A Mouse Aortic Interposition Model to Study Alloimmune-Induced Vascular Rejection

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

Use Balb/cYJ (H2^d) donor mice and C57Bl/6 (H2^b) recipient mice to examine allogeneic reactions. Mice are used for experiments between the ages of 8 to 12 weeks. Use either female or male mice. Syngeneic controls consist of aortic segments from C57Bl/6 donors into C57Bl/6 recipients.

1. Donor and Recipient Preparation

NOTE: Both the donor and recipient are anesthetized and prepared before the surgery to minimize ischemia of the graft. Injectable anesthetics are used in the protocol to prevent obstruction of the animal by equipment needed for the delivery of inhaled anesthetics. However, if desired, an inhaled anesthetic is an appropriate alternative. From the initial injection of the anesthetics, the entire procedure takes approximately 90 min to complete. The ischemic time of the graft is less than 30 min.

1. Anesthetize the mouse with intraperitoneal injections of ketamine (100 mg/kg; 10 mg/ml) and xylazine (10 mg/kg; 1 mg/ml). The mouse will be sedated within 10 to 15 min. Assess the depth of anesthesia by pinching the fatty part of the animal foot pad. Administer 1/3 of the original dose of the anesthetic cocktail, as needed, until the animal does not exhibit a withdrawal reflex. Monitor the respiratory rate closely after every cocktail is administered. During surgery, assess the level of anesthesia every 15 min by pinching the anterior abdominal wall with a pair of forceps. The mice should remain deeply anesthetized for 60 to 90 min.
2. Lubricate the mouse's eyes with an ophthalmic ointment to prevent dryness while under anesthesia.
3. Shave hair as close to the skin as possible on the abdominal ventral region from the mid-thorax to the pubis. Be particularly careful not to nick any skin. Do not use depilatory cream because it may be absorbed into the skin and could be inflammatory.
4. Place the animal supine on a towel above the heating table or pad.
5. Clean the surgical site with a preliminary scrub with 2% chlorohexidine. Prepare a large working area to maximize the surgical field. Start scrubbing at the center of the surgical site and move to the outside linearly or circularly. Dispose of the gauze. Repeat this procedure at least 5 more times. Apply an alcohol prep pad to the same area. Once the alcohol is dry, prepare the clean area with Betadine solution and drape it with sterile gauze.

2. End-to-end Anastomosis Procedure

1. Donor Operation
   1. Maintain an aseptic technique throughout the operation. Clean all countertop and surgical table surfaces with 0.5% accelerated hydrogen peroxide solution before use. Wrap and autoclave all surgical instruments, gauzes, drapes, and gowns before use. Verify the sterility of the instruments with a steam sterilizer indicator strip placed in each pack. Sterile surgical gloves are used and disposed of between surgeries. For multiple surgeries, sterilize the surgical instruments between uses with the hot glass bead sterilizer.
   2. Place the donor mouse supine on a clean, thin Plexiglas board wrapped with a sterile drape under the operating microscope at 8-30X magnification.
   3. After ensuring adequate surgical anesthesia as outlined in section 1.1, proceed with surgery.
   4. Using sterile scissors, make a single midline lower longitudinal abdominal incision, from the pubis to the xyphoid process.
   5. open the abdominal walls to expose the cavity using a small retractor.
   6. Using sterile cotton-tipped applicators, gently retract the intestines superiorly to the animal's left and cover with gauze moistened with saline solution. Move the reproductive organs inferiorly and locate the infrarenal aorta and the inferior vena cava (IVC). Moisten the exposed tissues periodically with saline solution.
   7. Using the medical No. 5 forceps, separate the aorta from the IVC, from the level of the left renal artery to the bifurcation. Use 10-0 polyamide monofilament sutures to ligate the small branches near the aorta.
   8. Once the donor's aorta has been separated from the IVC, saturate the vessel with saline, cover the exposed cavity with moistened gauze, and set the donor aside in a sterile area. Check the status of the donor (respiratory and cardiovascular function and depth of anesthesia) every 15 min. Begin operating on the recipient, isolating the aorta as described below (Steps 2.2.1 to 2.2.7).
   9. Once the recipient aorta has been separated from the IVC and set aside, return the donor under the microscope. Cross clamp (proximal and distal to the segment of interest) the donor aorta, approximately 5 mm apart, with two 4 mm microvascular clamps.
   10. Using the Vannas-Tubingen microscissors, transect a small graft segment (3 to 4 mm long) of the abdominal aorta.
   11. Using a 25 G 5/8 needle attached to a syringe, flush the excised aorta with heparinized (100 U/ml) saline solution. Ensure the needle tip does not come into contact with the vessel.
   12. Place the aorta in heparinized (100 U/ml) saline solution on ice and set aside. While still under deep anesthesia, release the microvascular clamps. Euthanize the donor by exsanguination.
   13. Implant the donor vessel within 30 min of excision. Although it is possible to use one donor vessel for multiple recipients by excising a larger length of the aorta, keep the ischemic time to less than 30 min.
2. Recipient Operation
   1. Maintain an aseptic technique throughout the operation, as in donor operation.
   2. Place the recipient mouse supine on a thin Plexiglas board wrapped with a sterile drape under the operating microscope at 8-30X magnification.
   3. After ensuring adequate anesthesia, proceed with surgery when the animal does not exhibit withdrawal reflex.
   4. Using sterile scissors, make a single midline lower longitudinal abdominal incision, from the pubis to the xyphoid process.
   5. Open the abdominal walls to expose the cavity using a small retractor.
   6. Using sterile cotton-tipped applicators, gently retract the intestines superiorly to the animal's left and cover with gauze moistened with saline solution. Move the reproductive organs inferiorly and locate the infrarenal aorta and the IVC. Moisten the exposed tissues periodically with saline solution.
   7. Separate the aorta from the IVC, from the level of the left renal artery to the bifurcation. Use 10-0 polyamide monofilament sutures to ligate the small branches near the aorta if necessary.
   8. Cross clamp (proximal and distal to the segment of interest) the aorta, approximately 5 mm apart, with two 4 mm microvascular clamps.
   9. Using the Vannas-Tubingen microscissors, make a single horizontal aortotomy and resect a small segment (no more than 0.5 mm) of the abdominal aorta to accommodate the donor aortic graft.
   10. Flush the excised aorta with heparinized (100 U/ml) saline solution. The donor aortic graft should be of appropriate length to connect the recipient's transected aortic ends.
   11. Place the donor aortic graft in the orthotopic position and anastomose the donor's graft end to the recipient's end, matching the respective graft anatomical orientation with the recipient's.
   12. Gently grasp the tunica externa of the vessel and evert it slightly using the medical No.5 forceps. Using the forceps, drive the needle attached to the 10-0 polyamide monofilament suture through the full thickness of the vessel wall to secure the donor aortic graft to the recipient's resected vessel. Ensure that the vessel opening is not closed off due to inadvertent stitching of the vessel's back wall.
   13. For continuous stitches, place stay sutures at 9 o'clock in both the upper and lower ends of the graft. Starting at the upper end of the graft from 3 o'clock, anastomose the resected ends with 2 running sutures and secure the suture to the stay suture.
       1. Flip the graft over and continue the running suture to the dorsal part of the vessel, meeting the origin stay suture. Secure without applying much pressure on the vessel. Repeat the suturing for the lower anastomosis.
          Note: The interrupted sutures start the same way as the continuous sutures, except that the vessel is anastomosed with three separate stitches between the stay sutures. Anastomosis time is usually 20 min.
   14. Once the anastomosis is complete, release the distal microvascular clamp to allow retrograde blood to flow and check for leakage of the anastomosed sites. If there is a leakage, immediately place a stitch to close the defective site. If there is no bleeding at the sites, release the proximal clamp.
   15. Examine the transplant and check that there is no blood obstruction in the graft and the proximal and distal portions of the recipient's vessel. A vigorous pulse pattern in the donor's and recipient's vessels indicates that the blood flows freely. Using the pair of forceps, gently grasp one end of the stay sutures and slightly evert the vessel to inspect the back wall of the vessel.
       NOTE: There should be no puckering of the vessel wall at both ends of the anastomosed sites. Poor blood flow after removal of the clamps is a sign of thrombosis.
   16. Return the intestines into the abdominal cavity using cotton-tipped applicators.
   17. With the Castroviejo needle holder and Graefe forceps, close the abdominal wall with 5-0 polypropylene sutures using continuous stitching. Close the skin layer with the same sutures using subcuticular closure.
   18. Administer Torbugesic (1 mg/kg) i.m. immediately upon completion of the transplant.
   19. Give immediately, in the order, Atipamezole (1 mg/kg), ketoprofen (5 mg/kg), and warmed Lactated Ringer's solution subcutaneously.
   20. Immediately after surgery, place mice in a heated cage under a water blanket overnight (12 hr). During the anesthetic-recovery period, place the animals alone in a clean, dry, unobstructed area. Line the cage (autoclaved) with clean paper towels and adjust the temperature of the water blanket to approximately 20 to 22 °C. Provide dry and wet kibbles ad libitum on the cage floor.
       NOTE: An essential component of post–surgical care is observing the animal and appropriate intervention, as required, during recovery from anesthesia and surgery. The necessary monitoring intensity will vary with the animal and might be greater during the immediate anesthetic recovery period than later in postoperative recovery.
   21. Continually monitor animals that have undergone anesthesia until they recover completely. The animal must be able to maintain unassisted sternal recumbency, and it must appear calm and free of pain before being left unattended.
   22. Give Buprenorphine (0.1 mg/kg BID) and ketoprofen (5 mg/kg SID) for three days, both subcutaneously.
   23. Monitor cardiovascular and respiratory function, body temperature, and postoperative pain or discomfort during recovery from anesthesia for at least three days. Additional care may be required, such as administering analgesics and other drugs and parenteral fluids to minimize dehydration and electrolyte loss.
   24. Assess the success of the transplant surgery by observing the motor function of the hind limbs. The complete success of the transplant involves unobstructed blood flow to the hind limb and tail, which should be immediate upon recovery of the animal, and full recovery with no paralysis of the hind limbs on the second day.