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><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&amp;uuml;rklin Elektronik</td><td>89 F 232</td><td></td></tr><tr><td>Ground screws</td><td>Screws &amp;amp; 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>Multimeter</td><td>Uni-T</td><td>UT61C</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>9.78012E+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&amp;amp;M</td><td></td></tr><tr><td>Carprofen</td><td>Zoetis</td><td>Rimadyl 50mg/ml</td><td></td></tr><tr><td>Recording amplifier</td><td>Intan Technologies</td><td>C3323</td><td></td></tr><tr><td>USB acquisition board</td><td>Intan Technologies</td><td>C3004</td><td></td></tr><tr><td>Recording cables</td><td>Intan Technologies</td><td>C3216</td><td></td></tr><tr><td>Electrical commutator</td><td>Doric lenses</td><td>HRJ-OE_FC_12_HARW</td><td></td></tr><tr><td>Acquisition software</td><td>OpenEphys (www.open-ephys.org)</td><td>GUI</td><td>allows platform-independent data acquisition</td></tr><tr><td>Computer for data acquisition</td><td></td><td></td><td></td></tr><tr><td>Analysis environment</td><td>Python (www.python.org)</td><td></td><td>allows platform-independent data analysis</td></tr><tr><td>Urethane</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>Vibratome</td><td>Leica</td><td>VT1000</td><td></td></tr><tr><td>Microscope slides</td><td>Carl Roth</td><td>H868.1</td><td></td></tr><tr><td>Cover slips</td><td>Carl Roth</td><td>H878.2</td><td></td></tr><tr><td>Embedding medium</td><td>Sigma-Aldrich</td><td>81381-50G</td><td></td></tr><tr><td>Distilled water</td><td>Millipore</td><td>Milli Q</td><td>Table-top machine for the production of distilled water</td></tr><tr><td>Tergazyme</td><td>Alconox</td><td>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

Universidad de Cartagena UDEC

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

Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit

This protocol outlines the procedures for preparing and recording from the isolated whole hippocampus, of WT and transgenic mice, along with recent improvements in methodologies and applications for the study of theta oscillations. A simple characterization of the isolated hippocampal preparation is presented whereby the relationship between internal hippocampal theta oscillators is examined together with the activity of pyramidal cells, and GABAergic interneurons, of the cornu ammonis-1 (CA1) and subiculum (SUB) areas. Overall, we show that the isolated hippocampus is capable of generating intrinsic theta oscillations in vitro and that rhythmicity generated within the hippocampus can be precisely manipulated by optogenetic stimulation of parvalbumin-positive (PV) interneurons. The in vitro isolated hippocampal preparation offers a unique opportunity to use simultaneous field and intracellular patch-clamp recordings from visually-identified neurons to better understand the mechanisms underlying theta rhythm generation.

Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit

Here, we present a protocol for recording rhythmic neuronal network theta and gamma oscillations from an isolated whole hippocampal preparation. We describe the experimental steps from extraction of the hippocampus to details of field, unitary and whole-cell patch clamp recordings as well as optogenetic pacing of the theta rhythm.

This protocol outlines the procedures for preparing and recording from the isolated whole hippocampus, of WT and transgenic mice, along with recent ...

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 ...

Acute Mouse Brain Slicing to Investigate Spontaneous Hippocampal Network Activity

Acute rodent brain slicing offers a tractable experimental approach to gain insight into the organization and function of neural circuits with single-cell resolution using electrophysiology, microscopy, and pharmacology. However, a major consideration in the design of in vitro experiments is the extent to which different slice preparations recapitulate naturalistic patterns of neural activity as observed in vivo. In the intact brain, the hippocampal network generates highly synchronized population activity reflective of the behavioral state of the animal, as exemplified by the sharp-wave ripple complexes (SWRs) that occur during waking consummatory states or non-REM sleep. SWRs and other forms of network activity can emerge spontaneously in isolated hippocampal slices under appropriate conditions. In order to apply the powerful brain slice toolkit to the investigation of hippocampal network activity, it is necessary to utilize an approach that optimizes tissue health and the preservation of functional connectivity within the hippocampal network. Mice are transcardially perfused with cold sucrose-based artificial cerebrospinal fluid. Horizontal slices containing the hippocampus are cut at a thickness of 450 &#956;m to preserve synaptic connectivity. Slices recover in an interface-style chamber and are transferred to a submerged chamber for recordings. The recording chamber is designed for dual surface superfusion of artificial cerebrospinal fluid at a high flow rate to improve oxygenation of the slice. This protocol yields healthy tissue suitable for the investigation of complex and spontaneous network activity in vitro.

This protocol describes the preparation of horizontal hippocampal-entorhinal cortex (HEC) slices from mice exhibiting spontaneous sharp-wave ripple activity. Slices are incubated in a simplified interface holding chamber and recordings are performed under submerged conditions with fast-flowing artificial cerebrospinal fluid to promote tissue oxygenation and the spontaneous emergence of network-level activity.

Acute rodent brain slicing offers a tractable experimental approach to gain insight into the organization and function of neural circuits with single-cell ...

Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit functions in systems neuroscience. However, high absorbance and scattering of light in mammalian tissue limit intravital imaging mostly to superficial brain regions, leaving deep-brain areas, such as the hippocampus, out of reach for optical microscopy. In this video, we show the preparation and implantation of the custom-made imaging window to enable chronic in vivo imaging of the dorsal hippocampal CA1 region in head-fixed behaving mice. The custom-made window is supplemented with an infusion cannula that allows targeted delivery of viral vectors and drugs to the imaging area. By combining this preparation with wide-field imaging, we performed a long-term recording of neuronal activity using a fluorescent calcium indicator from large subsets of neurons in behaving mice over several weeks. We also demonstrated the applicability of this preparation for voltage imaging with single-spike resolution. High-performance genetically encoded indicators of neuronal activity and scientific CMOS cameras allowed the recurrent visualization of subcellular morphological details of single neurons at high temporal resolution. We also discuss the advantages and potential limitations of the described method and its compatibility with other imaging techniques.

This article demonstrates the preparation of a custom-made imaging window supplemented with infusion cannula and its implantation onto the CA1 region of the hippocampus in mice.

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit ...

LFP Recording in Mouse Brain for Cognitive Research and Drug Analysis

Dual Extracellular Recordings in the Mouse Hippocampus and Prefrontal Cortex

The technique of recording local field potentials (LFPs) is an electrophysiological method used to measure the electrical activity of localized neuronal populations. It serves as a crucial tool in cognitive research, particularly in brain regions like the hippocampus and prefrontal cortex. Dual LFP recordings between these areas are of particular interest as they allow the exploration of interregional signal communication. However, methods for performing these recordings are rarely described, and most commercial recording devices are either expensive or lack adaptability to accommodate specific experimental designs. This study presents a comprehensive protocol for performing dual-electrode LFP recordings in the mouse hippocampus and the prefrontal cortex to investigate the effects of antipsychotic drugs and potassium channel modulators on LFP properties in these areas. The technique enables the measurement of LFP properties, including power spectra within each brain region and coherence between the two. Additionally, a low-cost, custom-designed recording device has been developed for these experiments. In summary, this protocol provides a means to record signals with high signal-to-noise ratios in different brain regions, facilitating the investigation of interregional information communication within the brain.

Author Spotlight: Studying Drug Impacts on Brain Signals Using Dual LFP Recording Protocol in Mice

This protocol outlines the use of a custom-designed recording device and electrodes to record local field potentials and investigate information flow in the mouse hippocampus and prefrontal cortex.

The technique of recording local field potentials (LFPs) is an electrophysiological method used to measure the electrical activity of localized neuronal ...

Recording Neuronal Activity During Odor Plume Navigation with Miniscope Imaging

Miniscope Recording Calcium Signals at Hippocampus of Mice Navigating an Odor Plume

Mice navigate an odor plume with a complex spatiotemporal structure in the dark to find the source of odorants. This article describes a protocol to monitor behavior and record Ca2+ transients in dorsal CA1 stratum pyramidale neurons in the hippocampus (dCA1) in mice navigating an odor plume in a 50 cm x 50 cm x 25 cm odor arena. An epifluorescence miniscope focused through a gradient-index (GRIN) lens imaged Ca2+ transients in dCA1 neurons expressing the calcium sensor GCaMP6f in Thy1-GCaMP6f mice. The paper describes the behavioral protocol to train the mice to perform this odor plume navigation task in an automated odor arena. The methods include a step-by-step procedure for the surgery for GRIN lens implantation and baseplate placement for imaging GCaMP6f in CA1. The article provides information on real-time tracking of the mouse position to automate the start of the trials and delivery of a water reward. In addition, the protocol includes information on using an interface board to synchronize metadata describing the automation of the odor navigation task and frame times for the miniscope and a digital camera tracking mouse position. Moreover, the methods delineate the pipeline used to process GCaMP6f fluorescence movies by motion correction using the NorMCorre algorithm followed by identification of regions of interest with EXTRACT. Finally, the paper describes an artificial neural network approach to decode spatial paths from CA1 neural ensemble activity to predict mouse navigation of the odor plume.

Author Spotlight: Understanding Processing of Olfactory and Spatial Information by Brain with Real-Time Behavioral Analysis

This protocol investigates the brain-behavior relationship in hippocampal CA1 in mice navigating an odor plume. We provide a step-by-step protocol, including surgery to access imaging of the hippocampus, behavioral training, miniscope GCaMP6f recording and processing of the brain, and behavioral data to decode the mouse position from ROI neural activity.

Mice navigate an odor plume with a complex spatiotemporal structure in the dark to find the source of odorants. This article describes a protocol to ...

Selective Manipulation of Theta Oscillations in a Mouse Model

Source: Bender, F. et al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/57349">Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice.</a> J. Vis. Exp (2018)
This video demonstrates a protocol to modulate theta oscillations in a genetically engineered mouse model with light-sensitive ion channels in its GABAergic medial septum neurons. By delivering light pulses to the hippocampus, the protocol aims to synchronize neuronal activity and generate theta oscillations, which are recorded as local field potentials.

Selektywna manipulacja oscylacjami theta w modelu mysim

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

JoVE Encyclopedia of Experiments

Neurophysiology

Stimulation and Modulation Techniques

Implanting an Electrode Array and Optic Fiber into the Mouse Hippocampus

Source: Bender, F., et. al., <a href="https://appjove.unicartagenaproxy.elogim.com/v/57349">Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice.</a> J. Vis. Exp. (2018)
This video demonstrates the stereotactic implantation of a wire electrode array and an optic fiber in the hippocampus of a mouse model.

Wszczepianie układu elektrod i światłowodu do hipokampa myszy

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

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

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.

Rejestrowanie aktywności elektrycznej mózgu u zachowujących się myszy za pomocą liniowych sond krzemowych z wieloma zębami

Electrophysiology Techniques

Recording of Local Field Potential in Mouse Hippocampal-Entorhinal Cortex Slices

Source: Whitebirch, A. C. <a href="https://appjove.unicartagenaproxy.elogim.com/v/61704">Acute Mouse Brain Slicing to Investigate Spontaneous Hippocampal Network Activity.</a> J. Vis. Exp. (2020)
The video demonstrates a procedure to record the local field potential (LFP) in mouse hippocampal-entorhinal cortex slices. Electrical stimulation is provided at the CA3 stratum radiatum of the hippocampus, which causes changes in the postsynaptic potential at the CA1 stratum pyramidale. The combined change in the membrane potential of CA1 neurons, termed the local field potential, is recorded.

Rejestracja potencjału pola lokalnego w wycinkach kory hipokampowo-śródwęchowej myszy

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Transcardial perfusion

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Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

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Rozdziały w tym wideo

Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Acute Mouse Brain Slicing to Investigate Spontaneous Hippocampal Network Activity

Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice

Dual Extracellular Recordings in the Mouse Hippocampus and Prefrontal Cortex

Miniscope Recording Calcium Signals at Hippocampus of Mice Navigating an Odor Plume

Selektywna manipulacja oscylacjami theta w modelu mysim

Wszczepianie układu elektrod i światłowodu do hipokampa myszy

Rejestrowanie aktywności elektrycznej mózgu u zachowujących się myszy za pomocą liniowych sond krzemowych z wieloma zębami

Rejestracja potencjału pola lokalnego w wycinkach kory hipokampowo-śródwęchowej myszy