Methods for Intravenous Self Administration in a Mouse Model

Hello, my name's Elizabeth Kotick and I'm here from Dr.Katherine Gill's lab at the addictions unit of the McGill University Health Center. Today I am gonna be showing you how to perform the catheterization surgery necessary for conducting intravenous self-administration studies. I'm gonna show you how to insert the catheter as well as some tips for maintaining catheter patency.

During nicotine intravenous self-administration sessions, the animals are hooked up to Tai on plastic tubing and nicotine infusions are delivered by the attached infusion pumps. Operant boxes were purchased from med associates. Each box is equipped with two ultrasensitive mouse levers.

There are a few items that need to be prepared prior to surgery in order to insert the catheter into the vein. A 20 gauge needle needs to be adapted to slide the tubing down into the vein. Shave down about two centimeters of the top part of the shaft of the needle, creating a channel that will be used to guide the catheter tubing into the vein.

Next, prepare the syringes for flushing and checking the catheter. 12 centimeters of tubing is attached at one end to the 26 gauge needle of the syringe while the other end is stretched over a 23 gauge needle. In order to make the tubing easier to affix to the catheter cannula, you will need two syringes like this one filled with the heparinized T carcin solution, the other with saline.

Lastly, prepare the catheter cannula cap. The catheter cannula needs to be capped in order to prevent debris from entering. These caps are made by stretching plastic tubing over 23 gauge needles.

The tubing is cut one centimeter from the needle tip and the open end is melted To seal the tubing, saline is pushed through the needle to make sure that there are no leaks in the caps. Mouse catheters were obtained from Cam Cs in the uk. Cut the excess tubing from the catheter so that there is 1.2 centimeters from the end of the catheter bulb.

Next, attach the syringe containing saline to the exterior metal cannula of the catheter and flush to inspect for leaks. Mis are anesthetized using isof fluorine gas mixed with oxygen and are maintained under anesthesia for the duration of the surgery. Using a breathing tube, levels of both oxygen and isof fluorine are set based on the body weight of the animal and are adjusted as needed during surgery, make sure the animal is properly anesthetized by pinching its back legs with your forceps and monitor the mouse throughout surgery for respiratory distress and depth of anesthesia.

Since surgery lasts between 20 and 30 minutes, apply an eye lubricant to both eyes. The mouse is shaved prior to surgery. Prepare a space for the catheter base on the animal's back.

Gently pull up the skin on the animal's back and make a small mid scapular longitudinal incision about two centimeters in length. Starting midway on the back and extending to just below the neck. Force apart connective tissue to make space for the catheter base and apply saline to the incision.

Gently pull the skin on both sides of the incision together and flip the animal over. The right tegular vein will be found superficially under the skin of the neck and can be exposed by making a S shallow incision. Make a diagonal cut about one to two centimeters in length, extending from around the right clavicle, going upward towards the animal's jaw.

The tubing from the base of the catheter needs to be brought through the ventral incision in the neck to be accessible for insertion into the vein. Guide the forceps under the skin of the shoulders and break through connective tissue. To reach the back incision, pull the tubing through the skin to the ventral opening around the animal's neck.

Grip the end of the tubing with artery clamps and leave it clamped by the animal's right side. Apply saline to the incision, isolate the right jugular vein. Very gently move away any connective and adipose tissue that may be obstructing your view and locate the vein.

Gently lift the vein around the top and use the curve forceps to break apart adipose and to connective tissue around and under the vein. Place the curved forceps under the vein and open them with your free hand. Pass a sterile plastic bar under the vein.

Once the vein is elevated, remove just enough fat to give you a clear unobstructed view of the vein. Periodically drip saline onto the exposed vein to keep it wet. Prepare suture knots around the vein to be able to secure the catheter tubing in place once it is inserted into the jugular Place.

Sutures underneath the vein at both extremities and make a single very loose open knot at each end. Thread the catheter tubing through the top knot and loop it over the suture thread to the rest over the right shoulder. Push saline through the catheter to weigh the tubing down to insert the catheter tubing into the jugular vein.

The modified 20 gauge needle is used to pierce the vein, wet both the vein and the insertion needle with saline to reduce friction during the insertion, gently pull back on the plastic bar and insert the needle closer to the bottom part of the vein. Just enough so that a half centimeter of the needle is in. Slide the catheter tubing down the needle and into the vein.

There should be no resistance. Resistance would indicate that you are in connective tissue. To be sure that you're within the vein, try pulling back on the catheter syringe to see if you're able to draw blood.

Once the tubing is in place, remove the needle and push the catheter bulb to the insertion point. Tie the bottom knot and make sure the catheter tubing is flush against the bar before tying the second knot right above the bulb. This will secure the catheter in place.

Remove the bar and make a second knot at each end. Tuck the catheter tubing and suture threads down under The skin prior to suturing. Use the artery clamp to guide the suture needle through the skin and close the incision.

The number of stitches will vary with the Size of the incision With the animal on its stomach. Position the catheter base on the animal's back. Apply saline to the incision and slide the catheter base under the skin.

Make sure that the tubing is minimally looped and under the base to minimize the chances of it being pierced. Stitch the skin around the catheter base. The number of stitches will depend on the size of the incision.

Usually three to four stitches are sufficient. Make sure that blood can still be drawn up from the catheter. Finally, flush the catheter with the heparinized tein solution to prevent infection and blood clots from forming.

Loosen the tubing connected To the catheter's cannula. Next, remove the saline tube and replace it quickly with tubing from the syringe containing the heparinized T Carin solution. Flush the catheter with 0.03 to 0.05 ccs of solution.

Again, loosen the tubing connected to the cannula. Grip the catheter cap with the forceps, and in once with motion, remove the tubing from the cannula and replace it with the catheter cap. This step must be done quickly to prevent blood from coming up.

Screw on the white catheter cover, making sure that the cover is secure without piercing the cannula tip. Give the mouse subcutaneous injections of analgesic and antibiotic. Inject the analgesic ketoprofen on one side at a dose of five milligrams per kilogram and the antibiotic amikacin on the other side at a dose of 10 milligrams per kilogram.

After discontinuing anesthesia, allow the mouse to recover in a clean cage with easy access to food and water. Reliable intravenous self-administration data can be obtained by following the techniques presented in this video. However, it is important to maintain catheter patency.

This is achieved through careful placement of the catheter into the right atrium, daily flushing of the catheter before and after opera sessions, and making sure that the catheter cannula is constantly capped to prevent the entry of debris.