The authors thank Nick Kathman for suggestions and help at preparing for the manuscript. This technique was developed in conjunction with work supported by the AFOSR under grant FA9550-10-1-0054 and the National Science Foundation under Grant No. IOS-1120305 to RER.

1. Preparation of Tetrode Wires
<ol>
	<li>Pull out a very thin nichrome wire (12 &#956;m diameter, PAC coating) of about 1.1 m length. Attach a tape tag to each end. Hang the wire over a horizontal threaded rod such that the two ends are at the same height near the benchtop.</li>
	<li>Repeat step 1.1 for a second wire, making two more ends for a total of 4, and place it next to the first wire (about 1 cm in between).</li>
	<li>Stick the four ends together with a tape tag and attach the tag to a motorized rotating winding device. This device can be made from an inexpensive dc motor.</li>
	<li>Wind the tetrode in one direction for 2 min (60 rpm) and unwind it in the opposite direction for 30 sec.</li>
	<li>Use a heat gun to fuse the wires together. Do not touch the wires with the gun. Use three up and down passes from alternating directions, with each pass taking about 10 sec.</li>
	<li>Cut the top and the bottom of the wound wires. The four wires are twisted and fused together at one end but separate at the other.</li>
	<li>Add the supporting tube. Cut a 30 cm length of polyethylene tubing (diameter: inside 0.28 mm, outside 0.61 mm). Thread the tetrode very slowly and carefully into the supporting tube so that it does not kink.</li>
	<li>Once the fused end appears out the other side, pull it through so that there is an equal length of wire at both ends of the guide tube.</li>
	<li>Grab the separate end of each wire with a forceps. Using the base of the flame of a gas burner, carefully burn the insulation off of the last 2 or 3 mm of each wire. Heat the wire until it glows, but does not curl.</li>
	<li>Connect the tetrode with a male-female IC socket adaptor that fits your recording device. Put the deinsulated end of each wire into a different socket of the adaptor with a forceps.&#160;Stabilize the wire in the socket with a small brass pin. Use a fine point soldering iron and fill the socket with the melted solder. Be careful to not contact the fragile wire with the soldering iron.</li>
	<li>Check the impedance of each wire and the inter impedance of each pair of wires.
	<ol>
		<li>Place the fused, twisted end into a container of saline and connect a copper wire conductor from the saline to the ohm meter.</li>
		<li>Connect the other end of the meter to the socket pin containing the wire. The impedance of each wire should be below 3 M&#8486;.&#160;</li>
		<li>If the above values are not attained, reattempt the solder connections.</li>
		<li>Remove the wires from the saline, rinse the tips with water, and test the inter wire impedance for each pairing (n=6). The inter impedance should be above 5 M&#8486;.</li>
		<li>If the above values are not attained, slice a small amount of the tip off at the fused end and retest.&#160;</li>
		<li>Discard any wire set that does not meet both of the impedance requirements for all of the wires.</li>
	</ol>
	</li>
	<li>Secure the tetrode.
	<ol>
		<li>Fold a small rectangular paper box slightly larger than the socket adaptor.</li>
		<li>Transfer the adaptor into the box with the male side at the bottom. Penetrate the box such that all the pins of the male side are outside the box while the rest of the adaptor is inside the box.</li>
		<li>Tape the corners of the box on the outside. Use small pieces of double sided sticky tape on the inside of the box to stabilize any individual strands of wire. The wire should be fused as it exits the box.</li>
		<li>Mix fast set 2 part epoxy and pour into the box to secure the adaptor and all the wires.</li>
		<li>Attach the near end of the guide tubing to one side of the box with dental wax but leave the tubing open such that the tetrode can be pulled through freely at both ends.</li>
	</ol>
	</li>
	<li>Sharpen the tetrode.
	<ol>
		<li>Before each experiment, cut the tip of the tetrode with a sharp scalpel blade, not scissors. This prevents crushing and splaying of the wire ends while providing a clean flat edge for the next step.
		<ol>
			<li>Use a small rotary tool mounted vertically with medium and fine grit sanding disks (these can be combined on one platform) to polish the tetrode and remove some tip insulation. Hold the bundle near its end with forceps. Tilt the wire set end to a 45&#176; angle relative to the sanding disk and gently touch it to the moderate speed spinning disk for about 1 or 2 sec each on the medium and then the fine grits. Repeat this three more times, axially rotating the bundle 90&#176; each time. It is critical that the direction of spin of the sanding disks is away from the shallow angle of the wire ends, otherwise separation of the wires may occur.</li>
			<li>The desired result transforms the bundle end from a straight edge to a pointed tip with small amounts of insulation removed from the end of each wire. Verify the point using a dissecting microscope before plating the tetrode. If any fraying occurs at the tip, recut and repolish.</li>
			<li>If impedance testing during the subsequent plating step shows extremely low inter wire values (less than 4 M&#8486;), it indicates too much material removed during the polishing step. Recut and repolish the tetrode.</li>
		</ol>
		</li>
		<li>Plate the tetrode. Put the tip of the tetrode into a saturated copper sulfate solution (85 ml water, 5 ml sulfuric acid, 50 g copper sulfate). Plate each wire with a current of 2.5 &#956;A with a stimulus isolator. Inject the current for 1 sec, pause for 1 sec and repeat this process 4x.</li>
		<li>Check the impedance of each wire and the interimpedance of each pair of wires. The impedance of each wire should be between 0.5-1 M&#8486; and the inter impedance should be above 4 M&#8486;.</li>
		<li>Mount the adaptor onto the headstage of a multichannel recording system.</li>
		<li>Attach a bent insect pin to a micromanipulator. Attach the tip of the tetrode to the insect pin with dental wax</li>
	</ol>
	</li>
</ol>
2. Animal Preparation
<ol>
	<li>Anesthetize the cockroach with ice.</li>
	<li>After the cockroach stops moving, restrain the cockroach vertically against a flat cork surface with large saddle pins that straddle the insect but do not penetrate any part of its body.</li>
	<li>Transfer the preparation into a plastic container and place ice around the animal to minimize blood flow and body movements.</li>
	<li>Position a plastic collar at the neck to support the head and place dental wax around the head to stabilize it.</li>
	<li>Cut a small window between the ocelli with a razor blade and remove the cuticle from the head.</li>
	<li>Remove connective tissues and fat with a forceps to expose the brain.</li>
	<li>Place some cockroach saline into the head capsule to cover the brain tissue.</li>
	<li>To desheath the brain, use a fine forceps to gently grab the sheath on top of the brain and use another fine forceps to tear the sheath apart in the wire implanted area.</li>
	<li>Open a small hole in the head capsule anterior to the brain with an insect pin. Insert a braid of three larger diameters (56 &#181;m) insulated copper wires into the hole to serve as a reference/ground electrode.</li>
	<li>Lower the tip of the tetrode to the brain surface with the micromanipulator and position it near the brain region of interest.</li>
	<li>Carefully place two small pieces of thin acetate sheet (2 mm x&#160;1 mm), slightly larger than the hole in the head capsule, anterior and posterior to the tetrode.</li>
	<li>Turn on the recording system.</li>
	<li>Slowly lower the tetrode 150-250 &#181;m below the brain surface depending on the recording quality.</li>
	<li>Turn off the recording system.</li>
	<li>Move the two pieces of acetate sheet as close to the tetrode as possible without touching it (Figure 1A).</li>
	<li>Heat a small spatula or flattened hypodermic needle and put it into dental wax such that there is liquid wax at the tip of the spatula. Carefully touch the far end of each piece of acetate sheet from the tetrode with the spatula so that liquid wax can flow onto each piece and seal the gap between it and the head cuticle.</li>
	<li>Repeat step 2.16.&#160;Drop a small amount of liquid wax onto the acetate sheet each time. Start the process far away from the tetrode and move gradually towards it. Eventually the tetrode will be anchored by dental wax. Avoid getting hot wax into the cavity and onto the brain.</li>
	<li>Use the same method as steps 2.16&#160;and 2.17&#160;to anchor the reference/ground electrode with wax.</li>
	<li>Heat the wax that attaches the tetrode to the micromanipulator to release the tetrode from it.</li>
	<li>Loop the tetrode into the dental wax on the head to provide a strain relief (Figure 1B).</li>
	<li>Cover the strain relief loop with dental wax (Figure 1C).</li>
	<li>Carefully remove the constraints and transfer the preparation onto a Petri dish. Restrain the preparation dorsal side up with large saddle pins.</li>
	<li>Attach a rod to the pronotum using a glue gun. This is a wooden stick which extends from the pronotum over the abdomen.</li>
	<li>Attach the tip of the tetrode tubing to the posterior end of the rod with dental wax.</li>
	<li>Anchor the tetrode and the reference/ground electrode to the anterior end of the rod with dental wax.</li>
	<li>Pull the tetrode from the socket end of the tubing as much as possible, but do not tug on it, in order to eliminate the chance that the animal may damage the portion of the tetrode outside the tubing&#160;(Figure 1D).</li>
	<li>Remove all the constraints. Attach the reference/ground electrode to the tetrode tubing with dental wax.</li>
	<li>Wait at least 60 min for the animal to recover from the ice anesthesia before any experiment.</li>
</ol>
3. Experimental Procedures
<ol>
	<li>Connect a PC with both the recording system and an LED light using a USB to serial port cable.</li>
	<li>Start neural recordings.</li>
	<li>Start video recordings at 20 frames per second for walking experiments using the Motmot image acquisition package16 or 120 fps for climbing experiments using a high speed camera.</li>
	<li>Place the cockroach into a 40 cm x&#160;40 cm Plexiglas arena for walking experiments or a 58 cm long, 5 cm wide, and 5 cm high arena for climbing experiments. The walking arena has a transparent barrier extending from the middle of the right wall to the center of the arena, above which the headstage is located. The barrier is used to prevent animals from walking in areas where the camera view is blocked by the headstage. The climbing arena has an acrylic block (either 1.2 cm or 1.8 cm high, and 5 cm wide) or a shelf located at a comparable height in the center.</li>
	<li>Generate a TTL pulse from the PC using a customized MATLAB command. (s=serial(&#39;COM4&#39;); fopen(s); s.RequestToSend=&#39;off&#39;/s.RequestToSend=&#39;on&#39;/; fclose(s); delete(s);). The TTL pulse generates a timestamp for the recording system and either turns on or turns off the LED light.&#160;</li>
	<li>Allow the cockroach to explore the arena until it stops moving for more than 30 seconds for walking experiments. Allow the cockroach to either climb over the block/shelf or tunnel through the shelf for climbing experiments.&#160;</li>
	<li>Stop video recordings.</li>
	<li>Stop neural recordings.</li>
	<li>Write down the timestamp generated by the TTL pulse.</li>
	<li>Remove the cockroach from the arena and wait at least 3 min.</li>
	<li>Repeat steps 3.2-3.10&#160;for the next trial.</li>
	<li>Once all recordings have been completed, pass 5 sec of 5 &#181;A DC current through one of the wire tips (anode) and the reference electrode (cathode) to deposit copper into the brain at the wire tip.</li>
</ol>
4. Offline Analysis
<ol>
	<li>Synchronize video and neural data by linking the frame where the LED light is switched and the timestamp recorded by the recording system at that moment.</li>
	<li>Mark wire tip locations. Use Timms intensification procedures to precipitate and observe the copper in 12 &#181;m serial sections17. Prominent deposits should be visible in 3-8 adjacent sections (about 18-48% of the length of the dorsal ventral plane of the area we record from) (Figure 2).</li>
	<li>Correlate specific electrical impulses to the activity of single neurons. Follow spike sorting procedures laid out in detail elsewhere10,14,18. Use the program KlustaKwik (version 1.5, author K. Harris, Rutgers University) to generate initial, automated clustering. Import them into the program MClust (version 3.5, authors A.D. Redish et al., University of Minnesota) for further refinement and analysis (Figure 3).</li>
	<li>Track the cockroach&#8217;s movements. For walking experiments, extract the position of the cockroach&#8217;s (visual) center of mass and its body orientation in each frame of the video recordings using the Caltech Multiple Fly Tracker (version 0.1.5.6; http://ctrax.sourceforge.net/) and the associated FixErrors toolbox for MATLAB19. For climbing experiments, extract the position of the block and the cockroach&#8217;s head and pronotum in each frame of the video using motion analysis software package.</li>
</ol>

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The authors declare no conflicts of interest.&#160;

While previous electrophysiological studies on the CC or other regions of the insect brain have provided us with insights into the central control of behavior, most of them were performed in either restrained preparations9,11 or tethered ones10,14. As a result, the animal&#8217;s sensory experience and physiological state could be very different from those in a natural setting. Furthermore, the behavioral tasks that the animal can perform are limited to one plane under those situations. Here we pres...

This article describes a successful system for recording from neurons within the central complex (CC) of the cockroach, Blaberus discoidalis, &#160;as the insect walks in an arena and deals with objects that cause it to turn around, tunnel under, or climb over obstacles. The wires can also be connected to a stimulator to evoke activity in the surrounding neuropil with consequent behavioral changes.

Over the last decade considerable attention has been directed at the roles played by various brain regions in controlling insect behavior. Much of this focus has been directed toward the midline brain neuropils that are collectively referred to as the central complex (CC). Progress has been made as a result of wide varieties of techniques targeting questions about the role of the CC in behavior. Those techniques range from neurogenetic manipulations, primarily in Drosophila, coupled with behavioral analysis1-3, to electrophysiological techniques that monitor neural activity within the CC and attempt to relate that activity to behaviorally relevant parameters.&#160; &#160;

Electrophysiological&#160; techniques include intracellular recording from individual identified neurons4-9 and extracellular recording, often with multi channel probes10,11. These two techniques are complimentary. Intracellular recording with sharp electrodes or whole cell patch provides very detailed data on identified neurons, but is limited to one or two cells at once, requires limited or no movement, and can be maintained for relatively short periods of time. Extracellular recordings can be easily set up, do not require restraint, and can be maintained for hours. With multi channel tetrodes and cluster cutting, fairly large populations of neurons can be analyzed simultaneously9,12. While whole cell patch has been successfully used in tethered insects13, we feel that there is also a need for techniques that allow us to record neural activity in the brain for long periods of time in freely behaving insects as they deal with barriers to forward movement.

The need to record as the insect moves and bounces up and down pushed us toward extracellular recording methods. We have had good success recording in restrained preparations with commercially available 16 channels silicon probes11, however the small size of even large cockroaches means that the probes have to be mounted off the body. That, coupled with the delicacy of the probe tines, made them inappropriate for a free walking preparation. In two previous projects, we used bundles of fine wires forming a tetrode to accomplish similar recording properties but in a more robust arrangement. These tetrode bundles allowed us to record from tethered cockroaches and relate CC unit activity to changes in walking speed14 and turning behavior resulting from antennal contact with a rod10.

As useful as these tethered preparations have been and will continue to be, they do present some limitations. First, the behaviors that the insect can perform are limited to one plane. That is, we could readily evoke changes in walking speed or turning, but climbing and tunneling actions were not possible, at least with the typical tether arrangement. Second, our tethered preparations are &#8220;open loop&#8221;. That is, they do not allow for normal movement related feedback to the system. Thus, as the cockroach turned on our tether, its visual world was not altered accordingly. It is possible to build closed loop tether systems to introduce this kind of feedback. However, they are limited by the complexity of the programming and hardware of the simulated visual environment. Nevertheless, we felt that we could improve upon our existing tethered recording methods by recording from the animal as it walked freely in an arena or track and encountered objects as it would in its natural surroundings.

Although wireless systems for recording brain activity15 would be ideal, current systems have limitations in the number of recording channels, time of data acquisition, battery life and weight. We, therefore, opted to try to adapt our tethered recording system for use in freely moving preparations. As better wireless systems become available, this technique can be readily adapted to such devices. The system that is described in this article is light weight, works very well and appears to have little deleterious effect on the cockroach&#8217;s behavior. With an inexpensive high speed camera and cluster cutting software, activity in individual brain neurons can be related to movement. Here we describe the preparation of the tetrode wires and their implantation into the insect&#8217;s brain as well as recording techniques for electrical activity and motion and how those data can be brought together for subsequent analysis.

<table border="0" cellpadding="0" cellspacing="0" width="728">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:207px;">Nichrome wire&#160;</td>
			<td style="width:179px;">Sandvik Heating Technology</td>
			<td style="width:137px;">Kanthal RO-800</td>
			<td style="width:205px;">Use for tetrode</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Biomedical polyethylene tubing</td>
			<td>A-M Systems</td>
			<td>800700</td>
			<td>Use for tetrode tubing</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Lynx-8</td>
			<td>Neuralynx</td>
			<td></td>
			<td>Use for multi-unit recording</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cheetah 32</td>
			<td>Neuralynx</td>
			<td></td>
			<td>Use for multi-unit recording</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">High speed camera</td>
			<td>Basler</td>
			<td>A602f</td>
			<td>Use fir video recording for walking experiments</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">High speed camera</td>
			<td>Casio</td>
			<td>EX-FC150&#160;</td>
			<td>Use for video recording for climbing experiments</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">WINanalyze</td>
			<td>Winanalyze</td>
			<td>version 1.4 3D</td>
			<td>Use for video tracking&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Matlab</td>
			<td>MathWorks</td>
			<td>MATLAB R2012b</td>
			<td>Use for TTL pulse generation and off-line data analysis</td>
		</tr>
	</tbody>
</table>

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 recorded the neural activity of 50 units from the CC in 27 preparations for walking experiments. For 15 of those preparations (23 units), climbing experiments were also performed. Individual units are named according to preparation and unit numbers (e.g. &#699;unit 1-2&#700; indicates preparation 1, unit 2).
Snapshots of the video of one climbing trial are shown in Figure 4. The entire video is available in supplemental Video 1 (The sound is from u...

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Extracellular Wire Tetrode Recording in Brain of Freely Walking 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 ...

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Research

JoVE Journal

Neuroscience

16.4K Views.  Case Western Reserve University. 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.

Video: Extracellular Wire Tetrode Recording in Brain of Freely Walking Insects

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

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

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

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

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

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.

Recording Single Neurons&#039; Action Potentials from Freely Moving Pigeons Across Three Stages of 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 ...

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 ...

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

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

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

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

Construction of an Improved Multi-Tetrode Hyperdrive for Large-Scale Neural Recording in Behaving Rats

Monitoring the activity patterns of a large population of neurons over many days in awake animals is a valuable technique in the field of systems neuroscience. One key component of this technique consists of the precise placement of multiple electrodes into desired brain regions and the maintenance of their stability. Here, we describe a protocol for the construction of a 3D-printable hyperdrive, which includes eighteen independently adjustable tetrodes, and is specifically designed for in vivo extracellular neural recording in freely behaving rats. The tetrodes attached to the microdrives can either be individually advanced into multiple brain regions along the track, or can be used to place an array of electrodes into a smaller area. The multiple tetrodes allow for simultaneous examination of action potentials from dozens of individual neurons, as well as local field potentials from populations of neurons in the brain during active behavior. In addition, the design provides for simpler 3D drafting software that can easily be modified for differing experimental needs.

We present the construction of a 3D-printable hyperdrive with eighteen independently adjustable tetrodes. The hyperdrive is designed to record brain activity in freely behaving rats over a period of several weeks.

Monitoring the activity patterns of a large population of neurons over many days in awake animals is a valuable technique in the field of systems ...

Hybrid Microdrive System with Recoverable Opto-Silicon Probe and Tetrode for Dual-Site High Density Recording in Freely Moving Mice

Multi-regional neural recordings can provide crucial information to understanding fine-timescale interactions between multiple brain regions. However, conventional microdrive designs often only allow use of one type of electrode to record from single or multiple regions, limiting the yield of single-unit or depth profile recordings. It also often limits the ability to combine electrode recordings with optogenetic tools to target pathway and/or cell type specific activity. Presented here is a hybrid microdrive array for freely moving mice to optimize yield and a description of its fabrication and reuse of the microdrive array. The current design employs nine tetrodes and one opto-silicon probe implanted in two different brain areas simultaneously in freely moving mice. The tetrodes and the opto-silicon probe are independently adjustable along the dorsoventral axis in the brain to maximize the yield of unit and oscillatory activities. This microdrive array also incorporates a set-up for light, mediating optogenetic manipulation to investigate the regional- or cell type-specific responses and functions of long-range neural circuits. In addition, the opto-silicon probe can be safely recovered and reused after each experiment. Because the microdrive array consists of 3D-printed parts, the design of microdrives can be easily modified to accommodate various settings. First described is the design of the microdrive array and how to attach the optical fiber to a silicon probe for optogenetics experiments, followed by fabrication of the tetrode bundle and implantation of the array into a mouse brain. The recording of local field potentials and unit spiking combined with optogenetic stimulation also demonstrate feasibility of the microdrive array system in freely moving mice.

This protocol describes the construction of a hybrid microdrive array that allows implantation of nine independently adjustable tetrodes and one adjustable opto-silicon probe in two brain regions in freely moving mice. Also demonstrated is a method for safely recovering and reusing the opto-silicon probe for multiple purposes.

Multi-regional neural recordings can provide crucial information to understanding fine-timescale interactions between multiple brain regions. However, ...

Multi-Fiber Photometry to Record Neural Activity in Freely-Moving Animals

Recording the activity of a group of neurons in a freely-moving animal is a challenging undertaking. Moreover, as the brain is dissected into smaller and smaller functional subgroups, it becomes paramount to record from projections and/or genetically-defined subpopulations of neurons. Fiber photometry is an accessible and powerful approach that can overcome these challenges. By combining optical and genetic methodologies, neural activity can be measured in deep brain structures by expressing genetically-encoded calcium indicators, which translate neural activity into an optical signal that can be easily measured. The current protocol details the components of a multi-fiber photometry system, how to access deep brain structures to deliver and collect light, a method to account for motion artifacts, and how to process and analyze fluorescent signals. The protocol details experimental considerations when performing single and dual color imaging, from either single or multiple implanted optic fibers.

This protocol details how to implement and perform multi-fiber photometry recordings, how to correct for calcium-independent artifacts, and important considerations for dual-color photometry imaging.

Recording the activity of a group of neurons in a freely-moving animal is a challenging undertaking. Moreover, as the brain is dissected into smaller and ...

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

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

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

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

Recording Brain Activity with Ear-Electroencephalography

The c-grid (ear-electroencephalography, sold under the name cEEGrid) is an unobtrusive and comfortable electrode array that can be used for investigating brain activity after affixing around the ear. The c-grid is suitable for use outside of the laboratory for long durations, even for the whole day. Various cognitive processes can be studied using these grids, as shown by previous research, including research beyond the lab. To record high-quality ear-EEG data, careful preparation is necessary. In this protocol, we explain the steps needed for its successful implementation. First, how to test the functionality of the grid prior to a recording is shown. Second, a description is provided on how to prepare the participant and how to fit the c-grid, which is the most important step for recording high-quality data. Third, an outline is provided on how to connect the grids to an amplifier and how to check the signal quality. In this protocol, we list best practice recommendations and tips that make c-grid recordings successful. If researchers follow this protocol, they are comprehensively equipped for experimenting with the c-grid both in and beyond the lab.

Presented here is the procedure for using the c-grid (ear-electroencephalography, sold under the name cEEGrid) for recording brain activity in and beyond the lab for extended durations. This protocol describes how to set up these arrays and how to record brain activity using them.

The c-grid (ear-electroencephalography, sold under the name cEEGrid) is an unobtrusive and comfortable electrode array that can be used for investigating ...