University Animal Care Committee Standard Operating Procedure
Document No: 10.11
Subject: Rodent Stereotaxic Surgery (Rat)
Date Issued: March 23, 2011
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.
- 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
- 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
- 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.
- 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.