Feb 25, 2020
Headbar implantation
- 1Janelia Research Campus
External link: https://www.ncbi.nlm.nih.gov/pubmed/24520413; https://www.ncbi.nlm.nih.gov/pubmed/23727820
Protocol Citation: Liu Liu, Arseny Finkelstein, Susu Chen, Nuo Li, Karel Svoboda 2020. Headbar implantation . protocols.io https://dx.doi.org/10.17504/protocols.io.bcrsiv6e
Manuscript citation:
Guo, Zengcai V., S. Andrew Hires, Nuo Li, Daniel H. O’Connor, Takaki Komiyama, Eran Ophir, Daniel Huber, et al. “Procedures for Behavioral Experiments in Head-Fixed Mice.” PloS One 9, no. 2 (2014): e88678. https://doi.org/10.1371/journal.pone.0088678.
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 19, 2020
Last Modified: February 25, 2020
Protocol Integer ID: 33298
Keywords: Mouse, Headbar, Surgery, Implant, Photostimulation, Clear skull, Electrophysiology, Behavior, MAP
Abstract
This protocol is to prepare a mouse for head-restrained electrophysiological recordings or behavior by attaching a headbar (small titanium bar) to the skull. Standard antiseptic surgery procedures are used. Animals are anesthetized with isofluorane (2-4 %). Following deep anesthesia, a flap of skin approximately 1cm2 covering the skull is removed. Liquid marcaine is applied to the wound margins for topical anesthesia. A custom machined titanium bar (22 x 3 mm) is glued and cemented to the animal's head using cyanoacrylate and dental acrylic.
Guidelines
Standard sterile procedures for surgeries apply.
Materials
MATERIALS
Petri DishP212121Catalog #LI-PD01100
Microcentrifuge
1 ml syringes (or U-100 Insulin Syringe)BD BiosciencesCatalog #329461
Anesthetic (Isoflurane)Fisher ScientificCatalog #NC9259743
ethanol
Betadine solutionVWR international LtdCatalog #AJ159701
Bead sterilizer (Germinator 500)Braintree ScientificCatalog #GER-5287-120V
StereomicroscopeMicroscope DepotCatalog #GC01650
GlovesAnsellCatalog #PK20782
Heating Pad GaymarCatalog #TP22B
Clippers Oster MiniMax trimmerPatterson VeterinaryCatalog #07-842-4245
KetoprofenPatterson VeterinaryCatalog #07-803-7389
Buprenorphine (Buprenex)Midwest Veterinary SUpplyCatalog # 191.26890.3
Applicators, Applicator stick; Length: 6 in.; Cotton-tippedThermo FisherCatalog #23400101
Shandon™ Disposable Scalpel No. 10, Sterile, Individually Wrapped, 5.75 (14.6cm)Thermo FisherCatalog #3120032
Shandon™ Iris Scissors, Probe/Point, Angular, Premium, 4.5 in. (11.4cm)Thermo FisherCatalog #71906
Dumont Forceps (Cover Slip Forceps)Fine Science ToolsCatalog #11251-33
Glass PipettesDrummond Scientific
Light Source
Stereotax
Drill
Eye lubricant
Dental Cement
Cortex buffer
Marcaine
Gelfoam
Krazy glue
Headbar
Acrylic Polishing KitCatalog #S23-5056
Kwik-CastWorld Precision Instruments
Bur CarbideHenry Schein Animal HealthCatalog #100-7205
silicone mixing potsHenry Schein Animal HealthCatalog #3840753
Before start
Sterilize surgery tools in hot bead sterilizer before surgery.
Spray stereotax and surrounding bench top area with Virkon 1% solution (antimicrobial agent). Wipe down all surfaces. Wipe down the metal with 70% ethanol after Virkon to remove all residues.
Turn on all machines including the self-regulating heating pad and light source.
Check Oxygen and Isoflurane levels and refill if needed.
Ensure that heat source is set between 37-38 °C.
Place animal into the anesthesia induction chamber with an O2 flow rate of 1L/min and 2.5%-3.0% Isoflurane for approximately 3-4 minutes, until the animal’s breathing slows to about one breath per second.
Flush the induction chamber for 15-30 seconds with O2 before opening the chamber, then switch the route of anesthesia from induction to nose-cone.
Move animal to the nose cone on the stereotax and gently shift the tongue to the side with plastic hemostats. Place the upper teeth into the hole on the tooth-bar and slide the nose cone over the animal’s face, locking it in place. Decrease anesthesia to maintenance level of ~1.75-2% Isoflurane with an O2 flow rate of 0.5-0.8L/min.
Note
*MONITOR ANIMAL AT ALL TIMES AND ADJUST ISOFLURANE LEVELS AS NECESSARY.*
a. Respiration rate should be at 1 breath every 1-2 seconds.
b. Mucous Membranes should be pink in color (monitor color of ears and toes).
c. Loss of muscle tone.
d. Loss of Pedal Reflex (see subsequent steps).
Fix animal into the head-bars and ensure that the head is stable, straight, and perpendicular to the nose cone.
Administer Buprenorphine (0.1mg/kg for mice) and Ketoprofen (5mg/kg) at this time according to the animals' weight.
Using a cotton tip applicator, apply a coat of eye lubricant to each eye.
Shave the head, allowing enough skin to be removed. Wipe away any excess hair from the head and surrounding areas.
Using a cotton tip applicator or gauze, cleanse the surgical area by alternating with ethanol and betadine 3 times each, starting in the center of the site and being careful to avoid the eyes.
Use the toe-pinch method (pedal reflex) to ensure animal is in a surgical plane of anesthesia.
Pinch both hind feet to observe any reaction and that breathing remains stable.
If needed, increase Isoflurane by 0.25-0.5% before testing again after 30 seconds.
Inject 0.05 mL Marcaine 0.5 % volume under the scalp around where the incision is to be made for the next step.
Hold the middle of the scalp, using the iris scissor, make a cut from between the ears to between the eyes to remove just the right amount of skin to expose the entire skull.
Use forceps to clean the periosteum underneath the skin, and cut any protruding hairs with small scissors.
A sterile cotton tip applicator can be used to clear the periosteum and other tissue or debris.
The bone is cleaned with cortex buffer and dried with sterile cotton-tip applicators.
Scratch the skull where you plan to attach the headbar and on areas close to the skin with a scalpel blade. This is to create a rough surface for the Krazy glue to fix.
Apply Krazy glue on the edge of the incision. Gently push back the skin to the sides and back to seal off the wound and create more suface area on the skull.
Note
Ensure the Krazy glue forms a seal around the skull. Creating a dry and clean surface for the headbar and other procedures. This also keep the skin from retracting and exposing the skull after the surgery.
Wait for the Krazy glue to dry 00:10:00 and level the animal's head while you wait.
Attach a sharp needle to the micromanipulator.
Gently lower the needle tip until it touches the Bregma of the skull.
Bregma examples in blue,
Image source: Modified from http://www.trailofpapers.net/2015/01/wheres-bregma.html
Zero the manipulator reading and move the needle tip to Lambda.
If the difference between Bregma and Lambda in the DV axis is more than 50 μm, then tilt the head accordingly to adjust. Repeat steps 21.2 and 21.3 until the head is leveled between Bregma and Lambda.
After the head is leveled in the AP axis, move the needle tip to 2mm left of Bregma.
Zero the manipulator reading and move the needle tip to 2mm right of bregma.
If the difference between Bregma ± 2mm in the DV axis is more than 40 μm, then tilt the head around the AP axis accordingly to adjust. Repeat steps 21.5 and 21.6 until the head is leveled between Bregma ± 2mm.
Under the microscope, mark Bregma and Lambda with a marker. These points will be used as fiducials later.
Use the micromanipulator and needle tip to find other fiducial markers on the skull, and mark those with a permanent marker.
If applicable, perform the virus injection protocol here.
Assemble the headbar holder as below,
Headbar in the headbar holder.
J001375 KS style headpost with three epoxy holes Rev -.pdf 
stereotax.holder.ipt Put the headbar in the holder, and fasten the screws on the two side parts.
Spray the headbar with 70% ethanol to disinfect.
Clamp the headbar holder onto the manipulator arm. The orientation of the holder should be parallel to the earbars.
Using a sterile cotton tip applicator that is broken in half to form a spatula, apply Krazy glue to the part of the skull where the headbar is to be attached.
Place headbar onto the skull, and apply a thick layer of Krazy glue over the headbar.
Wait for the glue to dry 00:10:00 , and release the headbar from the headbar holder.
Use dental cement to build a recording well surrounding the skull. Sculpt using thin layers of dental acrylic. The dental acrylic should cover the glue around the skull and the headbar.
Note
To avoid bubbles in cement, use thin layers of dental acrylic and build up one layer at a time.
The well protects the probe from the mouse's reach and houses the cortex buffer and ground wire during the recording.
Seal the inside of the well and cover the skull underneath with single use Krazy glue.
Note
Use single use Krazy glue as it is less viscous and therefore easier to fill any gaps in cement under the headbar or between the walls of the well and the skull. This prevents any holes in the well for it to leak. The glue will cure very quickly when in contact with wet dental acrylic.
Well and headbar implant.
Fill the well with Kwik-Cast, and cover the well with a thin layer of dental cement.
Note
The thin layer of dental cement prevents the Kwik-Cast from falling off. The layer should be thin enough so that it could be quickly removed with forceps when needed.
Turn off Isoflurane and remove mouse from ear-bars. You may leave the oxygen on for a short period of time while the animal is still unconscious (especially if surgery has lasted more than 1 hour).
Remove animal from stereotax and allow animal to recover on heating pad.
Turn off oxygen.
Administer warm fluids (saline) subcutaneously between the shoulder blades if surgery lasted longer than 60 minutes.
Administer Postop Ketoprofen (5mg/kg) daily for 2 days during recovery.