Marie Curie International Outgoing Fellowship (European Union's FP/2007-2013 Grant Agreements #221834 and 254780), J.D. McDonnell Foundation, NSF Grant SBE 0542013, National Institutes of Health Grant NS034994, National Institute of Mental Health Grant MH5467 and the Howard Hughes Medical Institute (Janelia Farm Research Campus grant).

1. Construction of the microdrive
All drives are made from the same basic elements: a moving part, which carries the electrode and a fixed part, which is anchored to the skull. An ideal microdrive allows smooth but long enough travel of the electrode in multiple small steps, is sturdy enough to prevent accidental movement of the electrode, easy to manipulate by the experimenter without interfering with the animal's behavior, small in size and light in weight. As a result of these competing requirements, different drives suite different applications.
Only 4 parts are needed to build our basic drive: a brass flat head screw, a matching nut, a plastic bridge prepared from a single row pin header and two custom-cut brass plates.
<ol>
 <li>Break a 3-pin piece from the header</li>
 <li>Gently pull out the middle pin.</li>
 <li>Enlarge the hole by drilling through it with a drill bit size #55.</li>
 <li>Cut a thread using the 00-90 tap.</li>
 <li>Cut two pieces out of the brass plate. </li>
 <li>File the edges of the plates with a Dremmel.</li>
 <li>Drill a hole in the middle of both pieces using a drill bit size #65. </li>
 <li>Assemble the drive pieces so that the brass plates are touching the pins. To accomplish this, insert the brass screw through, successively, the brass plate, the threaded pin header hole, the second brass plate, and the nut. Tighten the screw gently so that the assembly becomes stable.</li>
 <li>Solder the pin ends to the brass plates.</li>
 <li>File the protruding end of the screw.</li>
 <li>Solder the nut to the screw. Be careful not to solder the nut to the brass plate.</li>
 <li>Test the movement of the drive: turn the screw clock-wise to elevate the plastic bridge. </li>
</ol>
2. Preparing the silicon probe
Before fixing the probe to the drive, add extra insulation to the bonding area of the probe to prevent cerebrospinal fluid (CSF) or humidity from producing short-circuits:
<ol>
 <li>Weigh and mix the Sylgard Elastomer components in a 10:1 ratio.</li>
 <li>Using a sharpened cotton applicator, apply the Sylgard to the top end of the probe.</li>
 <li>Let it dry in an oven preheated at 60&deg;C for 2 hours.</li>
</ol>
To ensure that the recording sites are devoid of any debris, the probe tips need to be cleaned:
<ol start="4">
 <li>Prepare a 4 % dilution of Contrad detergent.</li>
 <li>Let the probe soak in the detergent at 63&deg;C for at least 2 hours.</li>
 <li>Rinse the detergent off by dipping the probe repeatedly in distilled water.</li>
</ol>
Before fixing the probe to the drive, the impedance of each recording site should be checked:
<ol start="7">
 <li>Dip the probe in 0.9% saline, and connect it to an impedance-meter. If too many recording sites have incorrect impedance, repeat steps 2.4-2.6 or consider using a different probe. Here we use an Impedance Conditioning Module from Frederick Haer, Co. (FHC), combined with a homemade channel selector. Alternatively, a niPOD by NeuroNexus, Inc. or NanoZ by Neuralynx, Inc. allows the monitoring of the impedance of all the probe channels simultaneously.</li>
</ol>
3. Affixing the probe to the microdrive 
<ol>
 <li>Using a razor blade, cut multiple grooves into the bridge to create a rugged surface.</li>
 <li>Appose the probe to the bridge of the drive. This procedure is best done under an operating microscope, by holding the drive with a clamp and adjusting the probe by a micromanipulator so that the shanks are perfectly parallel with the drive screw. This ensures that during advancement, the probe shanks move into brain tissue without 'cutting' through it. The exact depth of the probe tips relative to the base of the drive should be determined at this stage, taking into consideration the depth of the target structure from the surface of the skull.</li>
 <li>The probe is then fixed to the bridge with grip cement.</li>
 <li>Optional: for visualizing the probe track in the brain, DiI solution (1-2% diluted in ethanol) can be applied to the back of the probe at this stage.</li>
</ol>
4. Preparing the skull
Prior to surgery, the reference and ground electrodes, and the parts of the on-head Faraday cage are prepared:
<ol>
 <li>Cut two 2"-long pieces of copper wire, and solder one end of each insulated copper wire for about 1 mm.</li>
 <li>Using a needle, scrape the head of a 00-90, 1/8" stainless steel screw and solder one piece of copper wire to it. Soldering such stainless steel ground screw-electrodes requires an appropriate flux (e.g., N-3 All purpose flux from La-Co) and high soldering tip temperatures. Carefully prevent any solder from flowing into the groove of the screw. This will be used as the ground electrode. Repeat with another screw and copper wire to prepare the reference electrode.</li>
 <li>Cut trapeziodal pieces from copper mesh. These pieces will be assembled to protect the headstage.</li>
</ol>
Surgical instruments and preparation are the same as used in many small animal surgeries. The entire surgery is done under deep isoflurane anesthesia, using aseptic conditions, according to NIH approved guidelines. Please note that the (mock) surgery shown in this video is for demonstration purposes only. For appropriate visibility and filming purposes, several preparatory steps, surgical precautions and postoperative procedures are not shown/visible or discussed.
Prior to surgery, all components and supplies should be sterilized, following appropriate procedures (see Guidelines for Survival Rodent Surgery; http://oacu.od.nih.gov/ARAC/surguide.pdf). During surgery, a sterile field on the skull is prepared and isolated by sterile drapes. At the end the surgery, a broad spectrum antibiotics is applied locally and a long-acting pain killer is given intramuscularly (e.g., buprenorphine, [Buprenex] 0.05 mg/kg). In addition, painkiller (e.g., Ibuprofen) is provided in the drinking water at approximately 60 mg/kg/24 hrs for 5 days. For proper surgical and anesthesia procedures, consult appropriate sources12.
<ol start="4">
 <li>Install the animal in the stereotaxic apparatus, shave and clean the scalp13.</li>
 <li>Cut the skin along the midline and push aside the scalp. Remove the periosteum, clean and dry the skull. </li>
 <li>Measure the location and distance between bregma and lambda, and determine the x and y coordinates of the probe implantation site accordingly using a stereotaxic atlas14. Mark the site by scraping a cross on the skull with a scalpel.</li>
 <li>Drill the skull using a round head drill bit (size &frac14;) and drive support screws (stainless-steel, 000-120, 1/16") halfway into the bone, on different bone plates on the top and on the side of the skull. The screws will provide anchors to securely bond the headgear to the skull.</li>
 <li>Drill holes above the cerebellum and insert the ground and reference electrodes prepared in step 4.2. For recording local field potentials (LFP), the choice of the reference site is critical. This site is chosen because cerebellar LFP is the smallest of all cortical regions and muscle artifacts are minimal at this midline location.</li>
 <li>Apply dentin activator (Metabond kit) using a tiny brush over the entire surface of the skull. Rinse it with 0.9% saline.</li>
 <li>Apply dental cement (Metabond kit; follow manufacturer's instructions for mixing) on the skull, carefully covering anchor screws and ground and reference electrodes, but leaving the probe implantation site clear.</li>
 <li>Secure the four copper mesh flaps (prepared in step 4.3) to the skull. For this, cement the narrow base of each of them to the anterior, left, right, and posterior sides of the skull. The copper should never be in direct contact with the bone but always separated by a layer of cement.</li>
</ol>
5. Preparing the brain surface
<ol>
 <li>Using a round head drill bit, drill around the implantation site in multiple stages, while frequently irrigating the bone with saline. </li>
 <li>Carefully remove the bone flap and irrigate the brain surface.</li>
 <li>For inserting a multiple-shank probe, a large strip of dura is removed. Two tools are needed to remove the dura: a scalpel and a hook prepared from an insect needle (alternatively, a standard tungsten microelectrode). Bend the tip of the needle by pushing against a hard surface (e.g. glass microscope slide), and attach it to a handle (here, a piece of wooden Q-tip; alternatively a microdissecting needle holder).</li>
 <li>Lift the dura with the hook, and cut it with a scalpel. Special care is taken to avoid damaging the pia, vessels and the surface of the neocortex. Small bleeding can be resolved by saline irrigation. If major bleeding occurs or the neocortex is compromised in any way, one should consider terminating the surgery and preparing another animal. </li>
</ol>
6. Implanting the probe
At this stage, the density and orientation of cortical surface vessels are carefully evaluated. Stereotaxic coordinates should be adjusted, because the probe has to penetrate the brain in an area free from larger vessels.
For implantation, the drive assembly can be held with an alligator clip attached to the stereotaxic holder. Uninterrupted visibility of the brain surface and the tips of the probe are critical for successful penetration. 
<ol>
 <li>Slowly lower the probe down to approximately 1 mm above the intended target, while constantly irrigating the craniotomy with saline. For neocortical recording, the probe tips are lowered into the cortex approximately 0.5 mm and lifted back near the surface.</li>
 <li>Seal the craniotomy by applying a warm melted mixture of wax and paraffin oil through a needle (10-20g of wax in 10 mL paraffin oil, heated at 65&deg;C). Prior to application, cool the mix to 30&deg;C and test the density. It should be soft enough to allow easy probe movement). To facilitate complete coverage, the mixture may be melted in situ by approaching the hardened wax with the tip of a micro-cauterizer.</li>
 <li>Attach the bottom of the drive to the skull with grip cement, being careful to leave the nut free to turn. It is of utmost importance to avoid any accidental "bump" of the drive at this stage, otherwise the probe will damage the cortex. After the drive is secured to the skull, smooth movement of the probe should be verified.</li>
 <li>Cement the connector part of the probe to the skull.</li>
</ol>

7. Building the on-head Faraday cage
<ol>
 <li>Pull up and assemble the copper mesh flaps into a protective cylinder around the probe and drive. The cylinder also serves as an electrical shield against environmental noise and the slow wave artifacts produced by the charged whiskers in the behaving animal.</li>
 <li>Adjust the cylinder height by cutting away excess material so the copper mesh is level with the top of the probe connector.</li>
 <li>Solder the wires from the reference and ground screws to the appropriate pins of the connector. Also solder adjacent copper mesh flaps together to ensure their electrical continuity, and solder the ground wire to the copper mesh.</li>
 <li>Apply a layer of grip cement on the copper mesh to reinforce it and to prevent any direct contact between the metal and the animal&#x2019;s skin. Optionally, apply a layer of epoxy resin to further reinforce the headgear.</li>
 <li>Test the movement of the drive screw.</li>
 <li>Cover the top of the headgear with a piece cut from a rubber glove.</li>
</ol>
8. Recording in the freely moving animal
<ol>
 <li>After appropriate post-operative care, connect the animal to the recording system using a high impedance headstage and a lightweight, ultraflexible multi-strand cable. Counterbalance the weight of the headgear.</li>
 <li>Test the quality of recording each day in the homecage. The position of the recording sites is judged by both unit firing patterns and the shape of the local field potentials. Lower the probe gradually by turning the screw by small increments (typically 1/8 to 1/4 turn per day, i.e. 35-70 micrometers) until the target structure is reached.</li>
</ol>
9. Representative Results
Electrophysiological signals (local field potential and unit activity) vary depending on the recorded structure and the current behavior of the animal. Figure 1 shows examples of 32-channel CA1 hippocampal recordings while the rat is exploring an open field. Note the prominent 8 Hz (theta band) oscillation of the local field potential during exploration with superimposed spiking on multiple shanks and sites (examples of spikes indicated by arrowheads). To analyze neuronal unit activity, spikes are detected and sorted into single units using cluster analysis of their waveforms15-16.
<img src="/files/ftp_upload/3568/3568fig1.jpg" alt="Figure 1" /> 
Figure 1. CA1 Hippocampal recordings in the behaving rat using a 4 shanks x 8 sites silicon probe. Recordings are wideband and sampled at 20 000 Hz, which allows to study both local field potential oscillations (e.g. "theta" band 8 Hz rhythm) and neuronal spiking activity.
<img src="/files/ftp_upload/3568/3568table1.jpg" alt="Table 1" /> 
Table 1. Alternatives to reagents and equipment used. Please <a href="https://www.jove.com/files/ftp_upload/3568/3568table1_lg.jpg">click here</a> to see a larger version of this figure.

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No conflicts of interest declared.

This movie illustrates the implantation procedure of silicon probes for chronic large-scale recordings in the behaving rat. Critical steps to ensure quality recordings of neuronal activity arise from the fragility of both biological (brain tissue) and technical (silicon probe) materials. Special care should be taken while handling the probe to avoid any contact of shanks with any remotely "hard" surface (for example, the shanks would break if one tried to implant them in the brain without removing the dura). Similarly, a...

<table border="1">
	<tbody>
		<tr>
			<td>Name</td>
			<td>Type</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Silicon probe Buzsaki32, 4 shanks x 8 sites. Packaging: flexible polyamide cable</td>
			<td>Material</td>
			<td>NeuroNexus</td>
			<td>Probe: buzsaki32 Packaging: HC32</td>
			<td>Recording probe</td>
		</tr>
		<tr>
			<td>Round Brass Screw, 00-90 x 1/2 Round Brass Screws</td>
			<td>Material</td>
			<td>JIMorris</td>
			<td>R0090B500</td>
			<td>Drive part</td>
		</tr>
		<tr>
			<td>Brass Hex Nut, 00-90</td>
			<td>Material</td>
			<td>JIMorris</td>
			<td>N0090B</td>
			<td>Drive part</td>
		</tr>
		<tr>
			<td>Brass C260 Strip, ASTM-B36 
			Thickness: 0.025", Length: 12", Width: 1/2"</td>
			<td>Material</td>
			<td>Small Parts</td>
			<td>B000FMYU72</td>
			<td>Drive part</td>
		</tr>
		<tr>
			<td>Connector Header, pitch 2mm, male, single row, straigt, 36 positions</td>
			<td>Material</td>
			<td>Digikey</td>
			<td>2163S-36-ND</td>
			<td>Drive part</td>
		</tr>
		<tr>
			<td>2-part Sylgard silicon Elastomer</td>
			<td>Material</td>
			<td>World Precision Instruments</td>
			<td>SYLG184</td>
			<td>To extra-insulate the probe</td>
		</tr>
		<tr>
			<td>Decon Contrad 70 Liquid Detergent</td>
			<td>Reagent</td>
			<td>Fisher Scientific</td>
			<td>04-355 
			Decon Laboratories 
			No.:1002</td>
			<td>To clean the recording sites</td>
		</tr>
		<tr>
			<td>Impedance Conditioning Module</td>
			<td>Equipment</td>
			<td>FHC Inc.</td>
			<td>55-70-0</td>
			<td>Impedance meter</td>
		</tr>
		<tr>
			<td>niPOD &#x2013; 32 channels</td>
			<td>Equipment</td>
			<td>Neuronexus</td>
			<td>niPOD -32</td>
			<td>Impedance meter</td>
		</tr>
		<tr>
			<td>Grip Cement Industrial Grade</td>
			<td>Material</td>
			<td>Caulk Dentsply</td>
			<td>675571 (powder) 
			675572 (solvent)</td>
			<td>Grip cement</td>
		</tr>
		<tr>
			<td>1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate ('DiI'; DiIC18(3))</td>
			<td>Reagent</td>
			<td>Invitrogen</td>
			<td>D282</td>
			<td>To stain the probe track in the brain</td>
		</tr>
		<tr>
			<td>Stainless Steel Machine Screw, Binding Head, Slotted Drive, #00-90, 1/8" </td>
			<td>Material</td>
			<td>Small Parts</td>
			<td>MX-0090-02B</td>
			<td>Ground and reference screws</td>
		</tr>
		<tr>
			<td>Magnet wire, 20G, nylon-polyurethane coating, MW80</td>
			<td>Material</td>
			<td>Small Parts</td>
			<td>B000IJYRP2</td>
			<td>Ground and reference wire</td>
		</tr>
		<tr>
			<td>Stainless Steel Machine Screw, Binding Head Slotted Drive, #000-120, 1/16"</td>
			<td>Material</td>
			<td>Small Parts</td>
			<td>MX-000120-01B</td>
			<td>Anchor screws</td>
		</tr>
		<tr>
			<td>N-3 All purpose Flux Liquid</td>
			<td>Reagent</td>
			<td>La-Co (Markal)</td>
			<td>23512</td>
			<td>Allows to solder stainless-steel</td>
		</tr>
		<tr>
			<td>MicroGrid Precision Expanded Copper</td>
			<td>Material</td>
			<td>Dexmet</td>
			<td>3 CU6-050 FA</td>
			<td>Copper mesh for on-head Faraday cage</td>
		</tr>
		<tr>
			<td>C&amp;B-METABOND Quick! Cement System - Dentin Activator</td>
			<td>Material</td>
			<td>Parkell</td>
			<td>S380</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>C&amp;B-METABOND Quick! Cement System &#x2013; Dental cement</td>
			<td>Material</td>
			<td>Parkell</td>
			<td>S380</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Sharp point tungsten needle and holder</td>
			<td>Tool</td>
			<td>Roboz Surgical instruments</td>
			<td>RS-6064 and RS-6061</td>
			<td>To make the hook to lift the dura</td>
		</tr>
		<tr>
			<td>Carbide Bur HP 1/4</td>
			<td>Tool</td>
			<td>Henry Schein</td>
			<td>9990013</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Paraffin (Granules)</td>
			<td>Material</td>
			<td>Fisher Scientific</td>
			<td>P31-500</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Mineral Oil, Light (NF/FCC)</td>
			<td>Material</td>
			<td>Fisher Scientific</td>
			<td>O121-1</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>GC ELECTRONICS 10-114 2-Part Epoxy Adhesive</td>
			<td>Material</td>
			<td>Newark</td>
			<td>00Z416</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Type 1 LITZ 21 AWG 40/36 Red Single Polyurethane-Nylon (MW80-C) TO 0.041"+/-0.002" OD</td>
			<td>Material</td>
			<td>New England Wire Technologies Corporation</td>
			<td>N28-36E-400-2</td>
			<td>To make the cable between the headstage and the amplifier</td>
		</tr>
		<tr>
			<td>32-channel Very Large Scale Integration headstage, 20x gain</td>
			<td>Equipment</td>
			<td>Plexon</td>
			<td>HST/32V-G20</td>
			<td>Headstage</td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Large-scale Recording of Neurons by Movable Silicon Probes in Behaving Rodents at JoVE.com

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

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 ...

large-scale-recording-neurons-movable-silicon-probes-behaving

Research

JoVE Journal

Neuroscience

34.6K vues.  University of New Jersey. Nous décrivons les méthodes à grande échelle d'enregistrement de plusieurs unités simples et le potentiel de champ local à se comporter les rongeurs avec des sondes de silicium. Disque de fabrication, de fixation de sonde sur le disque et les processus d'implantation de la sonde sont illustrés dans des détails suffisants pour faciliter la reproduction.

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

Recording Large-scale Neuronal Ensembles with Silicon Probes in the Anesthetized Rat

Large scale electrophysiological recordings from neuronal ensembles offer the opportunity to investigate how the brain orchestrates the wide variety of behaviors from the spiking activity of its neurons. One of the most effective methods to monitor spiking activity from a large number of neurons in multiple local neuronal circuits simultaneously is by using silicon electrode arrays1-3.
Action potentials produce large transmembrane voltage changes in the vicinity of cell somata. These output signals can be measured by placing a conductor in close proximity of a neuron. If there are many active (spiking) neurons in the vicinity of the tip, the electrode records combined signal from all of them, where contribution of a single neuron is weighted by its 'electrical distance'. Silicon probes are ideal recording electrodes to monitor multiple neurons because of a large number of recording sites (+64) and a small volume. Furthermore, multiple sites can be arranged over a distance of millimeters, thus allowing for the simultaneous recordings of neuronal activity in the various cortical layers or in multiple cortical columns (Fig. 1). Importantly, the geometrically precise distribution of the recording sites also allows for the determination of the spatial relationship of the isolated single neurons4. Here, we describe an acute, large-scale neuronal recording from the left and right forelimb somatosensory cortex simultaneously in an anesthetized rat with silicon probes (Fig. 2).

Extracellular recordings of neuronal activity using silicon probes in the anesthetized rat will be described. This technique allows information to be obtained across multiple brain areas from more than 100 neurons simultaneously. It provides information with single cell resolution about neuronal ensembles dynamics in multiple local circuits.

Large scale electrophysiological recordings from neuronal ensembles offer the opportunity to investigate how the brain orchestrates the wide variety of ...

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 ...

Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

Recording Gamma Band Oscillations in Pedunculopontine Nucleus Neurons

Synaptic efferents from the PPN are known to modulate the neuronal activity of several intralaminar thalamic regions (e.g., the centrolateral/parafascicular; Cl/Pf nucleus). The activation of either the PPN or Cl/Pf nuclei in vivo has been described to induce the arousal of the animal and an increment in gamma band activity in the cortical electroencephalogram (EEG). The cellular mechanisms for the generation of gamma band oscillations in Reticular Activating System (RAS) neurons are the same as those found to generate gamma band oscillations in other brains nuclei. During current-clamp recordings of PPN neurons (from parasagittal slices from 9 - 25 day-old rats), the use of depolarizing square steps rapidly activated voltage-dependent potassium channels that prevented PPN neurons from being depolarized beyond -25 mV.
Injecting 1 - 2 sec long depolarizing current ramps gradually depolarized PPN membrane potential resting values towards 0 mV. However, injecting depolarizing square pulses generated gamma-band oscillations of membrane potential that showed to be smaller in amplitude compared to the oscillations generated by ramps. All experiments were performed in the presence of voltage-gated sodium channels and fast synaptic receptors blockers. It has been shown that the activation of high-threshold voltage-dependent calcium channels underlie gamma-band oscillatory activity in PPN neurons. Specific methodological and pharmacological interventions are described here, providing the necessary tools to induce and sustain PPN subthreshold gamma band oscillation in vitro.

The pedunculopontine nucleus (PPN) is located in the brainstem and its neurons are maximally activated during waking and rapid eye movement (REM) sleep brain states. This work describes the experimental approach to record in vitro gamma band subthreshold membrane oscillation in PPN neurons.

Synaptic efferents from the PPN are known to modulate the neuronal activity of several intralaminar thalamic regions (e.g., the ...

Large-scale Reconstructions and Independent, Unbiased Clustering Based on Morphological Metrics to Classify Neurons in Selective Populations

This protocol outlines large-scale reconstructions of neurons combined with the use of independent and unbiased clustering analyses to create a comprehensive survey of the morphological characteristics observed among a selective neuronal population. Combination of these techniques constitutes a novel approach for the collection and analysis of neuroanatomical data. Together, these techniques enable large-scale, and therefore more comprehensive, sampling of selective neuronal populations and establish unbiased quantitative methods for describing morphologically unique neuronal classes within a population.
The protocol outlines the use of modified rabies virus to selectively label neurons. G-deleted rabies virus acts like a retrograde tracer following stereotaxic injection into a target brain structure of interest and serves as a vehicle for the delivery and expression of EGFP in neurons. Large numbers of neurons are infected using this technique and express GFP throughout their dendrites, producing &#34;Golgi-like&#34; complete fills of individual neurons. Accordingly, the virus-mediated retrograde tracing method improves upon traditional dye-based retrograde tracing techniques by producing complete intracellular fills.
Individual well-isolated neurons spanning all regions of the brain area under study are selected for reconstruction in order to obtain a representative sample of neurons. The protocol outlines procedures to reconstruct cell bodies and complete dendritic arborization patterns of labeled neurons spanning multiple tissue sections. Morphological data, including positions of each neuron within the brain structure, are extracted for further analysis. Standard programming functions were utilized to perform independent cluster analyses and cluster evaluations based on morphological metrics. To verify the utility of these analyses, statistical evaluation of a cluster analysis performed on 160 neurons reconstructed in the thalamic reticular nucleus of the thalamus (TRN) of the macaque monkey was made. Both the original cluster analysis and the statistical evaluations performed here indicate that TRN neurons are separated into three subpopulations, each with unique morphological characteristics.

This protocol describes large-scale reconstructions of selective neuronal populations, labeled following retrograde infection with a modified rabies virus expressing fluorescent markers, and independent, unbiased cluster analyses that enable comprehensive characterization of morphological metrics among distinct neuronal subclasses.

This protocol outlines large-scale reconstructions of neurons combined with the use of independent and unbiased clustering analyses to create a ...

Quantification of Endosome and Lysosome Motilities in Cultured Neurons Using Fluorescent Probes

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, survival, and glial communications. To date, numerous studies have reported that defects in these systems cause various neuronal diseases. Thus, understanding the mechanisms underlying vesicle dynamics may provide influential clues that could aid in the treatment of several neuronal disorders. Here, we describe a method for quantifying vesicle motilities, such as motility distance and rate of movement, using a software plug-in for the ImageJ platform. To obtain images for quantification, we labeled neuronal endosome-lysosome structures with EGFP-tagged vesicle marker proteins and observed the movement of vesicles using a time-lapse microscopy. This method is highly useful and simplify measuring vesicle motility in neurites, such as axons and dendrites, as well as in the soma of both neurons and glial cells. Furthermore, this method can be applied to other cell lines, such as fibroblasts and endothelial cells. This approach could provide a valuable advancement of our understanding of membrane trafficking.

The investigation of membrane trafficking is crucial for understanding neuronal functions. Here, we introduce a method for quantifying vesicle motility in neurons. This is a convenient method that can be adapted to the quantification of membrane trafficking in the nervous system.

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, ...

Implantation of Chronic Silicon Probes and Recording of Hippocampal Place Cells in an Enriched Treadmill Apparatus

An important requisite for understanding brain function is the identification of behavior and cell activity correlates. Silicon probes are advanced electrodes for large-scale electrophysiological recording of neuronal activity, but the procedures for their chronic implantation are still underdeveloped. The activity of hippocampal place cells is known to correlate with an animal&#39;s position in the environment, but the underlying mechanisms are still unclear. To investigate place cells, here we describe a set of techniques which range from the fabrication of devices for chronic silicon probe implants to the monitoring of place field activity in a cue-enriched treadmill apparatus. A micro-drive and a hat are built by fitting and fastening together 3D-printed plastic parts. A silicon probe is mounted on the micro-drive, cleaned, and coated with dye. A first surgery is performed to fix the hat on the skull of a mouse. Small landmarks are fabricated and attached to the belt of a treadmill. The mouse is trained to run head-fixed on the treadmill. A second surgery is performed to implant the silicon probe in the hippocampus, following which broadband electrophysiological signals are recorded. Finally, the silicon probe is recovered and cleaned for reuse. The analysis of place cell activity in the treadmill reveals a diversity of place field mechanisms, outlining the benefit of the approach.

We describe the diverse steps to implant chronic silicon probes and to record place cells in mice that are running head-fixed on a cue-enriched treadmill apparatus.

An important requisite for understanding brain function is the identification of behavior and cell activity correlates. Silicon probes are advanced ...

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized animals. Highly sensitive, genetically encoded fluorescent reporters of Ca2+ have revolutionized the recording of cell and synaptic activity using non-invasive optical approaches in behaving animals. When combined with genetic and optogenetic techniques, the molecular mechanisms that modulate cell and circuit activity during different behavior states can be identified.
Here we describe methods for ratiometric Ca2+ imaging of single neurons in freely behaving Caenorhabditis elegans worms. We demonstrate a simple mounting technique that gently overlays worms growing on a standard Nematode Growth Media (NGM) agar block with a glass coverslip, permitting animals to be recorded at high-resolution during unrestricted movement and behavior. With this technique, we use the sensitive Ca2+ reporter GCaMP5 to record changes in intracellular Ca2+ in the serotonergic Hermaphrodite Specific Neurons (HSNs) as they drive egg-laying behavior. By co-expressing mCherry, a Ca2+-insensitive fluorescent protein, we can track the position of the HSN within ~ 1 &#181;m and correct for fluctuations in fluorescence caused by changes in focus or movement. Simultaneous, infrared brightfield imaging allows for behavior recording and animal tracking using a motorized stage. By integrating these microscopic techniques and data streams, we can record Ca2+ activity in the C. elegans egg-laying circuit as it progresses between inactive and active behavior states over tens of minutes.

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

This protocol describes the use of genetically encoded Ca2+ reporters to record changes in neural activity in behaving Caenorhabditis elegans worms. &#8195;

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized ...

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 ...

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic tectum, is a popular model to study how neural circuits self-assemble. The ability to carry out whole cell patch clamp recordings from tectal neurons and to record RGC-evoked responses, either in vivo or using a whole brain preparation, has generated a large body of high-resolution data about the mechanisms underlying normal, and abnormal, circuit formation and function. Here we describe how to perform the in vivo preparation, the original whole brain preparation, and a more recently developed horizontal brain slice preparation for obtaining whole cell patch clamp recordings from tectal neurons. Each preparation has unique experimental advantages. The in vivo preparation enables the recording of the direct response of tectal neurons to visual stimuli projected onto the eye. The whole brain preparation allows for the RGC axons to be activated in a highly controlled manner, and the horizontal brain slice preparation allows recording from across all layers of the tectum.

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

In this paper, we discuss three brain preparations used for whole cell patch clamp recording to study the retinotectal circuit of Xenopus laevis tadpoles. Each preparation, with its own specific advantages, contributes to the experimental tractability of the Xenopus tadpole as a model to study neural circuit function.

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic ...