Jan 16, 2026
Acute LC Craniotomy and Skull Thinning for Neuropixels Recordings and Optogenetic Stimulation
- Margaret Lee1,
- Kanghoon Jung1,
- Zhixiao Su1
- 1Allen Institute for Neural Dynamics
- Allen Institute for Neural Dynamics
Protocol Citation: Margaret Lee, Kanghoon Jung, Zhixiao Su 2026. Acute LC Craniotomy and Skull Thinning for Neuropixels Recordings and Optogenetic Stimulation. protocols.io https://dx.doi.org/10.17504/protocols.io.eq2ly6o6qgx9/v1
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: February 26, 2025
Last Modified: January 16, 2026
Protocol Integer ID: 123471
Keywords: craniotomy, Neuropixels, electrophysiology, neuromodulation, surgery, mouse surgery, acute lc craniotomy, anterior portion of cerebellum, cranial window in both hemisphere, skull thinning for neuropixels recording, acute craniotomy procedure, cerebellum, cranial window, exposed inferior colliculus, skull thinning, antidromic laser stimulation, regions of cortex, optogenetic stimulation this protocol, optogenetic stimulation, anterior portion, neuropixel, neuropixels recording, sagittal sinus, hemisphere, thinning procedure, skull, cortex, surrounding vasculature, locus coeruleus
Abstract
This protocol details the acute craniotomy procedure to target locus coeruleus (LC) in vivo with a Neuropixels 2.0 single shank probe, and includes the skull thinning procedure to better expose regions of cortex for antidromic laser stimulation. Given that LC is a small structure that is difficult to target precisely, leaving the skull and surrounding vasculature intact allows for more accurate targeting. The surgery should result in a cranial window in both hemispheres with an exposed inferior colliculus, through which LC is targeted, as well as an anterior portion of cerebellum. The sagittal sinus should also be relatively clear in order to orient the probe in relation to all of the aforementioned structures.
Image Attribution
Gabriel Rodriguez, Allen Institute
Guidelines
This procedure should only be performed in accordance with IACUC and veterinary requirements.
Materials
Surgical Tools and Supplies:
| Tool/Supply | Manufacturer/Supplier | Part Number | |
| Dumont #5 45° forceps | Fine Science Tools | 11251-35 | |
| Vanna scissors | World Precision Instruments | 500260 | |
| 45° or 90° Duratomy probe | Fine Science Tools | 10066-15 | |
| Alcohol swabs | Becton Dickinson Medical Supplies | 326895 | |
| Hemostatic Agent Surgifoam | McKesson Corporation | 403360 | |
| Sterile gauze 3×3" | Patterson Veterinary | 07-893-8587 | |
| Kimwipes | Fisher Scientific | 06-666 | |
| Sugi pointed sterile swabs | Fine Science Tools | 18105-01 | |
| Insulin syringes, U-100, 1mL, 31G | Advantor | BD328418 | |
| Sterile Drill Bits FG4 (coarse drill bit) | NeoBurr | 1734214 | |
| Sterile Drill Bits FG1 (fine drill bit) | NeoBurr | 1734948 | |
| Artificial Cerebrospinal Fluid.V | Made in-house | ||
| Well with grounding pin and 3D printed cap | Made in-house | ||
| Duragel | Ellsworth Adhesives | DOW DOWSIL™ 3-4680 | |
| 5mm microruler | TDI International | TDI MRM100H | |
| C Universal 4-META Catalyst, 0.7mL | Parkell | S371 | |
| B Quick Base for Metabond, 10mL | Parkell | S398 | |
| Radiopaque L-Powder, white, 5g | Parkell | S396 | |
| Superglue | Krazy Glue | Amazon PK4 KG58248SN |
All tools/supplies can be substituted with their equivalent.
Equipment:
| A | B | C | D | |
| Small Animal Stereotaxic Instrument | Kopf | 1900 | ||
| Adjustable Stage Platform | Kopf | 901 | ||
| Stereo Microscope | Leica | M80 | ||
| Gooseneck Illumination | AM Scope | LED-6WA | ||
| On-axis Illumination | Leica | KL2500 LED | ||
| Small Animal Temperature Control System | CWE Inc. | TC-1000 | ||
| Large Heat pad | Lectro-Kennel | Outdoor Heated Pet Pad | ||
| Dental Drill | NSK | Pana-Max2 M4 | ||
| Oxygen Concentrator | Nidek Medical Products | Nuvo Lite Model 525 | ||
| Isoflurane with oxygen delivery system | Patterson Scientific | Tec 3 EX | ||
| Isoflurane induction chamber | Patterson Scientific | 78933385 | ||
| Headframe clamp | Made in-house | |||
Troubleshooting
Safety warnings
Personal Protective Equipment (PPE) should be worn at all times while operating this protocol.
Isoflurane warning: Acute over-exposure to waste anesthetic gases (WAG) may cause eye irritation, headache, nausea, drowsiness or dizziness. Repeated exposure may cause damage to cardiovascular system and central nervous system. Refer to MSDS for additional information. Consult the surgical workstation guide to ensure all parts of the dispensation rig are functioning properly.
Ethics statement
Research focused rodent neurosurgery must be conducted according to internationally-accepted standards and should always have prior approval from an Institutional Animal Care and Use Committee (IACUC) or equivalent ethics committee(s).
This protocol has been approved by the Allen Institute Animal Care and Use Committee (IACUC).
PHS Assurance: D16-00781
AAALAC: Unit 1854
Before start
This surgery is a secondary procedure that should only be performed after the headframe has been attached and necessary viruses have been injected during the Stereotactic Injections with Headframe Implant procedure. Allow the mouse at least 2 weeks to recover from initial surgery before attempting this procedure.
Preparation of surgical rig
Reference following protocol for surgical rig setup:
Begin at step 1 "Disinfect the surgical area"
Stop at step 3.5 "Remove autoclaved surgery tools"
Anesthetization and preparation of animal
Reference following protocol for anesthetization of animal:
Begin at step 25 "Prepare the anesthesia system"
Stop at step 30 "Administer any drugs"
Secure mouse to surgical rig via the headframe attached during initial surgery.
Swab the exterior of the clear Metabond with an Alcohol swab.
Skull thinning procedure
Perform skull thinning procedure to enable sufficient light transmission for later optogenetic antidromic stimulation. For all regions of interest (ROIs), thin the skull over both hemispheres.
Using a 5 mm micro ruler, measure the regions of interest (prelimbic cortex, primary somatosensory cortex, and primary visual cortex) from the bregma marking made in the previous surgery.
Note
These measurements indicate distance from bregma and are consistent across all mice.
Prelimbic cortex (PrL): AP: 1.8, ML: ±0.5
Primary somatosensory cortex (S1): AP: -0.5, ML: ±2.35
Primary visual cortex (V1): AP: -3.3, ML: ±2
Using a dental drill with the coarse drill bit, remove the layer of Metabond that was applied during the previous surgery.
Continue to drill a 1.5 mm area at all ROIs until the blood vessels are easily visible but ensure that the skull remains intact.
Fig. 1 The black dot in the image denotes bregma as it is marked in initial surgery. The gray dashed circles denotes the thinned skull regions.
Perform craniotomy
Perform acute craniotomy at LC on both hemispheres.
Note
It is recommended to flush the surface of the skull with ACSF before beginning the craniotomy for better visibility of surrounding vasculature (such as the sagittal sinus) and to remove the Metabond debris from previous steps. This will result in a surgical area that has better clarity as illustrated in Fig. 2.
Using a 5 mm micro ruler measure the location of LC from bregma and drill at measured location to thin the skull covering both hemispheres. Expose enough of the skull to see where the sagittal sinus is located. Most of the craniotomy should be over the inferior colliculus (IC) for accurate targeting and include enough of the cerebellum to see where the boundary of the IC ends.
Note
For male mice, coordinates from bregma are AP: -5.4, ML: ±0.85
For female mice, coordinates from bregma are AP: -5.2, ML: ±0.85
Fig. 2 The black dot in image denotes bregma as it is marked in initial surgery. The SS denotes the sagittal sinus that runs along the edge of where the craniotomy will take place.
Flush the area with several drops of ACSF such that the entire craniotomy is covered. Remove the ACSF with enough Sugi spears to soak up the liquid. This ensures all debris is removed before performing craniotomy.
Using a fine drill bit, drill a 1 mm skull island over both LC regions.
Once there are visible cracks around the skull island and some CSF has started leaking out, cover the entire area with ACSF and, using Dumont forceps, carefully pry up the skull at multiple points to ensure the skull island is sufficiently separated. If blood starts leaking out, stop immediately and staunch the bleeding with Sugi spears and surgifoam soaked in ACSF.
Remove the skull island using Dumont forceps and ensure all bone fragments are removed. To minimize bleeding, tilt the bone fragment away from the sagittal sinus when detaching from rest of skull.
Rinse the area with ACSF and gently swab area with surgifoam soaked in ACSF to clean the craniotomy.
Perform duratomy
Perform the duratomy.
Make an initial cut in the dura using a Duratomy probe or a sterile 1 mL Insulin syringe, such that there is a flap that can be peeled back using forceps or lifted off the brain with the Duratomy probe.
Lift up the flap of dura using Dumont forceps and pull with enough force to remove the dura covering the brain. Alternatively, trim away with Vanna scissors.
Fig. 3 In above image, IC denotes inferior colliculus and C denotes cerebellum.
Attach well
Place well onto the headframe.
Note
This step may be omitted if well was attached with the headframe.
Apply a thin layer of Superglue to bottom of the well and attach to the headframe.
Secure well with Metabond inside the headframe and around the outside of the headframe. Ensure the well does not leak by filling it with ACSF and checking for leaks.
Remove the ACSF with a Kimwipe or cotton tipped applicators and ensure inside of well is dry.
Using a fine drill bit, drill a small burr hole (<0.2 mm) until the brain surface is exposed and some CSF leaks out.
Note
Depending on what type of headframe is used, the location of the grounding hole will differ. For the whole hemisphere headframe, place the grounding hole at anterior right hemisphere and for the dual hemisphere headframe place the grounding hole at posterior left hemisphere. Drill the hole away from any major blood vessels or brain regions of interest.
Gently place grounding pin in the small burr hole such that it is secure but is not depressing the brain tissue or causing bleeding.
Fig. 4 Final product of this procedure with 6 skull thinned areas, 2 craniotomies, and the grounding pin (GP) inserted into the skull.
Mix the Duragel in a sterile weigh boat with the 1 mL insulin needle in a 1:2 ratio of Duragel A and Duragel B. Scrape up a drop of the mixed Duragel with the needle and apply a thin layer to both craniotomies, the thinned skull regions, and the hole for the grounding pin. Ensure there is no exposed brain tissue.
Attach a well cap to the well.
Take down of surgical rig
Turn off the isoflurane vaporizer and remove mouse from surgical rig.
Turn off vacuum and oxygen sources by switching the stopcocks to off position.
Obtain postoperative weight of animal and return mouse to cage. Keep the cage on the 37°C heating pad until mouse regains consciousness and resumes normal behavior. Return mouse to vivarium.
Note
Follow post-operative veterinary guidelines.
Take down the surgical rig once surgery is complete as described in General Setup and Takedown Procedures for Rodent Neurosurgery V.2