We are grateful to Karin Winterhalter and Kerstin Semmler for technical assistance. This work was supported by the cluster of excellence BrainLinks - BrainTools (EXC 1086) of the German Research Foundation.

All methods involving living animals have been approved by the Regierungspr&#228;sidium Freiburg in accordance with the German Animal Welfare Act.
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 atlas12.</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&#160;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.&#160;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>
5. Histology
<ol>
	<li>After completion of the recording, deeply anesthetize the animal (e.g. 2 g/kg body weight urethane injected intraperitoneally). Confirm the anesthetic state by lack of response to toe pinching.</li>
	<li>Perfuse the mouse transcardially with ice-cold phosphate-buffered saline (~1 min) followed by 4% paraformaldehyde (~10 min) using standard intracardial perfusion methods13. Before perfusion, electrolytic lesioning of the recording sites might by performed (e.g. by applying 10 - 20 V of constant voltage for up to 1 s). Alternatively, fluorescent dyes applied to the shank tips before implantation can be used to track identification. Testing the various methods for the identification of electrode positions to obtain optimal results with different types of silicon probes is recommended.</li>
	<li>Cut brain sections (~100 &#181;m) and stain the slices with 4&#39;-6-diamidino-2-phenylindole (DAPI, 1 &#181;g/mL) followed by three washing steps in PB (each 10 min at room temperature).</li>
	<li>Place the sections on a microscope slide, apply a drop of embedding medium and cover the section with a cover slip. Let the embedding medium dry overnight at room temperature.</li>
	<li>Using an epifluorescence or confocal laser-scanning microscope, identify the location of the recording sites.</li>
	<li>To attempt recovery of the silicon probe for further use, hold the probe with a crocodile clamp and release the probe from the scull by carefully melting the dental cement with a soldering iron (400 &#176;C). Take care not to touch the probe shanks during this procedure!</li>
	<li>Wash the probe in hot distilled water (~80 &#176;C, 15 min) followed by enzymatic solution (1% Tergazyme in distilled water, 30 min at room temperature) and another washing step in distilled water (15 min). Note that the success rate of probe recovery is low.</li>
</ol>
6. CSD Analysis
<ol>
	<li>Using a suitable analysis environment (e.g. Python), convert LFP data of an individual shank to CSD by approximating the second spatial derivative along the shank as 
	<img alt="Equation" src="/files/ftp_upload/57714/57714eq1.jpg" /> 
	where LFPn,t is the LFP signal on the nth electrode at time t and &#916;z is the inter-electrode spacing. Note that due to the n-1 and n+1 operations, the CSD of the first and last electrodes of the shank cannot be estimated, which has to be taken under consideration during probe placement. Implement the approximation formula using a short segment of code that computes the CSD signal for each electrode while iterating over time (see Supplemental Code File).</li>
	<li>Use the obtained CSD signal for further analysis (e.g., studying specific frequency bands of brain oscillations by applying band-pass filters).</li>
</ol>

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The authors have nothing to disclose.

Increasing evidences indicate that brain oscillations in hippocampal neuronal circuits occur in discrete spatial domains10,11,16. CSD analysis drastically reduces the influence of volume conduction, a crucial prerequisite for the study of local oscillation events. With this video, we provide a guide to implanting silicon probes into the mouse hippocampus for the analysis of CSD data. We show representative examples of CSD signal...

Oscillations in the LFP are critically involved in information processing by neuronal circuits. They cover a wide spectrum of frequencies, ranging from slow waves (~1 Hz) to fast ripple oscillations (~200 Hz)1. Distinct frequency bands are associated with cognitive functions including memory, emotional processing, and navigation2,3,4,5,6,7. Current flow across neuronal membranes constitutes the largest part of the LFP signal8. Cations entering the cell (e.g. via activation of glutamatergic excitatory synapses) represent an active current sink (as charge leaves the extracellular medium). In contrast, net flow of positive charge to the extracellular medium, for instance by activation of GABAergic inhibitory synapses, depicts an active current source at that location. In neuronal dipoles, current sinks are paired with passive sources and vice versa due to compensating currents affecting membrane charge at distant sites.
The electrical field produced by remote neural processes can also result in considerable voltage deflections on a recording electrode and might thus be falsely considered as a local event. This volume conduction poses a serious challenge to the interpretation of LFP signals. CSD analysis provides information about local current sinks and sources underlying LFP signals and therefore comprises a means to reduce the impact of volume conduction8. In laminated structures like the hippocampus, one-dimensional CSD signals can be obtained by the second spatial derivative of the LFP recorded from equidistant electrodes arranged perpendicular to the laminar planes9. The advent of commercially available linear silicon probes has allowed researchers to utilize the CSD method for the study of local oscillation activity in the hippocampus. For example, it has been demonstrated that distinct gamma oscillations emerge in a layer-specific manner in the CA1 area10. Furthermore, CSD analysis has identified independent hot spots of gamma activity in the principal cell layer of the dentate gyrus11. Importantly, these findings were only apparent in local CSD but not in LFP signals. CSD analysis thus provides a powerful tool to gain insight in the microcircuit operations of the hippocampus.
In this protocol, we provide a comprehensive guide to obtain one-dimensional CSD signals with silicon probes. These methods will enable users to investigate localized oscillation events in the hippocampus of behaving mice.

<table border="0" cellpadding="0" cellspacing="0" width="1106">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Crocodile clamp with stand</td>
			<td style="width:381px;">Reichelt Elektronik</td>
			<td style="width:151px;">HALTER ZD-10D</td>
			<td style="width:376px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Silicon probe</td>
			<td style="width:381px;">Cambridge Neurotech</td>
			<td style="width:151px;">P-series 32</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Stereoscope</td>
			<td style="width:381px;">Olympus</td>
			<td style="width:151px;">SZ51</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Varnish-insulated copper wire</td>
			<td style="width:381px;">B&#252;rklin Elektronik</td>
			<td style="width:151px;">89 F 232</td>
			<td></td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:199px;">Ground screws</td>
			<td style="width:381px;">Screws &#38; More GmbH (screwsandmore.de)</td>
			<td style="width:151px;">DIN 84 A2 M1x2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Flux</td>
			<td style="width:381px;">Stannol</td>
			<td style="width:151px;">114018</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Ceramic-tipped forceps</td>
			<td style="width:381px;">Fine Science Tools</td>
			<td style="width:151px;">11210-60</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Paraffine Wax</td>
			<td style="width:381px;">Sigma-Aldrich</td>
			<td style="width:151px;">327204</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Cauterizer</td>
			<td style="width:381px;">Fine Science Tools</td>
			<td style="width:151px;">18010-00</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Soldering iron</td>
			<td style="width:381px;">Kurtz Ersa</td>
			<td style="width:151px;">OIC1300</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Multimeter</td>
			<td style="width:381px;">Uni-T</td>
			<td style="width:151px;">UT61C</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Ethanol</td>
			<td style="width:381px;">Carl Roth</td>
			<td style="width:151px;">9065.1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Pasteur pipettes</td>
			<td style="width:381px;">Carl Roth</td>
			<td style="width:151px;">EA65.1</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Heat sterilizer</td>
			<td style="width:381px;">Fine Science Tools</td>
			<td style="width:151px;">18000-45</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Stereotaxic frame</td>
			<td style="width:381px;">David Kopf</td>
			<td style="width:151px;">Model 1900</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Stereotaxic electrode holder</td>
			<td style="width:381px;">David Kopf</td>
			<td style="width:151px;">Model 1900</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Isoflurane</td>
			<td style="width:381px;">Abbvie</td>
			<td style="width:151px;">B506</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Oxygen concentrator</td>
			<td style="width:381px;">Respironix</td>
			<td style="width:151px;">1020007</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Buprenorphine</td>
			<td style="width:381px;">Indivior UK Limited</td>
			<td style="width:151px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Electrical shaver</td>
			<td style="width:381px;">Tondeo</td>
			<td style="width:151px;">Eco-XS</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Heating pad</td>
			<td style="width:381px;">Thermolux</td>
			<td style="width:151px;">463265/-67</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Surgical clamps</td>
			<td style="width:381px;">Fine Science Tools</td>
			<td style="width:151px;">18050-28</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Hydrogen peroxide</td>
			<td style="width:381px;">Sigma-Aldrich</td>
			<td style="width:151px;">H1009</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Sterile cotton wipes</td>
			<td style="width:381px;">Carl Roth</td>
			<td style="width:151px;">EH12.1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Drill</td>
			<td style="width:381px;">Proxxon</td>
			<td style="width:151px;">Micromot 230/E</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">21G injection needle</td>
			<td style="width:381px;">B. Braun</td>
			<td style="width:151px;">4657527</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Phosphate buffer/phosphate buffered saline</td>
			<td style="width:381px;"></td>
			<td style="width:151px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Stereotaxic atlas</td>
			<td style="width:381px;">Elsevier</td>
			<td style="width:151px;">9.78012E+12</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Surgical scissors</td>
			<td style="width:381px;">Fine Science Tools</td>
			<td style="width:151px;">14094-11</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Surgical forceps</td>
			<td style="width:381px;">Fine Science Tools</td>
			<td style="width:151px;">11272-40</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">27G injection needles</td>
			<td style="width:381px;">B. Braun</td>
			<td style="width:151px;">4657705</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Vaseline</td>
			<td style="width:381px;"></td>
			<td style="width:151px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Dental cement</td>
			<td style="width:381px;">Sun Medical</td>
			<td style="width:151px;">SuperBond T&#38;M</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Carprofen</td>
			<td style="width:381px;">Zoetis</td>
			<td style="width:151px;">Rimadyl 50mg/ml</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Recording amplifier</td>
			<td style="width:381px;">Intan Technologies</td>
			<td style="width:151px;">C3323</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">USB acquisition board</td>
			<td style="width:381px;">Intan Technologies</td>
			<td style="width:151px;">C3004</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Recording cables</td>
			<td style="width:381px;">Intan Technologies</td>
			<td style="width:151px;">C3216</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Electrical commutator</td>
			<td style="width:381px;">Doric lenses</td>
			<td style="width:151px;">HRJ-OE_FC_12_HARW</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Acquisition software</td>
			<td>OpenEphys (www.open-ephys.org)</td>
			<td style="width:151px;">GUI</td>
			<td>allows platform-independent data acquisition</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:199px;">Computer for data acquisition</td>
			<td style="width:381px;"></td>
			<td style="width:151px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Analysis environment</td>
			<td>Python (www.python.org)</td>
			<td style="width:151px;"></td>
			<td>allows platform-independent data analysis</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Urethane</td>
			<td style="width:381px;">Sigma-Aldrich</td>
			<td style="width:151px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Vibratome</td>
			<td style="width:381px;">Leica</td>
			<td style="width:151px;">VT1000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Microscope slides</td>
			<td style="width:381px;">Carl Roth</td>
			<td style="width:151px;">H868.1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Cover slips</td>
			<td style="width:381px;">Carl Roth</td>
			<td style="width:151px;">H878.2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Embedding medium</td>
			<td style="width:381px;">Sigma-Aldrich</td>
			<td style="width:151px;">81381-50G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Distilled water</td>
			<td style="width:381px;">Millipore</td>
			<td style="width:151px;">Milli Q</td>
			<td>Table-top machine for the production of distilled water</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:199px;">Tergazyme</td>
			<td style="width:381px;">Alconox</td>
			<td style="width:151px;">Tergazyme</td>
			<td></td>
		</tr>
	</tbody>
</table>

recording spatially restricted oscillations in the hippocampus of behaving mice

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed electrical field of a large volume of brain tissue, extracting information about local activity is challenging. Studying neuronal microcircuits, however, requires a reliable distinction between truly local events and volume-conducted signals originating in distant brain areas. Current source density (CSD) analysis offers a solution for this problem by providing information about current sinks and sources in the vicinity of the electrodes. In brain areas with laminar cytoarchitecture such as the hippocampus, one-dimensional CSD can be obtained by estimating the second spatial derivative of the LFP. Here, we describe a method to record multilaminar LFPs using linear silicon probes implanted into the dorsal hippocampus. CSD traces are computed along individual shanks of the probe. This protocol thus describes a procedure to resolve spatially restricted neuronal network oscillations in the hippocampus of freely moving mice.

Figure 1 illustrates the insertion tool used for the implantation of silicon probes. Recordings from chronically implanted silicon probes targeting the CA1 area and the granule cell layer of the dentate gyrus are shown in Figure 2. We recorded LFPs from the probe shanks during free movement in the homecage. To minimize the effect of volume conduction, the obtained signals were converted to CSD along each shank of the probe (<stro...

Watch this Scientific Journal Video about Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice at JoVE.com

Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

This protocol describes the recording of local field potentials with multi-shank linear silicon probes. Conversion of the signals using current source density analysis allows the reconstruction of local electrical activity in the mouse hippocampus. With this technique, spatially restricted brain oscillations can be studied in freely moving mice.

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed ...

recording-spatially-restricted-oscillations-hippocampus-behaving

Research

JoVE Journal

Neuroscience

8.7K Views.  University of Freiburg. This protocol describes the recording of local field potentials with multi-shank linear silicon probes. Conversion of the signals using current source density analysis allows the reconstruction of local electrical activity in the mouse hippocampus. With this technique, spatially restricted brain oscillations can be studied in freely moving mice.

Video: Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

Derivation of Glial Restricted Precursors from E13 mice

This is a protocol for derivation of glial restricted precursor (GRP) cells from the spinal cord of E13 mouse fetuses. These cells are early precursors within the oligodendrocytic cell lineage. Recently, these cells have been studied as potential source for restorative therapies in white matter diseases. Periventricular leukomalacia (PVL) is the leading cause of non-genetic white matter disease in childhood and affects up to 50% of extremely premature infants. The data suggest a heightened susceptibility of the developing brain to hypoxia-ischemia, oxidative stress and excitotoxicity that selectively targets nascent white matter. Glial restricted precursors (GRP), oligodendrocyte progenitor cells (OPC) and immature oligodendrocytes (preOL) seem to be key players in the development of PVL and are the subject of continuing studies. Furthermore, previous studies have identified a subset of CNS tissue that has increased susceptibility to glutamate excitotoxicity as well as a developmental pattern to this susceptibility. Our laboratory is currently investigating the role of oligodendrocyte progenitors in PVL and use cells at the GRP stage of development. We utilize these derived GRP cells in several experimental paradigms to test their response to select stresses consistent with PVL. GRP cells can be manipulated in vitro into OPCs and preOL for transplantation experiments with mouse PVL models and in vitro models of PVL-like insults including hypoxia-ischemia. By using cultured cells and in vitro studies there would be reduced variability between experiments which facilitates interpretation of the data. Cultured cells also allows for enrichment of the GRP population while minimizing the impact of contaminating cells of non-GRP phenotype.

This protocol outlines the derivation of Glial Restricted Precursors from fetal spinal cords and maintained in vitro either for transplantation or for the study of oligodendrocytic lineage.

This is a protocol for derivation of glial restricted precursor (GRP) cells from the spinal cord of E13 mouse fetuses. These cells are early precursors ...

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

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

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

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

Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

Alzheimer&#8217;s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the over-production of amyloid-beta and the deposition of amyloid-beta plaques in brain regions of the afflicted individuals. Throughout the years scientists have generated numerous Alzheimer&#8217;s disease mouse models that attempt to replicate the amyloid-beta pathology. Unfortunately, the mouse models only selectively mimic the disease features. Neuronal death, a prominent effect in the brains of Alzheimer&#8217;s disease patients, is noticeably lacking in these mice. Hence, we and others have employed a method of directly infusing soluble oligomeric species of amyloid-beta - forms of amyloid-beta that have been proven to be most toxic to neurons - stereotaxically into the brain. In this report we utilize male C57BL/6J mice to document this surgical technique of increasing amyloid-beta levels in a select brain region. The infusion target is the dentate gyrus of the hippocampus because this brain structure, along with the basal forebrain that is connected by the cholinergic circuit, represents one of the areas of degeneration in the disease. The results of elevating amyloid-beta in the dentate gyrus via stereotaxic infusion reveal increases in neuron loss in the dentate gyrus within 1 week, while there is a concomitant increase in cell death and cholinergic neuron loss in the vertical limb of the diagonal band of Broca of the basal forebrain. These effects are observed up to 2 weeks. Our data suggests that the current amyloid-beta infusion model provides an alternative mouse model to address region specific neuron death in a short-term basis. The advantage of this model is that amyloid-beta can be elevated in a spatial and temporal manner.

Here, we present a protocol for direct stereotaxic brain infusion of amyloid-beta. This methodology provides an alternative in vivo mouse model to address the short-term effects of amyloid-beta on brain neurons.

Alzheimer&#8217;s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the ...

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 ...

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools to selectively manipulate brain rhythms. Here we describe an approach combining projection-specific optogenetics with extracellular electrophysiology for high-fidelity control of hippocampal theta oscillations (5-10 Hz) in behaving mice. The specificity of the optogenetic entrainment is achieved by targeting channelrhodopsin-2 (ChR2) to the GABAergic population of medial septal cells, crucially involved in the generation of hippocampal theta oscillations, and a local synchronized activation of a subset of inhibitory septal afferents in the hippocampus. The efficacy of the optogenetic rhythm control is verified by a simultaneous monitoring of the local field potential (LFP) across lamina of the CA1 area and/or of neuronal discharge. Using this readily implementable preparation we show efficacy of various optogenetic stimulation protocols for induction of theta oscillations and for the manipulation of their frequency and regularity. Finally, a combination of the theta rhythm control with projection-specific inhibition addresses the readout of particular aspects of the hippocampal synchronization by efferent regions.

We describe the use of optogenetics and electrophysiological recordings for selective manipulations of hippocampal theta oscillations (5-10 Hz) in behaving mice. The efficacy of the rhythm entrainment is monitored using local field potential. A combination of opto- and pharmacogenetic inhibition addresses the efferent readout of hippocampal synchronization.

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools ...

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 ...

Long-term Sensory Conflict in Freely Behaving Mice

Long-term sensory conflict protocols are a valuable means of studying motor learning. The presented protocol produces a persistent sensory conflict for experiments aimed at studying long-term learning in mice. By permanently wearing a device fixed on their heads, mice are continuously exposed to a sensory mismatch between visual and vestibular inputs while freely moving in home cages. Therefore, this protocol readily enables the study of the visual system and multisensory interactions over an extended timeframe that would not be accessible otherwise. In addition to lowering the experimental costs of long-term sensory learning in naturally behaving mice, this approach accommodates the combination of in vivo and in vitro experiments. In the reported example, video-oculography is performed to quantify the vestibulo-ocular reflex (VOR) and optokinetic reflex (OKR) before and after learning. Mice exposed to this long-term sensory conflict between visual and vestibular inputs presented a strong VOR gain decrease but exhibited few OKR changes. Detailed steps of device assembly, animal care, and reflex measurements are hereby reported.

The presented protocol produces a persistent sensory conflict for experiments aimed at studying long-term learning. By permanently wearing a fixed device on their heads, mice are continuously exposed to a sensory mismatch between visual and vestibular inputs while freely moving in home cages.

Long-term sensory conflict protocols are a valuable means of studying motor learning. The presented protocol produces a persistent sensory conflict for ...

Investigating Long-term Synaptic Plasticity in Interlamellar Hippocampus CA1 by Electrophysiological Field Recording

The study of synaptic plasticity in the hippocampus has focused on the use of the CA3-CA1 lamellar network. Less attention has been given to the longitudinal interlamellar CA1-CA1 network. Recently however, an associational connection between CA1-CA1 pyramidal neurons has been shown. Therefore, there is the need to investigate whether the longitudinal interlamellar CA1-CA1 network of the hippocampus supports synaptic plasticity.
We designed a protocol to investigate the presence or absence of long-term synaptic plasticity in the interlamellar hippocampal CA1 network using electrophysiological field recordings both in vivo and in vitro. For in vivo extracellular field recordings, the recording and stimulation electrodes were placed in a septal-temporal axis of the dorsal hippocampus at a longitudinal angle, to evoke field excitatory postsynaptic potentials. For in vitro extracellular field recordings, hippocampal longitudinal slices were cut parallel to the septal-temporal plane. Recording and stimulation electrodes were placed in the stratum oriens (S.O) and the stratum radiatum (S.R) of the hippocampus along the longitudinal axis. This enabled us to investigate the directional and layer specificity of evoked excitatory postsynaptic potentials. Already established protocols were used to induce long-term potentiation (LTP) and long-term depression (LTD) both in vivo and in vitro. Our results demonstrated that the longitudinal interlamellar CA1 network supports N-methyl-D-aspartate (NMDA) receptor-dependent long-term potentiation (LTP) with no directional or layer specificity. The interlamellar network, however, in contrast to the transverse lamellar network, did not present with any significant long-term depression (LTD).

We used recording and stimulation electrodes in longitudinal hippocampal brain slices and longitudinally positioned recording and stimulation electrodes in the dorsal hippocampus in vivo to evoke extracellular postsynaptic potentials and demonstrate long-term synaptic plasticity along the longitudinal interlamellar CA1.

The study of synaptic plasticity in the hippocampus has focused on the use of the CA3-CA1 lamellar network. Less attention has been given to the ...

Electrocorticographic Recording of Cerebral Cortex Areas Manipulated Using an Adeno-Associated Virus Targeting Cofilin in Mice

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. Adeno-associated viruses (AAVs) are increasingly used to improve understanding of brain circuits and their functions. The AAV-mediated manipulation of specific cell populations and/or of precise molecular components has been tremendously useful to identify new sleep regulatory circuits/molecules and key proteins contributing to the adverse effects of sleep loss. For instance, inhibiting activity of the filamentous actin-severing protein&#160;cofilin&#160;using AAV prevents sleep deprivation-induced memory impairment. Here, a protocol is&#160;described that combines the manipulation of cofilin function in a cerebral cortex area with the recording of ECoG activity to examine whether cortical cofilin modulates the wakefulness and sleep ECoG signals. AAV injection is performed during the same surgical procedure as the implantation of ECoG and electromyographic (EMG) electrodes in adult male and female mice. Mice are anesthetized, and their heads are shaved. After skin cleaning and incision, stereotaxic coordinates of the motor cortex are determined, and the skull is pierced at this location. A cannula prefilled with an AAV expressing cofilinS3D, an inactive form of cofilin, is slowly positioned in the cortical tissue. After AAV infusion, gold-covered screws (ECoG electrodes) are screwed through the skull and cemented to the skull with gold wires inserted in the neck muscles (EMG electrodes). The animals are allowed three weeks to recover and to ensure sufficient expression of cofilinS3D. The infected area and cell type are verified using immunohistochemistry, and the ECoG is analyzed using visual identification of vigilance states and spectral analysis. In summary, this combined methodological approach allows the investigation of the precise contribution of molecular components regulating neuronal morphology and connectivity to the regulation of synchronized cerebral cortex activity during wakefulness and sleep.

This article describes a protocol for the manipulation of molecular targets in the cerebral cortex using adeno-associated viruses and for monitoring the effects of this manipulation during wakefulness and sleep using electrocorticographic recordings.

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. ...

Preparation of Acute Slices from Dorsal Hippocampus for Whole-Cell Recording and Neuronal Reconstruction in the Dentate Gyrus of Adult Mice

Although the general architecture of the hippocampus is similar along its longitudinal axis, recent studies have revealed prominent differences in molecular, anatomical and functional criteria suggesting a division into different sub-circuits along its rostro-caudal extent. Owing to differential connectivity and function the most fundamental distinction is made between the dorsal and the ventral hippocampus, which are preferentially involved in spatial and emotional processing, respectively. Accordingly, in vivo work regarding spatial memory formation has focused on the dorsal hippocampus.
In contrast, electro-physiological in vitro recordings have been preferentially performed on intermediate-ventral hippocampus, largely motivated by factors like slice viability and circuit integrity. To allow for direct correlation of in vivo data on spatial processing with in vitro data we have adapted previous sectioning methods to obtain highly viable transverse brain slices from the dorsal-intermediate hippocampus for long-term recordings of principal cells and interneurons in the dentate gyrus. As spatial behavior is routinely analyzed in adult mice, we have combined this transversal slicing procedure with the use of protective solutions to enhance viability of brain tissue from mature animals. We use this approach for mice of about 3 months of age. The method offers a good alternative to the coronal preparation which is frequently used for in vitro studies on dorsal hippocampus. We compare these two preparations in terms of quality of recordings and preservation of morphological features of recorded neurons.

We present a protocol to prepare acute-slices from the dorsal-intermediate hippocampus of mice. We compare this transversal preparation with coronal slicing in terms of quality of recordings and preservation of morphological features of recorded neurons.