1.1 The method of cardiac transplantation in mice

1.1.1 Introduction

The mouse heterotopic fully vascularized cardiac transplantation model has come into widespread use since its introduction by Drs. Robert Corry and Paul S. Russell in 1973 (1). This model is of particular value because it allows the fate of fully vascularized organ grafts to be evaluated in a species where a large number of genetically well-characterized strains and newer transgenic and gene knockout animals are available and for which there exists a very large number of immunological reagents.


In this model the donor ascending aorta is anastomosed end-to-side to the recipient abdominal aorta and the donor pulmonary artery is sutured to the recipients inferior vena cava (IVC). Hearts transplanted heterotopically behave functionally as aorto-caval fistulae. Blood enters the donor ascending aorta from the recipient abdominal aorta and is diverted into the coronary arteries by the closed aortic valve. After the myocardium is perfused, venous blood drains into the right atrium through the coronary sinus and is pumped back into the recipients IVC by the right ventricle.


The procedure is carried out using ideally an operating microscope (such as Carl Zeiss OPMI1-FC or OPMI6-CH) at a magnification of between 4 x - 25 x. Operating time is less than one hour and overall operative mortality is less than 5% in experienced hands.


Details of the procedure are as follows: Anaesthesia and Analgesia
Hypnormィ (Jensen Pharmaceutical Ltd., UK) contains fentanyl citrate and fluanisone. The Hypnormィ stock solution is diluted with sterile saline to a working solution containing 10オg/ml fentanyl and 330オg/ml fluanisone. Midazolam (Hypnovalィ, Roche Products Ltd., UK) is diluted from the stock solution with sterile saline to give a working solution of 5000オg/ml. Recipient mice are injected subcutaneously with 0.3ml of both Hypnormィ and midazolam working solutions before procedure. Donor mice receive 1ml of each anaesthetic. Recipient preparation
Recipient mouse is induced to urinate by placing gentle pressure on the lower abdomen before being anaesthetised. A midline abdominal incision is made from the pubic symphysis to the xyphoid. If the urinary bladder is still filled, it is emptied using a 25 gauge needle and 1 ml syringe. A self-retaining retractor is placed to expose the abdominal contents. Using two sterile cotton buds the small intestine is gently reflected superiorly onto a piece of moistened sterile gauze on the thoracic cavity. Testes are reflected inferiorly in the same fashion. The testes and intestines are covered with the gauze, and are kept moist throughout the procedure with sterile saline. The meso-sigmoid is divided allowing a strip of moist gauze to pass under the sigmoid colon. The strip of gauze is now used to reflect the sigmoid colon to the animal's left side.


Using two cotton buds the abdominal aorta and IVC are dissected free from the surrounding tissues in the retroperitoneum exposing the lumber arteries and veins which lie directly posterior coming from the great vessels in two or three groups between the renal vessels and the bifurcations inferiorly. The two or three groups of lumber vessels are ligated individually with 7-0 silk ties. If the recipient is male, you find two vessels on the IVC. These vessels are isolated from the IVC. Scoville-Lewis clamp (Downs Surgical Ltd., UK) is positioned at the proximal side of bifurcation interrupting flow in both the aorta and IVC. Another clamp is positioned just below the renal vessels in a similar fashion. Clamping in this order ensures partially filled vessels which will facilitate the aortotomy and venotomy. If you find the IVC contimues to fill after cross clamping, the ligation of lumber vessels shoud be checked. Under ideal conditions cross-clamping for as long as two hours is acceptable. Donor operation
The donor's abdominal cavity is opened with a large longitudinal incision. Abdominal contents are reflected to the left side of the animal exposing the IVC. Using 1 ml syringe and 25 gauge needle, the IVC is canulated and as much as blood possible is aspirated. At this point, 1 ml of cold heparin (Injection BP, LEO Laboratories Ltd., UK, diluted to 18,000U/ml final concentration with sterile saline) is administered via the IVC using another 1ml syringe and 25 gauge needle. The thoracic cavity is opened by cutting through the ribs along both sides of the thoracic spine all the way to the thoracic inlet. The anterior chest wall is reflected superiorly and held in place with a piece of adhesive tape. The IVC, azygous vein and the superior vena cava are ligated with 7-0 silk ties (Figure 1). The ascending aorta is cut below the brachiocephalic artery (Figure 1). The main pulmonary artery is cut proximal to its bifurcation (Figure 1). Anastomosis of the donor to recipient vessels is made considerably easier if the connective tissue between the acsending aorta and pumonary artery is gently teased awa at this stage. The pulmonary veins are ligated as a group with a single 7-0 silk tie (Figure 1). The donor heart is gently detached from the remaining connective tissues with blunt dissection and is placed in cold saline. Implantation of the vascularised heterotopic heart graft
The donor heart is placed into the recipients abdomen with the remnant of the ascending aorta lying towards the left side of the animal. An aortotomy is made in the abdominal aorta of the recipient using a 30 gauge needle. The aortotomy is then extended with micro-scissors superiorly to a length of equal to or slightly smaller than the opening of the donor ascending aorta. The donor ascending aorta is anastomosed to the recipient abdominal aorta using 10-0 Bear surgical suture on around-bodied 4mm (3/8) needle (Kyowa Precision Instruments Corp., Japan). A stay suture is placed at the proximal apex bringing the donor ascending aorta and the recipient abdominal aorta together (Figure 2-1). The ends of the stay suture are cut to a length of approximately 2mm and 5mm. Similarly, the distal apex of the donor aorta and the recipient aorta are brought together with a10-0 suture and tied leaving a 5mm length of suture on one end and the needle on the other end. The anastomosis is then completed with the same suture in a counter-clockwise direction with a continuous running stitch (Figure 2-2). The distance from the distal apex to the proximal apex is covered with four stitches. The stitch is tied to the proximal stay suture. The second side of the anastomosis is completed in a similar manner continuing from the proximal apex to the distal apex and is tied to the distal stay suture (Figure 2-3).


The donor pulmonary artery is now anastomosed to the recipient IVC. A venotomy is made in the same way as the aortotomy made at the level inferior to the aortotomy leaving a space between the two of approximately 3mm, which allows easier access for anastomosing the second side (Figure 2-4). In contrast to the recipient's aorta anastomosis, a stay suture is first placed at the distal apex bringing the donor pulmonary artery and the recipient IVC together. The ends of the stay suture are tied leaving a 5mm length of suture on one end and the needle on the other end (Figure 2-5). Similarly, the proximal apex of the donor pulmonary artery and the recipient IVC are brought together with a10-0 suture. The ends of the proximal stay suture are cut to a length of approximately 2mm and 5mm (Figure 2-6). Stay sutures in this order ensure the same length of both sides of donor pulmonary artery which makes the anastomosis more straightforward. The anastomosis is then completed with the distal stay suture in a clockwise direction with a continuous running stitch. The distance from the distal apex to the proximal apex is covered with four throws (Figure 2-7). The stitch is tied to the proximal stay suture. At this point the needle is passed between the two donor vessels and the donor heart is flipped from the left side of the animal to the right (Figure 2-7). The second side of the anastomosis is completed in a similar manner continuing from the proximal apex to the distal apex and is tied to the distal stay suture. The anastomosis is done similar to the aortic anastomosis except that the distal stay suture is done first and that a continuous running suture goes in clockwise fashion (Figure 2-8).


The distal clamp is removed first to check the anastomosis of the IVC. After appoximate 20 to 30 seconds the vessel will fill and the anastomosis will become competent. The small pieces of moistened haemostatic agent Spongostan (Johnson and Johnson Ltd., UK) are placed around each anastomosis. After ensuring adequate haemostasis the proximal clamp is slowly removed. Normally the donor heart will fill with blood immediately, become bright red in colour, and begin to contract. Provided that there is no further bleeding, the intestines and testes may be returned to the abdominal cavity. The abdominal incision is closed in two layers with 4-0 chromic catgut with needle (Davis and Geck, UK). Separately, the muscle and skin layers are closed with a continuous running stitch. Recovery
On completion of the procedure, the recipient is placed on soft dry bedding material under a warming lamp for 2 to 4 of hours with easy access to food and water.

1.1.3 Additional usuful points

(1) 4 or 5 stitches on each side excluding stay sutures are enough and sufficient. The next and last stitches of continuous running suture between donor and recipient aorta should be placed with a short bite and close to the stay suture to avoid stenosis of recipient aorta.
(2) Two stay sutures must be located on the opposite side. If the distal stay suture on IVC is first performed, you can recognise easily where the proximal stay suture should be.
(3) Some space between proximal end of venotomy and distal end of aortotomy should be prepared. Otherwise, the second side of IVC anastomosis can not be performed safely and confidently.
(4) The best length of venotomy and aortotomy is the same as the opening of pulmonary artery and ascending aorta of donor. However, smaller is much better than larger.
(5) Loose sutures are better than over-tight ones. It is much easier to control a small amount of bleeding than to attempt to open an occluded vessed.

1.1.4 Trouble shooting

(1) After release the distal clamp, check the bleeding from the anastomosis of recipient IVC and donor pulmonary artery. If the bleeding continues as the IVC fills, it can be controlled with haemostatic sponge. Defects in suturing may be repaired at this point.
(2) After releasing the proximal clamp, there is usually some bleeding from the anastomosis of the recipient and donor aorta. This usually resolves spontaneously. If bleeding continues, the proximal clamp can be replaced and the sutureing defect repaired.
(3) If the bleeding is from the atrium, it can be ligated at the base with a 7-0 silk tie.
(4) If you find that the bleeding is from a coronary vessel, haemastatic sponge can be applied. This is sometimes successful.
(5) If blood loss is severe, 1ml of warm saline shoulc be given intraperinearlly or subcutaneously.
(6) If you find both lower legs paralyzed after operation, there is no chance of survival because of occlusion of the abdominal aorta at the anastomosis. If you find one lower leg paralyzed, there is some chance of survival as this might be due to the air or fat embolism.

1.1.5 Assessment of graft survival

The function of the transplanted hearts is followed by abdominal palpation . Absence of a palpable heart beat is considered as rejection. This is normally easy to determine. However, in some cases the decision can be more difficult. In this case electrocardiogram (2) or direct visualisation of the graft is invaluable.

1.1.6 2nd cardiac allografts into the abdomen or neck

Replacement of the primary graft with a second heart is possible (3). A second heart can also be transplanted into the recipient's neck. Briefly, the donor ascending aorta is anastomosed end-to-side to the recipient carotid artery and the donor pulmonary artery is sutured end-to-side to the recipient external jugular vein. If a cuff technique is availabe, both anastomoses can be completed using 22 and 24 gauge cuffs for recipient vein and artery, respectively. This approach saves time and increases the success rate (4).

1.1.7 Non-vascularized cardiac allografts into the ear

Prior to the introduction of the vascularized cardiac graft model in the mouse, neonatal non-vascularized cardiac grafts were transplanted into the ear using the method originally described by Fulmer et al (5). Briefly, the heart is excised from newborn mice and implanted into the dorsal base of the pinna of the ear. The implant is subcutaneous and surgical incision is gently closed. The majority of grafts in normal syngeneic mice start to present contractile activity 7 to 10 days after transplantation.

1.2 The results of cardiac allografts in mice

1.2.1 MHC disparities

The rejection of heart grafts transplanted between MHC disparate mice is shown in Table 1. Usually allogeneic cardiac grafts in naive mice are rejected in approximately 7 to 12 days. It is clear that both MHC and non-MHC factors can play a role in the rejection of MHC incompatible heart grafts (6). Congenic strains have therefore been extremely useful in attempts to determine the relative importance of these factors in allograft rejection. It is important to note that in contrast to the mouse skin transplant model, time taken to reject a heart grafts is unpredictable. For example, in a small number of strain combinations, unmodified recipients accept MHC mismatched heart grafts indefinitely(6-8).

1.2.2 Limited MHC disparities

The survival of murine heart grafts transplanted across limited MHC barriers is shown in Table 2. Hearts transplanted across isolated class II incompatibilities (IA/IE) are usually rejected but have been reported to survive for more than 50 days in only 1 donor recipient combination(Table 2, groups 12-14). In contrast, hearts transplanted across isolated class I MHC barriers(K or D) survived for greater than 50 days in 7 of the 10 donor recipient combination tested(Table 2, groups 1-10). This suggests that in the mouse class II is the dominant MHC antigen barrier in unmodified recipients.

1.2.3 Non-MHC disparities

Initially it was thought that although transplantation across multiple non-MHC incompatibilities would result in the rejection of a primarily vascularized cardiac allograft, the rejection response elicited by multiple non-MHC antigens would not be as powerful as that evoked by a full MHC mismatch(12,13). However, it has been shown that multiple non-MHC disparities can induce acute rejection, both of tissue grafts(6) and vascularized heart grafts(6,10,14) and that the tempo of rejection is similar to that induced by full MHC incompatibilities (Table 3).

1.2.4 Transgenic and knockout or gene deleted mice

One of the advantages for using the mouse model is the availability of many types of knockout and transgenic animals. Campos et al.(16) indicated the importance of MHC class II locus products in the rejection of cardiac allografts by using MHC class I, MHC class II and both class I and II deficient mice(Table 4). Qian et al.(17) reported the importance of MHC class I in initiating second-set allograft rejection again using mice deficient in MHC class I and class II genes. A recent study by Krieger et al.(18) examined the survival of adult heart grafts in CD4, and CD8 knockout recipients. The data obtained indicated that CD4+ but not CD8+ T cells are required to initiate allograft rejection, supporting data from earlier studies(19,20).
Cytokine transgenic and knockout mice have also used on cardiac transplantation models(21-23).


Figure 1
Donor operation

Figure 2
Implantation of the vascularised heterotopic heart graft


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