Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

Podsumowanie

Mouse cardiac transplantation models represent valuable research tools for studying transplantation immunology. The present protocol details mouse heterotopic cervical cardiac transplantation that involves the placement of cuffs on the recipient's common carotid artery and the donor's pulmonary artery trunk to allow for laminar blood flow.

Streszczenie

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after transplantation, and the impact of immunomodulatory therapies on graft rejection. Heterotopic cervical heart transplantation in mice was first described in 1991 using sutured anastomoses and subsequently modified to include cuff techniques. This modification allowed for improved success rates, and since then, there have been multiple reports that have proposed further technical improvements. However, translation into more widespread utilization remains limited due to the technical difficulty associated with graft anastomoses, which requires precision to achieve adequate length and caliber of the cuffs to avoid vascular anastomotic twisting or excessive tension, which can result in damage to the graft. The present protocol describes a modified technique for performing heterotopic cervical cardiac transplantation in mice which involves cuff placement on the recipient's common carotid artery and the donor's pulmonary artery in alignment with the direction of the blood flow.

Wprowadzenie

Abbott et al. published¹ the first description of heterotopic abdominal heart transplantation in rats in 1964. These surgical techniques were refined and simplified by Ono et al. in 1969². Corry et al. first described a method for heterotopic abdominal heart transplantation in mice in 1973; similar to the previously reported rat models, this involved engraftment into the host's abdomen with revascularization by end-to-side anastomoses of the donor's pulmonary artery and ascending aorta to the recipient's inferior vena cava and abdominal aorta, respectively³. Heterotopic cervical heart transplantation in rats was described by Heron in 1971 using Teflon cuffs made from 16 G (1.6 mm outer diameter) intravenous catheters⁴. Chen⁵ and Matsuura et al.⁶ later reported heterotopic cervical heart transplantation in mice in 1991, whose techniques differed primarily in their method of re-anastomosis. Chen's approach involved sutured anastomoses of the donor's ascending aorta to the carotid artery of the recipient and the donor's pulmonary artery to the external jugular vein of the recipient⁵. Due to the advanced technical skill required for these microsurgical sutured anastomoses, a significant amount of time and experience was necessary to achieve a high success rate. Matsuura et al. described a method utilizing a non-suture cuff technique, similar to that used by Heron, which involved end-to-end anastomoses using the extra-luminal placement of cuffs. He fashioned Teflon cuffs from 22 G (0.8 mm outer diameter) and 24 G (0.67 mm outer diameter) intravenous catheters and placed them over the recipient's external jugular vein and common carotid artery, respectively⁶. These cuffs were then placed inside the donor's pulmonary artery and aorta and secured by tying a suture ligature around the connection. This approach translated into an improved success rate. Most importantly, it resulted in a shortening of the time required to complete both cervical anastomoses, thus reducing the warm ischemic time of the graft to less than one-third of that utilizing the abdominal suturing method. Furthermore, since the cuffs are placed around the external surface of the vessel, there is no foreign body exposed to the vessel lumen, which largely reduces the possibility of thrombosis after surgery⁷. Meanwhile, utilization of the cuff technique provides support around the vessels at the site of the anastomosis without requiring any suturing, which reduces the risk of bleeding after revascularization⁶.

Numerous revisions of this technique have been proposed. To accommodate the short length of the mouse common carotid artery (approximately 5 mm), Tomita et al.⁸ developed a modification of this technique with a smaller arterial cuff (0.6 mm outer diameter) while omitting holding sutures and pulling the artery directly through the cuff with fine forceps instead. Wang et al. further simplified this approach by placing 22 G and 24 G cuffs on the donor's right pulmonary artery and recipient's right common carotid artery, respectively⁹. Various reports have described modifications to these approaches, including the use of specialized cuffs, microsurgical clamps, vessel dilators, and cardioplegia^10,11,12. Notably, all of these methods involve the retrograde circulation of blood through the heart, with blood flowing from the recipient common carotid artery to the donor aorta, the coronary arteries, the coronary sinus, then emptying into the right atrium and exiting from the pulmonary artery into the recipient external jugular vein.

Compared to engraftment in the abdomen, cervical cardiac transplantation offers multiple advantages. As previously mentioned, cervical exposure allows for quicker revascularization and shorter warm ischemic times⁶. The cervical method is also less invasive and is associated with shorter postoperative recovery times as it avoids a laparotomy⁶. Importantly, end-to-end anastomoses with cuffs can be performed instead of end-to-side anastomoses, which decreases the risk of complications such as anastomotic bleeding. The abdominal approach also poses an increased risk of developing thrombotic complications in the abdominal aorta or inferior vena cava, leading to spinal cord ischemia and hindlimb paralysis. The superficial cervical location of the transplant allows for easy access to graft viability assessment by palpation, electrocardiography, and invasive or non-invasive imaging. Although the cervical grafts resume spontaneous cardiac activity following reperfusion, they do not significantly impact the systolic and diastolic parameters of the recipient. This model provides valuable insight for studying cellular responses following transplantation, such as ischemia-reperfusion injury and graft rejection. Furthermore, this model offers an ideal approach to allow for post-transplant imaging, such as intravital two-photon microscopy or positron emission tomography (PET) imaging. To this end, our laboratory has previously reported methods to image moving tissues and organs in the mouse, including beating murine hearts and aortic arch grafts following heterotopic cervical transplantation to visualize leukocyte trafficking during ischemia-reperfusion injury and within atherosclerotic plaques, respectively^13,14,15. Additionally, due to its superficial location and ease of exposure, this model is suitable for cardiac re-transplantation¹⁶.

This report describes a technique that allows for laminar blood flow with the external placement of the vascular cuffs on the vessels from which blood flow originates. This allows for a smooth transition of blood flow from one vessel to the next, avoiding the exposure of the distal vessel edge into the vascular lumen. Additionally, the technique utilizes a larger 20 G cuff, instead of previously used 22 G cuffs, for the donor pulmonary artery to ensure ample return of blood flow to the recipient.

Protokół

All animal handling procedures were conducted in compliance with the NIH Care and Use of Laboratory Animals guidelines and approved by the Animal Studies Committee at Washington University School of Medicine. Hearts from C57BL/6 (B6) and BALB/c mice (weighing 20-25 g) were transplanted into gender-matched B6 recipients (6-8 weeks of age). The mice were obtained from commercial sources (see Table of Materials). Syngeneic transplants were performed to evaluate cellular responses related to ischemia-reperfusion injury, and allogeneic transplants were performed to investigate the immune mechanisms involved in graft tolerance and rejection. B6 lysozyme M-green fluorescent protein (LysM-GFP) reporter mice¹⁷, originally obtained from Klaus Ley of La Jolla Institute for Allergy and Immunology, La Jolla, CA, and subsequently bred in our facility, were used as recipients for selected experiments to visualize neutrophil infiltration into cardiac grafts. Survival surgery was performed using aseptic procedures. 

1. Donor procedure

1. Anesthetize the mice by injecting ketamine (80−100 mg/kg) and xylazine (8−10 mg/kg) (see Table of Materials) intraperitoneally into the donor mouse. Confirm surgical plane of anesthesia with toe and tail pinch.
2. Prepare the surgical area by shaving the hair from the chest and abdomen using an electric razor.
3. Administer 100 units of heparin (see Table of Materials) intravenously into the penile vein (males) or external jugular vein (males or females).
4. Place the mice in a supine position with forelimbs overhead. Secure the forelimbs and hindlimbs with surgical tape and disinfect the skin with three alternating scrubs of 0.75% iodine and 70% ethanol.
5. Perform an incision, median laparosternotomy, from umbilicus to sternal angle (3-4 cm), followed by a bilateral thoracotomy along each costal margin (2 cm bilaterally). Fold the anterior chest wall over the neck for full exposure of the mediastinum.
6. Excise the thymus and expose the intrathoracic inferior vena cava.
7. Transect across the width of the abdominal aorta for exsanguination.
8. For retrograde perfusion, inject 1.5 mL of 4 °C saline into the intrathoracic inferior vena cava with the needle oriented superiorly toward the graft, as previously described¹³.
9. Ligate the superior vena cava using an 8-0 silk suture and divide distally.
10. Repeat the retrograde perfusion by injecting another 1.5 mL of 4 °C saline via the inferior vena cava.
11. Ligate the inferior vena cava using an 8-0 silk suture and divide distally.
12. Dissect the aortic arch and pulmonary artery trunk for graft harvesting and transect both distally. Ligate the pulmonary veins on the posterior surface of the heart using a 6-0 silk suture and divide distally.
13. Perform graft preparation by removing the donor heart from the chest cavity. Place the excised heart in a plastic container filled with 4 °C heparinized saline for 1-2 min. Transfer the graft onto a sterile plastic flask filled with ice for cuff placement (Figure 1A).
    NOTE: The heart graft needs to remain on the flask for approximately 5 min to place the donor pulmonary artery cuff.
14. Place a 1 mm long 20 G angiocatheter (see Table of Materials) cuff over the pulmonary artery for the donor cuff. Using fine forceps, gently fold the edges of the artery back over the cuff. Secure the folded vessel to the cuff using a 10-0 nylon tie, as described previously¹⁸ (Figure 1B,C).
15. Store the donor heart in heparinized saline or another preservation solution at 4 °C.
    NOTE: While some may prefer specific preservation solutions (e.g., the University of Wisconsin solution) for prolonged ischemic preservation, it can be costly¹⁹. Saline may be a suitable alternative for short periods of ischemia (<1 h)²⁰. Ultimately, the choice of preservation solution depends on the experimental design²¹.

2. Recipient procedure

1. Inject ketamine (80−100 mg/kg) and xylazine (8−10 mg/kg) intraperitoneally into the recipient mouse for anesthesia. Inject sustained-release buprenorphine (0.5-1.0 mg/kg) subcutaneously for analgesia. Confirm surgical plane of anesthesia with toe and tail pinch.
2. Prepare the surgical area by shaving the hair from the cervical area using an electric razor. Apply sterile, non-medicated ophthalmic ointment to the eyes to prevent corneal drying.
3. Place the animal in a supine position with forelimbs adjacent to the body and the head turned slightly to the left. Secure the forelimbs and hindlimbs with surgical tape. Disinfect the skin with three alternating scrubs of 0.75% iodine and 70% ethanol.
4. Make a midline cervical incision from the lower mandible to the sternum.
5. Transect the right sternocleidomastoid muscle. Excise the right lobe of the submandibular gland to create space for graft implantation.
6. Tie a slipknot over the proximal external jugular vein using a 6-0 silk suture. Ligate the distal external jugular vein and adjacent branches using an 8-0 silk suture. Make a transverse incision across the anterior wall of the external jugular vein.
7. Place a 10-0 nylon suture through the edge of the proximal external jugular vein and the underlying tissue to secure the vein during cuff insertion (Figure 1D).
8. Ligate the distal right common carotid artery using an 8-0 silk suture just inferior to the carotid bifurcation. Tie a slipknot over the proximal common carotid artery using a 6-0 silk suture. Transect the artery distally between the sutures.
9. Similar to the donor cuff, place a 0.6 mm long 24 G angiocatheter cuff over the recipient's right common carotid artery. Using fine forceps, gently fold the edges of the artery back over the cuff. Secure the folded vessel to the cuff using a 10-0 nylon tie.
10. Place the donor heart superior to the right cervical area.
11. Drip cold saline onto the heart graft every few minutes during implantation.
12. Place a 10-0 nylon suture through the edge of the donor aorta and through a superficial bite of the underlying tissue to secure the graft in place (Figure 1E).
13. Flush the donor aorta with 0.5 mL of 0.9% heparinized saline.
14. Insert the recipient's common carotid artery cuff into the donor aorta. Secure the anastomosis with an 8-0 silk tie (Figure 1F). Remove the aortic anchor suture.
15. De-air the external jugular vein by flushing the recipient's external jugular vein with 0.5 mL of 0.9% heparinized saline.
16. Perform pulmonary artery anastomosis by inserting the donor pulmonary artery cuff into the recipient's external jugular vein and secure with an 8-0 silk tie (Figure 1G). Remove the external jugular vein anchor suture and transect the remaining posterior wall of the external jugular vein to free the graft from the underlying tissue. Ensure the graft is properly oriented without kinking or twisting of the anastomoses.
17. Release the slipknots on the recipient's external jugular vein followed by the common carotid artery to initiate heart graft reperfusion (Figure 1H).
18. Close the cervical skin incision using an interrupted 6-0 nylon suture.

3. Postoperative care

1. Place the recipient in a warm recovery chamber immediately following surgery and monitor closely until fully recovered from anesthesia (approximately 1 h).
2. Continue to closely monitor the animal (every 6-8 h) for at least 72 h after surgery for signs of abnormal behavior, such as lethargy, shaking, rapid breathing, or anorexia.
3. For pain control, inject carprofen (5 mg/kg) subcutaneously every 8-12 h for analgesia, in addition to subcutaneous buprenorphine (0.05 mg/kg) every 8-12 h for 24-48 h starting at the end of the surgery.

4. Intravital two-photon imaging of leukocyte trafficking in the heart graft

1. Inject ketamine (80-100 mg/kg) and xylazine (8-10 mg/kg) intraperitoneally into a B6 LysM-GFP recipient mouse¹⁷ 2 h after graft reperfusion for anesthesia.
2. Perform orotracheal intubation using a 20 G angiocatheter, as previously described¹⁸.
3. Connect the angiocatheter to tubing from a mouse mechanical ventilator and ventilate with room air at a rate of 120 breaths/min and a tidal volume of 0.5 mL¹⁸.
4. Inject 12 µL of 655 nm nontargeted Quantum dots (see Table of Materials), suspended in 50 µL of PBS intravenously, as described previously¹³.
5. Reopen the neck incision to expose the heart graft. Place the mouse in a stabilization chamber.
6. Secure a portion of the free wall of the left ventricle using a thin ring of tissue adhesive (see Table of Materials), applied to a glass coverslip attached to the upper chamber plate.
7. Place the chamber under the two-photon microscope objective to acquire images and videos, as described previously¹³.

Wyniki

This mouse cervical heterotopic cardiac transplantation model has been utilized to perform over 1,000 transplants in our laboratory, with a survival rate of approximately 97%. The success rate is slightly higher than previous reports using other cervical heterotopic heart transplantation techniques in mice^10,11,20. This could potentially be attributed to the larger 20 G cuff placed on the donor pulmonary artery to ensure ample r...

Dyskusje

Utilizing this technique, mouse heterotopic cervical cardiac transplantation can be performed in less than 40 min by an experienced microsurgeon and in approximately 60 min by an entry-level microsurgeon. While cervical heart transplantation has been studied in numerous animal models, a mouse model remains the gold standard due to multiple well-defined genetic strains, genetic alteration capabilities, and the availability of numerous reagents, including monoclonal antibodies²⁴. The technique descr...

Materiały

Name	Company	Catalog Number	Comments
6-0 braided silk ties	Henry Schein Inc	7718729
0.75% Providone iosine scrub	Priority Care Inc	NDC 57319-327-0
10-0 nylon suture	Surgical Specialties Corporation	AK-0106
655-nm nontargeted Q-dots	Invitrogen	Q21021MP
70% Ethanol	Pharmco Products Inc	111000140
8-0 braided silk ties	Henry Schein Inc	1005597
Adson forceps	Fine Science Tools Inc	91127-12
BALB/c and C57BL/6 mice (6-8 weeks)	Jackson Laboratories
Bipolar coagulator	Valleylab Inc	SurgII-20, E6008/E6008B
Carprofen (Rimadyl) injection	Transpharm	35844
Carprofen (Rimadyl) oral chewable tablet	Transpharm	38995/37919
Custom-built 2P microscope running ImageWarp acquisition software	A&B Software
Dumont no. 5 forceps	Fine Science Tools Inc	11251-20
Fine vannas style spring scissors	Fine Science Tools Inc	15000-03
GraphPad Prism 5.0	Sun Microsystems Inc.
Halsey needle holder	Fine Science Tools Inc	91201-13
Halsted-Mosquito clamp curved tip	Fine Science Tools Inc	91309-12
Harvard Apparatus mouse ventilator model 687	Harvard Apparatus	MA1 55-0001
Heparin solution (100 U/mL)	Abraxis Pharmaceutical Products	504031
Imaris	Bitplane
Ketamine (50 mg/kg)	Wyeth	206205-01
Microscope—Leica Wild M651 × 6–40 magnification	Leica Microsystems
Moria extra fine spring scissors	Fine Science Tools Inc	15396-00
Ohio isoflurane vaporizer	Parkland Scientific	V3000i
Qdots	ThermoFisher	1604036
S&T SuperGrip Forceps angled tip	Fine Science Tools Inc	00649-11
S&T SuperGrip Forceps straight tip	Fine Science Tools Inc	00632-11
Sterile normal saline (0.9% (wt/vol) sodium chloride	Hospira Inc	NDC 0409-4888-20
Sterile Q-tips (tapered mini cotton tipped 3-inch applicators)	Puritan Medical Company LLC	823-WC
Surflow 20 gauge 1/4-inch Teflon angiocatheter	Terumo Medical Corporation	SR-OX2032CA
Surflow 24 gauge 3/4-inch Teflon angiocatheter	Terumo Medical Corporation	R-OX2419CA
ThermoCare Small Animal ICU System (recovery settings 3 L/min O2, 80 °C, 40% humidity)	Thermocare Inc
VetBond	Santa Cruz Biotechnology SC361931	NC0846393
Xylazine (10 mg/kg)	Lloyd Laboratories	139-236