We thank Daniel Carpi for his help and early contributions to this project. We also thank Lucrecia Novoa for her assistance with artwork and images. This work was supported by NIH/NIAID program grant 5P01AI073693-03.

1. Tetrode Fabrication
<ol>
	<li>Start by using insulated 12.5 &mu;m (0.0005") diameter core platinum-iridium wire from California Fine Wire. The length of the wire should be cut to the appropriate length for the target structure. For example, cut the wire to at least 30 cm long for targeting the dorsal subiculum or hippocampus.</li>
	<li>Fold the wire over at the center so that there are two parallel wires which will be 15 cm in length. Drape the midpoint of this wire over a horizontal arm to form four parallel wires of 7.5 cm in length. Next attach the rubber-coated clip near the bottom of the draped wire, creating a bundle of four wires.</li>
	<li>Place the rubber clip into the motorized Tetrode Spinner, making sure that the wire is taut, but not too taut or bearing weight as it will break during the spinning process.</li>
	<li>Switch the Tetrode Spinner to "Manual" mode and push the joystick to "Right" to spin the wire in a clockwise direction. The spinner will rotate at approximately 2 Hz, creating a tight bundle of four wires to form the tetrode.</li>
	<li>Apply 80 clockwise rotations then stop by pushing "Up" on the joystick. This will pause the motorized spinner. Next, apply 20 counter-clockwise ("Left") rotations in order to release tension on the tetrode. The final number of rotations per length of wire should be 8 rotations per micron.</li>
	<li>Use the heating gun on the lower setting 1, which reaches a maximum of 400 degrees, in order to fuse the wires together by melting the VG bond coat. Hold the heat gun ~2 cm from the wire and run the gun up and down the straight length of the wire for about 5 sec from several different angles. Make sure to constantly sweep the heat gun and not hold it any single location as this will melt the HML insulation and cause the wires to fuse together within the bundle.</li>
	<li>Make a cut at the top of the tetrode (near the horizontal arm) and then release the tetrode from the clip at the bottom. Cut the single loop so that there are four separate wires on one end of the tetrode, these wires will be electrically connected to gold pins or a circuit board at a later step.</li>
	<li>Place the completed tetrode in a dust-free holding box for storage until the drive has been completed.</li>
</ol>
2. Custom Microdrive Assembly
<ol>
	<li>First construct the base that will hold the microdrive(s). The base of the implanted microdrive is generally most stable if it is secured and positioned along the midline of the skull. This protocol describes steps to build a base with a single microdrive to hold four polyamide tube carriers. Additional microdrives and tubes can be easily added as needed.</li>
	<li>Start with an approximately 20 mm square piece of Plexiglas acrylic (5 mm thick) and sand the acrylic into a shape that will allow the mouse to move freely with the drive after it is implanted on the head.</li>
	<li>Next, assemble the drive unit. Use customized 3.3 x 6.3 mm brass guides that will carry the drive screw. First, solder two brass guides together perpendicularly. The vertical brass guide will hold the drive screw and electrodes, while the horizontal piece will be glued into the acrylic base.</li>
	<li>After soldering the brass pieces together, begin assembly of the drive itself by passing a fillister head brass screw through the top of the guide and into a Delrin plastic block. The square block is designed so that the thread hole is slightly off center (by 0.2 mm), resulting in one face of the block protruding very slightly from the guide. This is the side where the polyamide tubes carrying the electrodes will sit.</li>
	<li>With the Delrin block inside the guide, and the screw all the way through, thread a hex brass nut until the nut is nearly touching the bottom of the guide. Do not tighten the nut fully; instead melt a small amount of solder onto the end in order to join the nut and the screw but being careful not to solder anything to the guide. Now, rotating the screw should move the Delrin block up (clockwise) and down (counterclockwise) vertically along the screw thread. Cut off thread that protrudes past the soldered nut.</li>
	<li>Once the drive has been assembled, go back to the acrylic base and cut a 3 mm wide slot where the electrode drive will be. Pass the horizontal brass guide through the slot and then use cryanoacrylate glue to secure the piece to the base.</li>
	<li>Place the acrylic base in a vise to secure it in place. Place a electronic interface board (EIB) on top of the base and mark the locations of the two screw holes. EIBs are microchips that provide a signal connection between the electrode wires and a pre-amplifier headstage. Using a 1.5 mm tip drill bit, carefully drill holes at the marks for screws that will hold the EIB in place on top of the base. Place the EIB and thread two brass screws into the holes.</li>
	<li>Use micro dissecting scissors to cut four 7 mm long pieces of polyamide tubing. Line the four tubes next to each other on a piece of folded laboratory tape. Apply cyanoacrylate to the center to join them together but take caution not to get glue within the tubes themselves. Allow the joined tubes to dry fully.</li>
	<li>Rotate the drive base 90 degrees so that the EIB chip is vertical and the drive is positioned horizontally with the protruding Delrin block facing upward. Looking through a dissecting microscope, carefully dab a small amount of cyanoacrylate on the Delrin face then place the four joined tubes on the glue. Allow the glue to set completely before attempting to move the drive.</li>
	<li>Test that the polyamide tubes are securely attached and that the entire assembly moves smoothly without touching the guide or meeting any resistance.</li>
	<li>Next, prepare the ground screw and connect the ground wire to the EIB. Make a ground screw by taking a brass screw (3/32") and sanding down the threads until only 1-2 threads remain. This should be ~1 mm as this screw will sit within the skull and is not intended to penetrate brain tissue.</li>
	<li>Cut a 30 mm length of copper wire (the exact length will depend on where on the skull to place the animal ground). The copper wire should be 100 - 500 &mu;M (0.004-0.02") in diameter, this is roughly equivalent to 38 AWG thru 24 AWG wire. Apply solder flux to both ends of the copper wire. On one end, solder the ground screw to the wire. On the other end, solder an EIB gold pin. This ground wire can be set aside and connected to the EIB later during the implantation surgery.</li>
	<li>The next step is to guide the electrodes through the polyamide tubes and connect them to the channel holes on the EIB chip. Turn the drive screw fully clockwise so that the tubes are at their top position.</li>
	<li>For single electrodes, cut a 50 mm length of Stableohm 50 &mu;M wire and guide it through one polyamide tube, allowing it to extend at least 2.0 mm past the tube end (for targeting subiculum or hippocampus). Apply a small drop of cyanoacrylate at the top of the tube, affixing the wire to the tube and preventing any wire movement. Next, connect the loose end of the wire to an EIB channel hole using a gold pin. Trim off any excess wire with fine scissors. Repeat for other microelectrodes.</li>
	<li>For connecting tetrodes, take one completed tetrode out of the storage box. Guide the fused end of the tetrode through one polyamide tube and allow it to extend at least 2.0 mm past the tube end (for subiculum or hippocampus). Apply a small drop of cyanoacrylate at the top of the tube, affixing the tetrode to the tube and preventing any movement. Take the four loose wires at the other end of the tetrode and connect each wire to an EIB channel hole using a gold pin. Trim off any excess wire. Repeat for the other tetrodes.</li>
</ol>
3. VersaDrive Assembly
<ol>
	<li>Begin constructing a four tetrode VersaDrive; this consists of a base, enclosure, and cap pieces.</li>
	<li>Cut one polyamide tubing to 10 mm and guide it through the smallest hole on a tetrode carrier. Allow the tube to extend past the carrier very slightly (0.5 mm). Use 5-min epoxy to glue the polyamide tube in place, being careful not to allow the epoxy to go into the tube itself. Repeat this for three other tubes and carriers.</li>
	<li>After the epoxy has fully set, guide each polyamide tube through one of the four holes on the VersaDrive base. Once all four tubes are through their holes, push an insect pin through the outer hole; this will hold the tetrode carrier in line and serve as a rail for the carrier to travel on. Repeat this for the three other carriers.</li>
	<li>Take a cap and line it up with the four insect pins so that the cap covers the base and the tetrode carriers reside within the cap. Thread a 1 mm x 5 mm machine screw through the appropriate hole in the cap and into the tetrode carrier. This will be the drive screw for moving the carrier up and down. Repeat this for the other three screws.</li>
	<li>Turn all screws clockwise until tetrode carriers are at their top position and the polyamide tubes are visible through the cap opening. Using fine micro dissecting scissors, cut the tubing just below (1 mm) the base so that all four polyamide tubes are of the same length.</li>
	<li>Using a dissecting microscope, carefully thread one tetrode through a polyamide tube. It is important to keep the tetrode wire perfectly straight as it advances through the tube as any kinks or bends will make it very difficult to fully thread the tetrode through. Repeat for three other tetrodes.</li>
	<li>Once all the tetrodes are in their tubes, carefully apply a small drop of cyanoacrylate to the top of each tube, securing the tetrodes within their respective tubes. Take caution not to get any cyanoacrylate between the carriers or on the loose tetrode wires that are protruding through the cap.</li>
	<li>Cut the tetrodes so that they only extend past the tubes 2.0 mm (for subiculum or hippocampus). Then place the drive base (with the four insect pins inserted) into the VersaDrive jig. The other half of the jig will hold the VersaDrive cap that has all the receptacle holes for making the channel connections.</li>
	<li>Turn the drive screw completely counterclockwise so that the tetrodes are in their lowest position.</li>
	<li>Before connecting the tetrode wires to the gold receptacles, first connect the ground wires to the cap. The VersaDrive cap has two pin holes for ground connections at the center position of the two rows of holes. Cut a copper wire of at least 30 mm (depends where on the skull to place the ground) and guide it through one of these center holes. The copper wire should be 100 - 500 &mu;M (0.004-0.02") in diameter, this is roughly equivalent to 38 AWG thru 24 AWG wire. Push a gold receptacle through the hole to catch the copper wire in place and trim any excess wire. On the other end of the copper wire, apply flux and solder this wire end to a ground screw (see 2.11.). Repeat for the second ground wire.</li>
	<li>Next, guide all the loose tetrode wires (there should be sixteen in total) through their respective receptacle holes on the cap. It's best to start with one tetrode and thread the individual wires to the appropriate four holes that will end up directly above it. The individual tetrode wires should be handled with light pressure as they are fragile and can crimp easily if gripped too firmly. Install the cap by lining up the insect pins holes and press fitting to the base.</li>
	<li>With the tetrode wires protruding through the cap, press fit the gold receptacles to capture the tetrode wires in place and make the electrical connections. Approximately 50% of the wires will be clipped off (above the cap) once the gold receptacle is pushed down. Trim any excess wire that remains protruding from the top of the cap. In rare instances (less than 5%), pushing the down gold receptacle will crush the wire and break it below receptacle, resulting in a disconnected channel. This disconnection may not be realized until the impedance testing and electroplating steps (see 4.7).</li>
	<li>Repeat the press-fitting process for the three other tetrodes. Turn the drive screws clockwise to move them back to the top and ensure that the drive movement is smooth.</li>
</ol>
4. Gold-plating of Electrode Tips
<ol>
	<li>Regardless of what type of microelectrodes being used, the tips of the electrodes should be gold-plated in order to reduce tip impedance. This will maximize the ability to reliably record and discriminate single-unit action potentials. Test the electrode impedance using the Neuralynx nanoZ device. The nanoZ is a computer-based device that measures impedance and allows for automated electroplating.</li>
	<li>First turn the microdrive screws down (counter-clockwise) to their lowest position. Then securely mount the microdrive on a clamp that will allow lowering of the electrode tips into the gold plating solution.</li>
	<li>Fill one Delrin tower with SIFCO Gold solution and the other tower with distilled water. Lower the electrode tips into the gold solution.</li>
	<li>Plug the nanoZ USB cable into a Windows-based computer and then open the nanoZ program. This program will give impedance readings and perform gold-plating on each connected channel of the microdrive.</li>
	<li>Go to the Device drop-down and select the nanoZ, after which it will show "Connection established" at the bottom of the window. Next, select the appropriate adaptor for testing in the drop-down menu. Click on "Test impedances" and set the test frequency to 1004 Hz (40 cycles, 0 msec pause). Click on "Test probe", which will open the "Probe Report" window that shows all the available channels with their M&Omega; readings. Save these impedance values by clicking on the disk icon or by selecting "File", then "Save report".</li>
	<li>Next, click on "DC Electroplate" and assign the following values: Mode = match impedances, Plating current = -1.0 &mu;A, Target = 350 k&Omega; at 1,004 Hz, 5 runs, 5 sec interval, 2 sec pause.</li>
	<li>Click "Autoplate". The program will first read the impedance of each channel, then apply the specified current to that channel, re-test the impedance and apply current as needed until the Target impedance (or a lower value) is reached. While the goal is to reduce electrode impedance, it is possible that channels will electroplate below values of 100 k&Omega;. In such cases, it is possible that neighboring wires on the tetrode have been shorted together. If this happens, reverse the current polarity (+ 1.0 &mu;A) to remove excess gold particles, re-test the impedance of that channel, and then repeat the electroplating. Typical final impedance values on a bundle of four 12.5 &mu;M wires range from 150 - 325 k&Omega;.</li>
	<li>If there is any single channel that has not plated below 350 k&Omega;, repeat the electroplating process. The program will skip over channels that have already reached the Target and will only plate channels which have not.</li>
	<li>Once all channels have been plated to an acceptable impedance, close the nanoZ program and disconnect the device. Raise the electrodes out of the plating solution and lower the tips into the distilled water Delrin tower in order to rinse off excess gold particles.</li>
	<li>Turn the drive screws clockwise until the electrodes are raised to their top position. Now the microdrive and electrodes are ready for implantation.</li>
</ol>

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The authors declare that they have no competing financial interests.

We have described a set of techniques for constructing light and compact microdrives for the recording of extracellular unit and field potential activity in mice. By building custom microdrives with bases fashioned from acrylic glass (methyl methacrylate), the core system can be easily adapted for multiple drives and for the targeting of a wide array of neural regions. We have successfully modified the system for recording from multiple brain targets and with larger arrays for recordings in mice. With further modif...

The use of the microelectrode technique for recording extracellular neural signals in vivo has a long and valued tradition in neuroscience 1, 2. The ability to record electrical activity from many brain regions in freely behaving animals is, however, a more recent technology that is becoming increasingly common as the software packages for the acquisition, analysis and discrimination of neural signals becomes more sophisticated and user-friendly 3, 4. The technological advances on the software side have also been accompanied by reductions in the weight and bulk of the implantable devices, which have been scaled down sufficiently for recording in small mammals, such as mice. By using lightweight (mostly plastic) components, researchers are able to construct microdrives that allow for independent positioning of electrodes or tetrodes to target a wide variety of brain regions 5-7. Even deep brain structures, such as the amygdala 6 and the striatum 5, can be routinely targeted with the selection of an appropriately long drive screw. These recording techniques allow researchers to obtain high-fidelity neural signals and are in register with the electrical activity of single neurons recorded intracellularly 8, 9. Using these types of microdrives, we have successfully recorded single-units from mice for up to two months after implantation 10. In addition, the lightweight nature of the devices (approximately 1.5-2.0 g) has resulted in behavioral performance that is comparable to non-implanted mice in many behavioral tasks. In particular, we have demonstrated that implanted mice exhibit normal performance in the novel object recognition task 10 and the object place task (unpublished data).	
	The use of microdrives coupled to multiple tetrodes allows researchers to monitor and analyze neural activity at the network level while also recording from multiple single-units within the brain. Recording with these tetrodes has several major advantages for unit identification purposes and enables the high accuracy acquisition and discrimination of multiple single-units 11. We describe how to fabricate and gold-plate tetrode bundles and then subsequently load them into driveable electrode carriers. One type of drive carrier we describe is commercially available and the other is a simple, but easily expandable, drive design that can accommodate multiple carriers and tetrode arrangements without a significant investment of resources.

<table border="1">
		<tbody>
		 <tr>
			<td>Name of Reagent/Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		 </tr>
		 <tr>
			<td>0.0005" (12.5 &mu;M) diameter Platinum-Iridium wire </td>
			<td>California Fine Wire</td>
			<td>CFW#100-167</td>
			<td>HML VG insulated <a href="http://www.calfinewire.com" target="_blank">www.calfinewire.com</a></td>
		 </tr>
		 <tr>
			<td>0.002" (50 &mu;M) diameter Stableohm 675 wire</td>
			<td>California Fine Wire</td>
			<td>CFW# 100-188</td>
			<td>HML insulated Ni-Cr</td>
		 </tr>
		 <tr>
			<td>polyamide tubing</td>
			<td>Polymicro Technologies</td>
			<td>1068150020</td>
			<td>99 micron I.D., 166 micron O.D. <a href="http://www.polymicro.com" target="_blank">www.polymicro.com</a></td>
		 </tr>
		 <tr>
			<td>brass guides</td>
			<td>World Plastics Inc</td>
			<td></td>
			<td>3.3 x 6.6 mm</td>
		 </tr>
		 <tr>
			<td>Delrin blocks</td>
			<td>World Plastics Inc</td>
			<td></td>
			<td>3.13 x 2.5 mm</td>
		 </tr>
		 <tr>
			<td>Fillister head brass screws</td>
			<td>J.I. Morris Co.</td>
			<td>00-90 x 1/2</td>
			<td>drive screw <a href="http://www.jimorrisco.com" target="_blank">www.jimorrisco.com</a></td>
		 </tr>
		 <tr>
			<td>hex brass nuts</td>
			<td>J.I. Morris Co.</td>
			<td>00-90</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Fillister head brass screws</td>
			<td>J.I. Morris Co.</td>
			<td>000-120 x 3/32</td>
			<td>EIB mount and ground screw</td>
		 </tr>
		 <tr>
			<td>plexiglass acrylic</td>
			<td>Canal Street Plastics</td>
			<td></td>
			<td>5 mm thick, clear, <a href="http://www.cpcnyc.com" target="_blank"> www.cpcnyc.com</a></td>
		 </tr>
		 <tr>
			<td>cyanoacrylate</td>
			<td>Krazy Glue</td>
			<td></td>
			<td>2 g tube</td>
		 </tr>
		 <tr>
			<td>electronic interface board</td>
			<td>Neuralynx</td>
			<td>EIB-18</td>
			<td><a href="http://www.neuralynx.com" target="_blank">www.neuralynx.com</a></td>
		 </tr>
		 <tr>
			<td>non-cyanide gold solution</td>
			<td>SIFCO</td>
			<td>SIFCO 5355</td>
			<td><a href="http://www.sifcoasc.com" target="_blank">www.sifcoasc.com</a></td>
		 </tr>
		 <tr>
			<td>VersaDrive 4</td>
			<td>Neuralynx</td>
			<td></td>
			<td>four tetrode model</td>
		 </tr>
		 <tr>
			<td>tetrode assembly station</td>
			<td>Neuralynx</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>motorized tetrode spinner</td>
			<td>Neuralynx</td>
			<td>tetrode spinner 2.0</td>
			<td></td>
		 </tr>
		 <tr>
			<td>VersaDrive jig</td>
			<td>Neuralynx </td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>soldering iron</td>
			<td>Radio Shack </td>
			<td>64-2802B</td>
			<td><a href="http://www.radioshack.com" target="_blank">www.radioshack.com</a></td>
		 </tr>
		 <tr>
			<td>nanoZ</td>
			<td>Neuralynx</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>small bit drill/driver</td>
			<td>Ram Products </td>
			<td>Rampower 35</td>
			<td>with footpedal controller, <a href="http://www.ramprodinc.com" target="_blank">www.ramprodinc.com</a></td>
		 </tr>
		 <tr>
			<td>drill bits</td>
			<td>Small Parts, Inc.</td>
			<td></td>
			<td>3/32" bits, <a href="http://www.smallpartsinc.com" target="_blank">www.smallpartsinc.com</a></td>
		 </tr>
		 <tr>
			<td>dissecting microscope</td>
			<td>Olympus </td>
			<td>SZ-60</td>
			<td><a href="http://www.olympusamerica.com" target="_blank">www.olympusamerica.com</a></td>
		 </tr>
		 <tr>
			<td>heat gun</td>
			<td>Alphawire</td>
			<td>Fit gun 3</td>
			<td>use setting "1" only, <a href="http://www.alphawire.com" target="_blank">www.alphawire.com</a></td>
		 </tr>
		 <tr>
			<td>26 AWG copper wire</td>
			<td>Arcor Electronics</td>
			<td>F26</td>
			<td>for ground wires, <a href="http://www.arcorelectronics.com" target="_blank"> www.arcorelectronics.com</a></td>
		 </tr>
		 <tr>
			<td>soldering flux</td>
			<td>Eagle</td>
			<td>2 oz, #205</td>
			<td></td>
		 </tr>
		 <tr>
			<td>0.02" diameter solder</td>
			<td>Kester</td>
			<td>24-6337-0010</td>
			<td><a href="http://www.kester.com" target="_blank"> www.kester.com</a></td>
		 </tr>
		 <tr>
			<td>benchtop vise</td>
			<td>Vacu-Vise</td>
			<td>Model 300</td>
			<td></td>
		 </tr>
		 <tr>
			<td>fiber optic light</td>
			<td>Nikon </td>
			<td>MKII</td>
			<td>dual light arms, <a href="http://www.nikon.com" target="_blank"> www.nikon.com</a></td>
		 </tr>
		 <tr>
			<td>5-min epoxy</td>
			<td>Allied Electronics</td>
			<td></td>
			<td>25 ml, <a href="http://www.alliedelec.com" target="_blank"> www.alliedelec.com</a></td>
		 </tr>
		 <tr>
			<td>fine tweezers</td>
			<td>Roboz Surgical Instrument Co.</td>
			<td>RS-4907, RS-5010</td>
			<td>INOX material, <a href="http://www.roboz.com" target="_blank"> www.roboz.com</a></td>
		 </tr>
		 <tr>
			<td>micro dissecting scissors</td>
			<td>Roboz Surgical Instrument Co.</td>
			<td>RS-5880</td>
			<td></td>
		 </tr>
		</tbody>
 </table>
 Table 1. Materials and reagents used for constructing tetrodes and microdrives.

construction of microdrive arrays for chronic neural recordings in awake behaving mice

State-of-the-art electrophysiological recordings from the brains of freely behaving animals allow researchers to simultaneously examine local field potentials (LFPs) from populations of neurons and action potentials from individual cells, as the animal engages in experimentally relevant tasks. Chronically implanted microdrives allow for brain recordings to last over periods of several weeks. Miniaturized drives and lightweight components allow for these long-term recordings to occur in small mammals, such as mice. By using tetrodes, which consist of tightly braided bundles of four electrodes in which each wire has a diameter of 12.5 &mu;m, it is possible to isolate physiologically active neurons in superficial brain regions such as the cerebral cortex, dorsal hippocampus, and subiculum, as well as deeper regions such as the striatum and the amygdala. Moreover, this technique insures stable, high-fidelity neural recordings as the animal is challenged with a variety of behavioral tasks. This manuscript describes several techniques that have been optimized to record from the mouse brain. First, we show how to fabricate tetrodes, load them into driveable tubes, and gold-plate their tips in order to reduce their impedance from M&Omega; to K&Omega; range. Second, we show how to construct a custom microdrive assembly for carrying and moving the tetrodes vertically, with the use of inexpensive materials. Third, we show the steps for assembling a commercially available microdrive (Neuralynx VersaDrive) that is designed to carry independently movable tetrodes. Finally, we present representative results of local field potentials and single-unit signals obtained in the dorsal subiculum of mice. These techniques can be easily modified to accommodate different types of electrode arrays and recording schemes in the mouse brain.

After implanting the microdrive and lowering the electrodes to the intended brain targets, an amplified data acquisition system, such as a Neuralynx Lynx-8, is needed for recording neural signals. Representative neural recordings of local field potentials (LFPs) and single-unit action potentials (often termed "spikes") from the mouse dorsal subiculum are shown in Figure 2. LFP signals were sampled at 3 kHz and band-pass filtered between 0.1-500 Hz (Figure 2A and 2B...

Watch this Scientific Journal Video about Construction of Microdrive Arrays for Chronic Neural Recordings in Awake Behaving Mice at JoVE.com

Construction of Microdrive Arrays for Chronic Neural Recordings in Awake Behaving Mice

The design and assembly of microdrives for in vivo electrophysiological recordings of brain signals from the mouse is described. By attaching microelectrode bundles to sturdy driveable carriers, these techniques allow for long-term and stable neural recordings. The lightweight design allows for unrestricted behavioral performance by the animal following drive implantation.

State-of-the-art electrophysiological recordings from the brains of freely behaving animals allow researchers to simultaneously examine local field ...

construction-microdrive-arrays-for-chronic-neural-recordings-awake

Research

JoVE Journal

Behavior

20.0K Views.  North Shore LIJ Health System. The design and assembly of microdrives for in vivo electrophysiological recordings of brain signals from the mouse is described. By attaching microelectrode bundles to sturdy driveable carriers, these techniques allow for long-term and stable neural recordings. The lightweight design allows for unrestricted behavioral performance by the animal following drive implantation.

Video: Construction of Microdrive Arrays for Chronic Neural Recordings in Awake Behaving Mice

Long-term Potentiation of Perforant Pathway-dentate Gyrus Synapse in Freely Behaving Mice

Studies of long-term potentiation of synaptic efficacy, an activity-dependent synaptic phenomenon having properties that make it attractive as a potential cellular mechanism underlying learning and information storage, have long been used to elucidate the physiology of various neuronal circuits in the hippocampus, amygdala, and other limbic and cortical structures. With this in mind, transgenic mouse models of neurological diseases represent useful platforms to conduct long-term&#160;potentiation (LTP) studies to develop a greater understanding of the role of genes in normal and abnormal synaptic communication in neuronal networks involved in learning, emotion and information processing. This article describes methodologies for reliably inducing LTP in the freely behaving mouse. These methodologies can be used in studies of transgenic and knockout freely&#160;behaving mouse models of neurodegenerative diseases.

Transgenic and knockout mouse models of neurological diseases are useful for studying the role of genes in normal and abnormal neurophysiology. This article describes methodologies which can be used to study long-term potentiation, a cellular mechanism which may underlie learning and memory, in transgenic and knockout freely&#160;behaving mouse models of neuropathology.

Studies of long-term potentiation of synaptic efficacy, an activity-dependent synaptic phenomenon having properties that make it attractive as a potential ...

Automated, Quantitative Cognitive/Behavioral Screening of Mice: For Genetics, Pharmacology, Animal Cognition and Undergraduate Instruction

We describe a high-throughput, high-volume, fully automated, live-in 24/7 behavioral testing system for assessing the effects of genetic and pharmacological manipulations on basic mechanisms of cognition and learning in mice. A standard polypropylene mouse housing tub is connected through an acrylic tube to a standard commercial mouse test box. The test box has 3 hoppers, 2 of which are connected to pellet feeders. All are internally illuminable with an LED and monitored for head entries by infrared (IR) beams. Mice live in the environment, which eliminates handling during screening. They obtain their food during two or more daily feeding periods by performing in operant (instrumental) and Pavlovian (classical) protocols, for which we have written protocol-control software and quasi-real-time data analysis and graphing software. The data analysis and graphing routines are written in a MATLAB-based language created to simplify greatly the analysis of large time-stamped behavioral and physiological event records and to preserve a full data trail from raw data through all intermediate analyses to the published graphs and statistics within a single data structure. The data-analysis code harvests the data several times a day and subjects it to statistical and graphical analyses, which are automatically stored in the &#34;cloud&#34; and on in-lab computers. Thus, the progress of individual mice is visualized and quantified daily. The data-analysis code talks to the protocol-control code, permitting the automated advance from protocol to protocol of individual subjects. The behavioral protocols implemented are matching, autoshaping, timed hopper-switching, risk assessment in timed hopper-switching, impulsivity measurement, and the circadian anticipation of food availability. Open-source protocol-control and data-analysis code makes the addition of new protocols simple. Eight test environments fit in a 48 in x 24 in x 78 in&#160;cabinet; two such cabinets (16 environments) may be controlled by one computer.

Fully automated system for measuring physiologically meaningful properties of the mechanisms mediating spatial localization, temporal localization, duration, rate&#160;and probability estimation, risk assessment, impulsivity, and the accuracy and precision of memory, in order to assess the effects of genetic and pharmacological manipulations on foundational mechanisms of cognition in mice.

We describe a high-throughput, high-volume, fully automated, live-in 24/7 behavioral testing system for assessing the effects of genetic and ...

Cortical Source Analysis of High-Density EEG Recordings in Children

EEG is traditionally described as a neuroimaging technique with high temporal and low spatial resolution. Recent advances in biophysical modelling and signal processing make it possible to exploit information from other imaging modalities like structural MRI that provide high spatial resolution to overcome this constraint1. This is especially useful for investigations that require high resolution in the temporal as well as spatial domain. In addition, due to the easy application and low cost of EEG recordings, EEG is often the method of choice when working with populations, such as young children, that do not tolerate functional MRI scans well. However, in order to investigate which neural substrates are involved, anatomical information from structural MRI is still needed. Most EEG analysis packages work with standard head models that are based on adult anatomy. The accuracy of these models when used for children is limited2, because the composition and spatial configuration of head tissues changes dramatically over development3.&#160;
In the present paper, we provide an overview of our recent work in utilizing head models based on individual structural MRI scans or age specific head models to reconstruct the cortical generators of high density EEG. This article describes how&#160;EEG recordings are acquired, processed, and analyzed with pediatric populations at the London Baby Lab, including laboratory setup, task design, EEG preprocessing, MRI processing, and EEG channel level and source analysis.&#160;

In recent years, there has been increasing interest in estimating the cortical sources of scalp measured electrical activity for cognitive neuroscience experiments. This article describes how high density EEG is acquired and how recordings are processed for cortical source estimation in children from the age of 2 years at the London Baby Lab.

EEG is traditionally described as a neuroimaging technique with high temporal and low spatial resolution. Recent advances in biophysical modelling and ...

A Novel Experimental and Analytical Approach to the Multimodal Neural Decoding of Intent During Social Interaction in Freely-behaving Human Infants

Understanding typical and atypical development remains one of the fundamental questions in developmental human neuroscience. Traditionally, experimental paradigms and analysis tools have been limited to constrained laboratory tasks and contexts due to technical limitations imposed by the available set of measuring and analysis techniques and the age of the subjects. These limitations severely limit the study of developmental neural dynamics and associated neural networks engaged in cognition, perception and action in infants performing &#8220;in action and in context&#8221;. This protocol presents a novel approach to study infants and young children as they freely organize their own behavior, and its consequences in a complex, partly unpredictable and highly dynamic environment. The proposed methodology integrates synchronized high-density active scalp electroencephalography (EEG), inertial measurement units (IMUs), video recording and behavioral analysis to capture brain activity and movement non-invasively in freely-behaving infants. This setup allows for the study of neural network dynamics in the developing brain, in action and context, as these networks are recruited during goal-oriented, exploration and social interaction tasks.

This protocol presents a novel methodology for the neural decoding of intent from freely-behaving infants during unscripted social interaction with an actor. Neural activity is acquired using non-invasive high-density active scalp electroencephalography (EEG). Kinematic data is collected with inertial measurement units and supplemented with synchronized video recording.

Understanding typical and atypical development remains one of the fundamental questions in developmental human neuroscience. Traditionally, experimental ...

A Pressure Injection System for Investigating the Neuropharmacology of Information Processing in Awake Behaving Macaque Monkey Cortex

The top-down modulation of feed-forward cortical information processing is functionally important for many cognitive processes, including the modulation of sensory information processing by attention. However, little is known about which neurotransmitter systems are involved in such modulations. A practical way to address this question is to combine single-cell recording with local and temporary neuropharmacological manipulation in a suitable animal model. Here we demonstrate a technique combining acute single-cell recordings with the injection of neuropharmacological agents in the direct vicinity of the recording electrode. The video shows the preparation of the pressure injection/recording system, including preparation of the substance to be injected. We show a rhesus monkey performing a visual attention task and the procedure of single-unit recording with block-wise pharmacological manipulations.

Here, we show the pressure injection of neuropharmacological substances during single-cell recording in an awake, behaving macaque monkey. This procedure allows pharmacological manipulation in the direct vicinity of a cortical recording site.

The top-down modulation of feed-forward cortical information processing is functionally important for many cognitive processes, including the modulation ...

Using Laser Doppler Imaging and Monitoring to Analyze Spinal Cord Microcirculation in Rat

Laser Doppler flowmetry (LDF) is a noninvasive method for blood flow (BF) measurement, which makes it preferable for measuring microcirculatory alterations of the spinal cord. In this article, our goal was to use both Laser Doppler imaging and monitoring to analyze the change of BF after spinal cord injury. Both the laser Doppler image scanner and the probe/monitor were being employed to obtain each readout. The data of LDPI provided a local distribution of BF, which gave an overview of perfusion around the injury site and made it accessible for comparative analysis of BF among different locations. By intensely measuring the probing area over a period of time, a combined probe was used to simultaneously measure the BF and oxygen saturation of the spinal cord, showing overall spinal cord perfusion and oxygen supply. LDF itself has a few limitations, such as relative flux, sensitivity to movement, and biological zero signal. However, the technology has been applied in clinical and experimental study due to its simple setup and rapid measurement of BF.

Here we present a combination of laser Doppler perfusion imaging (LDPI) and laser Doppler perfusion monitoring (LDPM) to measure spinal cord local blood flows and oxygen saturation (SO2), as well as a standardized procedure for introducing spinal cord trauma on rat.

Laser Doppler flowmetry (LDF) is a noninvasive method for blood flow (BF) measurement, which makes it preferable for measuring microcirculatory ...

A Wireless, Bidirectional Interface for In Vivo Recording and Stimulation of Neural Activity in Freely Behaving Rats

In vivo electrophysiology is a powerful technique to investigate the relationship between brain activity and behavior at a millisecond and micrometer scale. However, current methods mostly rely on tethered cable recordings or only use unidirectional systems, allowing either recording or stimulation of neural activity, but not at the same time or same target. Here, a new wireless, bidirectional device for simultaneous multichannel recording and stimulation of neural activity in freely behaving rats is described. The system operates through a single portable head stage that both transmits recorded activity and can be targeted in real-time for brain stimulation using a telemetry-based multichannel software. The head stage is equipped with a preamplifier and a rechargeable battery, allowing stable long-term recordings or stimulation for up to 1 h. Importantly, the head stage is compact, weighs 12 g (including battery) and thus has minimal impact on the animal&#180;s behavioral repertoire, making the method applicable to a broad set of behavioral tasks. Moreover, the method has the major advantage that the effect of brain stimulation on neural activity and behavior can be measured simultaneously, providing a tool to assess the causal relationships between specific brain activation patterns and behavior. This feature makes the method particularly valuable for the field of deep brain stimulation, allowing precise assessment, monitoring, and adjustment of stimulation parameters during long-term behavioral experiments. The applicability of the system has been validated using the inferior colliculus as a model structure.

A Wireless, Bidirectional Interface for In Vivo Recording and Stimulation of Neural Activity in Freely Behaving Rats

A wireless, bidirectional system for multi-channel neural recordings and stimulation in freely behaving rats is introduced. The system is light and compact, thus having minimal impact on the animal&#180;s behavioral repertoire. Moreover, this bidirectional system provides a sophisticated tool to assess causal relationships between brain activation patterns and behavior.

In vivo electrophysiology is a powerful technique to investigate the relationship between brain activity and behavior at a millisecond and micrometer ...

A Behavioral Assay for Investigating the Role of Spatial Memory During Instinctive Defense in Mice

Evolution has selected a repertoire of defensive behaviors that are essential for survival across all animal species. These behaviors are often stereotyped actions elicited in response to innately aversive sensory stimuli, but their success requires enough flexibility for adapting to different spatial environments, which can change rapidly. Here, we describe a behavioral assay to evaluate the influence of learned spatial knowledge on defensive behaviors in mice. We have adapted the widely used Barnes maze spatial memory assay to investigate how mice navigate to a shelter during escape responses to innately aversive sensory stimuli in a novel environment, and how they adapt to acute changes in the environment. This new assay is an ethological paradigm that does not require training and exploits the natural exploration patterns and navigation strategies in mice. We propose that the set of protocols described here are a powerful means of studying goal-directed behaviors and stimulus-triggered navigation, which should be of interest to both the fields of instinctive behaviors and spatial memory.

This protocol describes a behavioral assay, based on the Barnes maze test, to study how instinctive defensive actions are modified by the knowledge of spatial environment.

Evolution has selected a repertoire of defensive behaviors that are essential for survival across all animal species. These behaviors are often ...

The Unpredictable Chronic Mild Stress Protocol for Inducing Anhedonia in Mice

Depression is a highly prevalent and debilitating condition, only partially addressed by current pharmacotherapies. The lack of response to treatment by many patients prompts the need to develop new therapeutic alternatives and to better understand the etiology of the disorder. Pre-clinical models with translational merits are rudimentary for this task. Here we present a protocol for the unpredictable chronic mild stress (UCMS) method in mice. In this protocol, adolescent mice are chronically exposed to interchanging unpredictable mild stressors. Resembling the pathogenesis of depression in humans, stress exposure during the sensitive period of mice adolescence instigates a depressive-like phenotype evident in adulthood. UCMS can be used for screenings of antidepressants on the variety of depressive-like behaviors and neuromolecular indices. Among the more prominent tests to assess depressive-like behavior in rodents is the sucrose preference test (SPT), which reflects anhedonia (core symptom of depression). The SPT will also be presented in this protocol. The ability of UCMS to induce anhedonia, instigate long-term behavioral deficits and enable reversal of these deficits via chronic (but not acute) treatment with antidepressants strengthens the protocol&#39;s validity compared to other animal protocols for inducing depressive-like behaviors.

Here we present the unpredictable chronic mild stress protocol in mice. This protocol induces a long-term depressive-like phenotype and enables to assess the efficacy of putative antidepressants in reversing the behavioral and neuromolecular depressive-like deficits.

Depression is a highly prevalent and debilitating condition, only partially addressed by current pharmacotherapies. The lack of response to treatment by ...

Novel Object Recognition and Object Location Behavioral Testing in Mice on a Budget

Ethologically relevant behavioral testing is a critical component of any study that uses mouse models to study the cognitive effects of various physiological or pathological changes. The object location task (OLT) and the novel object recognition task (NORT) are two effective behavioral tasks commonly used to reveal the function and relative health of specific brain regions involved in memory. While both of these tests exploit the inherent preference of mice for the novelty to reveal memory for previously encountered objects, the OLT primarily evaluates spatial learning, which relies heavily on hippocampal activity. The NORT, in contrast, evaluates non-spatial learning of object identity, which relies on multiple brain regions. Both tasks require an open-field-testing arena, objects with equivalent intrinsic value to mice, appropriate environmental cues, and video recording equipment and the software. Commercially available systems, while convenient, can be costly. This manuscript details a simple, cost-effective method for building the arenas and setting up the equipment necessary to perform the OLT and NORT. Furthermore, the manuscript describes an efficient testing protocol that incorporates both OLT and NORT and provides typical methods for data acquisition and analysis, as well as representative results. Successful completion of these tests can provide valuable insight into the memory function of various mouse model systems and appraise the underlying neural regions that support these functions.

Here we provide a protocol which includes comprehensive instructions for the economical establishment of murine object location and novel object recognition behavioral testing, including the design, cost, and construction of required equipment as well as execution of behavioral testing, data collection, and analysis.