Mar 21, 2024

General Setup and Takedown Procedures for Rodent Neurosurgery V.1

DOI
https://dx.doi.org/10.17504/protocols.io.kqdg392o7g25/v1
General Setup and Takedown Procedures for Rodent Neurosurgery
  • 1Allen Institute;
  • 2Allen Institute for Neural Dynamics
  • Allen Institute for Neural Dynamics
Ali Williford
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DOI: https://dx.doi.org/10.17504/protocols.io.kqdg392o7g25/v1
Protocol Citation: Avalon Amaya, Jackie Swapp, Ali Williford, Robert Howard 2024. General Setup and Takedown Procedures for Rodent Neurosurgery. protocols.io https://dx.doi.org/10.17504/protocols.io.kqdg392o7g25/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: December 07, 2022
Last Modified: June 19, 2024
Protocol Integer ID: 73620
Keywords: Neurosurgery, Rodent Neurosurgery, Craniotomy, Stereotaxic Injection, Headpost, takedown procedures for rodent neurosurgery, rodent stereotaxic neurosurgical procedure, rodent neurosurgery, takedown procedure, procedure
Abstract
This protocol describes the pre-operative setup and post-operative take-down procedures utilized for rodent stereotaxic neurosurgical procedures.
Guidelines
Only perform this procedure in accordance with IACUC and veterinary requirements.
Materials
**Please note, not all supplies and equipment are used for every surgical procedure

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

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

1 g AtropinebiorbytCatalog #orb322218

1 g CarprofenbiorbytCatalog #orb321211


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 / Supplier Part 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
HemostatsFine Science Tools12004-16
large iris forcepsFisher Scientific13-820-073
Bulldog clampFine Science Tools18053-28
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 wellAdvantor29443-952
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)Advantor89133-810
Sugi pointed sterile swabsFine Science Tools18105-01
Insulin syringes, U-100, 0.3 ml, 31GAdvantorBD328438
Insulin syringes, U-100, 1 ml, 31G AdvantorBD328418
Luer-Lock Syringe, 20 ml ORAdvantor53548-025
Luer-Lock Syringe, 10ml Advantor75846-756
25G 5/8-inch needleAdvantor89134-134
32 mm Syringe Filter 0.2 µm Supor MembraneAdvantor75846-756
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 ORAdvantor21909-378
Sterile Scalpel blades, #11Advantor21909-380
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
Silicone implant coating, SORTA-Clear 18Renolds Advanced MaterialsSORTA-Clear 18
Loctite 4305Henkel303389
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
ParafilmAdvantor52858-000
Lightweight Mineral OilSigma-AldrichM8410
30 gauge, 2" Backfilling NeedleDrummond Scientific3-000-027
Sterile Bone WaxCentral Infusion Alliance, Lukens CIA2160287, 901
5-0 Monofilament suture with 17mm 1/2C taper needle attachedPenn Veterinary SupplyMonomend MT
Sterilization pouchesAdvantor89140-804
Fiber Optic Cannulae, 200 um fiber core diameter, Black ceramic ferruleNeurophotometrics FOC_BF_200um/1.25mm
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:

Equipment Manufacturer / Supplier Part Number
Small Animal Stereotaxic InstrumentKopf1900
Adjustable Stage PlatformKopf901
Stereo MicroscopeLeciaM80
Gooseneck IlluminationAM ScopeLED-6WA
On-axis IlluminationLeciaKL2500 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
Xlite LED Curing Light Independent DentalFlight Xlight2-CUR
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
5mm Cranial Window (two 5mm stacked with single 7mm circular cover glass lip)Tower Optical 18687-2
3mm Cranial Window (3mm coverslip with single 4mm circular cover glass lip). Tower Optical
CAM Well0160-200-10
Mesoscope Well0160-200-20
Neuropixel Well0160-200-45
Surgical Implant for Whole Hemisphere Craniotomy0251-110-42
Titanium 42 Headpost0160-100-42
Titanium AI Straight Bar1365-6428-001
Titanium VisCtx Headpost0160-100-10
Titanium LC / Brainstem Headpost 0160-100-52
Whole Hemisphere Well0160-055-08
2p Whole Hemisphere Headpost0160-100-45
Well Cap0160-055-09
Bregma Stylus0251-900-04
Lambda Stylus0111-300-01
Dovetail Clamp0111-200-00
Whole Hemisphere Craniotomy Clamp Tracer0251-119-00
Whole Hemisphere Craniotomy Hand Tracer0251-110-45
-6⁰ Ear bar Headframe Clamp0155-100-00
-0⁰ Ear bar Headframe Clamp0155-110-00
Prober Holder0155-200-00
Titanium MotorCtx Headpost 0160-100-54
Laser Leveling Tool0111-500-00
All equipment can be substituted with their equivalent.




Personal Protective Equipment (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
Notice:
Refer to sections via table of contents to view surgery specific setup. Skip any section titled with "Setup specific..." if not applicable.
Prepare Surgical Station for Surgery (all procedures)
Disinfect the surgical area.

Spray area for the surgical drape with PREempt and let sit for at least 00:05:00 .

5m
Spray all other surfaces - surgical rig, induction chamber, station tools, knobs buttons, and switches you touch during the procedure with 70% Ethanol reapplying after 5 min, so you have a minimum contact time of 00:10:00 .

10m
Using a non-sterile Kimwipe wipe up any residual PREempt and 70% Ethanol.
Cover heating pad on surgical rig with a layer of press ‘n’ seal.
Prepare the surgical drape with supplies.
Open a fresh sterile drape, touching only the blue side to ensure sterility, place it white side up and blue side down, on the area disinfected with PREempt.
Open sterile packages surgical supplies (Cotton swabs, kimwipes, gauze, and sugi absorbent spears), pouring the items onto the surgical drape to preserve sterility.
Fill a 10mL or 20mL syringe with ACSF (Artificial Cerebrospinal Fluid), then attach 0.2um syringe filter and 25G 5/8” needle.
Prepare peri-operative drugs.

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

Remove autoclaved surgery tools from the sterilization tray and place on the sterile drape, taking care to not touch the instrument tips.
Obtain titanium headpost with associated well, spray with 70% ethanol and place on drape with the side that will interface with the skull up. 
Prepare additional 0.3ml insulin syringe for vetbond application.

Note
Do not fill syringe until use as it may become clogged.

Obtain #10 Scalpel blade for skin incision.
Proceed to desired procedure "Specific Setup" section.
Setup Specific to Headpost Only Procedures (no craniotomy)

Prepare the 24-well plate with supplies.
12345
A
Betadine
ACSF
ACSF
B
Nair
ACSF
C
D
1
A
70% Ethanol
B
70% Ethanol
C
D


Fill one well with Betadine and one well with Nair.
Fill three wells with ACSF.

Note
Two wells of ACSF minimum for rinsing 70% Ethanol, 1 well of ACSF for soaking Surgifoam.

Fill two wells with 70% ethanol.

Note
70% Ethanol can be used for disinfecting any non-sterile supplies or tools used in the surgery typically: Coverslip, Tracers, Stylus, Fiber Implants.

Soak three cotton swabs in the Betadine well for betadine application to incision site.
Use sterile forceps to tear off pieces of sterile surgifoam and place in well with ACSF to soak.
Place stylus in one of 70% ethanol wells.
Setup Specific to Headpost and Craniotomy Procedures
Prepare the 24-well plate:
12345
A
Betadine
ACSF
ACSF
B
Nair
ACSF
ACSF
C
ACSF
D
1
A
70% Ethanol
B
70% Ethanol
C
70% Ethanol
D



Fill one well with Betadine and one well with Nair.
Fill three to four wells with ACSF.

Note
Two wells of ACSF minimum for rinsing 70% Ethanol, 1 well of ACSF for soaking Surgifoam.

Fill three wells with 70% ethanol.

Note
70% Ethanol can be used for disinfecting any non-sterile supplies or tools used in the surgery typically: Coverslip, Tracers, Stylus, Fiber Implants.

Soak three cotton swabs in the Betadine well for betadine application to incision site.
Use sterile forceps to tear off pieces of sterile surgifoam and place in well with ACSF to soak.
For 5mm or 3mm craniotomy procedures: use forceps to place a stacked coverslip in one of the three 70% ethanol wells in the well plate.

Note
Skip this step if the coverslip is coated in silicone.

Place the craniotomy tracer in 70% Ethanol well.
For 5mm or 3mm craniotomies: place glass coverslip in well.
For whole hemisphere craniotomies: place the WHC tracer in the well, ensuring the portion that will come in contact with the skull is submerged.
Place stylus in one of 70% ethanol wells.
Setup Specific to Iontophoretic Injections (with or without headpost)
Prepare 24-well plate:

12345
A
Betadine
ACSF
ACSF
B
Nair
ASCF
C
D
1
A
70% Ethanol
B
70% Ethanol
C
D


Fill one well with Betadine and one well with Nair.
Fill three wells with ACSF.

Note
Two wells of ACSF minimum for rinsing 70% Ethanol, 1 well of ACSF for soaking Surgifoam.

Fill 1-2 wells with 70% Ethanol.

Note
70% Ethanol can be used for disinfecting any non-sterile supplies or tools used in the surgery typically: Coverslip, Tracers, Stylus, Fiber Implants.

Soak three cotton swabs in the Betadine well for application to incision site.
Prepare procedure-specific additional supplies:
  • 5-0 Monofilament suture
  • Parafilm square
  • Bulldog clamp
  • Silver wire
Remove one aliquot of virus from -80ºC freezer, thaw at Room temperature , and spin down in the mini centrifuge.
Obtain Iontophoretic-specific pipette.
Setup Specific to Nanoject III Injection (with or without headpost)
Prepare 24-well plate:
12345
A
Betadine
ACSF
ACSF
B
Nair
ASCF
C
D
1
A
70% Ethanol
B
C
D




Fill one well with Betadine and one well with Nair.
Fill three wells with ACSF.

Note
Two wells of ACSF minimum for rinsing 70% Ethanol, 1 well of ACSF for soaking Surgifoam.

Fill two wells with 70% ethanol.

Note
70% Ethanol can be used for disinfecting any non-sterile supplies or tools used in the surgery typically: Coverslip, Tracers, Stylus, Fiber Implants.

Soak three cotton swabs in the Betadine well for application to incision site.
Prepare procedure-specific additional supplies:
  • 5-0 Monofilament suture
  • Parafilm square
Remove one aliquot of virus from -80ºC freezer, thaw at Room temperature , and spin down in the mini centrifuge.
Obtain Nanoject-specific pipette.
Prepare nanoject-specific pipette.
Using a 30g, 2” backfilling Hamilton syringe filled with mineral oil, backfill the pipette.
Insert the tip of the Hamilton syringe into the pulled pipette, all the way to the shoulder, and slowly depress the plunger on the Hamilton syringe, filling the pipette with oil. Be sure not to introduce bubbles into the system or the injection may not be successful.
Once the pipette is backfilled with oil, loosen the collet on the end of the injector. The wire plunger should be 2-3mm past the collet.

Nanoject III Injection Standard Collet/Chuck/Green Gasket.

Gently slide the pipette over the wire plunger and push it through the chuck and seating it in the green rubber seal.

Nanoject III with pipette.

Tighten the collet.
Load the virus into the pipette
Once the pipette is secured to the collet, press, and hold the ‘EMPTY’ button on the control box until an audible beep is heard. This will drive the wire plunger out forcing oil to the tip of the pipette. Any excess oil will be expelled. Expel about 4-5 drops and work out any air bubbles.
Take virus aliquot and spin down for 10-15 seconds.
Use the P20 micropipette with a microfil tip to draw up ~2 µl of virus.
Using the surgical bed as a platform, aspirate the virus sample onto a clean piece of Parafilm.
Using the stereotaxic apparatus, lower the tip of the pipette (secured in the Nanoject) into the sample. Be careful to not “bottom out”.
Press the ‘FILL’ button and draw up the desired amount of virus (button will change to red). Press the button again to stop the filling of the pipette when finished.

Note
Do not introduce air bubbles into the system as this may result in inaccurate injection volumes. Bubbles will be visible.


Carefully set injector aside where pipette will not be disturbed.
Setup Specific to Optic Fiber Implants
Prepare 24-well plate:


12345
A
Betadine
ACSF
ACSF
B
Nair
ACSF
ACSF
C
ACSF
D
1
A
70% Ethanol
B
70% Ethanol
C
70% Ethanol
D


Fill one well with Betadine and one well with Nair.
Fill 3-5 wells with ACSF.

Note
Two wells of ACSF minimum for rinsing 70% Ethanol, 1 well of ACSF for soaking Surgifoam.

Fill three wells with 70% ethanol.

Note
70% Ethanol can be used for disinfecting any non-sterile supplies or tools used in the surgery typically: Coverslip, Tracers, Stylus, Fiber Implants.

Soak three cotton swabs in the Betadine well for application to incision site.
Use sterile forceps to tear off pieces of sterile surgifoam and place in well with ACSF to soak.
Gather fiber optic implants and soak them in the 70% ethanol wells.
If using a headframe that is not already sterilized, soak headframe in 70% ethanol.
Prepare the Anesthesia System and Anesthetize the Mouse (all procedures)
Prepare the anesthesia system.
Follow anesthesia system's manufacturer guidelines and your facilities environmental health and safety committee regarding the use of an anesthesia system for rodent neurosurgery.
Turn on the oxygenator and ensure your facilities vacuum lines are functioning properly (i.e vacuum lines are on and gauge displays psi).
Ensure that all tubes are connected securely and that you can feel the vacuum suction in the scoop underneath the nose cone.

Ensure the Isoflurane line to the induction chamber is open and the line to the nose cone is closed via their designated stopcocks.
Anesthetize the mouse.
Remove the animal from its experimental cage, obtain a preoperative weight.
Place the mouse into the induction chamber, open the vacuum valve (stopcock) and isoflurane line (stopcock) for the chamber, and then turn on the isoflurane vaporizer to 5%.
Once the mouse is fully unconscious, turn off the isoflurane and wait at least 10 seconds with the vacuum on to allow the chamber to clear before removing the animal.
Position mouse on surgical rig by placing maxillary incisors in the hole on the bite bar and securing head with ear bars.
Secure the nose cone over the mouse’s snout. Make sure the body of the mouse is on top of the heading pad, resting comfortably. Redirect the gas flow from the induction chamber to the surgical rig via line stopcocks.
Set the isoflurane to ~1.5-2%, turn off the vacuum line to the induction chamber and close the lid.
Monitor the mouse’s breathing throughout the process and adjust gas levels as necessary. 
Prepare the Mouse for Surgery (all procedures)
Apply Systane to the end of a cotton swab and use to push the whiskers away from the surgical field. 
Cover the mouse’s eyes with a generous amount of Systane. Additional Systane should be added as needed to prevent eye dryness and protection from the scope light. 
Administer any drugs that have a timing after induction or prior to incision.
Use black handled scissors to shorten hair from top of the head. Use caution when trimming hair around the eyes. Avoid cutting off whiskers.

Note
Hair removal is contingent on the type of surgery and placement of headframe, optic fibers, etc.

Apply Nair with the pointed end of a non-sterile cotton swab, gently swirling it down to the interface of the skin and hair.
Remove all Nair with several alcohol swabs. 
Disinfect the surgical site with 3 rounds of alternating Betadine-soaked sterile swabs and alcohol wipes. The last application of Betadine should not be wiped off.
Place Saran Wrap over the trunk of the animal and change into new gloves.
Take Down Steps When the Procedure is Complete
Dispose of used syringes, blades, drill bits, swabs and needles or anything that could puncture a plastic bag in the biohazard sharps container.
Dispose of all disposable materials that came into contact with blood in the biohazard waste container.
Spray down surgical tools with pH neutral surgical tool cleaner and wipe with a kimwipe or alcohol swab. Be sure to wipe any blood, skin, or cement residue off the tools. Use caution when wiping down Dumont’s.
Place tools into tool kit. Place tool kit into sterilization pouch and write your name, department initials and the date on the pack. Bring tool kit to the designated location to be autoclaved.
If the tools are to be used again the same day, sterilize using the hot bead dry sterilizer.
Disinfect ear bars with alcohol swab, then place back on the surgical rig.
Turn off the vacuum and oxygen sources by switching the stopcocks to the off position.
Release the air from the drill lines by pressing down on the pedals until the pressure reads 0 psi. If the air source is connected to a wall valve, then turn the valve to the off position.
Turn off heating pad, bead sterilizer, stereotax, and scope light.
Turn off compressed air, vacuum, and oxygen concentrator.
Ensure Isoflurane is turned off.
Spray down station with 70% cleaning up any debris and detritus from the surgery paying close attention to the nose cone, ear bars, vacuum scoop, and Induction chamber.