���˴�Ƭ

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.