Mar 20, 2024
Version 1
General Setup and Takedown Procedures for Rodent Neurosurgery V.1
- 1Allen Institute;
- 2Allen Institute for Neural Dynamics
- Allen Institute for Neural Dynamics
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 06, 2022
Last Modified: June 18, 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 / Supply | Manufacturer / Supplier | Part Number | |
| Black handle scissors, ToughCut | Fine Science Tools | 14058-11 | |
| Scalpel handle | Fine Science Tools | 10003-12 | |
| Iris forceps | Fine Science Tools | 11064-07 | |
| Dumont #5 45° forceps | Fine Science Tools | 11251-35 | |
| 45° Vanna scissors, 8cm | World Precision Instruments | 500260 | |
| 45° or 90° Durotomy probe | Fine Science Tools | 10066-15 | |
| Plastic sterilization container | Fine Science Tools | 20810-02 | |
| Hemostats | Fine Science Tools | 12004-16 | |
| large iris forceps | Fisher Scientific | 13-820-073 | |
| Bulldog clamp | Fine Science Tools | 18053-28 | |
| PREempt Disinfectant spray | McKesson Corporation | 21101 | |
| 70% Ethanol (Diluted in-house) | Sigma Aldrich | 459836 | |
| Alchohol wipes | Becton, Dickinson and Company | 326895 | |
| Sterile Surgical Drape, 18x26 | Fisher Scientific | NC9517505 | |
| Sterile Multi-well plate, 24 well | Advantor | 29443-952 | |
| Nair hair removal cream | Arm & Hammer | 40002957 | |
| Betadine Solution 10% | McKesson Corporation | 1073829 | |
| Hemostatic Agent Surgifoam | McKesson Corporation | 403360 | |
| Sterile Gauze, 3x3” squares, (autoclave sterilized) | Patterson Veterinary | 07-893-8587 | |
| Cotton swabs, double ended, (autoclave sterilized) | Advantor | 89133-810 | |
| Sugi pointed sterile swabs | Fine Science Tools | 18105-01 | |
| Insulin syringes, U-100, 0.3 ml, 31G | Advantor | BD328438 | |
| Insulin syringes, U-100, 1 ml, 31G | Advantor | BD328418 | |
| Luer-Lock Syringe, 20 ml OR | Advantor | 53548-025 | |
| Luer-Lock Syringe, 10ml | Advantor | 75846-756 | |
| 25G 5/8-inch needle | Advantor | 89134-134 | |
| 32 mm Syringe Filter 0.2 µm Supor Membrane | Advantor | 75846-756 | |
| Press ‘n’ Seal | Medline | CLO70441 | |
| Saran Wrap | GLAD | Amazon B015CLAVU | |
| Sterile Drill Bits, 0.5/0.4, FG1/4 AND/OR | NeoBurr | 1734948 | |
| Sterile Drill Bits, 1.4/1.1, FG4 AND/OR | NeoBurr | 1734214 | |
| Sterile Drill Bits 1.0/4.2, EF4 | NeoBurr | 1730012 | |
| Sterile Scalpel blades, #10 OR | Advantor | 21909-378 | |
| Sterile Scalpel blades, #11 | Advantor | 21909-380 | |
| Systane Eye Ointment | Systane | Amazon 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 | Parkell | S371 | |
| B Quick Base for MetaBond, 10 ml | Parkell | S398 | |
| Radiopaque L-Powder, white, 5 gm | Parkell | S396 | |
| Radiopaque L-Powder, clear 3 gm | Parkell | S399 | |
| Silicone implant coating, SORTA-Clear 18 | Renolds Advanced Materials | SORTA-Clear 18 | |
| Loctite 4305 | Henkel | 303389 | |
| CS-5R Coverslips, 5 mm | Warner Instruments | 64-0700 | |
| Vetbond Glue | Patterson Veterinary | 07-805-5031 | |
| Superglue, Singles | Krazy Glue | Amazon PK4 KG58248SN | |
| 3 ml transfer pipette, plastic | Avantor | 52947-970 | |
| Ortho-Jet BCA Liquid | Lang Dental Maufacturing Company | Ortho-Jet BCA Liquid | |
| Black cement (1) = 4 parts of Ortho-Jet BCA Powder (mixture) AND | Lang Dental Manufacturing Company | Ortho-Jet BCA Liquid | |
| Black cement (2) = 1 part of Powder tempura point, black | Jack Richeson & Co | 1# Black 62, Amazon B00JGZ8Q1A | |
| Kwik-Sil Sealant | World Precision Instruments | KWIK-SIL | |
| Kwik-Cast Sealant | World Precision Instruments | KWIK-CAST | |
| Heat-sterilized Glass pipettes AND/OR | Drummond Scientific | 3-000-203-G/X | |
| Heat-sterilized Glass pipettes | World Precision Instruments | 1B120F-4 | |
| “Marker” glass pipette, pulled, broken, and Sharpie mark for measuring coordinates | World Precision Instruments | 1B120F-4 | |
| Microcapillary Pipette tips | Eppendorf | 89009-310 | |
| Parafilm | Advantor | 52858-000 | |
| Lightweight Mineral Oil | Sigma-Aldrich | M8410 | |
| 30 gauge, 2" Backfilling Needle | Drummond Scientific | 3-000-027 | |
| Sterile Bone Wax | Central Infusion Alliance, Lukens | CIA2160287, 901 | |
| 5-0 Monofilament suture with 17mm 1/2C taper needle attached | Penn Veterinary Supply | Monomend MT | |
| Sterilization pouches | Advantor | 89140-804 | |
| Fiber Optic Cannulae, 200 um fiber core diameter, Black ceramic ferrule | Neurophotometrics | 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.
Equipment:
| Equipment | Manufacturer / Supplier | Part Number | |
| Small Animal Stereotaxic Instrument | Kopf | 1900 | |
| Adjustable Stage Platform | Kopf | 901 | |
| Stereo Microscope | Lecia | M80 | |
| Gooseneck Illumination | AM Scope | LED-6WA | |
| On-axis Illumination | Lecia | KL2500 LED | |
| Bead sterilizer | Sigma-Aldrich | Z378585 | |
| Small Animal Temperature Control System | CWE Inc. | TC-1000 | |
| Large Heat plate/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 | |
| Ear bars | Kopf | 1922 | |
| Ultra Fine Point Sharpie | Sharpie | 37001 | |
| Metabond ceramic mixing dish | Parkell | S387 | |
| Xlite LED Curing Light | Independent Dental | Flight Xlight2-CUR | |
| Electrode Holder | Kopf | 1970 | |
| Galaxy Mini Centrifuge | Avantor | 76269-066 | |
| P20 Pipettor | Gilson | F123600 | |
| Silver wire | Stoelting | 50880 | |
| Midgard Precision Current Source | Stoelting | 51595 | |
| Nanoject II Variable Volume (2.3 to 69 nL) Automatic Injector OR | Drummond Scientific | 3-000-204 | |
| Nanoject III Programmable Nanoliter Injector | Drummond Scientific | 3-000-207 |
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):
| Material | Part 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 Well | 0160-200-10 | |
| Mesoscope Well | 0160-200-20 | |
| Neuropixel Well | 0160-200-45 | |
| Surgical Implant for Whole Hemisphere Craniotomy | 0251-110-42 | |
| Titanium 42 Headpost | 0160-100-42 | |
| Titanium AI Straight Bar | 1365-6428-001 | |
| Titanium VisCtx Headpost | 0160-100-10 | |
| Titanium LC / Brainstem Headpost | 0160-100-52 | |
| Whole Hemisphere Well | 0160-055-08 | |
| 2p Whole Hemisphere Headpost | 0160-100-45 | |
| Well Cap | 0160-055-09 | |
| Bregma Stylus | 0251-900-04 | |
| Lambda Stylus | 0111-300-01 | |
| Dovetail Clamp | 0111-200-00 | |
| Whole Hemisphere Craniotomy Clamp Tracer | 0251-119-00 | |
| Whole Hemisphere Craniotomy Hand Tracer | 0251-110-45 | |
| -6⁰ Ear bar Headframe Clamp | 0155-100-00 | |
| -0⁰ Ear bar Headframe Clamp | 0155-110-00 | |
| Prober Holder | 0155-200-00 | |
| Titanium MotorCtx Headpost | 0160-100-54 | |
| Laser Leveling Tool | 0111-500-00 |
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.
| 1 | 2 | 3 | 4 | 5 | |
| 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:
| 1 | 2 | 3 | 4 | 5 | |
| 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:
| 1 | 2 | 3 | 4 | 5 | |
| 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:
| 1 | 2 | 3 | 4 | 5 | |
| 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:
| 1 | 2 | 3 | 4 | 5 | |
| 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.