Guide/Cranial window surgeries

Contributed by Megha Sehgal (Sehgal Lab, Ohio State University).

Please read before using this protocol. This protocol is shared as an initial reference to help you develop your own procedures — it is not a substitute for hands-on training. Adapt it to your own institution's IACUC-approved animal-use protocol, and follow your approved protocol and local veterinary guidance for all surgical technique, drugs, dosing, and postoperative monitoring. Neither the author nor the Miniscope project guarantees outcomes from following this guide.

Overview

This protocol describes the placement of a cranial window over the dorsal cortex. The standard preparation uses a 3 mm square cranial window spanning the sagittal sinus.

Our laboratory uses this protocol, with slight modifications, for:

One-photon imaging using a Miniscope
Two-photon functional imaging
Two-photon structural imaging
Cranial windows targeting other cortical regions

Important notes

Virus injections

This protocol includes stereotaxic virus injections. These steps can be omitted when using transgenic mice or when viral expression is not required.

Optimization for Miniscope imaging

For one-photon Miniscope imaging, the largest source of variability is typically viral expression. Virus parameters should be optimized at the beginning of a project and may require minor adjustment between virus batches. Important variables include:

Viral serotype
Promoter
Viral titer and dilution
Injection volume
Injection rate
Injection depth and DV coordinates

I have primarily used Addgene GCaMP viruses for Miniscope imaging. Transgenic animals and other viral preparations may also work, although expression levels should be independently validated.

Long surgeries and postoperative recovery

Depending on the number of viral injections, these surgeries can be long. Several precautions substantially improve recovery and postoperative health. The importance of these measures increases with surgery duration.

Minimize anesthesia exposure. After induction and stabilization, progressively lower the isoflurane concentration so that the mouse remains just above the minimum acceptable surgical plane of anesthesia.
Provide supplemental fluids. Administer approximately 0.5 mL saline subcutaneously for each hour of surgery. Additional saline may be appropriate if substantial bleeding occurs during skull removal.
Maintain body temperature during recovery. Allow the mouse to recover overnight with access to a heating pad. Even when mice regain locomotion quickly, overnight warmth generally helps them return to normal activity more rapidly.

Equipment and materials

Surgical and preparation supplies

Air duster (optional)
Hair removal cream, such as Nair
Artificial tear or ophthalmic ointment
Betadine
70% ethanol
Hydrogen peroxide
Sterile saline, ice-cold, and room temperature
Sterile cotton-tipped applicators
Kimwipes
Dumont No. 2 forceps (FST #11223-20)
Dumont No. 55 forceps (FST #11255-20)
Surgical scissors
Scalpel and blade
Dental drill and drill bit (1007205, Henry Schein Inc. – 206100250100)
10 mL syringe filled with saline
1 mL syringe filled with saline
3 mm glass coverslip
Pencil or fine-tip Sharpie
Biopsy tissue punch (optional; for round craniotomies)
Tapered wooden applicator or needle tip

Drugs

Carprofen or Rimadyl
Dexamethasone

Adhesives and dental cement

Krazy Glue (product link)
Cyanoacrylate accelerator (optional)
Metabond liquid (Parkell Products Inc, Patterson Item #75533492)
Metabond powder (Parkell Products Inc, Patterson Item #75533559)
Metabond catalyst (Parkell Products Inc, Patterson Item #75533500)

Procedure

1. Prepare the surgical area

Turn on the glass bead sterilizer. Allow approximately 20 minutes for it to reach operating temperature.
Turn on the heating pad. Adjust the system so that the mouse reaches approximately 37 °C when positioned with the temperature sensor.
Prepare all surgical instruments and supplies. Instruments should be autoclaved in advance.
Carefully attach the scalpel blade.
Place scissors, forceps, and other reusable instruments in the glass bead sterilizer for 10–15 seconds.
Weigh the mouse and record its preoperative weight.
Place a Kimwipe in the isoflurane induction chamber.
Turn on the SlowFlo isoflurane system.

Induction settings:

Flow setting: High
Flow rate: 400
Isoflurane: 4%

2. Anesthetize and position the mouse

Place the mouse in the induction chamber.
Wait until the mouse stops moving and its respiratory rate slows.
Transfer the isoflurane tubing from the induction chamber to the stereotaxic surgery frame.
Reduce the flow rate to the appropriate low-flow setting.
Position the mouse on the stereotaxic frame.
Using forceps as needed, place the mouse's mouth over the bite bar. Carefully move the tongue out of the airway. Make sure that the tongue is below the bite bar.
Slide the metal nose cylinder over the muzzle.
Gently position the ear bars on either side of the head.
As an initial position, align the 4 or 5 mark on the ear bar with 0 on the stereotaxic frame.
Confirm that the head is stable. The ear bars should securely hold the skull but should not be excessively tight.
Adjust the muzzle as necessary to ensure unobstructed breathing.
Apply ophthalmic ointment to both eyes.
Gently close the eyelids over the ointment to ensure complete coverage of the cornea.

3. Prepare the surgical site

Carefully shave the head and neck.
Use particular caution around the ears and whiskers.
Remove loose hair using an air duster, if desired.
Apply a small amount of hair removal cream, such as Nair, to remove residual hair.

Important: I perform hair removal carefully and exhaustively. Stray hair can enter the craniotomy during later steps and interfere with window placement or healing. They are also very annoying.

Apply Betadine to the shaved area using a sterile cotton-tipped applicator.
Apply 70% ethanol using a fresh cotton-tipped applicator.
Repeat the Betadine and ethanol preparation two additional times, for a total of three alternating applications.
Once the mouse is stable, progressively lower the isoflurane concentration while maintaining an appropriate surgical plane of anesthesia.

4. Expose and prepare the skull

Lift the skin at the back of the neck using forceps.
Make an initial incision using surgical scissors.
Extend the incision anteriorly toward the ears and around the front of the skull.
Make as few cuts as possible to create a clean skin margin.

Practical note: Do not assume that removing the smallest possible amount of skin produces the best implant. The implant should have sufficient skull surface area for stable attachment. I generally expose the skull as close to the ears as practical. Very small implants have less surface area for adhesion and are more likely to become unstable. This is especially important when the animal will later be baseplated for Miniscope imaging: small implant size prevents baseplates from sitting appropriately.

Apply hydrogen peroxide to the exposed skull using a cotton-tipped applicator. This helps remove residual fascia.
Remove the hydrogen peroxide with saline.
Continue rinsing until no bubbles remain.
Using a scalpel, gently scrape the surface of the skull to remove any remaining fascia. Clean skull = stable implant.
Wipe the skull clean using saline and a cotton-tipped applicator.
Using the scalpel, lightly score the exposed skull in a grid pattern to increase surface area for implant adhesion.
Avoid skull sutures.
Avoid planned virus injection sites.

Note: Scoring is optional but increases the available surface area for cyanoacrylate and may improve implant stability.

Using the scalpel, detach the neck muscles from the posterior skull from ear to ear.
Keep the scalpel blade closely aligned with the skull surface.

Important: Do not push downward. The goal is to detach the muscle from the skull, not to apply pressure to the underlying tissue.

5. Align the skull for stereotaxic injections

This section is only required when performing stereotaxic virus injections.

Adjust the stereotaxic muzzle approximately so that the skull appears level by eye.
Use an alignment tool and the stereotaxic manipulators to locate bregma and lambda.
Align the skull so that bregma and lambda are in the same DV plane.
Confirm ML alignment.
If the ML measurements differ by more than 50 µm, move the ear bars together to the left or right to correct the alignment.
Measure the DV coordinate at bregma and lambda.
Adjust the head pitch until the DV measurements are within approximately 50 µm of one another.
Align the coronal tilt to level the skull laterally to ensure the left and right hemispheres are aligned in the same horizontal plane.

6. Perform viral injections

A detailed viral injection protocol is not included here because injection parameters vary substantially depending on the brain region, virus, experimental goal, and injection system. The following describes our general approach.

Injection system

We use a Nanoject to deliver viruses. This system allows precise control over injection volume and delivery rate. The small glass capillary also minimizes mechanical damage to the injected brain region. Injection speed, volume, viral titer, and dilution can substantially alter viral spread and expression.

Considerations for Miniscope imaging

Transgenic reporter lines may work in principle but are often too dim for reliable Miniscope imaging.

One-photon imaging is generally restricted to tissue approximately 200–300 µm below the dura. Therefore, viral injections should target superficial cortical layers, typically layer 2/3 or upper layer 4/5a, depending on the cortical region.

For initial optimization, I recommend beginning with hSyn-GCaMP6f using a relatively high-expression viral preparation. Viral titer and dilution should be empirically optimized for the experiment and imaging system.

Example: retrosplenial cortex injections

Our approximate RSC coordinates are:

AP: −2.3 mm
ML: ±0.5 mm
DV: 0.6–0.8 mm from skull

For V4 Miniscope surgeries, we often perform two viral injections per hemisphere, for a total of four injections.

Example injection parameters:

Volume per injection: 500 nL
Injection rate: 50 nL/min
Injection duration: approximately 10 minutes per injection

I am happy to share a complete RSC viral injection protocol separately.

After completing the viral injections, administer:

0.1 mL dexamethasone
0.5 mL saline subcutaneously

7. Mark the cranial window

Remove all visible debris from the exposed skull using an air canister or sterile saline.
Clean the skull using a cotton-tipped applicator soaked in sterile saline.
Dry the skull using a Kimwipe or cotton-tipped applicator.
Place a 3 mm square coverslip on the skull.
Position the coverslip so that the virus injection sites or target imaging region are centered within the planned cranial window.
Mark the four corners of the coverslip using a pencil or a fine-tip Sharpie.
Remove the coverslip.
Using a scalpel, gently connect the four marks to etch the outline of the planned craniotomy. These shallow etched lines provide a guide for drilling and help prevent the drill bit from slipping.
Replace the coverslip and verify the window position and dimensions.

Important: The craniotomy should be only slightly larger than the 3 mm coverslip. A window that is too small increases the risk of bone fragments or skull flaps becoming trapped beneath the coverslip. These fragments can promote bone regrowth and interfere with imaging. An unnecessarily large window may be difficult to glue properly.

Round cranial windows

For a round craniotomy, use a biopsy tissue punch to mark the desired circular area. Tissue punches are available in multiple diameters.

8. Drill the craniotomy

Using a dental drill, follow the etched lines to create a square or circular track around the planned cranial window.
Complete one full pass around the craniotomy.
Remove bone debris.
Apply several drops of saline to briefly cool the skull and underlying tissue.
Remove the saline and dry the skull.
Repeat the drilling, cleaning, and cooling cycle.
Keep the drilling depth as uniform as possible around the entire track.
Do not drill in one location for an extended period.

Drilling over the sagittal sinus

Use particular caution when drilling over the sagittal sinus. The skull is often thicker in this region and may require additional drilling.

Drill control

We use an air-powered dental drill controlled with a foot pedal. This allows precise regulation of drilling speed through foot pressure and is particularly useful when drilling over the sagittal sinus or other sensitive areas.

Skull bleeding

If the skull begins to bleed, absorb the blood using the twisted corner of a Kimwipe.

Skull bleeding is more common in older animals, which often have more vascularized skulls. While the use of saline, especially chilled saline (or ACSF), is popular to prevent bleeding, I often found that bleeding over the sagittal sinus is better controlled when allowed to clot for a few seconds. Continuously flooding the area with saline prevents clotting and prolongs bleeding.

9. Determine when to stop drilling

Stop drilling only after the craniotomy meets the following criteria.

Criterion 1: cortical vasculature is clearly visible

Apply a drop of saline or ACSF over the thinned skull. The finest cortical blood vessels should become clearly visible. At this stage, the thinned bone may begin to crack. Once a section of the track cracks, stop drilling that area. Continue thinning the remaining portions of the track.

Criterion 2: the central skull piece moves with gentle pressure

Gently tap the central skull piece using forceps. The central piece should begin to depress with very light pressure. Once an area of the track begins to give way, stop drilling that region and continue thinning the remaining areas. Continue until the entire central skull piece moves with gentle pressure.

10. Soak the thinned skull

Cover the craniotomy with saline.
Allow the skull to soak for approximately 15–20 minutes.

A longer soak generally makes the skull easier to remove. However, balance the soaking period against total surgery duration and the overall condition of the mouse.

Remove most of the saline using a Kimwipe or cotton-tipped applicator.
Gently clean away any remaining blood or bone debris.
Add a small amount of fresh saline to the craniotomy track.

11. Remove the central skull piece

Using No. 55 forceps, gently push the lower forceps tip through one section of the thinned skull track.
Carefully position the tip beneath the central piece of skull, taking care to keep the angle of the forceps shallow so as not to touch the brain.
Remove the forceps.
Repeat this process at multiple points around the thinned track.

The goal is to create several small perforations and progressively release the central skull piece. Typically the skull is already so thin that it does not take much to break the skull along the drilled track.

Stabilize the forceps with your second hand if necessary.
Once the central skull piece begins to lift with very light upward pressure, gently grasp the bone with the forceps.
Slowly increase upward pressure.

Important: Continuously monitor the position of the forceps tip. Do not allow the forceps to penetrate or tear the dura. Watch the skull opposite the area you lift with the forceps and ensure it does not lever down into the brain.

Slowly lift the central skull piece vertically.
Ensure that the loose skull fragment does not rotate downward or tear the underlying brain tissue.
Once the skull piece is removed, immediately cover the exposed brain with saline.
If bleeding occurs:
- Gently absorb the blood using a Kimwipe.
- Reapply saline.
- Repeat as necessary until the surgical field is clean.

12. Place the coverslip

Prepare a clean 3 mm glass coverslip.
If needed, rinse the coverslip with sterile water and dry completely.
Remove all saline from the exposed brain immediately before coverslip placement.
Gently position the coverslip in the center of the craniotomy.
Confirm that the coverslip completely covers the craniotomy and is correctly aligned with the imaging region.
Hold the coverslip in position using forceps.
Place several drops of cyanoacrylate in a clean Petri dish.
Dip a tapered wooden applicator or needle tip into the cyanoacrylate.
Carefully apply cyanoacrylate to the gap surrounding the coverslip.

Do not apply cyanoacrylate directly from the bottle onto the skull or coverslip. It's too hard to apply the right amount.

Apply a small additional amount of cyanoacrylate around the coverslip edge to create a thin lip over the perimeter of the glass.
Gently apply steady downward pressure to the coverslip so that the glass sits level with, or slightly below, the surrounding skull. A small custom tool can be used to apply pressure; alternatively, carefully apply steady manual pressure.
Allow the cyanoacrylate to dry for 10–15 minutes.
A cyanoacrylate accelerator may be used to shorten the curing time.

Practical note: Coverslip placement requires fine motor control and becomes substantially easier with practice. Many positioning tools and additional stabilization steps become optional as surgical proficiency improves.

13. Secure the implant

Apply cyanoacrylate and then Metabond around the cranial window.

I prefer Metabond because I have not lost a cranial window secured with Metabond.

Extend the adhesive or dental cement across the exposed skull surrounding the cranial window.
Secure the implant to the surrounding skin and muscle as appropriate.
Seal the skin margins with cyanoacrylate. Carefully sealing these edges helps prevent bacteria from entering the surgical site and may reduce infection and unwanted tissue growth.
Allow the cement to dry completely.

14. Prepare the recovery cage

While the cement is drying, clean the surgical area and prepare the postoperative recovery cage. The cage should contain:

Soft food or pellet mush
Soft bedding
A clearly labeled cage card

The cage card should include:

Date of surgery
Number of viral injections
Surgeon initials

15. Complete the surgery and begin recovery

Administer:
- 0.5 mL saline subcutaneously
- 0.1–0.3 mL carprofen subcutaneously
Turn the isoflurane concentration to 0%.
Carefully remove the mouse from the stereotaxic frame.
Transfer the mouse to the heated recovery area.
Monitor the mouse during recovery according to the approved institutional animal care and postoperative monitoring procedures.
Maintain supplemental warmth overnight when appropriate, particularly following long surgeries.
Continue postoperative analgesia and monitoring according to the approved animal protocol.