunneling. Coupled with high speed video and cluster cutting, we can now relate brain activity to various parameters of the movement of freely behaving insects.

We previously developed a technique for implanting tetrode wires into the central complex of cockroach brains that allows us to monitor activity in individual units of tethered cockroaches. Here we present a modified version of that technique that allows us to also record brain activity in freely moving insects.

Increasing interest in the role of brain activity in insect motor control requires that we be able to monitor neural activity while insects perform ...

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the researcher to measure gustatory responses directly and quantitatively, reflecting the sensory input that the insect nervous system receives from taste stimuli in its environment. This protocol outlines all key steps in performing this technique. The critical steps in assembling an electrophysiology rig, such as selection of necessary equipment and a suitable environment for recording, are delineated. We also describe how to prepare for recording by making appropriate reference and recording electrodes, and tastant solutions. We describe in detail the method used for preparing the insect by insertion of a glass reference electrode into the fly in order to immobilize the proboscis. We show traces of the electrical impulses fired by taste neurons in response to a sugar and a bitter compound. Aspects of the protocol are technically challenging and we include an extensive description of some common technical challenges that may be encountered, such as lack of signal or excessive noise in the system, and potential solutions. The technique has limitations, such as the inability to deliver temporally complex stimuli, observe background firing immediately prior to stimulus delivery, or use water-insoluble taste compounds conveniently. Despite these limitations, this technique (including minor variations referenced in the protocol) is a standard, broadly accepted procedure for recording Drosophila neuronal responses to taste compounds.

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

This protocol describes extracellular recording of the action potential responses fired by labellar taste neurons in Drosophila.

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the ...

Recording EEG in Freely Moving Neonatal Rats Using a Novel Method

EEG is a useful method to detect electrical activity in the brain. Moreover, it is a widely used diagnostic tool for various neurological conditions, such as epilepsy and neurodegenerative disorders. However, it is technically difficult to obtain EEG recordings in neonates as it requires specialized handling and great care. Here, we present a novel method to record EEG in neonatal rat pups (P8-P15). We designed a simple and reliable electrode using computer pin loci; it can be easily implanted into the skull of a rat pup to record high-quality EEG signals in the normal and epileptic brain. Pups were given an intraperitoneal (i.p.) injection of the neurotoxin kainic acid (KA) to induce epileptic seizures. The surgical implantation performed in this procedure is less expensive than other EEG procedures for neonates. This method allows one to record high-quality and stable EEG signals for more than 1 week. Furthermore, this procedure can also be applied to adult rats and mice to study epilepsy or other neurological disorders.

Here, we introduce a novel technique designed to record electroencephalography (EEG) in freely moving neonatal epileptic pups and describe its procedures, features, and applications. This method allows one to record EEG for more than 1 week.

EEG is a useful method to detect electrical activity in the brain. Moreover, it is a widely used diagnostic tool for various neurological conditions, such ...