An Immunological Model for Heterotopic Heart and Cardiac Muscle Cell Transplantation in Rats

Summary

We describe a model of heterotopic abdominal heart transplantation in rats, implying modifications of current strategies, which lead to a simplified surgical approach. Additionally, we describe a novel rejection model by in-ear injection of vital cardiac muscle cells, allowing further transplant immunological analyses in rats.

Abstract

Heterotopic heart transplantation in rats has been a commonly used model for diverse immunological studies for more than 50 years. Several modifications have been reported since the first description in 1964. After 30 years of performing heterotopic heart transplantation in rats, we have developed a simplified surgical approach, which can be easily taught and performed without further surgical training or background.

After dissection of the ascending aorta and the pulmonary artery and ligation of superior and inferior caval and pulmonary veins, the donor heart is harvested and subsequently perfused with ice-cold saline solution supplemented with heparin. After clamping and incising the recipient abdominal vessels, the donor ascending aorta and pulmonary artery are anastomosed to the recipient abdominal aorta and inferior vena cava, respectively, using continuous running sutures.

Depending on different donor-recipient combinations, this model allows analyses of either acute or chronic rejection of allografts. The immunological significance of this model is further enhanced by a novel approach of in-ear injection of vital cardiac muscle cells and subsequent analysis of draining cervical lymphatic tissue.

Introduction

Heterotopic heart transplantation is a frequently used experimental model for different investigations regarding transplantation tolerance, acute and chronic allograft rejection, ischemia-reperfusion injury, machine perfusion or cardiac remodelling. Among other advantages, the graft function can be monitored noninvasively by palpation and graft failure does not lead to a vital impairment of the recipient in contrast to other organs, such as kidneys or livers.

In 1964, Abbott et al. initially described heterotopic abdominal heart transplantation in rats¹. Later, in 1966, the end-to-side technique for anastomoses was described by Tomita et al.². The groundwork for the currently used model was reported by Ono and Lindsey in 1969³. During the last decades, several modifications have been published to create different types of unloaded, partially loaded or loaded left ventricular heart grafts including combined heterotopic heart-lung transplantation^4,5,6. For immunological analyses a non-volume loaded heart graft transplantation is most commonly performed. In this case, the blood flow retrogradely enters the donor ascending aorta and subsequently the coronary arteries. The venous drainage occurs along the coronary sinus into the right atrium and ventricle (Figure 1A-B). Therefore, the left ventricle is excluded from blood flow, apart from marginal amounts of blood from Thebesian veins. This also makes it a useful model for studying the pathophysiological mechanisms during left ventricular assist device therapy⁷.

Heterotopic heart transplantation has been performed in various species including mice, rabbits, pigs and has even been used as a uni- or biventricular assist device in humans^8,9,10,11. The rat still represents a popular experimental animal for transplant models, especially since the graft survival times for different rat strain combinations have been well-defined in the past and a large number of immunological reagents are accessible^12,13. Unlike mice, rats are larger making surgery and access to lymphatic tissue for immunological analyses more feasible¹². Furthermore, the introduction of commercial cloning technologies in rats in recent years will most likely lead to a recurring interest in experimental rat models¹⁴.

In general, heterotopic heart grafts can be attached to the recipient vessels either by performing cervical or abdominal anastomosis. However, a few studies suggest that a femoral anastomosis facilitates improved monitoring due to better access for manual palpation or transfemoral echocardiography and thus allows a more precise detection of graft failure^15,16.

It has been shown that there is no difference regarding operation time, complication rate, outcome and graft survival time between both anastomosis techniques¹⁷. Clearly, the availability of a sufficient number of draining lymph nodes must be mentioned as a benefit of cervical anastomosis; however, longer training periods are required. In contrast, the abdominal anastomosis is less complicated and equally valuable for immunological investigations, especially when combined with results from a novel method of in-ear injection of allogenic cardiac muscle cells and subsequent cervical lymphadenectomy. A combination of both models offers a broad spectrum of post-interventional immunological analyses.

The following protocol refers to operating in pairs of surgeons in order to reduce ischemia time. However, all experiments can be performed by a single person. The setup of instruments and materials for heart explantation and implantation is displayed in Figure 2A-B.

Protocol

All animal experiences have been performed according to the guidelines of the local Ethics Animal Review Board of the regional authorities for consumer protection and food safety of Lower Saxony (LAVES, Oldenburg, Germany) with the approval IDs 12/0768 and 17/2472.

1. Heart explantation and perfusion

NOTE: As graft donors, female or male rats at an age of 7-22 weeks were used.

1. Anesthetize the donor rat by isoflurane inhalation (induction at 5% and maintenance at 3% with an O₂ flow of 1 L/min). Inject 5 mg of Carprofen subcutaneously per kg of bodyweight for perioperative analgesia and check for the absence of the toe pinch withdrawal reflex.
2. Apply eye lubricant and remove the abdominal and thoracic fur using a mechanical clipper.
3. Place the donor in a supine position, fix the limbs at the base of the operation table with elastic bands and sterilize the skin with 70% ethanol or another sufficient alternative.
4. Incise the skin in longitudinal direction and after application of local anesthetic (e.g., lidocaine 0.2%) perform a median laparotomy by using scissors.
5. Insert retractors, mobilize the intestine to the left of the donor, and expose the inferior vena cava with sterilized cotton swabs.
6. For anticoagulation, inject 500 I.U. of heparin dissolved in 1 mL of ice-cold isotonic saline solution intravenously by puncturing the inferior vena cava. Stop the bleeding at the puncture site by light compression with a cotton swab after retraction of the needle (Figure 3A).
7. Incise the diaphragm and perform lateral thoracotomy to both sides of the donor.
8. Pin the mobilized ventral wall of the thorax onto the operation table.
9. Remove the pericardium and the vagal nerve by blunt preparation using two micro-needle holders.
10. Perform transection of abdominal vessels in order to exsanguinate the donor and unload the heart.
11. Insert the blunt branch of a probe pointed scissors into the transvers pericardial sinus and separate the ascending aorta and pulmonary artery as distal as possible under light caudal traction of the heart with a wetted compress (Figure 3B).
12. Place a single 5-0 ligature around the superior and inferior vena cava and the pulmonary veins and tighten it as dorsal as possible (Figure 3C).
13. Sever the tissue dorsal to the ligature and extract the heart (Figure 3D).
14. Perfuse the explanted heart with an 18 G cannula from an intravenous catheter through the ascending aorta and the pulmonary artery with 30 mL of ice-cold, isotone saline solution supplemented with 1000 I.U. of heparin and place the heart in a 15 mL tube filled with saline solution on ice (Figure 3E-F).

2. Heart implantation

NOTE: As recipients, 10-14 weeks old female or male rats were used. Donors and recipients were approximately weight matched.

1. Perform anesthesia of the recipient rat by also using isoflurane inhalation (induction at 5% and maintenance at 1.5-2% with an O₂ flow of 1 L/min). Inject 5 mg of Carprofen subcutaneously per kg of bodyweight for perioperative analgesia and check for the absence of the toe pinch withdrawal reflex.
2. Apply eye lubricant, remove the abdominal fur, fix the limbs and sterilize the skin analogously to the donor preparation. For optimal postoperative outcome, perform the operation on a heating mat to prevent intraoperative hypothermia.
3. After longitudinal incision of the skin, apply a local anesthetic, such as lidocaine (0.2%), on the abdominal fascia. Open the abdominal cavity by median laparotomy and insert retractors.
4. Mobilize the intestine to the upper left side of the recipient and place it in a warm, wetted compress.
5. After mobilizing the duodenum and proximal jejunum, respectively, using the surgical microscope (or magnifying spectacles) with a 5-7x magnification, expose the abdominal aorta and inferior vena cava by blunt preparation with cotton swabs. Do not separate the abdominal vessels.
6. Elevate the abdominal vessels using two micro needle holders without injuring the lumbar veins and position the Cooley vascular clamp (Figure 4A).
7. Puncture the abdominal vessels with a 30-45° arched 27 G cannula (Figure 4B).
8. Enlarge the puncture site using Potts scissors to create a longitudinal incision that matches the size of the lumen of the donor vessels (Figure 4C-D) and perfuse the recipient vessels with saline solution in order to remove clots and prevent postoperative thrombosis.
9. Place the graft into the situs and fixate the donor ascending aorta to the recipient abdominal aorta by two simple interrupted stitches (8-0 monofilament non-resorbable suture) at the cranial and caudal corner of the longitudinal incision (Figure 4E).
10. Anastomose the ascending aorta of the donor with the abdominal aorta of the recipient by a running 8-0 monofilament suture in two steps: first, place the graft to the right of the recipient vessels and perform the first half of the anastomosis (Figure 4E). Subsequently, place the graft to the left of the recipient vessels and perform the second half of the anastomosis (Figure 4F).
11. Fixate the donor pulmonary artery to the inferior vena cava analogously to the aortal anastomosis (8-0 monofilament non-resorbable suture). Suture the first half of the venous anastomosis from the intraluminal side of the vessel (Figure 4G-H).
12. Flush the anastomoses with saline directly before tightening the knots to prevent peripheral embolism.
13. Place a hemostatic gauze around both anastomoses and carefully release the Cooley vascular clamp so that the reperfusion of the graft can begin. Handle bleeding along the anastomoses by light compression with sterilized cotton swabs.
    NOTE: The graft should start beating after around 60 s.
14. Replace the intestine in a meander like fashion. Make sure that there are no malrotations of the mesenteric radix to prevent intestinal necrosis or mechanical obstruction.
15. Close the abdominal muscles/fascia and skin separately using continuous 3-0 polyfilament running sutures.

3. Postoperative care

1. For postoperative analgesia, supply the recipients with an additional subcutaneous injection of 5 mg of Carprofen per kg of bodyweight on the first postoperative day (POD). Additionally, add 1 g of Metamizol to 500 mL of drinking water until the third POD.
2. Start monitoring the heart graft function by daily abdominal palpation on the third POD.
   NOTE: In case of graft failure before the third POD, a surgical rather than an immunological failure should be considered. However, this of course depends on the chosen strain combination and the respective immunological model (e.g., hyperacute rejection after prior immunization).
3. Following graft rejection, extract tissues like the draining retroperitoneal lymph nodes cranial of the anastomoses, the spleen, the blood, the thymus and the graft for further immunological analyses via flow cytometry or immunohistochemistry.

4. Enzymatic digestion of the heart and subcutaneous injection of heart cells in the ear

1. Perform heart explantation and perfusion analogously to heterotopic heart transplantation (see step 1).
2. Shred the heart in 3 mm x 3 mm blocks using a sterile scalpel or sterile scissors and incubate it for 30 min at 37 °C in culture medium containing 0.5 mg/mL collagenase.
   NOTE: It is important to use culture medium containing penicillin, streptomycin and glutamine without fetal calf serum (FCS) particularly as FCS inhibits collagenase digestion.
3. Add the digested tissue to a large-pored sieve, while removing the culture medium and mince thoroughly to get a suspension of vital cardiac muscle cells, mostly dead single heart cells and remaining blood cells. Wash the cell suspension twice with sterile isotonic saline solution.
   NOTE: Centrifugation settings: 10 min, 200 x g, 20 °C
4. Filter the suspension using a 40 µm cell strainer and collect the vital cell congeries by flushing the cell strainer with 5-10 mL of isotonic saline solution.
5. After centrifugation, resuspend the cardiac muscle cells in saline solution dissolved at a concentration of 5x10⁵ cells/mL and draw the cell solution up into a 1 mL syringe.
6. Perform anesthesia analogously to the protocol described for the recipient narcosis (see step 2) for heterotopic heart transplantation.
7. Place the recipient in a lateral position and fix the ear with a finger using double sided tape (Figure 5A).
8. Inject 20 µL of the cardiac muscle cell solution (containing 1 x 10⁴ cells) via a 27 G cannula s.c. close to the visual capillary vessels into the recipient's ear (Figure 5B).
9. After a defined observation period (depending on the chosen strain combination and strength of rejection), extract the draining cervical lymph nodes and perform further analyses such as flow cytometry or co-cultures (Figure 5C).
   NOTE: Furthermore, histological analysis of the pinna can be performed to determine cell infiltration.

Results

In the past, different immunological issues have been addressed on the basis of the model, which was validated in the work group by more than 500 transplants with a survival rate of more than 95%^{13,18,19,20,21,22,23,24}. Tota...

Discussion

The previously described method of heterotopic cardiac transplantation in rats is mainly based on the description of Ono and Lindsey in 1969³. Since then, several modifications have been introduced in various species leading to a wide diversity of this model. Combining several of these modifications and introducing our own experience resulting from over 30 years of performing heterotopic heart transplants in the laboratory, we created a feasible surgical approach, which does not require long train...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Anesthesia device (including isoflurane vaporizer)	Summit Anesthesia Solutions	No Catalog Number available
Cannula (27 G)	BD Microlance	302200
Carprofen	Pfizer	Rimadyl 50 mg/mL
Cellstar Tubes (15 mL)	GreinerBioOne	188271
Cell strainer (40 µm)	BD Falcon	2271680
Collagenase Type CLSII	Biochrome	C2-22
Compresses 5x5 cm	Fuhrmann	31501
Compresses 7.5x7.5 cm	Fuhrmann	31505
Cotton swabs	Heinz Herenz Medizinalbedarf	1032128
Dexpathenol (5 %)	Bayer	"Bepanthen"
DPBS BioWhittaker	Lonza	17-512F
Forceps	B. Braun	Aesculap BD557R
Forceps	B. Braun	Aesculap BD313R
Forceps	B. Braun	Aesculap BD35
Heating mat	Gaymar Industries	"T/Pump"
Hemostatic gauze	Ethicon	Tabotamp
Heparin-Natrium 25 000 I.E.	Ratiopharm	No Catalog Number available
Isofluran CP	CP-Pharma	No Catalog Number available
Large-pored sieve (stainless steel)	Forschungswerkstätten Hannover Medical School	No Catalog Number available
Lidocaine	Astra Zeneca	2 % Xylocain
Metamizol-Natrium	Ratiopharma	Novaminsulfon 500 mg/mL
Micro forceps	B. Braun	Aesculap BD3361
Micro needle holder	Codman, Johnson & Johnson Medical	Codmann 80-2003
Micro needle holder	B. Braun	Aesculap BD336R
Micro needle holder	B. Braun	Aesculap FD241R
Micro scissors	B. Braun	Aesculap FD101R
Micro scissors	B. Braun	Aesculap FM471R
Needle holder	B. Braun	Aesculap BM221R
Penicillin/Streptomycin/Glutamine (100x)	PAA	P11-010
Peripheral venous catheter (18 G)	B. Braun	4268334B
Peripheral venous catheter (22 G)	B. Braun	4268091B
Probe pointed scissors	B. Braun	Aesculap BC030R
Retractors	Forschungswerkstätten Hannover Medical School	No Catalog Number available
RPMI culture medium	Lonza	BE12-702F
Saline solution (NaCl 0.9 %)	Baxter	No Catalog Number available
Scissors	B. Braun	Aesculap BC414
Surgical microscope	Carl-Zeiss	OPMI-MDM
Sutures (anastomoses)	Catgut	Mariderm 8-0 monofil
Sutures (ligature)	Resorba	Silk 5-0 polyfil
Sutures (skin, fascia)	Ethicon	Mersilene 3-0
Syringe (1 mL)	B. Braun	9166017V
Syringe (10 mL)	B. Braun	4606108V
Syringe (20 mL)	B. Braun	4606205V
Vascular clamp	B. Braun	Aesculap FB708R