Figure 1, Figure 3, Figure 4, Figure 8 and Figure 9 were created with BioRender.com. We would like to thank Scott Kilianski for the help with MATLAB code and sharing his scripts. We thank Anna Grace Carns for the help with probe trajectory reconstruction.

All studies were approved by the Institutional Animal Care and Use Committee at the University of Virginia (Charlottesville, Virginia). Five male C57BL/6J&#160;mice, age 3-7 months, weighing 25-30 g,&#160;were used. The details of the reagents and the equipment used here are listed in the Table of Materials.
1. Headplate and headset implantation
<ol>
	<li>Make an ECoG headset by soldering perfluoroalkoxy (PFA) coated stainless steel wire to a 3-pin connector header (Figure 2A).</li>
	<li>Identify the insertion coordinates based on a mouse stereotaxic atlas17. Using the Pythagorean theorem to calculate the angle and depth of probe insertion -particularly when there is little information in the literature -is a reasonable starting point (Figure 1)18. 
	NOTE: Ultimately, coordinates will be adjusted by trial and error. To target the vlPAG the following coordinates were used in adult mice: anteroposterior (AP) -3.6 mm, mediolateral (ML) +0.5 mm, dorsoventral (DV) -4 mm. The probe was inserted at a 20&#176; angle (AP).</li>
	<li>Induce general anesthesia by placing the mouse in the induction chamber (1.5%-3% Isoflurane in oxygen). Once anesthetized, position the mouse in the stereotaxic frame, placing the animal&#39;s nose in the nose cone and stabilizing the head with head bars.</li>
	<li>Apply ophthalmic ointment to the eyes to prevent corneal damage. Use a temperature control system to maintain body temperature at 37 &#176;C.</li>
	<li>Apply hair removal cream or shave fur over the scalp, then disinfect with povidone iodine and 70% alcohol. Use the &#39;tips only&#39;&#160;aseptic technique to maintain a sterile surgical field.&#160;&#160;</li>
	<li>Perform a toe pinch test to check the depth of anesthesia.</li>
	<li>Administer analgesia: carprofen 2.5 mg/kg, given subcutaneously in the back.</li>
	<li>Make a 5 mm scalp incision to remove a circular patch of skin above the parietal and occipital bones. Gently remove the meninges by scraping them with the scalpel blade.</li>
	<li>Use the scalpel blade to cut muscle attachments and expose the parietal and occipital bones19. Apply hydrogen peroxide as needed to control bleeding and dry the skull surface. It is crucial to apply dental cement and resin on dry skull to achieve a strong bond.</li>
	<li>First, identify the bregma and lambda landmarks on the skull20. Then, adjust the nose cone position to level the anterior-posterior position of the skull, ensuring that no more than 100 &#181;m height difference exists between the two landmarks.</li>
	<li>To level the medial-lateral position of the skull, pick two opposite points between bregma and lambda, each one 1mm from the sagittal suture and check their level. If there&#39;s more than 100 &#181;m height difference between them, adjust the medial-lateral position of the skull by manipulating the head bars.</li>
	<li>Measure the distance between bregma and lambda and compare it to the distance reported in Franklin-Paxinos stereotaxic atlas (usually 4.2 mm)17. Use the difference between the measured and reported distances to scale your AP coordinate proportionally. Mark craniotomy coordinates on the skull with a sterilized pencil. 
	NOTE: If the measured bregma-lambda distance differs from 4.2 mm, then the coordinates need to be scaled proportionately. All AP coordinates reported in a stereotaxic atlas correspond to a standardized bregma-lambda distance. Because the size of a mouse skull is variable, it is important to adjust your coordinates accordingly.</li>
	<li>Using a stereotaxic micromanipulator, position the headplate directly on top of the lambda suture, and secure it to the skull by applying dental cement on the headplate and around it (Figure 3A,B). Allow at least 10 min for the cement to dry.</li>
	<li>Drill burr holes (0.5 mm in diameter) for two cortical electrodes (frontal and parietal cortex) and for one that will serve as a reference electrode (cerebellum). Place the stripped ends (0.5 mm) of coated silver wire electrodes within the burr holes and secure using ultraviolet light-activated resin.</li>
	<li>Completely cover the coated stainless-steel wires with dental cement so that no wire is exposed. Cover the underside and sides of the headset with dental cement so that it is firmly in place. Make sure bregma and lambda sutures remain visible once the headset is secured in place (Figure 2B).</li>
	<li>Let the mouse recover for a minimum of 7 days, examine the animal and surgical site daily for any irregularities. Administer analgesics (Carprofen 2.5 mg/kg, SC) as needed.</li>
</ol>
2. Silicon probe placement and recording
<ol>
	<li>Habituate the mouse to the recording rig and head fixation (at least 1.5 h on two separate days) (Figure 2D).</li>
	<li>On the day of the recording, place the mouse in the anesthesia chamber (Isoflurane 1.5%-3% in oxygen).</li>
	<li>Position the mouse in a stereotaxic frame, adjust nose cone and head bars as described in 1.3. Maintain anesthesia&#160;(Isoflurane 1.5%-3% in oxygen) throughout the stereotaxic surgery.&#160;Perform a toe pinch test to check the depth of anesthesia.</li>
	<li>Apply ophthalmic ointment to the eyes and maintain adequate body temperature.</li>
	<li>Identify bregma and lambda, ensuring that no more than 100 &#181;m height difference exists between the two anatomical landmarks.</li>
	<li>Find and mark the calculated coordinates on the skull with a sterile pencil, then create an outline of the 2 mm x 2 mm craniotomy window around the coordinates.</li>
	<li>Perform a toe pinch test to check anesthesia depth.</li>
	<li>Use a high-speed drill to create a 2 mm x 2 mm craniotomy window. Apply 0.5-1 mL of normal saline to prevent brain surface from drying. Remove the dura using a syringe needle and fine forceps. 
	NOTE: Watch for major vessels (superior sagittal sinus, transverse sinus) to avoid excessive bleeding (Figure 3).</li>
	<li>Use a high-speed drill to create a separate burr hole for the silicon probe&#39;s reference electrode, generally ~1-2 mm from the cranial window.</li>
	<li>Apply 0.2 mL low toxicity silicon adhesive on the skull to completely seal the craniotomy, using the attached applicator.</li>
	<li>Let the mouse recover for approximately 1 h.</li>
	<li>Affix the head of the mouse to the electrophysiology recording rig using the headplate and screws (Figure 2D).</li>
	<li>Coat the silicon probe shank with the fluorescent dye DiI (1:4 DiI:Ethanol) so that the probe trajectory can be reconstructed after the experiment.</li>
	<li>Mount the probe on the manipulator and set the desired angle. To target the vlPAG, an AP angle of 15-20&#176; was applied.</li>
	<li>Lower the recording probe to the brain surface within the center of the cranial window. First, manually insert the probe to a depth of ~300 &#181;m. Once inserted to this depth, slowly lower the probe automatically (e.g., 200 &#181;m/min) to the targeted depth to minimize tissue damage21 (Figure 2C). 
	NOTE: Manual insertion of the probe is initially recommended. Manual probe insertion ensures the ability to stop and retract the probe should it bend upon initial insertion. Once the probe fully penetrates the tissue, it is generally safe to continue descending the probe in automatic mode.</li>
	<li>Apply mineral oil to the brain surface within the craniotomy window to prevent from drying.</li>
	<li>Let the recording probe settle for 10 min after insertion.</li>
	<li>Record data from the silicon probe and ECoG at 30kHz using an Intan Recording Controller.</li>
	<li>1-2 h after recording, use the transcardial perfusion technique to fix the brain in 4% paraformaldehyde22. 
	NOTE: Because of the cement, it might be difficult to remove the rostral part of the skull. That&#39;s why it is preferable to excise the brain by removing the dorsal parts of the skull.</li>
	<li>Decapitate the mouse, cut the skin following the midline on the dorsal side, from the neck to the bottom jaw. Remove muscles and tissue attached to the skull, cut off the bottom jaw for easier access to the brain.</li>
	<li>If the perfusion is done&#160;correctly the brain should shrink a little,&#160;leaving enough space to insert&#160;fine&#160;scissors in the dorsal part of the&#160;foramen magnum.&#160;Make one medial cut, one lateral and one contralateral cut, around 1 mm in size, in the occipital bone.</li>
	<li>Carefully remove the dorsal part of the skull using ophthalmic forceps. Start at the occipital bone, remove all skull fragments until the whole dorsal part of the brain is exposed.</li>
	<li>Slide a micro spatula under the brain, starting at the olfactory bulbs to scoop the brain out.</li>
	<li>Once perfused and removed, the brain can be stored in 4% paraformaldehyde at 4 &#176;C for 24-48 h.</li>
</ol>
3. Histology for probe trajectory reconstruction
<ol>
	<li>Section the brain into 70 &#181;m coronal sections using a vibratome.</li>
	<li>Mount the sections on slides using a DAPI mounting medium that stains cell nuclei. Coverslip and seal the slides with clear nail polish.</li>
	<li>Image the slides on a fluorescence microscope. Adjust contrast/brightness so that probe tracks are clearly visible. Ensure the resultant tiff image file sizes are less than 10 MB, so that MATLAB codes run smoothly. Reconstruct the probe tracks using a MATLAB code23.</li>
</ol>
4. Electrophysiological data analysis
<ol>
	<li>Analyze recorded neural signals from the silicon probe using an off-line detection and automatic sorting software (Kilosort)24.</li>
	<li>Manually classify detected clusters with Phy as multi- or single units25. Classify clusters as single units when they have a physiological spike waveform shape, show a refractory period in the cross-correlogram, and a normal distribution of amplitude view.</li>
	<li>Import single unit data to MATLAB and analyze23.</li>
</ol>

Angled Silicon Multielectrode Recordings for Deep Brainstem Structures in Mice

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The authors have no competing financial interests or other conflicts of interest pursuant to this work.

Brainstem nuclei mediate fundamental functions such as breathing, consciousness, and sleep26,27,28. The brainstem&#39;s location (deep and posterior) presents a challenge in studying its neuronal activity in vivo using standard techniques. Here an angled anterior approach is presented to allow reproducible single unit recording in head-fixed mice.
Careful insertion of the multi-electrode sili...

Silicon multielectrode-based recordings are becoming increasingly popular to measure neuronal activity across many brain regions with single action potential resolution1,2,3,4. Over the last decade, high-density recording technology has grown considerably. Current silicon-based recording electrodes can accommodate high channel counts, optical fibers, and electrocorticography (ECoG) recording devices5,6. Moreover, chronic implantation of these electrodes allows for long-term recordings7,8.
Despite recent technological advancements, targeting deep caudal structures like the brainstem with multi-channel probes remains challenging. When targeting brainstem structures such as the ventrolateral periacqueductal gray (vlPAG), one significant obstacle is identifying a probe trajectory that avoids major blood vessels, e.g., the superior sagittal venous sinus and the transverse venous sinus. Injury to these large veins can cause extensive bleeding, damage to the underlying brain tissue, and even death9,10. We propose targeting brainstem structures from anterior coordinates at an angle, allowing the recording probe to penetrate the brain below such high-risk vascular structures (see Figure 1). This angled approach, compared to a vertical one, maximizes the number of brain regions accessible for recording. Additionally, in experimental circumstances wherein ECoG recordings are desired, the angled anterior approach affords more skull surface available for ECoG headset implantation, as the craniotomy window for probe insertion is positioned more anteriorly10,11.
Identifying the specific cell groups and circuits responsible for anesthesia-induced REM sleep changes remains a major goal of anesthesia research. Thus, the objective here was to reproducibly and reliably access the vlPAG - a brainstem region associated with REM sleep - to obtain single unit, multi-electrode recordings in head-fixed mice before and during sevoflurane anesthesia12,13. Previous studies have used electrophysiological local field potential (LFP) measurements of the vlPAG in awake mice to identify neural state changes associated with anesthesia14,15. However, LFP measurements are primarily sensitive to synaptic activity, not action potentials, within the recorded area16. Consequently, there remains a limited understanding of how anesthetics directly affect the neural activity patterns produced by vlPAG neurons. Here, a method is described to obtain high-density single-neuron recordings from the brainstem of head-fixed mice. This method can also be adapted to record single-neuron activity from various other deep and posterior brainstem structures.

<table><tbody><tr><td>1024 channel RHD Recording Controller</td><td>Intan Technologies, Los Angeles, California, USA</td><td>C3008</td><td>Silicon probe recording; recording hardware and software</td></tr><tr><td>24 mm x 50 mm No. 1.5 VWR coverslip</td><td>VWR, Radnor, Pennsylvania, USA</td><td>48393-081</td><td>Histology</td></tr><tr><td>4% PFA in PBS</td><td>ThermoFisher Scientific, Waltham, Massachusetts, USA</td><td>J61899.AK</td><td>Histology; perfusion solution</td></tr><tr><td>C&amp;amp;B metabond</td><td>Patterson Dental, Richmond, Virginia, USA</td><td>powder: 5533559, quick base: 5533492, catalyst: 55335007</td><td>Headplate &amp;amp;Headset Implantation</td></tr><tr><td>C57/6J mice 4-6 weeks, males</td><td>The Jackson Laboratory, Bar Harbor, Maine, USA</td><td>000664</td><td></td></tr><tr><td>Capnomac Ultima</td><td>Datex, Helsinki, Finland&amp;nbsp;</td><td>ULT-SVi-27-07</td><td>Gas Analyzer; discontinued; alternative gas analyzer can be purchased from Bionet America&amp;nbsp;</td></tr><tr><td>CM-DiI</td><td>ThermoFisher Scientific, Waltham, Massachusetts, USA</td><td>V22888</td><td>Red fluorescent dye for coating of the silicon probe</td></tr><tr><td>Connector Header</td><td>DigiKey, Thief River Falls, Minnesota, USA</td><td>1212-1788-ND</td><td>ECoG Headset</td></tr><tr><td>DAPI Fluoromount-G</td><td>SouthernBiotech, Birmingham, Alabama, USA</td><td>0100-20</td><td>Histology</td></tr><tr><td>iBOND Universal</td><td>Patterson Dental, Richmond, Virginia, USA</td><td>044-1113</td><td>Headplate &amp;amp;Headset Implantation; for&amp;nbsp; securing stainless steel wires to the skull</td></tr><tr><td>Low toxicity silicon adhesive</td><td>World Precision Instruments, Sarasota, Florida, USA</td><td>KWIK-SIL</td><td>Headplate</td></tr><tr><td>Micro-Manipulator System</td><td>New Scale Technologies, Victor, New York, USA</td><td>Multi-Probe Manipulator: XYZ Stage Assembly: 06464-0000, MPM System Kit: 06267-3-0001, MPM-Platform-360, MPM ring for MPM Manual Arms, MPM_Ring-72 DEG: 06262-3-0000</td><td>Silicon probe recording; inserting the probe into the brain</td></tr><tr><td>Microprobes</td><td>UCLA, Los Angeles, California, USA</td><td>256 ANS, 64M</td><td>Discontinued; alternative silicon probes can be purchased from Neuropixels</td></tr><tr><td>Mineral Oil</td><td>Sigma Aldrich, Saint Luis, Missouri, USA</td><td>M8410-100ML</td><td>Silicon probe recording; preventing the tissue from drying during the recording</td></tr><tr><td>Normal saline</td><td>ThermoFisher Scientific, Waltham, Massachusetts, USA</td><td>Z1376</td><td>Headplate &amp;amp;Headset Implantation; preventing the brain from drying during the surgery</td></tr><tr><td>PFA-Coated Stainless Steel Wire-Diameter 0.008 in. coated with striped ends</td><td>A-M systems, Sequim, Washington, USA</td><td>791400</td><td>ECoG Headset &amp;amp; reference electrode for ECoG&amp;nbsp;</td></tr><tr><td>Platinum wire 24AWG&amp;nbsp;</td><td>World Precision Instruments, Sarasota, Florida, USA</td><td>PTP201</td><td>Reference electrode for the silicon probe recording&amp;nbsp;</td></tr><tr><td>Shandon Colorfrost Plus microscope slides</td><td>ThermoFisher Scientific, Waltham, Massachusetts, USA</td><td>99-910-01</td><td>Histology</td></tr><tr><td>Stainless steel Headplate</td><td>Star Rapid, China</td><td>custom made part</td><td>Headplate &amp;amp;Headset Implantation; design available upon request</td></tr><tr><td>Stereotaxic apparatus</td><td>KOPF, Tujunga, California, USA</td><td>Model 940 Small Animal Stereotaxic Instrument with Digital Display Console</td><td>Headplate &amp;amp;Headset Implantation</td></tr></tbody></table>

acute single-unit multi-electrode recordings from the brainstem of head-fixed mice

Silicon multielectrode-based recordings are increasingly popular for studying neuronal activity at the temporal resolution of action potentials in many brain regions. However, recording neuronal activity from deep caudal structures like the brainstem using multi-channel probes remains challenging. A significant concern is finding a trajectory for probe insertion that avoids large blood vessels, such as the superior sagittal venous sinus and the transverse venous sinus. Injuring these large veins can cause extensive bleeding, damage to the underlying brain tissue, and potentially death. This approach describes targeting brainstem structures by coupling anterior coordinates with an angled approach, allowing the recording probe to penetrate the brain below high-risk vascular structures. Compared to a strictly vertical approach, the angled approach maximizes the number of brain regions that can be targeted. Using this strategy, the ventrolateral periaqueductal gray (vlPAG), a brainstem region associated with REM sleep, can be reproducibly and reliably accessed to obtain single-unit, multi-electrode recordings in head-fixed mice before and during sevoflurane anesthesia. The ability to record neuronal activity in the vlPAG and surrounding nuclei with high temporal resolution is a step forward in advancing the understanding of the relationship between REM sleep and anesthesia.

Five male C57BL/6J were implanted with an ECoG headset and headplate (Figure 4A). After recovery, mice were habituated to head-fixation and the electrophysiology recording rig during two 1.5 h sessions on separate days (Figure 4B). Next, a 2 mm x2 mm craniotomy window was created (Figure 4C) and a silicon probe was inserted with the mouse awake and head-fixed (Figure 4D). Two types of silicon UCLA probe...

Author Spotlight: Investigating Anesthesia-Induced Sleep Pathways and Neuronal Excitability in Mice

This protocol describes a method to obtain in vivo, high-density single-neuron recordings from the brainstem of head-fixed mice. This approach is deployed to measure the action potential firing of neurons in the ventrolateral periaqueductal gray - a brainstem region inactive during Rapid Eye Movement (REM) sleep - before and during general anesthesia.

Acute Single-Unit Multi-Electrode Recordings from the Brainstem of Head-Fixed Mice

Silicon multielectrode-based recordings are increasingly popular for studying neuronal activity at the temporal resolution of action potentials in many ...

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Neuroscience

640 Views.  University of Virginia. Ons laboratorium onderzoekt de effecten van veelgebruikte anesthetica op slaapgerelateerde neuronen en hun communicatiepaden. We streven ernaar beter te begrijpen hoe anesthetica de endogene slaappaden beïnvloeden en de perioperatieve slaap en cognitie beïnvloeden. Onlangs hebben we de anatomische substraten geïdentificeerd voor enkele van de slaapveranderingen die worden veroorzaakt door bepaalde soorten algemene anesthetica.We deden dit door genet...

Video: Auteur in de schijnwerpers: onderzoek naar door anesthesie geïnduceerde slaappaden en neuronale prikkelbaarheid bij muizen

High-density EEG Recordings of the Freely Moving Mice using Polyimide-based Microelectrode

Electroencephalogram (EEG) indicates the averaged electrical activity of the neuronal populations on a large-scale level. It is widely utilized as a noninvasive brain monitoring tool in cognitive neuroscience as well as a diagnostic tool for epilepsy and sleep disorders in neurology. However, the underlying mechanism of EEG rhythm generation is still under the veil. Recently introduced polyimide-based microelectrode (PBM-array) for high resolution mouse EEG1 is one of the trials to answer the neurophysiological questions on EEG signals based on a rich genetic resource that the mouse model contains for the analysis of complex EEG generation process. This application of nanofabricated PBM-array to mouse skull is an efficient tool for collecting large-scale brain activity of transgenic mice and accommodates to identify the neural correlates to certain EEG rhythms in conjunction with behavior. However its ultra-thin thickness and bifurcated structure cause a trouble in handling and implantation of PBM-array. In the presented video, the preparation and surgery steps for the implantation of PBM-array on a mouse skull are described step by step. Handling and surgery tips to help researchers succeed in implantation are also provided.

In this article, we described the surgery procedure and handling tips for implantation of ultra-thin polyimide-based microelectrode array (PBM-array) on the mouse skull for acquisition of high-density encephalography (EEG) in a mouse model.

Electroencephalogram (EEG) indicates the averaged electrical activity of the neuronal populations on a large-scale level. It is widely utilized as a ...

Preparation of an Awake Mouse for Recording Neural Responses and Injecting Tracers

It is well known that anesthesia alters neural response properties in various regions of the brain.13. In the auditory system, fundamental response properties of brainstem neurons including threshold, frequency specificity, and inhibitory sidebands are altered in significant ways under anesthesia1-2. These observations prompted physiologists to seek ways to record from single neurons without the contaminating effects of anesthesia. One result was a decerebrate preparation, where the brainstem was completely transected at the level of the midbrain4. The drawbacks of this preparation are a formidable surgery, the elimination of descending projections from the forebrain, and an inability to use sensory stimulation to examine structures above the midbrain. A different strategy has been to implant electrode arrays chronically to record from single neurons and multiunit clusters while the animal is awake and/or behaving5,6. These techniques however are not compatible with injecting tracer dyes after first electrophysiologically characterizing a brain structure. To avoid altering neural response properties with anesthetics while recording electrophysiological response properties from single neurons, we have adapted a head restraint technique long used in bats7-9 to mouse10-12. Using this method, we are able to conduct electrophysiological recordings over several days in the unanesthetized mouse. At the end of the recording sessions, we can then inject a dye to reconstruct electrode positions and recording sites or inject a tracer so that pathways to and from the recording loci can be determined. This method allows for well isolated single neuron recordings over multiple days without the use anesthetics.

Electrophysiological characterization of neuronal responses is important for understanding brain function and for guiding the placement of dyes for pathway tracing. However, many studies are performed in anesthetized animals. To understand brain function without anesthetics, we developed a method to record neuronal response properties and inject dyes in awake mouse.

It is well known that anesthesia alters neural response properties in various regions of the brain.13.  In the auditory system, fundamental response ...

Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and awake animals. Therefore, methods allowing the manipulation of synaptic systems in awake animals are required in order to determine the contribution of synaptic inputs to neuronal processing unaffected by anesthetics. Here, we present methodology for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. By combining these procedures, we performed microiontophoretic injections of gabazine to selectively block GABAA receptors in neurons of the inferior colliculus of head-restrained mice. Gabazine successfully modified neural response properties such as the frequency response area and stimulus-specific adaptation. Thus, we demonstrate that our methods are suitable for recording single-unit activity and for dissecting the role of specific neurotransmitter receptors in auditory processing.
The main limitation of the described procedure is the relatively short recording time (~3 hr), which is determined by the level of habituation of the animal to the recording sessions. On the other hand, multiple recording sessions can be performed in the same animal. The advantage of this technique over other experimental procedures used to manipulate the level of neurotransmission or neuromodulation (such as systemic injections or the use of optogenetic models), is that the drug effect is confined to the local synaptic inputs to the target neuron. In addition, the custom-manufacture of electrodes allows adjustment of specific parameters according to the neural structure and type of neuron of interest (such as the tip resistance for improving the signal-to-noise ratio of the recordings).

We present methods for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. This technique allows the detailed analysis of putative local synaptic inputs to the neuron of interest.

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and ...

High-density Electroencephalographic Acquisition in a Rodent Model Using Low-cost and Open-source Resources

Advanced electroencephalographic analysis techniques requiring high spatial resolution, including electrical source imaging and measures of network connectivity, are applicable to an expanding variety of questions in neuroscience. Performing these kinds of analyses in a rodent model requires higher electrode density than traditional screw electrodes can accomplish. While higher-density electroencephalographic montages for rodents exist, they are of limited availability to most researchers, are not robust enough for repeated experiments over an extended period of time, or are limited to use in anesthetized rodents.1-3 A proposed low-cost method for constructing a durable, high-count, transcranial electrode array, consisting of bilaterally implantable headpieces is investigated as a means to perform advanced electroencephalogram analyses in mice or rats.

Procedures for headpiece fabrication and surgical implantation necessary to produce high signal to noise, low-impedance electroencephalographic and electromyographic signals are presented. While the methodology is useful in both rats and mice, this manuscript focuses on the more challenging implementation for the smaller mouse skull. Freely moving mice are only tethered to cables via a common adapter during recording. One version of this electrode system that includes 26 electroencephalographic channels and 4 electromyographic channels is described below.

Instructions for the low-cost construction and surgical implantation of a chronic transcranial high-density electroencephalographic montage into mice are provided. Signal recording, extraction, and processing techniques are also described.

Advanced electroencephalographic analysis techniques requiring high spatial resolution, including electrical source imaging and measures of network ...

Computer-based Multitaper Spectrogram Program for Electroencephalographic Data

Current web resources provide limited, user friendly tools to compute spectrograms for visualizing and quantifying electroencephalographic (EEG) data. This paper describes a Windows-based, open source code for creating EEG multitaper spectrograms. The compiled program is accessible to Windows users without software licensing. For Macintosh users, the program is limited to those with a MATLAB software license. The program is illustrated via EEG spectrograms that vary as a function of states of sleep and wakefulness, and opiate-induced alterations in those states. The EEGs of C57BL/6J mice were wirelessly recorded for 4 h after intraperitoneal injection of saline (vehicle control) and antinociceptive doses of morphine, buprenorphine, and fentanyl. Spectrograms showed that buprenorphine and morphine caused similar changes in EEG power at 1&#8722;3 Hz and 8&#8722;9 Hz. Spectrograms after administration of fentanyl revealed maximal average power bands at 3 Hz and 7 Hz. The spectrograms unmasked differential opiate effects on EEG frequency and power. These computer-based methods are generalizable across drug classes and can be readily modified to quantify and display a wide range of rhythmic biological signals.

This protocol provides an open source, compiled MATLAB program that generates multitaper spectrograms for electroencephalographic data.

Current web resources provide limited, user friendly tools to compute spectrograms for visualizing and quantifying electroencephalographic (EEG) data. ...

Extracellular Electrophysiological Recording in the Motor Cortex of Mice

Multichannel Extracellular Recording in Freely Moving Mice

The protocol aims to uncover the properties of neuronal firing and network local field potentials (LFPs) in behaving mice carrying out specific tasks by correlating the electrophysiological signals with spontaneous and/or specific behavior. This technique represents a valuable tool in studying the neuronal network activity underlying these behaviors. The article provides a detailed and complete procedure for electrode implantation and consequent extracellular recording in free-moving conscious mice. The study includes a detailed method for implanting the microelectrode arrays, capturing the LFP and neuronal spiking signals in the motor cortex (MC) using a multichannel system, and the subsequent offline data analysis. The advantage of multichannel recording in conscious animals is that a greater number of spiking neurons and neuronal subtypes can be obtained and compared, which allows the evaluation of the relationship between a specific behavior and the associated electrophysiological signals. Notably, the multichannel extracellular recording technique and the data analysis procedure described in the present study can be applied to other brain areas when conducting experiments in behaving mice.

Author Spotlight: Advancements in Multichannel Extracellular Recording for Studying Neuronal Activity in Freely Moving Mice 

The protocol describes the methodology of extracellular recording in the motor cortex (MC) to reveal extracellular electrophysiological properties in freely moving conscious mice, as well as the data analysis of local field potentials (LFPs) and spikes, which is useful for evaluating the network neural activity underlying behaviors of interest.

The protocol aims to uncover the properties of neuronal firing and network local field potentials (LFPs) in behaving mice carrying out specific tasks by ...

Stimulating Mouse Brain with a Low-Cost EEG and Millimeter-Sized Coil

Low-Cost Electroencephalographic Recording System Combined with a Millimeter-Sized Coil to Transcranially Stimulate the Mouse Brain In Vivo

A low-cost electroencephalographic (EEG) recording system is proposed here to drive transcranial magnetic stimulation (TMS) of the mouse brain in vivo, utilizing a millimeter-sized coil. Using conventional screw electrodes combined with a custom-made, flexible, multielectrode array substrate, multi-site recording can be carried out from the mouse brain. In addition, we explain how a millimeter-sized coil is produced using low-cost equipment usually found in laboratories. Practical procedures for fabricating the flexible multielectrode array substrate and the surgical implantation technique for screw electrodes are also presented, which are necessary to produce low-noise EEG signals. Although the methodology is useful for recording from the brain of any small animal, the present report focuses on electrode implementation in an anesthetized mouse skull. Furthermore, this method can be easily extended to an awake small animal that is connected with tethered cables via a common adapter and fixed with a TMS device to the head during recording.The present version of the EEG-TMS system, which can include a maximum of 32 EEG channels (a device with 16 channels is presented as an example with fewer channels) and one TMS channel device, is described. Additionally, typical results obtained by the application of the EEG-TMS system to anesthetized mice are briefly reported.

Author Spotlight: Low-Cost Electroencephalographic Recording System Combined with a Millimeter-Sized Coil to Transcranially Stimulate the Mouse Brain In Vivo  

A low-cost electroencephalographic recording system combined with a millimeter-sized coil is proposed to drive transcranial magnetic stimulation of the mouse brain in vivo. Using conventional screw electrodes with a custom-made, flexible, multielectrode array substrate, multi-site recording can be carried out from the mouse brain in response to transcranial magnetic stimulation.

A low-cost electroencephalographic (EEG) recording system is proposed here to drive transcranial magnetic stimulation (TMS) of the mouse brain in vivo, ...

Fabrication and Surgical Implantation of Electromyographic Electrodes in Mice

Recording Forelimb Muscle Activity in Head&#45;Fixed Mice with Chronically Implanted EMG Electrodes

Powerful genetic and molecular tools available in mouse systems neuroscience research have enabled researchers to interrogate motor system function with unprecedented precision in head-fixed mice performing a variety of tasks. The small size of the mouse makes the measurement of motor output difficult, as the traditional method of electromyographic (EMG) recording of muscle activity was designed for larger animals like cats and primates. Pending commercially available EMG electrodes for mice, the current gold-standard method for recording muscle activity in mice is to make electrode sets in-house. This article describes a refinement of established procedures for hand fabrication of an electrode set, implantation of electrodes in the same surgery as headplate implantation, fixation of a connector on the headplate, and post-operative recovery care. Following recovery, millisecond-resolution EMG recordings can be obtained during head-fixed behavior for several weeks without noticeable changes in signal quality. These recordings enable precise measurement of forelimb muscle activity alongside in vivo neural recording and/or perturbation to probe mechanisms of motor control in mice.

Author Spotlight: Investigating Mouse Motor Cortex Interactions from Muscle Activity to Neural Dynamics

This protocol describes the hand fabrication and surgical implantation of electromyographic (EMG) electrodes in the forelimb muscles of mice to record muscle activity during head-fixed behavior experiments.

Powerful genetic and molecular tools available in mouse systems neuroscience research have enabled researchers to interrogate motor system function with ...

Injectable Mesh Electronics for Stable Single-Neuron Recordings in a Mouse Brain

Source: Schuhmann Jr., T. G., et al. <a href="https://app.jove.com/v/58003">Syringe-injectable Mesh Electronics for Stable Chronic Rodent Electrophysiology.</a> J. Vis. Exp. (2018).
The video demonstrates the injection of mesh electronics probes into a mouse brain for stable long-term recordings at a single-neuron level. An anesthetized mouse with an exposed skull is secured on a stereotaxic frame, and the probe&#39;s mesh device region is injected into the brain through drilled holes. The probe&#39;s input-output pads are connected to a circuit, and recordings are obtained using a data acquisition system.

1. Stereotaxic Injection of Mesh Electronics into Live Mouse Brain
NOTE: Mice were anesthetized by intraperitoneal injection with a mixture of 75 mg/kg ...

JoVE Encyclopedia of Experiments

Neurophysiology

Stimulation and Modulation Techniques

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

Source: Sauer, J., et al. <a href="https://app.jove.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.

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

Acute Single-Unit Multi-Electrode Recordings from the Brainstem of Head-Fixed Mice

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