High Precision Stereotaxic Surgery in Mice - Wiley Online Library

ANIMAL TECHNIQUES

APPENDIX 4

High Precision Stereotaxic Surgery in Mice

APPENDIX 4A

Stereotaxic surgery is a well-established technique for localizing specific structures within the brains of living animals. Among other applications, this technique can include the direct introduction of fluids or the chronic implantation of cannulae within these structures. These surgeries allow the assessment of the effects of blood-brain barrier-impermeable substances on the central nervous system or the structure-specific effects of a substance within the brain. The development of ever-increasing numbers of transgenic and knockout strains of mice has made them highly desirable as experimental subjects. While stereotaxic surgeries have been carried out in mice for some time (e.g., see UNIT 7.2), the relatively recent development of higher precision stereotaxic frames capable of 10- or even 1-µm precision has greatly improved the accuracy and usefulness of these techniques. This unit describes the use of the higher precision machines. The protocols described herein use bony features on the surface of the skull as reference points for determining the location of brain structures within. These features are bregma and lambda, the intersections of the sagittal suture with the coronal and lambdoidal sutures, respectively. A mouse stereotaxic atlas (e.g., Slotnick and Leonard, 1975) allows the determination of the anterior-posterior (AP), medial-lateral (ML), and dorsal-ventral (DV) coordinates of brain structures from measurements of these skull features. These coordinates are used to inject substances into target structures within the brains of anesthetized mice (see Basic Protocol) or implant cannulae, which allow the injection of substances in awake, behaving mice (see Alternate Protocol). While cannulae are commercially available, relatively simple protocols for the construction of cannulae, wire plugs, and injection needles are presented as well (see Support Protocol). NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals. SITE-SPECIFIC CENTRAL MICROINJECTION IN THE ANESTHETIZED MOUSE

BASIC PROTOCOL

For applications in which the mouse does not need to be awake and behaving (e.g., biochemical or molecular biological assays) or where the injected substance has a relatively long duration of action (e.g., antisense oligonucleotides or viral vectors), this protocol describes the delivery of substances to specific regions within the brains of mice. First, mice are anesthetized and mounted to the stereotaxic frame. The skull is exposed and measurements of the skull features are taken. Finally, a hole is drilled in the skull above the target structure, the injection is carried out, the animal is sutured and allowed to recover. The entire protocol takes ∼30 min per animal and does not require the construction of cannulae, wire plugs, or injection needles, making this a less time consuming technique than cannulation. However, this protocol induces stress in the animals and introduces a fresh trauma to the brain simultaneous with the injection. Materials Ketamine/xylazine cocktail (see recipe) Hair shaver or depilatory cream (optional) Ophthalmic ointment or mineral oil Contributed by Jaime Athos and Daniel R. Storm Current Protocols in Neuroscience (2001) A.4A.1-A.4A.9 Copyright © 2001 by John Wiley & Sons, Inc.

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Betadine Bone wax Mouse stereotaxic frame (Cartesian Research) equipped with: Sighting scope Drill with #74 bit Monoinjector with 24-G Hamilton syringe (model #8800 for injections of up to 5 µl) Heating pad Forceps Scalpel Surgical scissors Agricola-style retractors (Fine Science Tools) Cotton swabs Sutures Anesthetize mouse 1. Anesthetize the mouse with an intraperitoneal injection of an appropriate dose of ketamine/xylazine cocktail. After the animal has reached full anesthesia, remove the fur from the top of the skull with a shaver or depilatory cream or by plucking with fingertips or a pair of forceps. A dose of ∼20 µl/g body weight is generally appropriate, however, mice of different strains, ages, and adiposities will have differing sensitivities to the anesthetic. The total volume of the injection should be limited to 750 µl to prevent discomfort to the animals. For large or obese mouse strains, this may require that the concentration of the anesthetic be increased to permit smaller volume injections. The correct level of anesthesia has been reached when the mouse shows no palpebral or tail-pinch reflexes, generally within ∼5 min of the injection. A surgical level of anesthesia should persist for 45 min to 1 hr.

Mount mouse to stereotaxic frame 2. Swing the nose clamp and incisor bar down and out of the way, and then lock one of the ear bars at ∼3.5 mm from center. Set a heating pad on low (∼35°C) and place it on the stereotaxic frame to maintain body temperature and raise the animal to the level of the ear bars. While supporting the animal’s head from beneath, guide the tip of the locked ear bar into the external auditory meatus and hold firmly. Slide the other ear bar into the opposite ear and apply steadily increasing pressure until small popping sounds are heard; this popping indicates that the ear bars are properly inserted to the tympanic membranes. The animals will show some hearing loss immediately after the surgery, but hearing seems normal after one week of post-operative recovery. For experiments where normal hearing is critical, non-rupture ear bars may be preferred. However, the head will not be held as stably with this style of ear bar, and the accuracy and precision of the surgery may suffer as a result. If mounting an adult mouse and the total distance between the ear bars is 2 mm in either direction, ensure that the ear bars are properly seated within the sockets and apply more pressure.


3. Swing the incisor bar up and slide it just into the mouth while holding the jaw open with a pair of blunt forceps. Apply downward pressure to the bridge of the nose while sliding the incisor bar back into the mouth. Stop when the incisors drop into the hole in the bar. Swing the nose clamp into position on the bridge of the nose and tighten firmly. Optionally apply ophthalmic ointment or mineral oil to the eyes to keep them from drying out and protect them from accidental spills.


Current Protocols in Neuroscience

Perform surgery and injection 4. Disinfect the scalp with Betadine and make an incision along the midline with a scalpel. Using surgical scissors, extend this incision forward to the back of the eyes and backward to between the ears. Use caution to avoid damaging the muscles that insert on the external occipital crest on the back of the skull. Set Agricola-style retractors in the incision to expose the skull, and scrape away the transparent pericranial tissues with a cotton swab. 5. Fit the monoinjector to the stereotaxic frame and take DV measurements at bregma and lambda by lowering the tip of the injection needle until it is just touching these structures. Raise or lower the incisor bar/nose clamp to bring the bregma and lambda DV coordinates within 0.1 mm of each other. Zero the DV scale at bregma (see Fig. 3.10.1). Fit the sighting scope to the stereotaxic frame, align the crosshairs with bregma and zero the AP and ML scales. 6. Fit the drill to the stereotaxic frame, set it at the target AP and ML coordinates and drill a hole through the skull at a rate of ∼25 to 50 µm/sec. The diameter of the hold will depend upon the size of the cannula or monoinjector needle used. If a 24-G cannula or monoinjector needle is used, then a diameter of ∼0.57 mm works well. As a general rule, the depth of the hole should be ∼200 µm from the surface of the skull. However, the thickness of the skull varies and, depending on the stereotaxic coordinates being utilized, the depth may have to be adjusted to ensure that the hole is completely through the skull while minimizing damage to the brain. More accurate injections can be achieved by using a correction factor for determining the target coordinates. To get this correction factor, divide the bregma-to-lambda distance of each animal by the bregma-to-lambda distance from a stereotaxic atlas. Multiply the coordinates given by the atlas by this number to determine the corrected coordinates.

7. Fit the monoinjector to the stereotaxic frame. Lower the needle into the hole to the target DV coordinate and inject. Leave the injection needle in place for at least 30 sec after completion of the injection and before removal of the needle. Injections directly into brain tissue should not be applied at >0.5 µl/min, however, intracerebroventricular injections can be carried out a faster rate (1 µm/min).

8. Remove the monoinjector and seal the hole with bone wax. Suture the incision and remove the animal from the stereotaxic frame. Keep the animal warm during recovery by placing its cage on a heating pad set on low (∼35°C). House animals individually until the incision has healed to prevent them from chewing each other’s stitches. CANNULA IMPLANTATION AND CENTRAL MICROINJECTION IN THE BEHAVING MOUSE

ALTERNATE PROTOCOL

For applications in which behavior is to be assessed during or soon after injection, this protocol describes the implantation of cannulae and subsequent central injection in awake, behaving mice. First the animals are anesthetized and mounted on the stereotaxic frame. The skull is exposed and measurements of the skull features are taken. A hole is drilled in the skull and a cannula is lowered into the brain and affixed to the skull. Finally, after a recovery period, the animals are injected. The surgery takes ∼45 min and requires the construction of cannulae, wire plugs, and injection needles, making it a more time consuming technique than central microinjection in anesthetized mice. Further, the injection requires the help of an assistant. However, this technique has the benefit of allowing central microinjection in behaving mice and allowing a recovery period for healing of the trauma introduced to the brain during the surgery. Animal Techniques

A.4A.3 Current Protocols in Neuroscience

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Additional Materials (also see Basic Protocol) Filter-sterilized food coloring (dark colors such as blue, black, or green work best) Cyanoacrylate glue and accelerator (hobby supplies store) Dental acrylic Injectate Cannula and wire plug (see Support Protocol) Cannula holder Injection needle (see Support Protocol) Polyethylene tubing (0.58-mm i.d.) Micro-syringe Syringe pump Anesthetize mouse and prepare cannula holder 1. Anesthetize the mouse with an intraperitoneal injection of an appropriate dose of ketamine/xylazine cocktail (see Basic Protocol, step 1). 2. Prior to the surgery, slip a wire plug into a cannula and lightly clamp them into the cannula holder. Be sure that the wire plug is fully seated within the cannula and that the clamp securely holds both. Alternatively, the cannula can be implanted without the wire plug. In this case, the cannula should be implanted 1 mm dorsal to the target and the wire plug inserted immediately after the surgery.

Mount mouse to stereotaxic frame 3. Prop the animal up on a heating pad set on low (∼35°C). Lock ear bars firmly into ear sockets of the stereotaxic frame (see Basic Protocol, step 2). 4. Slide incisor bar into place and tighten nose clamp firmly on the bridge of the nose (see Basic Protocol, step 3). Perform surgery 5. Disinfect the scalp with Betadine and make an incision along the midline with a scalpel (see Basic Protocol, step 4). Set retractors in the incision to expose the skull. Score the surface of the skull with a scalpel, use caution to avoid the skull sutures. Dab the skull with filter-sterilized food coloring and scrape away the pericranial tissues with a cotton swab. The food coloring dyes the transparent pericranial tissues, making complete removal easier. This is critical for proper glue adhesion to the skull. Remove the dye with an ethanol-soaked swab, leaving the skull clean and dry and the skull sutures stained and distinct.

6. Fit the cannula holder to the stereotaxic frame and, using the tip of the wire plug, take DV measurements at bregma and lambda. Level the head so these DV measurements are within 0.1 mm of each other. Zero the AP, ML, and DV scales at bregma (see Basic Protocol, step 5). 7. Put 2 or 3 drops of cyanoacrylate glue on the skull and spread to cover the exposed bone. Add 2 drops of glue accelerator and allow it to dry. Score the dried surface of the glue with a scalpel. The glue is used here to increase the rigidity of the skull as well as to improve the adhesion of the dental acrylic to the skull. High Precision Stereotaxic Surgery in Mice

While it generally is not necessary, the adhesion of the dental acrylic (step 10) can be improved through the use of anchoring screws. The mouse skull is relatively thin and fragile,



but the cyanoacrylate glue reinforces it sufficiently that a pair of small jeweler’s screws can be anchored approximately 5 to 6 mm apart. Small guide holes should be drilled for the screws to avoid splitting the skull, and care should be taken to only advance the screws to about 1 mm depth. This should firmly anchor the screws without depressing the meninges and the surface of the brain. The screws should be placed so as to avoid the weaker areas of the skull near its sutures and not interfere with the positioning of the cannulae.

8. Fit the drill to the stereotaxic frame, set it at the target AP and ML coordinates and drill a hole through the glue and skull. Use a correction factor for determining precise target coordinates (see Basic Protocol, step 6).

9. Fit the cannula holder to the stereotaxic frame. Lower the wire plug/cannula assembly into the hole to the target DV coordinate. Clean the surface of the glue thoroughly with an ethanol-soaked swab and allow to dry completely. 10. Mix the dental acrylic and apply in a small mound around the cannula, taking care to avoid cementing the cannula holder to the skull or creating sharp edges that may interfere with post-operative healing. Allow the cement to dry fully before releasing the cannula from the holder and removing the cannula holder from the stereotaxic frame. Fit a small length of plastic tubing over the plug cap and exposed portion of the cannula to act as a retaining sleeve. 11. Suture the incision behind the mound of cement, and remove the animal from the stereotaxic frame. Keep the animal warm while it recovers from the anesthetic by placing its cage on a heating pad set on low (∼35°C). House the animals individually after surgery to prevent them from chewing each other’s stitches or removing each other’s wire plugs. While the animal can be injected at any time after the surgery, the authors recommend allowing the animals to recover for a week. This ensures that the incision will have healed and the animal will experience a minimum of stress and discomfort during restraint and injection.

Inject the behaving animal 12. Prepare for injection by attaching an injection needle (see Support Protocol) to a length of 0.58-mm i.d. polyethylene tubing. Fill the tubing with water, making sure to purge any air bubbles. Attach the free end of the tubing to a micro-syringe mounted on a syringe pump. Draw 1 µl of air into the tubing, then an appropriate amount of injectate. The air bubble prevents dilution of the injectate. Also, by marking the position of the leading edge of the air bubble before injecting and noting its movement during injection, one can be sure that the injection was successful and not impeded by an obstruction in the needle.

13. Remove the wire plug with forceps while an assistant restrains the animal. Fit the injection needle into the cannula and inject while allowing the animal to roam freely. After the injection, restrain the animal, remove the injection needle and replace the wire plug. A small length of tubing fit over the exposed portion of the cannula and the injection needle helps to keep the needle in place during injection. Give the animal enough slack so that it will not pull the needle out, but not enough so that it can chew the tubing.

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SUPPORT PROTOCOL

CANNULA, WIRE PLUG, AND INJECTION NEEDLE MANUFACTURE While cannulae, wire plugs, and injection needles are available commercially, they can be relatively expensive. Protocols for inexpensive and simple construction of 8-mm cannulae and 9-mm wire plugs, and injection needles, which are appropriate for target structures in the dorsal part of the brain, are described. However, the lengths and diameters of the tubing from which they are constructed can be modified for specific applications. It is important that the wire plugs and injection needles fit within the interior diameter of the cannulae, that they extend beyond the tips of the cannulae to maintain patency, and that the cannulae are long enough to reach the target brain structure while long enough to protrude above the skull to be securely affixed with dental acrylic. Materials Ethanol Acid flux (welding supply store) Dremel tool with abrasive wheel and cutting disk (Small Parts) Small bench vise 24-G thin-walled stainless steel hypodermic tubing (Small Parts) 30-G stainless steel hypodermic tubing and/or solid core wire (Small Parts) Hemostats Soldering iron (40-W minimum) Silver bearing solder (welding supply store) Outside micrometer (Small Parts) NOTE: To further reduce the damage to the brain, smaller gauge cannulae, wire plugs and injectors can be used. For example, 28-G thin-walled tubing and 33-G tubing can be substituted for the 24-G thin-walled tubing and 30-G tubing recommended here. The smaller tubing is more prone to bending and occlusion, however, and will increase the difficulty of parts manufacture and animal infusion. Construct cannula (see Figure A.4A.1A) 1a. Clamp the Dremel tool in the bench vise. Using the cutting disk, cut 24-G stainless steel hypodermic tubing to 8-mm lengths as measured with the outside micrometer. 2a. Slide the 8-mm lengths onto a longer piece of 30-G stainless steel hypodermic tubing or a 30-G syringe needle. Using the abrasive wheel on the Dremel tool, rough up the outside of the 8-mm lengths as they spin freely. Roughing up the outside of the cannulae allows for better adhesion of the dental acrylic used in implantation.

3a. Place the finished cannulae in a small container of ethanol and vortex well to remove loose metal particles. Remove from ethanol and allow to air dry before use. Construct wire plug (see Figure A.4A.1B) 1b. Cut ∼15-mm lengths of 30-G stainless steel hypodermic tubing or wire and an equal number of 2- to 3-mm lengths of 24-G stainless steel hypodermic tubing. 2b. Clamp one of the finger loops of the hemostat into the bench vise. Slide the 24-G piece of stainless steel hypodermic tubing onto the 30-G piece of stainless steel hypodermic tubing, flush one end, and clamp in hemostat.




A

B 2-3 mm

8 mm

C

D ~3 mm

24-G

24-G

9 mm 15 mm 24-G 1 mm

9 mm

30-G

Figure A.4A.1 Diagram of the cannula, wire plug, and injector needle. (A) Cannula: stainless steel hypodermic tubing is cut to length and abraded with an abrasive wheel. (B) Wire plug: a short stainless steel cap is soldered onto a length of smaller diameter tubing or solid-core wire such that when it is seated within the cannula, (C) its tip protrudes. (D) Injection needle: a length of stainless steel tubing is soldered onto smaller diameter tubing. Like the wire plug, the tip of the injection needle extends beyond the end of the cannula. However, the smaller diameter tubing is left open so that it can be attached to a length of polyethylene tubing and fluids can be administered through it by means of a syringe pump.

3b. Add a drop of acid flux to the tubing and solder the 24-G piece of stainless steel hypodermic tubing into place with a soldering iron and silver bearing solder. As the flux is applied, it should wick up into the space between the 2 pieces of tubing. The melted solder should also wick into this space as well as occluding the open end of the 30-G tubing if solid-core wire is not used.

4b. Slide plug into a 9-mm piece of 24-G stainless steel hypodermic tubing and cut off the protruding excess. 5b. Place the finished wire plugs in a small container of ethanol and vortex well. Air dry before using. As a quicker alternative to soldering a cap onto the wire plug, the wires can be inserted into a 9 mm 24-G piece and the ends bent to an acute angle. However, as a plastic retaining

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sleeve cannot be used with this design, the time saved in manufacture may have to be spent replacing wire plugs that have fallen out of the animals.

Construct injection needle (see Figure A.4A.1D) 1c. Cut ∼30-mm lengths of 30-G tubing and an equal number of 15-mm lengths of 24-G tubing. 2c. Slide 24-G piece onto 30-G piece, ∼3 mm from one end, and clamp in hemostats. 3c. Add flux and solder 24-G piece, use caution not to block the open end of the 30-G piece, with a soldering iron and silver bearing solder. The solder should seal the space between the 24-G and 30-G pieces such that no fluid leaks through during injection.

4c. Slide plug into a 9-mm piece of 24-G tubing and cut off the protruding excess. 5c. Place the finished injection needles in a small container of ethanol and vortex well. Air dry before using. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Ketamine/xylazine cocktail, 10 ml 0.65 ml of 100 mg/ml ketamine stock solution 0.22 ml of 20 mg/ml xylazine stock solution 9.13 ml sterile isotonic saline This working solution is stored at room temperature and is stable until the expiration dates of its respective components. The recipe is adequate for anesthetizing ∼20 adult mice. NOTE: The Drug Enforcement Administration (DEA) has placed ketamine into Schedule III of the Controlled Substances Act. Therefore, registration with the DEA and your state board of pharmacy may be required in order to purchase and use this drug. COMMENTARY Background Information


The blood-brain barrier presents a major obstacle to many pharmacological investigations of central neurobiology. Many pharmacological agents are not able to cross this barrier passively, so must instead be presented to the brain directly. These agents, or other substances, can either be introduced to the brain of an anesthetized animal while it is mounted to a stereotaxic frame (see Basic Protocol) or to the brain of an awake, behaving mouse through an implanted cannula (see Alternate Protocol). Injections into the brains of anesthetized animals are relatively quick and involve a minimum of equipment and are most useful in applications in which long acting substances (e.g., antisense oligonucleotides, viral vectors, some pharmacological stimulators and inhibitors) are used. While more involved, the implantation of can-

nulae allows for injections of shorter acting substances in which effects on behavior are to be assessed. Also, when used in conjunction with the implantation of osmotic pumps, cannulation allows for chronic infusions lasting for several weeks.

Critical Parameters and Troubleshooting The most critical parameter with stereotaxic surgeries is accuracy. The easiest step in improving accuracy is to align the stereotaxic frame regularly. Also, since the exact coordinates of a target brain structure can vary with the strain and age of a mouse, pilot studies may be necessary in order to refine these coordinates by trial and error. Dye injections followed by histological examination can be used to determine the site of injection and the extent to which



the injectate spreads within the brain tissue. In applications where the restriction of this spread is critical, the volume of the injectate should be adjusted accordingly. Also, minimize damage to the brain tissue by limiting the volume of the injection or carrying the injection out over a longer period of time. This allows more time for the injectate to diffuse into the tissue and minimizes the pressure exerted on the tissue by the injection. Some damage to the brain tissue is inevitable with stereotaxic surgery. However, careful technique can minimize this damage. The dental acrylic used in the cannulation protocol is very neurotoxic, and its contact with the brain tissue should be minimized. Drilling with a bit that is very close to the size of the cannula that is being implanted will leave very little space through which the acrylic can drip down into the brain tissue. To further protect the brain, the hole around the cannula can be sealed with bone wax prior to application of the acrylic. Take care that any excess bone wax is cleaned from the surface of the skull before applying the dental acrylic, as this will interfere with adhesion. While these practices avoid the major trauma inflicted upon the cortex when using larger dental burrs and the subsequent neurotoxicity resulting from acrylic leaking through the burr hole around the cannula, a relatively small amount of gliosis will still be induced along the path of the cannula or injection needle. Typical histological sections from animals subjected to site-specific injection show a closed lesion surrounded by a column of gliosis with a diameter of ∼50 to 100 µm. Sections from cannulated animals show a lesion of approximately the size of the cannula surrounded by moderate gliosis that radiates for ∼50 to 100 µm on all sides. In contrast, animals cannulated using traditional methods generally show extensive neuron loss on the surface of the cortex that manifests as a cone-shaped pit of ≥500 µm in depth radiating for a millimeter or more outward from the lesion. Further, the gliosis induced using traditional methods will often radiate for a millimeter or more within the remaining cortical tissue. The most common problem with cannulation is weak adhesion of the glue and dental acrylic to the skull. Cleaning and drying the skull thoroughly prior to the application of these adhesives is absolutely critical. Another

common problem with cannulation is difficulty removing the wire plug prior to injection. Remove and replace the wire plug 1 week after the implantation and on a weekly basis thereafter. This helps to clean out fluids and tissues that can dry in the barrel of the cannula and seal the plug in place.

Time Considerations Site-specific microinjections in anesthetized mice take ∼30 min per animal. Anesthetize and shave/pluck the next animal while an animal is being injected to minimize downtime. Cannulation takes ∼45 min per animal. Allowing the dental acrylic to harden, which takes 8 to 10 min, is the rate-limiting step in this procedure. Anesthetize and pluck the next animal during this time to minimize downtime. Wire plugs are generally made in batches of 30 at ∼2 hr per batch. They can be sterilized and reused if they have not been bent or otherwise damaged. Cannulae cannot be reused and are made in batches of 30 at ∼1 hr per batch. Injection needles can also be reused, and it takes ∼5 min to make an injection needle. Very few injection needles are generally needed. The stereotaxic frame should be aligned on a weekly basis when used regularly. This takes ∼15 min.

Literature Cited Slotnick, B.M. and Leonard, C.M. 1975. A Stereotaxic Atlas of the Albino Mouse Forebrain. DHEW Publication (ADM) 75-100, U.S. Government Printing Office, Rockville, Md.

Key References Flecknell, P.A. 1996. Laboratory Animal Anaesthesia: A Practical Introduction for Research Workers and Technicians. Academic Press, San Diego. A good discussion of alternative methods of anesthesia including Avertin, pentobarbital, and halothane. Slotnick and Leonard, 1975. See above. In the authors’ experience, the most accurate mouse stereotaxic atlas.

Contributed by Jaime Athos and Daniel R. Storm University of Washington Seattle, Washington

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