recording brain electrical activity in behaving mice using multi-shank linear silicon probes

Recording Brain Electrical Activity in Behaving Mice Using Multi-Shank Linear Silicon Probes

Mount the electrode insertion tool on a stereotaxic holder, and zero the probe on bregma. Move the probe to the stereotaxic coordinates over the craniotomy, and slowly penetrate the brain surface. Make sure the probe shafts do not bend and avoid implanting through blood vessels. Then, slowly lower the probe until about 200 micrometers above the desired depth. Next, cover the craniotomy and shanks of the silicone probe with sterilized vaseline for protection.
Subsequently, apply dental cement to fix the base of the probe to the anchoring screw in the skull. Right after cement application, slowly move the probe to the target depth. Advance the last 200 micrometers after applying the cement to reduce lateral movement of the probe and ensure minimal tissue damage in the target area.
After the cement has cured, release the probe from the insertion tool by melting the wax with a cauterizer. Then, release the connector board from the insertion device and position it at a suitable place on the skull using a crocodile clamp. Fix the connector board to the skull using dental cement.
In the case of probe implantation in the hippocampus, place the connector board on the contralateral parietal bone. Then, solder the ground and reference wires of the connector board to the wires attached to the two screws over the cerebellum. Trim the protective cover to the correct height and place it over the silicone probe.
Afterward, fix the cover of the connector board and skull using dental cement, avoiding the skin around the exposed skull. After the surgery, apply appropriate analgesic treatment to the mouse for at least two days. Single-house the mouse to prevent damage to the implant.

recording-brain-electrical-activity-behaving-mice-using-multi-shank

Universidad de Cartagena UDEC

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JoVE Encyclopedia of Experiments

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Electrophysiology Techniques

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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Preparations
<ol>
	<li>Design and build an appropriate insertion tool transiently carrying the silicon probe and the electrode connector during the process of implantation. See Figure 1 for an example custom built insertion tool.</li>
	<li>Carefully release the silicon probe and electrode connector from its packaging using ceramic-tipped forceps.</li>
	<li>Lift the connector board and securely fix it with a crocodile clamp attached to a stand.</li>
	<li>Using a stereoscope, align the probe with the insertion tool with ceramic-tipped forceps. Apply a ~2 mm layer of paraffin wax melted with a cauterizer to glue the probe to the insertion tool. Take care not to touch the probe shanks during this procedure.</li>
	<li>Fix the electrode connector to the shaft of the insertion tool using standard adhesive tape. Note that depending on the manufacturer, ground wires might need to be soldered to the electrode connector board prior to implantation. Remove the insulation from two short pieces of varnish-insulated copper wire using tin-solder applied with a soldering iron (400 &#176;C). Solder the ground wires to the appropriate slots in the electrode connector board.</li>
	<li>Remove the insulation of two additional pieces of copper wire. Wrap each bare copper wire three times around a stainless steel screw (1 mm diameter, 2 mm length). Apply flux suitable for soldering steel and solder the copper wire to the bottom of the screw cap. Make sure that the bottom half of the screw thread remains free of tin-solder.</li>
	<li>Use a standard multimeter to check for electrical contact between wire and screw.</li>
	<li>Disinfect the shanks of the silicon probe and the ground screws by immersion in 70% ethanol (10 s).</li>
	<li>Prepare a protective cover for the probe implant by cutting the head of a plastic Pasteur pipette in half.</li>
</ol>
2. Implantation Surgery
<ol>
	<li>Sterilize surgical instruments (scissors, fine-tipped forceps, surgical clamps) with a hot bead sterilizer. Wipe all surfaces with 70% ethanol.</li>
	<li>Induce anesthesia with 3% isoflurane in oxygen delivered at ~1 L/min.
	<ol>
		<li>For maintenance, use 1 - 1.5% isoflurane. Note that the isoflurane concentration required to achieve surgical tolerance can vary from animal to animal.</li>
		<li>Stable surgical tolerance is achieved when the animal fails to respond to toe-pinching. Monitor the breathing rate of the mouse and adjust the concentration of isoflurane if necessary.</li>
		<li>Apply ointment to the animal&#39;s eyes to prevent drying out.</li>
	</ol>
	</li>
	<li>Mount the mouse in a stereotaxic frame by gently inserting ear bars into the ear canal. Once the head of the mouse is stabilized by the ear bars, place a mouth piece over the snout for continuous isoflurane delivery. Place the mouse on towel or pad over a heating pad and inject buprenorphine subcutaneously (0.05 - 0.1 mg/kg body weight) to ensure postoperative analgesia.</li>
	<li>Shave the head with a standard shaver and disinfect the skin with 70% ethanol. Using surgical scissors, make an incision into the skin along the midline of the skull and open the skin using surgical clamps.</li>
	<li>Align the head of the animal with the aid of a stereotaxic alignment tool to level bregma and lambda. There should be less than 50 &#181;m of height offset between bregma and lambda. Furthermore, level the head along the mediolateral axis by measuring the depth from bregma at the skull surface at defined distances left and right (e.g., 1 mm left and right of bregma). Adjust the tilt of the head if necessary.</li>
	<li>Clean the head with 3% hydrogen peroxide and wipe dry with sterile cotton wipes.</li>
	<li>Determine the location of the craniotomy relative to bregma using an appropriate stereotaxic atlas.</li>
	<li>Using a 0.9 mm drill head, drill two screw holes in the bone over the cerebellum to place ground and reference screws. Additionally, 1 - 3 holes for anchoring screws are desirable to stabilize the implant. The location of anchor screws will depend on the location of the craniotomy. For implantation into the hippocampus, place anchor screws over the contralateral parietal and ipsilateral frontal cortex. Insert the screws in the bone using a suitable screwdriver. Take care not to penetrate into the brain.</li>
	<li>Perform the craniotomy by slowly thinning the skull with the drill in a rectangular area around the implantation side. Frequently moisten the bone with sterilized phosphate buffer (PB). The remaining thinned skull can be gently pierced and removed with the aid of a fine (27G) injection needle and a pair of tweezers.</li>
	<li>Carefully pierce the dura mater with a thin (27G) injection needle. Form a small hook by bending the tip of the needle with a pair of tweezers and pull the dura for removal. Apply PB to prevent the brain surface from drying out.</li>
	<li>Mount the electrode insertion tool on a stereotaxic holder, zero the probe on bregma, and move the probe to the stereotaxic coordinates over the craniotomy. Slowly penetrate the brain surface. Make sure the probe shafts do not bend. Avoid implanting through blood vessels.</li>
	<li>Slowly lower the probe until ~200 &#181;m above the desired depth. Cover the craniotomy and shanks of the silicon probe with sterilized Vaseline for protection. Apply dental cement to fix the base of the probe to the anchoring screw in the skull.</li>
	<li>Right after cement application, slowly move the probe to the target depth. Advancing the last ~200 &#181;m after application of the cement reduces lateral movement of the probe and ensures minimal tissue damage in the target area. Note that the curing time of the cement used can affect this step of the protocol. With quickly curing cement, omit this step and directly implant the probe to the target depth in order to avoid damage to the silicon probe.</li>
	<li>After the cement has cured, release the probe from the insertion tool by melting the wax with a cauterizer.</li>
	<li>Release the connector board from the insertion device and position it at a suitable place on the skull using a crocodile clamp attached to the insertion handle. In case of probe implantation into the hippocampus, place the connector board on the contralateral parietal bone. Fix the connector board to the skull using dental cement.</li>
	<li>Solder the ground and reference wires of the connector board to the wires attached to the two screws over the cerebellum.</li>
	<li>Trim the protective cover to the correct height and place it over the silicon probe. Fix the cover to the connector board and skull using dental cement, avoiding the skin around the exposed skull. Suturing the skin around the implantation site is usually not required.</li>
</ol>
3. Recovery After Surgery
<ol>
	<li>Apply appropriate analgesic treatment for at least 2 days (e.g. subcutaneous injections of buprenorphine every 6 h during daytime and in the drinking water overnight combined with carprofen (4 - 5 mg/kg body weight) subcutaneously every 24 h). Single housing is recommended to prevent damage to the implant.</li>
	<li>Allow at least one week for recovery. Consult with local animal welfare guidelines.</li>
</ol>
4. Data Acquisition
<ol>
	<li>Record LFPs from freely moving mice using a suitable data acquisition system connected through a commutator. To acquire LFPs, use a sampling frequency of 1 - 5 kHz. Higher sampling rates (20 - 30 kHz) are required if single-unit discharges are to be recorded along with the LFP.</li>
	<li>Store raw recording files of the individual channels for offline analysis.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Crocodile clamp with stand</td>
			<td>Reichelt Elektronik</td>
			<td>HALTER ZD-10D</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicon probe</td>
			<td>Cambridge Neurotech</td>
			<td>P-series 32</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereoscope</td>
			<td>Olympus</td>
			<td>SZ51</td>
			<td></td>
		</tr>
		<tr>
			<td>Varnish-insulated copper wire</td>
			<td>B&#252;rklin Elektronik</td>
			<td>89 F 232</td>
			<td></td>
		</tr>
		<tr>
			<td>Ground screws</td>
			<td>Screws &#38; More GmbH (screwsandmore.de)</td>
			<td>DIN 84 A2 M1x2</td>
			<td></td>
		</tr>
		<tr>
			<td>Flux</td>
			<td>Stannol</td>
			<td>114018</td>
			<td></td>
		</tr>
		<tr>
			<td>Ceramic-tipped forceps</td>
			<td>Fine Science Tools</td>
			<td>11210-60</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraffine Wax</td>
			<td>Sigma-Aldrich</td>
			<td>327204</td>
			<td></td>
		</tr>
		<tr>
			<td>Cauterizer</td>
			<td>Fine Science Tools</td>
			<td>18010-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Soldering iron</td>
			<td>Kurtz Ersa</td>
			<td>OIC1300</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Carl Roth</td>
			<td>9065.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Pasteur pipettes</td>
			<td>Carl Roth</td>
			<td>EA65.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Heat sterilizer</td>
			<td>Fine Science Tools</td>
			<td>18000-45</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic frame</td>
			<td>David Kopf</td>
			<td>Model 1900</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic electrode holder</td>
			<td>David Kopf</td>
			<td>Model 1900</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Abbvie</td>
			<td>B506</td>
			<td></td>
		</tr>
		<tr>
			<td>Oxygen concentrator</td>
			<td>Respironix</td>
			<td>1020007</td>
			<td></td>
		</tr>
		<tr>
			<td>Buprenorphine</td>
			<td>Indivior UK Limited</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Electrical shaver</td>
			<td>Tondeo</td>
			<td>Eco-XS</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating pad</td>
			<td>Thermolux</td>
			<td>463265/-67</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical clamps</td>
			<td>Fine Science Tools</td>
			<td>18050-28</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrogen peroxide</td>
			<td>Sigma-Aldrich</td>
			<td>H1009</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile cotton wipes</td>
			<td>Carl Roth</td>
			<td>EH12.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Drill</td>
			<td>Proxxon</td>
			<td>Micromot 230/E</td>
			<td></td>
		</tr>
		<tr>
			<td>21G injection needle</td>
			<td>B. Braun</td>
			<td>4657527</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffer/phosphate buffered saline</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic atlas</td>
			<td>Elsevier</td>
			<td>9.78E+12</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical scissors</td>
			<td>Fine Science Tools</td>
			<td>14094-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical forceps</td>
			<td>Fine Science Tools</td>
			<td>11272-40</td>
			<td></td>
		</tr>
		<tr>
			<td>27G injection needles</td>
			<td>B. Braun</td>
			<td>4657705</td>
			<td></td>
		</tr>
		<tr>
			<td>Vaseline</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dental cement</td>
			<td>Sun Medical</td>
			<td>SuperBond T&#38;M</td>
			<td></td>
		</tr>
		<tr>
			<td>Carprofen</td>
			<td>Zoetis</td>
			<td>Rimadyl 50mg/ml</td>
			<td></td>
		</tr>
		<tr>
			<td>Recording amplifier&#160;</td>
			<td>Intan Technologies&#160;</td>
			<td>C3323</td>
			<td></td>
		</tr>
		<tr>
			<td>USB acquisition board&#160;</td>
			<td>Intan Technologies&#160;</td>
			<td>C3004</td>
			<td></td>
		</tr>
		<tr>
			<td>Recording cables</td>
			<td>Intan Technologies&#160;</td>
			<td>C3216</td>
			<td></td>
		</tr>
		<tr>
			<td>Electrical commutator&#160;</td>
			<td>Doric lenses&#160;</td>
			<td>HRJ-OE_FC_12_HARW</td>
			<td></td>
		</tr>
	</tbody>
</table>

Source: Sauer, J., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/57714">Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice.</a> J. Vis. Exp. (2018).
This video outlines a protocol for implanting a multi-shank linear silicon probe into the mouse brain to measure local field potentials. The procedure involves securing the probe to a stereotaxic unit, positioning it accurately over the target brain region, inserting it into the brain, and securing it with dental cement. After implantation, the assembly is protected with a casing. Following recovery, the setup allows for recording local field potentials, providing insights into neuronal activity.

<img alt="Figure 1" src="/files/ftp_upload/22754/22754_Figure1.jpg" /> 
Figure 1: Image of the insertion tool. The silicon probe is attached to an insect pin glued to the base of a crocodile clamp. The shaft of the holder can be inserted in a stereotaxic micro manipulator for implantation.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Take an anesthetized mouse secured to a stereotaxic frame with its skull exposed.
The skull features implanted ground, reference, and anchoring screws, as well as a craniotomy to expose the brain.
Take a silicon probe fixed to an insertion tool and mounted on a holder.
The probe features shanks with linearly arranged electrodes.
Insert the probe into the brain slightly above the desired depth and fix it to the anchoring screws with dental cement.
&#160;Then, lower it further to reach the target depth.
Release the insertion tool, then release the connector board and fix it on the skull.
Connect the connector board&#39;s ground and reference wires to the corresponding wires on the skull for a stable electrical connection.
Cover the assembly with a protective casing.
Allow the mouse to recover and start recording.
The electrodes extracellularly record the neuronal activity at the implanted site, called the local field potential.

24 Aufrufe. Aufzeichnung der elektrischen Aktivität des Gehirns bei sich verhaltenden Mäusen mit linearen Siliziumsonden mit mehreren Schäften

Serie: Aufzeichnung der elektrischen Aktivität des Gehirns bei sich verhaltenden Mäusen mit linearen Siliziumsonden mit mehreren Schäften - Experiment

Acute In Vivo Electrophysiological Recordings of Local Field Potentials and Multi-unit Activity from the Hyperdirect Pathway in Anesthetized Rats

Converging evidence shows that many neuropsychiatric diseases should be understood as disorders of large-scale neuronal networks. To better understand the pathophysiological basis of these diseases, it is necessary to precisely characterize in which way the processing of information is disturbed between the different neuronal parts of the circuit. Using extracellular in vivo electrophysiological recordings, it is possible to accurately delineate neuronal activity within a neuronal network. The application of this method has several advantages over alternative techniques, e.g., functional magnetic resonance imaging and calcium imaging, as it allows a unique temporal and spatial resolution and does not rely on genetically engineered organisms. However, the use of extracellular in vivo recordings is limited since it is an invasive technique that cannot be universally applied. In this article, a simple and easy to use method is presented with which it is possible to simultaneously record extracellular potentials such as local field potentials and multiunit activity at multiple sites of a network. It is detailed how a precise targeting of subcortical nuclei can be achieved using a combination of stereotactic surgery and online analysis of multi-unit recordings. Thus, it is demonstrated, how a complete network such as the hyperdirect cortico-basal ganglia loop can be studied in anesthetized animals in vivo.

Acute In Vivo Electrophysiological Recordings of Local Field Potentials and Multi-unit Activity from the Hyperdirect Pathway in Anesthetized Rats

In this study, the methodology is presented on how to perform multi-site in vivo electrophysiological recordings from the hyperdirect pathway under urethane anesthesia.

Akut<em&gt; In vivo</em&gt; Elektrophysiologische Aufzeichnungen von lokalen Feldpotentialen und Multi-Unit-Aktivitäten von der Hyperdirect-Pathway in anästhesierten Ratten

Converging evidence shows that many neuropsychiatric diseases should be understood as disorders of large-scale neuronal networks. To better understand the ...

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Neurowissenschaften

Simultaneous Recordings of Cortical Local Field Potentials, Electrocardiogram, Electromyogram, and Breathing Rhythm from a Freely Moving Rat

Monitoring the physiological dynamics of the brain and peripheral tissues is necessary for addressing a number of questions about how the brain controls body functions and internal organ rhythms when animals are exposed to emotional challenges and changes in their living environments. In general experiments, signals from different organs, such as the brain and the heart, are recorded by independent recording systems that require multiple recording devices and different procedures for processing the data files. This study describes a new method that can simultaneously monitor electrical biosignals, including tens of local field potentials in multiple brain regions, electrocardiograms that represent the cardiac rhythm, electromyograms that represent awake/sleep-related muscle contraction, and breathing signals, in a freely moving rat. The recording configuration of this method is based on a conventional micro-drive array for cortical local field potential recordings in which tens of electrodes are accommodated, and the signals obtained from these electrodes are integrated into a single electrical board mounted on the animal's head. Here, this recording system was improved so that signals from the peripheral organs are also transferred to an electrical interface board. In a single surgery, electrodes are first separately implanted into the appropriate body parts and the target brain areas. The open ends of all of these electrodes are then soldered to individual channels of the electrical board above the animal's head so that all of the signals can be integrated into the single electrical board. Connecting this board to a recording device allows for the collection of all of the signals into a single device, which reduces experimental costs and simplifies data processing, because all data can be handled in the same data file. This technique will aid the understanding of the neurophysiological correlates of the associations between central and peripheral organs.

This study introduces a method for the simultaneous recording of local field potentials in the brain, electrocardiograms, electromyograms, and breathing signals of a freely moving rat. This technique, which reduces experimental costs and simplifies data analysis, will contribute to the understanding of the interactions between the brain and peripheral organs.

Gleichzeitige Aufnahmen der kortikalen lokales Feld Potentiale, Elektrokardiogramm, Elektromyogramm und Atemrhythmus aus einem frei beweglichen Ratte

Monitoring the physiological dynamics of the brain and peripheral tissues is necessary for addressing a number of questions about how the brain controls ...

Construction of Local Field Potential Microelectrodes for in vivo Recordings from Multiple Brain Structures Simultaneously

Researchers often need to record local field potentials (LFPs) simultaneously from several brain structures. Recording from multiple desired brain regions requires different microelectrode designs, but commercially available microelectrode arrays often do not offer such flexibility. Here, the present protocol outlines the straightforward design of custom-made microelectrode arrays to record LFPs from multiple brain structures simultaneously at different depths. This work describes the construction of the bilateral cortical, striatal, ventrolateral thalamic, and nigral microelectrodes as an example. The outlined design principle offers flexibility, and the microelectrodes can be modified and customized to record LFPs from any structure by calculating stereotaxic coordinates and quickly changing the construction accordingly to target different brain regions in either freely moving or anesthetized mice. The microelectrode assembly requires standard tools and supplies. These custom microelectrode arrays allow investigators to easily design microelectrode arrays in any configuration to track neuronal activity, providing LFP recordings with millisecond resolution.

Construction of Local Field Potential Microelectrodes for in vivo Recordings from Multiple Brain Structures Simultaneously

The present protocol describes the construction of custom-made microelectrode arrays to record local field potentials in vivo from multiple brain structures simultaneously.

Aufbau von lokalen Feldpotential-Mikroelektroden für in vivo Aufnahmen aus mehreren Gehirnstrukturen gleichzeitig

Researchers often need to record local field potentials (LFPs) simultaneously from several brain structures. Recording from multiple desired brain regions ...

Recording Brain Electrical Activity in Behaving Mice Using Multi-Shank Linear Silicon Probes

Transkript

Recording Brain Electrical Activity in Behaving Mice Using Multi-Shank Linear Silicon Probes