Blood Circuit Reconstruction in an Abdominal Mouse Heart Transplantation Model

Summary

A novel technique for blood circuit reconstruction in a heterotopic abdominal mouse heart transplantation model is demonstrated.

Abstract

The surgical technique of heterotopic abdominal heart transplantation in mice is a standard model for research in transplantation immunology. Here, the established technique for a modified blood circuit reconstruction in a heterotopic abdominal heart transplantation model is presented. This method uses the intrathoracic inferior vena cava (IIVC) instead of the pulmonary artery of the donor heart for the anastomosis to the inferior vena cava of the recipient. It is facilitating and improving success rates for abdominal heart transplantation in mice.

Introduction

The surgical technique of heterotopic abdominal heart transplantation in mice represents a standard model for research in transplantation immunology^1,2,3. However, it is very challenging to perform and this implicates a restriction to the widespread use of this model^4,5.

In traditional mouse heart transplantation (THTx), the donor aorta and the recipient abdominal aorta are anastomosed while the pulmonary artery is anastomosed to the recipient inferior vena cava^6,7,8.

In this modified mouse heart transplantation technique, the donor aorta is anastomosed to the recipient abdominal aorta and the donor IIVC is anastomosed to the recipient inferior vena cava^(3,4,6) (Figure 2 and Figure 3).

Protocol

All animal experiments were conducted following the guidelines from the directive 2010/63/EU of the European Parliament on protection of animals used for scientific purposes (Ethic committee approved, #G1071/09).

NOTE: Preliminary preparation, anesthesia, post-operative care and monitoring work are the same as performed in traditional surgical methods^1,2,4. BALB/c mice served as heart donors and C57BL/6J as transplant recipients. Mice were aged 8-12 weeks, weighed~30 g at transplantation and were housed under standard conditions.

1. Preparatory steps

1. For anesthesia, give mice inhalative isoflurane (2%) until they fall asleep, followed by intraperitoneal injections of ketamine (100 mg/kg) + xylazine (10 mg/kg) + acepromazine (2 mg/kg). For postoperative analgesia, apply Metamizol (200 mg/kg) p.o. and Carprofen (5 mg/kg) s.c..
   NOTE: The application of antibiotics was abstained on purpose as these substances may influence immunological responses.
2. For surgery, use a set of microscopic instruments including a micro-scissor, micro-forceps, a needle holder and micro hemostatic clamps. An electrosurgical pen is also necessary. Perform sutures using 7/0er, 10/0er and 4/0er nylon types.
3. Place the mouse in a box for isoflurane inhalation (2%) for 40-60 seconds. Determine the depth of anesthesia by squeezing the paw with tweezers. If there is a complete lack of response for this stimulus, go to the next step.
4. Once the mouse has fallen asleep, weigh the mouse.
5. Apply an intraperitoneal injection of ketamine (100 mg/kg) + xylazine (10 mg/kg) + acepromazine (2 mg/kg) to the anesthetized mouse.
6. Clip the abdominal fur and place the mouse on the operation table. Perform disinfection using povidone iodide for 3 times, then properly drape the mouse using a fenestrated surgical towel.

2. Donor operation procedure

1. Use scissors to cut the skin from the neck to the lower abdomen, and peel off the full layer of the skin to the midline of both axillae.
2. Use scissors to cut the muscles of the abdominal wall and gently move the viscera to the left (from the operator's view). Wrap away the viscera with a saline imbibed gauze to safely expose the inferior vena cava.
3. Use a 1 mL syringe to inject 0.4 mL of the heparin solution (contains 500 U heparin) slowly into the lower vena cava and wait for 1 minute before pulling out the needle.
4. Pull out the needle and use a micro-scissor to cut both the inferior vena cava and the abdominal aorta to accelerate exsanguination.
5. Use scissors to open the chest cavity by performing a u-shaped cut; completely expose the heart, lungs and all chest blood vessels.
   1. Expose the thoracic aorta, cut 1/2 of the lumen, and then cut the pulmonary vein to facilitate irrigation and drainage.
   2. Insert an irrigation tube into the opening of the thoracic aorta, inject at least 2 mL of 4 °C cold histidine-tryptophan-ketoglutarate cardioplegia solution (Custodiol HTK solution)⁹ until the pulmonary vein outflow is completely clear and the heart completely stops beating.
6. Pull out the irrigation tube and detach the sternum.
7. Use micro scissors to remove the thymus and to slightly strip the fat around the aortic arch.
8. Use straight and curved forceps to expose and ligate the trunk of the arteria pulmonalis (on the right side of the aortic arch) with a 10/0 suture.
9. Use micro forceps to separate the fat and connective tissue attached to the IIVC, expose and ligate the superior vena cava (on the left side of the aortic arch) with a 7/0 suture and use micro scissors to cut it behind the ligation.
10. Make a 7/0 suture around the base of the heart underneath the aortic arch, the IIVC, and both auricles. Then ligate the pulmonary artery branches and venous lung vessels.
11. Use micro scissors to transect the aortic arch as distal as possible, the lung vessels underneath the ligature and the IIVC near the diaphragm. Remove the heart out of the chest.
12. Place the explanted donor's heart into 4 °C cold HTK cardioplegia solution and preserve temporarily.

3. Recipient operation procedure

NOTE: The initial operation steps are similar to those previously shown for the donor mouse, including anesthesia and disinfection.

1. Perform the abdominal skin cut in a transverse manner, cover the abdominal organs with a wet gauze using saline solution.
2. Use micro forceps to expose the inferior vena cava and abdominal aorta and free them from surrounding fat tissue.
3. Use micro forceps to ligate or electrocauterize side branch vessels (lateral or under the vein/aorta) below the renal vessels.
4. Use clip applicator forceps to position two micro hemostatic clamps at the abdominal part of vein/aorta coming from the right leaving more than 1 cm distance for both aorta/vein to ensure space for the construction of the anastomosis in between them.
5. Use micro scissors to make an incision into the aorta a little nearer to the lower clamp than to the upper clamp. Alternatively, use a 30 G needle to make a small hole and open it up with micro scissors.
6. Position the recipient mouse so that the aorta is facing the operator with the vena cava on the other side. Then place the heart into the abdominal cavity and cover it with a small wet gauze pad.
7. Use a 10/0 suture to adapt and stitch the donor aorta to the recipient aorta starting caudally, make a knot and proceed with a running suture to the top of the incision (about 4-5 stitches). Next, flip the heart over to the right (from the subject's view), cover it again and continue the suture on the left side until reaching the caudal end and knot it.
8. Use an irrigation tube to inject at least 0.5 mL of 4 °C HTK cardioplegia solution to flush the donor's IIVC.
9. Use micro-scissors to cut a round hole on the abdominal inferior vena cava of the recipient, which should have the same size of the donors IIVC lumen. The incision should be located above the aortic anastomotic opening. Make the vein incision larger than the aortic incision.
10. Use a 10/0 suture to sew the donor IIVC to the recipient vena cava starting caudally. Tie a knot and perform a running suture until the top of the incision is reached. Use five stitches and continue the suture on the left. Finally, tie a knot in the tail corner, and carefully tighten (be careful not to pull too tight).
11. Place the small parts of the hemostatic sponge around the vein and aortic anastomoses.
12. Use clip applicator forceps to remove first the lower and then the upper micro-hemostatic clamps and rinse the abdominal cavity with 38.0°C tempered 0.9% sodium chloride.
13. Use micro forceps to take away the hemostatic sponge.
14. Observe the heartbeat of the transplanted heart.
15. Use forceps to put the intestines back into the abdominal cavity and two-layer sutures (abdominal muscles followed by skin) to close the abdominal wound with a 4/0 suture.
16. Put the mice into an oxygen and temperature control workstation chamber (e.g., INVIVO2-400) to provide a warm and oxygen rich environment for the transplanted mice to recover, wait for the mice to wake up.
17. For postoperative analgesia, directly give Metamizol 200 mg/kg per os after operation. Four and 16 hours after operation give Metamizol 200 mg/kg per os+ Carprofen (5mg/kg) s.c. In the further followup, give Carprofen (5 mg/kg) s.c to the transplanted mice every 24 hours for three consecutive days after operation.

Results

Here, a modified technique of heterotopic abdominal heart transplantation in mice that has been previously developed in our laboratory and has proven useful for the last 16 years is presented. Previously, it was reported that in overall 40 cases of vena cava to vena cava (V-V group) compared to 40 cases of the traditional pulmonary artery to vena cava (P-V group) anastomosis procedure⁴ (Table 1) the vessel anastomosis took 20.8±1.3 min in the V-V group, which was si...

Discussion

The surgical technique of heterotopic abdominal heart transplantation in mice is very challenging and this implicates a restriction to the widespread use of this model.

One of the disadvantages of the conventional technique is the limiting length of the donor's pulmonary artery (PA). It is usually of about 2 mm of length, whereas the length of the IIVC of the donor heart used in our model is generally about 1 cm (Figure 2). That means that in the modified mode...

Materials

Name	Company	Catalog Number	Comments
30G-needles	Braun	456300
acepromazine	CP Pharma	Tranquisol P
BALB/c AnNCrl mice	Charles River. Germany	no catalog number
Bepanthen eye ointment	Haus-Apotheke	PZN 01578675
Bonn Micro Forceps	FST	11083-07
Box for insulation and oxygen supply device	RUSKINN	INVIV
C57BL/6J  mice	Charles River. Germany	no catalog number
Carprofen	Zoetis	Rimadyl 50 mg/ml
CATHETER-FEP 26G	TERUMO	Surflo-W
Clip Applicator Forceps Style	FST	18057-14
Curved forceps	WPI	14114-G
custodiol/HTK	Dr. Franz Köhler Chemie	no catalog numer
Cutasept skin disinfection	VWR	BODL980365
electrosurgical pen	Bovie	CHANGE-A-TIP
gauze pads, cotton swabs	Lohmann-Rauscher	13353
Heating mat	THERMO MAT PRO 30W	HTP-30
Hemostatic sponge	CuraSpon	J1276A
heparine-solution	Haus-Apotheke	PZN 03029820
Ice box	PETZ	No Catalog Number available
Inhalation anesthesia device	GROPPLER	BKGM 0616
insulation and oxygen supply device	RUSKINN	INVIV
isoflurane	CP Pharma	Isofluran CP 1 ml/ml
ketamine	Zoetis	no catalog numer
metamizole	WDT	no catalog numer
Micro scissors	FST	15000-00,15000-10
Micro Serrefine ( Clamp ) Angled / 16 mm	FST	18055-06
Microscope	Leica	LEICAMZ6
Microscope light	SCHOTT	KL2500LED
Saline solution (NaCl 0.9%)	Haus-Apotheke	PZN 06178437
Scissors	Peha Instruments	991083/4
small Petri dish	Sarstedt	833900
Straight forceps	WPI	14113-G
surgical tape	BSN	4120
Suture Tying Forceps - 10 cm	FST	18025-10
Sutures(10-0)	Medtronic	N2540
Sutures(4-0)	ETHILON	V4940H
Sutures(7-0)	ETHILON	1647H
Syringe (0.3 mL)	BD	324826
Syringe (1 mL)	BD	320801
xylazine	Bayer	Rompun