Jun 26, 2026

Whole Hemisphere Craniotomy for Electrophysiology

DOI
https://dx.doi.org/10.17504/protocols.io.6qpvr1o8pgmk/v1
Whole Hemisphere Craniotomy for Electrophysiology
  • 1Allen Institute / Neural Dynamics;
  • 2Columbia University
  • Allen Institute for Neural Dynamics
Ali Williford
Icon indicating open access to content
QR code linking to this content
DOI: https://dx.doi.org/10.17504/protocols.io.6qpvr1o8pgmk/v1
Protocol Citation: Avalon Amaya, Corbett Bennett, Benjamin Ouellette, Tamina K Ramirez, Ali Williford, Robyn Naidoo 2026. Whole Hemisphere Craniotomy for Electrophysiology . protocols.io https://dx.doi.org/10.17504/protocols.io.6qpvr1o8pgmk/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: June 16, 2026
Last Modified: June 26, 2026
Protocol  Integer ID: 319282
Keywords: Neurosurgery, Rodent neurosurgery, Craniotomy, Electrophysiology, Cranial implant, Cranial Windowing, SHIELD, In Vivo, injections within the cranial window, whole hemisphere craniotomy for electrophysiology, cranial windowing surgery, whole hemisphere craniotomy, cranial window over the left whole hemisphere, cranial window, craniotomy, stereotactic injection, variations of stereotactic injection, cortical damage, surgical procedure, timing of injection, left whole hemisphere, injection, mouse cortex, stabilizing headframe, electrophysiology, procedure, neural imaging
Abstract
This protocol describes the surgical procedure, instrumentation, and reagents necessary for the installation of a stabilizing headframe and cranial window over the left whole hemisphere of an adult mouse. This protocol should be utilized for neural imaging and experimentation regarding the mouse cortex.

This procedure accommodates for 2 variations of stereotactic injections (Optional 1 and Optional 2). Specifically, this protocol accommodates for injections before and after the craniotomy and durotomy. Timing of injections during cranial windowing surgeries is typically dependent on their coordinates. Injections outside of the cranial window should be performed prior to the craniotomy and durotomy to allow for adequate burr hole sealing. Injections within the cranial window should be performed after the craniotomy and durotomy to minimize cortical damage.
Image Attribution
Gabriel Rodriguez, Allen Institute.
Guidelines
Only perform this procedure in accordance with IACUC and veterinary requirements.

This protocol has been written for review purposes only, and should not be utilized for training. Users of this protocol should be familiar with common vocabulary and technique for rodent neurosurgery.
Materials
Anesthesia and other drugs

IsofluranePatterson VeterinaryCatalog #07-890-8115

1 g DexamethasonebiorbytCatalog #orb134330

Ceftriaxone Injection AmerisourceBergen MWI Animal HealthCatalog # 094311

Lactated Ringers Injection, USP, Preservative-Free, BaxterHenry Schein Animal HealthCatalog #059380

1 g AtropinebiorbytCatalog #orb322218

1 g CarprofenbiorbytCatalog #orb321211

Ethiqa XR Buprenorphine Extended-Release Injectable Suspension for Mice and Rats 1.3mg/mL, 3mLFidelis PharmaceuticalsCatalog #099114

Note
Drugs should only be administered in accordance with IACUC and veterinary requirements. Ensure timing, dosage, and route of administration are accounted for.


Surgical tools and supplies

Tool / SupplyManufacturer / SupplierPart Number
Black handle scissors, ToughCutFine Science Tools14058-11
Scalpel handleFine Science Tools10003-12
Iris forcepsFine Science Tools11064-07
Dumont #5 45° forcepsFine Science Tools11251-35
45° Vanna scissors, 8cmWorld Precision Instruments500260
45° or 90° Durotomy probeFine Science Tools10066-15
Plastic sterilization containerFine Science Tools20810-02
large iris forcepsFisher Scientific13-820-073
PREempt Disinfectant sprayMcKesson Corporation21101
70% Ethanol (Diluted in-house)Sigma Aldrich 459836
Alchohol wipesBecton, Dickinson and Company326895
Sterile Surgical Drape, 18x26Fisher ScientificNC9517505
Sterile Multi-well plate, 24 wellAvantor29443-952
Nair hair removal creamArm & Hammer40002957
Betadine Solution 10%McKesson Corporation1073829
Hemostatic Agent Surgifoam McKesson Corporation403360
Sterile Gauze, 3x3” squares, (autoclave sterilized)Patterson Veterinary07-893-8587
Cotton swabs, double ended, (autoclave sterilized)Avantor89133-810
Sugi Absorption SpearFine Science Tools18105-01
Insulin syringes, U-100, 0.3 ml, 31GAvantorBD328438
Insulin syringes, U-100, 1 ml, 31G AvantorBD328418
Luer-Lock Syringe, 20 ml ORAvantor53548-025
Luer-Lock Syringe, 10ml Avantor75846-756
25G 5/8-inch needleAvantor89134-134
32 mm Syringe Filter 0.2 µm Supor MembraneAvantor75846-756
Press ‘n’ SealMedlineCLO70441
Saran WrapGLADAmazon B015CLAVU
Sterile Drill Bits, 0.5/0.4, FG1/4 AND/ORNeoBurr1734948
Sterile Drill Bits, 1.4/1.1, FG4 AND/ORNeoBurr1734214
Sterile Drill Bits 1.0/4.2, EF4NeoBurr1730012
Sterile Scalpel blades, #10 ORAvantor21909-378
Sterile Scalpel blades, #11Avantor21909-380
Systane Eye OintmentSystaneAmazon ALCON293787
Artificial Cerebrospinal Fluid.V*Made in-house. Protocol referenced.http://dx.doi.org/10.17504/protocols.io.besjjecn
C Universal 4-META Catalyst, 0.7 ml ParkellS371
B Quick Base for MetaBond, 10 mlParkellS398
Radiopaque L-Powder, white, 5 gmParkellS396
Radiopaque L-Powder, clear 3 gmParkellS399
CS-5R Coverslips, 5 mmWarner Instruments64-0700
Vetbond GluePatterson Veterinary07-805-5031
Superglue, SinglesKrazy GlueAmazon PK4 KG58248SN
3 ml transfer pipette, plasticAvantor52947-970
Ortho-Jet BCA LiquidLang Dental Maufacturing CompanyOrtho-Jet BCA Liquid
Black cement (1) = 4 parts of Ortho-Jet BCA Powder (mixture) ANDLang Dental Manufacturing CompanyOrtho-Jet BCA Liquid
Black cement (2) = 1 part of Powder tempura point, blackJack Richeson & Co 1# Black 62, Amazon B00JGZ8Q1A
Kwik-Sil SealantWorld Precision InstrumentsKWIK-SIL
Kwik-Cast SealantWorld Precision InstrumentsKWIK-CAST
Heat-sterilized Glass pipettes AND/ORDrummond Scientific3-000-203-G/X
Heat-sterilized Glass pipettesWorld Precision Instruments1B120F-4
“Marker” glass pipette, pulled, broken, and Sharpie mark for measuring coordinatesWorld Precision Instruments1B120F-4
Microcapillary Pipette tipsEppendorf89009-310
ParafilmAvantor52858-000
Lightweight Mineral OilSigma-AldrichM8410
30 gauge, 2" Backfilling NeedleDrummond Scientific3-000-027
Sterile Bone WaxCentral Infusion Alliance, Lukens CIA2160287, 901
Sterilization pouchesAvantor89140-804
Loctite 4305Henkel Adhesives303389
All tools / supplies can be substituted with their equivalent.

Key:
AND = Including the tool/supply in row below.
OR = Can use tool/supply in row below instead.
Autoclaved sterilized = Sterilized in-house.
mixture = Mix with tool/supply in row below.
* = Artificial Cerebrospinal Fluid.V


Equipment

Tool / SupplyManufacturer / SupplierPart Number
Small Animal Stereotaxic InstrumentKopf1900
Adjustable Stage PlatformKopf901
Stereo MicroscopeLeciaM80
Gooseneck IlluminationAM ScopeLED-6WA
On-axis IlluminationLeicaKL2500 LED
Bead sterilizerSigma-AldrichZ378585
Small Animal Temperature Control SystemCWE Inc.TC-1000
Large Heat plate/padLectro-KennelOutdoor Heated Pet Pad
Dental DrillNSKPana-Max2 M4
Oxygen ConcentratorNidek Medical ProductsNuvo Lite Model 525
Isoflurane with oxygen delivery systemPatterson ScientificTec 3 EX
Isoflurane induction chamberPatterson Scientific78933385
Ear barsKopf1922
Ultra Fine Point SharpieSharpie37001
Metabond ceramic mixing dishParkellS387
Stylus Pro USB UV PenlightStreamlight66149
Electrode Holder Kopf1970
Galaxy Mini CentrifugeAvantor76269-066
P20 PipettorGilsonF123600
Silver wireStoelting50880
Midgard Precision Current SourceStoelting51595
Nanoject II Variable Volume (2.3 to 69 nL) Automatic Injector ORDrummond Scientific3-000-204
Nanoject III Programmable Nanoliter InjectorDrummond Scientific3-000-207
All equipment can be substituted with their equivalent.

Key:
OR = Can use equipment in row below instead.

Materials/Equipment designed/made in-house (CAD available upon request)

MaterialPart Number
Well Cap0160-055-09
Bregma Stylus0251-900-04
Dovetail Clamp0111-200-00
-0⁰ Ear bar Headframe Clamp0155-110-00
SHIELD Implant Tracer0251-119-00
SHIELD Well0160-055-08
Probe Holder0155-200-00
SHIELD Implant0251-110-42
SHIELD Titanium Headpost0160-073-00
All equipment can be substituted with their equivalent.
https://portal.brain-map.org/explore/toolkit/hardware


Personal Protective Procedures (PPE):

Suggested PPE
Gloves
Disposable lab coat
Disposable face mask
Shoe covers / surgery shoes
Scrubs
Surgical cap
Biohazard sharps disposal container
Biohazard waste disposal container
Blue light blocking glasses
Utilize PPE in accordance with IACUC and veterinary requirements. Ensure sterility when necessary.


Safety warnings
  • Personal Protective Equipment (PPE) should be used 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.

  • Blue-light filter safety goggles must be worn while using LED curing light.
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
Reference General Setup and Takedown Procedures for Rodent Neurosurgery for all setup and takedown procedures.

Headpost and Headframe will be used interchangeably throughout this protocol.
Graphical Overview of Procedure

Map diagram of protocol workflow.

Expose and Prepare the Skull Surface
10m
After hair removal and disinfection, create midline incision with a scalpel blade from approximately behind the eyes to the front of the ears.

Illustration of initial skin incision on mouse head.
Mouse fixed to stereotactic instrument base via bite bar and ear bars. Fur from top of head has been removed and skin has been sterilized (see General Setup and Takedown Procedures).

Using Vanna scissors, cut a round teardrop shape of skin away.

Illustration of skin removal on mouse head.

On the left and right side remove enough skin to see the interface of the skull and cheek muscle. On the posterior side stop as soon as the neck muscle is exposed. Anteriorly, stop at around 1mm past the rostral rhinal vein.
Tear or cut the periosteum and ensure removal at surgical site. Rehydrate with Artificial Cerebral Spinal Fluid (ACSF) if needed to finish removal.

Note
Utilize a 10mL or 20mL syringe, 25G 5/8-inch needle, and a syringe filter to store and dispense ACSF.

Detach muscles to allow more surface area for headframe securing.


Illustration of skin removal boundaries and muscle removal extent.

Detach the cheek muscle on the left side of the skull by poking into the interface between the muscle and skull, and then drag the Dumont forceps along the length of the cheek. Detach the neck muscles with Dumont forceps by scraping the muscle attachments.
On the left side of the skull, pull separated muscle and skin away from the skull as shown in the illustration. At the posterior edge, tuck the detached neck muscles underneath the skin.
Seal all along incision site with Vetbond. Use a Sugi Absorption Spear to absorb any excess fluid either prior or during. Extend Vetbond seal 1-2mm past incision site along healthy skin.
Align The Skull
10m
Locate Bregma and Lambda landmarks with Dovetail Clamp and Bregma Stylus, and use them to level the skull in the anterior-posterior axis within 0.1mm.

Illustration of Bregma and Lambda landmarks.

If Lambda-Bregma offset is greater than 0.1mm in X, use the yaw adjustment on stereotactic alignment system to adjust the yaw to within 0.1mm.

At midline, approximately midway between Bregma and Lambda, measure 2mm laterally on both the left and right hemisphere, ensuring that the skull is level in the medial-lateral axis within 0.15mm.
Optional (1): Perform Stereotactic Injections Outside of the Cranial Window
Optional 1​
This optional section accommodates injections prior to performing the craniotomy and durotomy. Typically, this is done for injection coordinates that do not lie within the craniotomy window and could be obstructed by placement of the headframe. Injection coordinates within the craniotomy window are performed after the craniotomy and durotomy as it will minimize cortical damage. ​
​
Skip this section if you will not be performing injections prior to the craniotomy and durotomy.

Reference protocol Stereotactic Injection by Nanoject Protocol V.6 for performing stereotactic injections via nanoject:
dx.doi.org/10.17504/protocols.io.bp2l6nr7kgqe/v6

Begin at step 8.2.6 "Mark the Injection Site".
Stop at step 8.5 "Suturing".
Reference protocol tereotactic Surgery for Delivery of Tracers by Iontophoresis V.6 for performing stereotactic injections via Iontophoresis:
dx.doi.org/10.17504/protocols.io.14egn8ewzg5d/v6

Begin at step 8.2.6 "Mark the Injection Site".
Stop at step 8.5 "Suturing".
Once completed with the injection, fill burr hole with bone wax with the broken end of a cotton swab.

Note
Do not leave excess wax residue on the skull. Excess wax can contribute to having an insufficient seal between the skull and the headframe.

Optional (2): Mark Stereotactic Injection Coordinates for After Craniotomy and Durotomy
Optional 2​
This optional section accommodates injections that will be performed after the craniotomy and durotomy. This is the first section out of three required. This first section is specifically for identifying and marking injection coordinates while the skull is level. These markings will be referenced in the second optional section (Optional 2.1).​
​
Skip this section if not marking injection coordinates for injections after the craniotomy and durotomy.
Considerations:

For injections within the cranial window, it is important to take the injection depth into consideration. Avoid targeting subcortical brain structures, as deeper insertion of the pipette into the brain tissue can increase risk of edema.

Additionally, maintain Bregma as the fiducial point. Avoid the use of multiple viruses/pipettes as pipettes are not made identically (variance of up to 0.8mm difference).
Reference protocol Stereotactic Injection by Nanoject Protocol V.6 for performing stereotactic injections via nanoject:
dx.doi.org/10.17504/protocols.io.bp2l6nr7kgqe/v6

Begin at step 8.2.6 "Mark the Injection Site".
Stop before step 8.2.7 "Drilling the Burr Hole".
Reference protocol Stereotactic Surgery for Delivery of Tracers by Iontophoresis V.6 for performing stereotactic injections via Iontophoresis:
dx.doi.org/10.17504/protocols.io.14egn8ewzg5d/v6

Begin at step 8.2.6 "Mark the Injection Site".
Stop at step 8.5 "Suturing".
Ensure injection coordinate is marked as precisely as possible. This is best done with a fine point sharpie mark made from the marker pipette tip.
Etch Craniotomy and Secure Headframe
10m
Ensure skull is level.
If performing an angled injection, return mouse back to level state.
Using the dovetail clamp, zero Bregma stylus over Bregma, and replace with the SHIELD Implant Tracer.
Using the SHIELD Implant Tracer (Bennett et al., 2024) as a guide, etch the shape of the craniotomy onto the skull using Dumont forceps or #11 scalpel.



Remove the dovetail clamp from the stereotactic arm with the tracer still connected. Do not move the X or Y axis.
Using the FG1/4 or EF4 drill bit, drill a shallow trench over the etch.

Cropped photo of craniotomy etch over hemisphere.

Return dovetail clamp with the SHIELD Implant Tracer back onto the stereotactic arm.

Note
Raise the stereotactic arm using the Z axis to prevent the tracer from touching the mouse's skull during this process.

Again, confirm that the tracer aligns with the etch. Make any adjustments necessary.
Secure the Headframe
10m
With dovetail clamp still centered over bregma, raise the Z axis and then replace the SHIELD Implant Tracer with the SHIELD Titanium Headframe.
Lower the Z axis until the headframe comes in contact with the skull.

Note
The contact point for this headframe will be between the eyes, however the fixed skin by the eyes will likely prevent the headframe from making direct contact with the skull. This is ok; however, the headframe should be placed as close to the skull as possible, without compressing the fixed skin into the eye or disrupting the level plane of the skull.

Note
Lowering the headframe too far down onto the mouse's skull can potentially crack a cranial suture line. If this occurs, headframe will not attach to the skull due to Cerebrospinal fluid leakage.

Prepare and apply the first round of Metabond.
Apply Metabond inside the headframe and around the outside where it's accessible. Place Metabond so that the majority of the skull within the well remains exposed. Let dry ( ~ 00:05:00 ).
5m
Turn off the isoflurane, remove the ear bars, and loosen the nose cone slightly.
Carefully detach the headframe from the dovetail clamp.
Pull back nose cone and gently remove mouse from the bite bar.

Apply Metabond outside the headframe to cover all exposed muscle and skin margins.
Reposition animal and clamp headframe into custom tool that adapts to fit in the ear bar slot of the stereotactic apparatus, reposition nose cone and turn on isoflurane.

Headframe clamp secured in the ear bar slot of the stereotactic apparatus.


Optional (2.1): Align Injection Device for Stereotactic Injections After Craniotomy and Durotomy
Optional 2.1​
This optional section accommodates injections that will be performed after the craniotomy and durotomy. This is the second section out of three required. This second section is specifically for aligning the injection device with the desired injection coordinates that were marked in Optional 2. Since the craniotomy and/or durotomy can obscure the coordinate fiducials we need to align the injection device to the precise X and Y (ML and AP) coordinates. After the craniotomy in Optional 2.2, the Z axis (DV) will be manipulated to perform the injections. If there are multiple coordinates, identify one of them and zero the coordinates creating that point as a fiducial to the remaining coordinates.


Skip this section if not marking injection coordinates for injections after the craniotomy and durotomy.
Ensure skull is level.
If headframe has been secured, set the roll and pitch of the mouse to zero degrees.
Attach injection device with loaded pipette onto stereotactic arm and center over injection location. This stereotactic coordinate should be premarked from
Place the injection device onto the stereotax arm.
Zero the X and Y and coordinates over marked injection coordinate.
Remove injection device from stereotax arm.
DO NOT move the stereotax arm out of the way. It will stay in this position during the craniotomy and durotomy.

Note
Nanoject/Iontophoresis pipettes should not be changed after setting up the stereotax arm. The variation in pipette straightness can cause the injection location to move up to 0.8mm.

Perform Craniotomy
30m
Using drill and FG1/4 drill bit, drill away any excess metabond that would interfere with the implant.
Using drill and FG1/4 or EF4 drill bit, return to the previously drilled etch/trench and drill until a crack forms all around the craniotomy.

Note
The drilling process will take 20-40 minutes.

Rinse with ACSF to remove debris or blood during the craniotomy. Be aware that soaking the skull in ACSF will cause it to become soft and can cause the drill to take off more layers than anticipated.
Stop all bleeds with Hemostatic Agent Surgifoam, ACSF rinses, and or sterile Sugi absorbent spears. Make sure there is always ACSF covering the brain surface.
Rinse the well using copious amounts of ACSF and use sterile gauze/kimwipes to absorb debris filled fluid from the headframe well before removing the skull island.
Add ACSF to the well to ensure the brain is not exposed to air after removing the skull island. Using Dumont forceps, check how loose the skull island is by gently prying up the skull at multiple points to ensure it is sufficiently separated.

Note
If the skull island is still attached (i.e. bone bridge, not cracked), remove ACSF and drill around the skull island until it is loose but not moving independently from the rest of the skull. This is especially crucial on the posterior edge. Rinse with ACSF before removal.

Prior to removing the skull island, place Hemostatic Agent Surgifoam within reach for easy bleed management.

Note
Place Hemostatic Agent Surgifoam within the well if anticipating bleeds. The whole hemisphere craniotomy is designed to avoid major sinuses. However, it is possible bleeding will occur either at the sagittal sinus, transverse sinus or rostral rhinal vein. Anyone with little experience with this craniotomy should utilize anticipatory Surgifoam within the window before removing the skull island.

When the skull island appears ready to be removed (sufficiently detached), use Dumont forceps to carefully remove the skull island, being careful to minimize pressure on the brain (i.e. pressure from skull island or tools).
With Dumont forceps, gently grab the skull island from the edge. Slowly pull the skull upwards. Watch for any unnoticed bone bridges, the adherence of the dura or sagittal sinus to the skull, bleeding, and bending of the skull island at the skull sutures.

Note
Avoid any rapid movements to prevent the skull from dipping down and damaging the cortex or the ripping/tearing of blood vessels or sinuses. If the skull island is very difficult to remove, consider removing the ACSF and drilling any remaining bone bridges, especially those on the posterior edge. Rinse the well after drilling to remove any debris before removing the skull island. If the dura is adhering to the skull island, try letting the skull island soak in ASCF for ~5 minutes to loosen the connection. Note where any bleeds start while removing the skull island and be sure to add Hemostatic Agent Surgifoam to the area as soon as the skull island is removed from the well. If the skull island starts to bend at the sutures, very gently lower the skull island down and loosen the skull island on other side of the sutures until the whole skull island can be lifted as one piece.

Stop all bleeds with Hemostatic Agent Surgifoam, ACSF rinses, and/or sterile Sugi absorbent spears. Make sure there is always ACSF covering the brain surface.

Note
Ensure all bleeds are stopped as rapidly and efficiently as possible. Excess blood loss can result in mouse death.

Perform a Durotomy
20m
Make the initial incision using a durotomy probe.
Using angled Vanna scissors, cut the dura back to the bone line. Remove dura either by cutting along the bone line or using Dumont forceps to pull flaps over the bone line. No dura should remain within the window.
Remove dura by using Dumont forceps to hold the dura taught and the angled Vanna scissors to cut.
Use caution when cutting the dura away near the sagittal sinus and large cranial vessels. The dura is connected to the sagittal sinus as well as many large vessels branching off of the sinus. Excess tugging or pressure can compromise the vessels and cause bleeding.
Stop all bleeds with Hemostatic Agent Surgifoam, ACSF rinses, and/or sterile Sugi absorbent spears. Make sure there is always ACSF covering the brain surface.

Note
Ensure all bleeds are stopped as rapidly and efficiently as possible. Excess blood loss can result in mouse death.

Clear craniotomy of any debris with ACSF.

Cropped image of the brain after craniotomy and durotomy.

Optional (2.2): Perform Stereotactic Injections Within the Cranial Window
Optional 2.2
This optional section accommodates injections that will be performed after the craniotomy and durotomy. This is the last section out of the three required. This second section is specifically for injecting the desired material at the injection coordinates that were marked in Optional 2.

Skip to next section ( ) if not performing stereotactic injections within the cranial window.
Ensure mouse's skull is level.
Set the roll and pitch of the mouse to zero degrees.

Note
Since the headframe was placed on a level skull, having the pitch and roll set to zero with mouse in the ear bar clamp creates a level skull.
Reminder: the visual cortex headframe and ear bar clamp have a pitch of -6° and 6° respectively.

Reference protocol Stereotactic Injection by Nanoject Protocol V.6 for performing stereotactic injections via nanoject:
dx.doi.org/10.17504/protocols.io.bp2l6nr7kgqe/v6

Begin at step 8.3 " If Injecting with Nanoject II" or 8.4 " If Injecting with Nanoject III".
Stop at step 8.5 "Suturing".
Reference protocol Stereotactic Surgery for Delivery of Tracers by Iontophoresis V.6 for performing stereotactic injections via Iontophoresis:
dx.doi.org/10.17504/protocols.io.14egn8ewzg5d/v6

Begin at step 8.4 "Inject Virus with Iontophoresis"
Stop at step 8.5 "Suturing".
Raise and remove injection device from stereotax arm.
Reference protocol Stereotactic Surgery for Delivery of Tracers by Iontophoresis V.3 for performing stereotactic injections via Iontophoresis:
dx.doi.org/10.17504/protocols.io.bgpvjvn6

Begin at step 8.4.4 "Zero the Z coordinate...".
Stop at step 8.5 "Suture and Recovery".
Raise and remove injection device from stereotactic arm.
Apply and Seal the Implant
20m
Remove SHIELD Implant (Bennet et al., 2024) from packaging and rinse in a well of ACSF. This protocol uses the custom designed SHIELD Implant to seal the craniotomy, but it is not required to seal the craniotomy. Other implants of a different shape or size may be used, if desired; however, be sure the shape and size of the tracer used to outline the borders of the craniotomy matches the implant used to seal it.
If there are visible air bubbles, rinse again and/or use the ACSF syringe or a sterile swab soaked in ACSF to detach bubbles.

Note
This is to prevent the addition of air into/onto the brain. Air bubbles on the implant occasionally arise from surface tension when implant is being soaked and/or removed from ACSF well.

Use Dumont forceps to place the implant so the inside edge sits over the craniotomy and slowly lower the silicone coated probe onto the center of the implant.

Note
If the implant is placed too forcefully or dropped, cortex damage may occur.
It is best to have a significant amount of ACSF covering the cortex to cushion it's placement.

Gently compress until all edges of the implant lip are contacting the full perimeter of the craniotomy. The implant is fully compressed when pushing down on the lip of the implant does not displace it from it's position.

Note
Excess pressure on the implant can cause detrimental brain compression.

Apply Vetbond to the perimeter of the implant that is contacting the skull. Be careful not to cover any holes in the implant.
Reinforce Vetbond seal and cover any additional exposed skull with Metabond.
Apply Metabond within the well halfway up the implant lip with the wooden end of a cotton swab. Be careful not to cover any holes in the implant with Metabond.

Note
If Metabond does get on the implant, it is best to wipe it away immediately with the cotton end of the swap. The Sorta-Clear coating of the SHIELD implant easily scuffs.

Apply the black cement to top of the white Metabond in the well being sure not to cover any holes in the implant. Allow to dry completely (roughly 00:05:00 ).
5m
Add equal parts Jet Fast Curing Acrylic Resin Liquid and black cement powder to an empty well of the 24 well plate.
Use the broken end of a swab to layer the black cement.

Note
Work quickly as black cement hardens rapidly. Note that the black cement does not provide any seal to the SHEILD implant, it is used solely for the purpose of blocking light from entering the well and reflection from the white Metabond.




Completed Whole Hemisphere Craniotomy with SHEILD implant.



Attach well cap to protect the implant from gathering dirt and debris in cage.
Recover Mouse and Takedown
10m
Remove mouse from stereotactic instrument base once Metabond and black cement are fully dry.
Turn off isoflurane.
Detach headframe clamp from headframe carefully.
Remove earbars.
Remove mouse teeth from bitebar by lightly scruffing mouse in a way that raises the teeth out of the bite bar.
Obtain the mouse’s postoperative weight.
Place the mouse back in a recovery cage and put the cage on the 37 °C heat plate.
Reference General Setup and Takedown Procedures for Rodent Neurosurgery protocol for takedown procedures.
Protocol
General Setup and Takedown Procedures for Rodent Neurosurgery
NAME
General Setup and Takedown Procedures for Rodent Neurosurgery
CREATED BY
Avalon Amaya

Protocol references
SHIELD: Skull-shaped hemispheric implants enabling large-scale electrophysiology datasets in the mouse brain. Bennett, Corbett et al. Neuron, Volume 112, Issue 17, 2869 - 2885.e8 doi: https://doi.org/10.1016/j.neuron.2024.06.015

Allen Institute for Brain Science. (2023). Stereotaxic Injection by Nanoject Protocol V.4. protocols.io. dx.doi.org/10.17504/protocols.io.bp2l6nr7kgqe/v4

Allen Institute for Brain Science. (2024). Stereotaxic Injection by Iontophoresis V.4. protocols.io. dx.doi.org/10.17504/protocols.io.14egn8ewzg5d/v4