SOP 10.11 Rodent Stereotaxic Surgery (Rat)

University Animal Care Committee Standard Operating Procedure

Document No: 10.11

Subject: Rodent Stereotaxic Surgery (Rat)

Date Issued: March 23, 2011

Revision: 3

Location: Queen’s University

Responsibility: Principal Investigators (PI), Research Staff, Veterinary Staff

Purpose: The purpose of this Standard Operating Procedure (SOP) is to describe rodent stereotaxic surgery.

1. Introduction and Definitions:

A stereotaxic instrument is a device which permits the precise location of a moveable object in space. It enables an investigator to place an electrode or cannula accurately in a small brain structure even though that structure is hidden from view deep inside the skull and brain. This makes it possible to stimulate or destroy such a structure without directly disturbing overlying brain tissue. The basis of stereotaxic surgical technique is that the various regions of the brain have a definite and predictable location with respect to the surrounding parts of the skull.

2. Materials:

  • Isoflurane
  • Xylocaine cream
  • Weigh scale
  • Analgesics (refer to SOP 10.1 Pain Management in Rats, or AUP)
  • Sterile syringes (multiple sizes)
  • Sterile needles (multiple sizes)
  • PPE (cap, gloves, mask, clean lab coat or surgical gown)
  • Standard sterile surgical gloves OR autoclaved nitrile exam gloves
  • Clippers
  • Polystyrene weigh boats
  • 2% chlorhexidine surgical scrub
  • Povidone-iodine solution
  • 70 % isopropyl alcohol
  • Heating pad
  • Sterile surgical towels
  • Sterile paper towels
  • Sterile ophthalmic ointment
  • Stereotaxic unit and ear bars
  • Sterile drill bits, cannula guide, screwdriver and screws
  • Sterile cannulas
  • Sterile cannula dummies
  • Sterile surgical kit
  • Sterile gauze
  • Sterile swab applicators
  • Suture material
  • 0.9% sodium chloride
  • Lactated Ringers solution
  • Dental acrylic
  • Heat lamp
  • Antibiotic ointment


3. Procedures

  • Use a decontamination spray to clean the drill.
  • Prep surgical area according to SOP 10.3 “Aseptic Surgical Techniques (Rat)”.
  • Open the surgical packs.
  • Line the bottom of the induction chamber with clean paper towel.
  • Weigh the rat and place in the induction chamber.
  • Connect the isoflurane to the induction chamber and turn on the O2 flow to a flow rate of 1-1.5L of oxygen/min.
  • Turn the isoflurane vaporizer to 4%.
  • Once anesthetized, remove from the induction chamber.
  • Administer analgesics as per the approved Animal Use Protocol.
  • Shave the top of the rats’ head; this should be performed away from the surgical area.
  • Administer bupivacaine subcutaneously.
  • If the animal is showing signs of regaining consciousness, place back in the induction chamber.
  • Apply a small amount of xylocaine gel to the ear bars and position the animal on the ear bars.
  • Alternatively, a small dollop of gel can be applied directly in the ears.
  • Open the jaw and move the tongue to the side to avoid its teeth from getting caught on the bite bar.
  • Place the bite bar in between the upper and lower jaws.
  • Administer sterile ophthalmic ointment and reapply as required.
  • Assess corneal/blinking reflex.
  • Test for a pedal reflex to ensure the rat has reached a surgical plane of anaesthesia.
  • Don standard sterile surgical gloves or autoclaved nitrile exam gloves using aseptic technique.
  • Incise the skin over the skull using a scalpel blade and retract skin ensuring that it does not go beyond the start of the eyes.
  • Gently remove the periosteum with the dental scraper.
  • Retract the membrane in four locations.
  • Wipe the top of the skull with a sterile cotton swab until bregmais visualized.
  • Ensure that the left arm of the stereotax is at zero degrees.
  • Insert the drill bit, ensuring it is tight and attach the drill to stereotax arm.
  • Find the anterior-posterior line (vertical) line of best fit and take the anterior-posterior
  • measurement (bottom, flat bar).
  • Find the medial-lateral line (horizontal) of best fit and take the medial-lateral measurement (top, horizontal bar).
  • Make your calculations. Place drill where required.
  • Drill the cannula hole(s), and 4 screw holes. Gently assess depth with the dental scraper. Drill only once for the screw holes.
  • Instill the screws. Hold them with tweezers or haemostats and use the screwdrivers. DO NOT
  • screw them in fully, leave some head space.
  • If filtered air is accessible, use an air hose to dry around the screws.
  • Use a straight edge to ensure the cannula guides are straight and at 90 degrees.
  • Remove the drill and replace with the cannula guide holder. Put the cannula guides on and recheck for straightness.
  • Lower the cannula guides until they are in the center of the cannula hole. Place a drop of sterile saline in the cannula hole – slowly lower down to the skull to check placement of cannula guide (liquid should displace).
  • Take dorsal-ventral (DV) measurements (the top, vertical bar).
  • Calculate the cannula depth from the DV measurement and cannula depth.
  • Lower the cannula to the calculated depth.
  • Mix the dental acrylic in a weigh boat and transfer slowly to a disposable pipette. The solution should not be too runny as that will increase the exothermic reaction and may potentially cause tissue irritation.
  • The scalp must be dry and free of any tissue for maximum adherence.
  • Express the acrylic from the pipette and use the small sculptor to form it into a skullcap.
  • Ensure all screws are completely covered, and that the cannulas are not protruding excessively. The cannula should have no more than a 1 mm projection.
  • Prepare and add more acrylic cement, as necessary.
  • Wait 3-5 minutes, or until the acrylic is firm, then withdraw the cannula guide holder.
  • Put the pins in carefully.
  • Administer analgesics as per approved protocol.
  • Administer 5-10mL Lactated Ringers subcutaneously.
  • Clean around the skull cap with a swab and sterile saline.
  • Remove any acrylic cement from the surrounding skin.
  • Apply a small amount of antibiotic ointment around the skull cap with a sterile swab.
  • Remove the membrane clamps – cut off any dead skin if necessary.
  • Remove the rat from the ear bars. Place in a clean cage lined with paper towel under a heat lamp and
  • recover animals as per SOP 10.4 “Rodent Post-operative Care (Rat)”.
  • Record the surgical procedure, injections, and rat identity number on the cage card. Appropriately identify the animal (ear punch, tail markings, etc.).
  • All rats must be ambulatory with a strong righting reflex before being left alone in the recovery room.

  1. LeMoine, D., Bergdall, V., & Freed, C. (2015) Performance Analysis of Exam Gloves Used for Aseptic Rodent Surgery. Journal of the American Association for Laboratory Animal Science, Vol 54, 311–316.

Date New Version
03/23/2011 SOP Created
11/26/2015 Revised
11/16/2017 Revised
10/22/2020 Revised


SOP 10.11 - Rodent Stereotaxic Surgery (Rat)

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