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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>
			<td></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>
			<td></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>
			<td></td>
		</tr>
		<tr>
			<td>C&#38;B metabond</td>
			<td>Patterson Dental, Richmond, Virginia, USA</td>
			<td>powder: 5533559, quick base: 5533492, catalyst: 55335007</td>
			<td>Headplate &#38;Headset Implantation</td>
			<td></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>
			<td></td>
		</tr>
		<tr>
			<td>Capnomac Ultima</td>
			<td>Datex, Helsinki, Finland&#160;</td>
			<td>ULT-SVi-27-07</td>
			<td>Gas Analyzer; discontinued; alternative gas analyzer can be purchased from Bionet America&#160;</td>
			<td></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>
			<td></td>
		</tr>
		<tr>
			<td>Connector Header</td>
			<td>DigiKey, Thief River Falls, Minnesota, USA</td>
			<td>1212-1788-ND</td>
			<td>ECoG Headset</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI Fluoromount-G</td>
			<td>SouthernBiotech, Birmingham, Alabama, USA</td>
			<td>0100-20</td>
			<td>Histology</td>
			<td></td>
		</tr>
		<tr>
			<td>iBOND Universal</td>
			<td>Patterson Dental, Richmond, Virginia, USA</td>
			<td>044-1113</td>
			<td>Headplate &#38;Headset Implantation; for&#160; securing stainless steel wires to the skull</td>
			<td></td>
		</tr>
		<tr>
			<td>Low toxicity silicon adhesive</td>
			<td>World Precision Instruments, Sarasota, Florida, USA</td>
			<td>KWIK-SIL</td>
			<td>Headplate</td>
			<td></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>
			<td></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>
			<td></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>
			<td></td>
		</tr>
		<tr>
			<td>Normal saline</td>
			<td>ThermoFisher Scientific, Waltham, Massachusetts, USA</td>
			<td>Z1376</td>
			<td>Headplate &#38;Headset Implantation; preventing the brain from drying during the surgery</td>
			<td></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 &#38; reference electrode for ECoG&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Platinum wire 24AWG&#160;</td>
			<td>World Precision Instruments, Sarasota, Florida, USA</td>
			<td>PTP201</td>
			<td>Reference electrode for the silicon probe recording&#160;</td>
			<td></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>
			<td></td>
		</tr>
		<tr>
			<td>Stainless steel Headplate</td>
			<td>Star Rapid, China</td>
			<td>custom made part</td>
			<td>Headplate &#38;Headset Implantation; design available upon request</td>
			<td></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 &#38;Headset Implantation</td>
			<td></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.

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

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

Our laboratory investigates the effects of commonly used anesthetics on sleep-related neurons and their communication pathways. We aim to better understand how anesthetics impact endogenous sleep pathways and affect perioperative sleep and cognition. Recently we identified the anatomical substrates for some of the sleep changes caused by certain types of general anesthetics.We did so by using genetically modified mice in which there is endogenous expression of c-Fos, a marker of neuronal activation. However, using this new technique, we will be able to directly measure cellular excitability in groups of neurons in the brainstem that are associated with sleep. This is a major advantage over simply quantifying c-Fos expression as a surrogate marker of neuronal activation.Our next step will be to pursue the cellular and molecular mechanisms underlying the sleep changes induced by certain types of general anesthetics. This will entail measuring cellular excitability in brainstem neurons associated with sleep using neuropixel probes, as well as using spatial transcriptomic approaches to quantify and map the expression of genes related to sleep in these areas.

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...

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.

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

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Research

JoVE Journal

Neuroscience

Electrocorticography Headplate Implantation and Electrode Placement for Neuronal Activity Recording

To begin, make an electrocorticography or ECoG headset by soldering perfluoroalkoxy-coated stainless steel wire to a three-pin connector header. After identifying the insertion coordinates based on a mouse stereotaxic atlas, position the anesthetized mouse in the stereotaxic frame. Place the animal's nose in the nose cone and stabilize the head with head bars.Then perform a toe pinch test to check the depth of anesthesia. After making a five-millimeter scalp incision above the parietal and occipital bones, scrape the meninges with the scalpel blade to remove them. Use the scalpel blade to cut muscle attachments and expose the parietal and occipital bones.Apply hydrogen peroxide to control bleeding and dry the skull surface. Next, identify the bregma and lambda landmarks on the skull. Then adjust the nose cone position to level the anterior-posterior position of the skull.To level the medial lateral position of the skull, pick two opposite points between bregma and lambda and check their level. Now, measure the distance between bregma and lambda and compare it to the distance reported in the Franklin Paxinos Stereotaxic Atlas. Use the difference between the measured and reported distances to scale the anterior-posterior coordinate proportionally.Mark craniotomy coordinates on the skull with a sterilized pencil. Using a stereotaxic micro manipulator, position the headplate directly on top of the lambda suture. Apply dental cement to the headplate and around it to secure it to the skull and allow it to dry for 10 minutes.Then drill burr holes for two cortical electrodes and one reference electrode. Place the stripped ends of coated silver wire electrodes within the burr holes and secure them using ultraviolet light activated resin. 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 to ensure it is firmly in place and allow the mouse to recover for seven days.

Silicon Probe Placement and Single Unit Recording from the Brainstem of Head-Fixed Mice

Position the anesthetized mouse in a stereotaxic frame. Identify bregma and lambda, ensuring no more than a 100 micrometer height difference between the two landmarks. Find and mark the calculated coordinates on the skull with a sterile pencil.Around the coordinates, create an outline of the two millimeter by two millimeter craniotomy window. After checking the depth of anesthesia, use a high-speed drill to create a two millimeter by two millimeter craniotomy window. Apply 0.5 to 1 milliliter of normal saline to prevent the brain surface from drying.Remove the dura using a syringe needle and fine forceps. Next, use a high-speed drill to create a separate burr hole for the silicon probes reference electrode, generally one to two millimeters from the cranial window. Apply 0.2 milliliters of low toxicity silicon adhesive on the skull to completely seal the craniotomy.Using the head plate and screws, affix the head of the mouse to the electrophysiology recording rig. Then coat the silicon probe shank with fluorescent dye so that the probe trajectory can be reconstructed after the experiment. After that, mount the probe on the manipulator and set the desired angle.To lower the recording probe to the brain surface within the center of the cranial window, manually insert the probe to a depth of approximately 300 micrometers. Once inserted to this depth, slowly lower the probe automatically to 200 micrometers per minute to the targeted depth to minimize tissue damage. Finally, apply mineral oil to the brain surface within the craniotomy window to prevent drying.Then record data from the silicon probe and ECOG at 30 kilohertz using an Intan recording controller. Most recorded neurons decreased their firing during sevoflurane anesthesia. VLPAG firing significantly decreased from baseline to during sevoflurane.This decrease was consistent across all VLPAG neurons.

402 Views.  University of Virginia.  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.

Angled Silicon Multielectrode Recordings for Deep Brainstem Structures in Mice

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