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

Summary

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.

Abstract

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.

Introduction

Silicon multielectrode-based recordings are becoming increasingly popular to measure neuronal activity across many brain regions with single action potential resolution^1,2,3,4. 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 devices^5,6. Moreover, chronic implantation of these electrodes allows for long-term recordings^7,8.

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 death^9,10. 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 anteriorly^10,11.

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 anesthesia^12,13. Previous studies have used electrophysiological local field potential (LFP) measurements of the vlPAG in awake mice to identify neural state changes associated with anesthesia^14,15. 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.

Protocol

All studies were approved by the Institutional Animal Care and Use Committee at the University of Virginia (Charlottesville, Virginia). Five male C57BL/6J mice, age 3-7 months, weighing 25-30 g, were used. The details of the reagents and the equipment used here are listed in the Table of Materials.

1. Headplate and headset implantation

1. Make an ECoG headset by soldering perfluoroalkoxy (PFA) coated stainless steel wire to a 3-pin connector header (Figure 2A).
2. 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° angle (AP).
3. 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's nose in the nose cone and stabilizing the head with head bars.
4. Apply ophthalmic ointment to the eyes to prevent corneal damage. Use a temperature control system to maintain body temperature at 37 °C.
5. Apply hair removal cream or shave fur over the scalp, then disinfect with povidone iodine and 70% alcohol. Use the 'tips only' aseptic technique to maintain a sterile surgical field.  
6. Perform a toe pinch test to check the depth of anesthesia.
7. Administer analgesia: carprofen 2.5 mg/kg, given subcutaneously in the back.
8. 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.
9. 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.
10. 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 µm height difference exists between the two landmarks.
11. 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's more than 100 µm height difference between them, adjust the medial-lateral position of the skull by manipulating the head bars.
12. 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.
13. 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.
14. 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.
15. 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).
16. 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.

2. Silicon probe placement and recording

1. Habituate the mouse to the recording rig and head fixation (at least 1.5 h on two separate days) (Figure 2D).
2. On the day of the recording, place the mouse in the anesthesia chamber (Isoflurane 1.5%-3% in oxygen).
3. Position the mouse in a stereotaxic frame, adjust nose cone and head bars as described in 1.3. Maintain anesthesia (Isoflurane 1.5%-3% in oxygen) throughout the stereotaxic surgery. Perform a toe pinch test to check the depth of anesthesia.
4. Apply ophthalmic ointment to the eyes and maintain adequate body temperature.
5. Identify bregma and lambda, ensuring that no more than 100 µm height difference exists between the two anatomical landmarks.
6. 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.
7. Perform a toe pinch test to check anesthesia depth.
8. 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).
9. Use a high-speed drill to create a separate burr hole for the silicon probe's reference electrode, generally ~1-2 mm from the cranial window.
10. Apply 0.2 mL low toxicity silicon adhesive on the skull to completely seal the craniotomy, using the attached applicator.
11. Let the mouse recover for approximately 1 h.
12. Affix the head of the mouse to the electrophysiology recording rig using the headplate and screws (Figure 2D).
13. 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.
14. Mount the probe on the manipulator and set the desired angle. To target the vlPAG, an AP angle of 15-20° was applied.
15. 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 µm. Once inserted to this depth, slowly lower the probe automatically (e.g., 200 µm/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.
16. Apply mineral oil to the brain surface within the craniotomy window to prevent from drying.
17. Let the recording probe settle for 10 min after insertion.
18. Record data from the silicon probe and ECoG at 30kHz using an Intan Recording Controller.
19. 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's why it is preferable to excise the brain by removing the dorsal parts of the skull.
20. 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.
21. If the perfusion is done correctly the brain should shrink a little, leaving enough space to insert fine scissors in the dorsal part of the foramen magnum. Make one medial cut, one lateral and one contralateral cut, around 1 mm in size, in the occipital bone.
22. 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.
23. Slide a micro spatula under the brain, starting at the olfactory bulbs to scoop the brain out.
24. Once perfused and removed, the brain can be stored in 4% paraformaldehyde at 4 °C for 24-48 h.

3. Histology for probe trajectory reconstruction

1. Section the brain into 70 µm coronal sections using a vibratome.
2. Mount the sections on slides using a DAPI mounting medium that stains cell nuclei. Coverslip and seal the slides with clear nail polish.
3. 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.

4. Electrophysiological data analysis

1. Analyze recorded neural signals from the silicon probe using an off-line detection and automatic sorting software (Kilosort)²⁴.
2. 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.
3. Import single unit data to MATLAB and analyze²³.

Results

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...

Discussion

Brainstem nuclei mediate fundamental functions such as breathing, consciousness, and sleep^26,27,28. The brainstem'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...

Materials

List of materials used in this article

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