En este trabajo se presenta un protocolo de evaluación de un corazón implantado heterotópicamente tras la preservación ex situ normotérmica en el modelo de rata.
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Artículo de método
Modelo de rata de trasplante cardíaco heterotópico con perfusión ex-situ normotémica
* These authors contributed equally
En este artículo
Aviso de erratum
Important: There has been an erratum issued for this article. View Erratum Notice
Resumen
Resumen
El trasplante de corazón es la terapia más eficaz para la insuficiencia cardíaca terminal. A pesar de las mejoras en los enfoques e intervenciones terapéuticas, el número de pacientes con insuficiencia cardíaca que esperan un trasplante sigue aumentando. La técnica de conservación ex situ normotérmica se ha establecido como un método comparable a la técnica convencional de almacenamiento estático en frío. La principal ventaja de esta técnica es que los corazones de los donantes pueden conservarse hasta 12 h en condiciones fisiológicas. Además, esta técnica permite la reanimación de los corazones del donante después de la muerte circulatoria y aplica las intervenciones farmacológicas necesarias para mejorar la función del donante después de la implantación. Se han establecido numerosos modelos animales para mejorar las técnicas de preservación normotérmica ex situ y eliminar las complicaciones relacionadas con la preservación. Aunque los modelos de animales grandes son fáciles de manejar en comparación con los modelos de animales pequeños, es costoso y desafiante. Presentamos un modelo de rata de preservación normotérmica ex situ del corazón de donante seguido de trasplante abdominal heterotópico. Este modelo es relativamente barato y puede ser realizado por un solo experimentador.
Introducción
El trasplante cardíaco sigue siendo la única terapia viable para la insuficiencia cardíaca refractaria 1,2,3,4. A pesar de un aumento constante en el número de pacientes que necesitan trasplante cardíaco, no se ha observado un aumento proporcional en la disponibilidad de órganos de donantes5. Para abordar este problema, se han desarrollado enfoques novedosos para preservar los corazones de los donantes con el objetivo de mejorar los desafíos y aumentar la disponibilidad de donantes 6,7,8,9.
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Protocolo
El comité de ética del Centro de Investigación con Animales de Laboratorio del Hospital Universitario Nacional de Chonnam (aprobación no. CNU IACUC - H - 2022-36) aprobó todos los experimentos con animales. Las ratas macho Sprague-Dawley (350-450 g), utilizadas en este estudio, recibieron cuidados de acuerdo con las directrices para el cuidado y uso de los animales de laboratorio. Las ratas fueron alojadas en habitaciones con temperatura controlada con un ciclo de luz-oscuridad de 12 horas, con comida y agua estándar disponibles.
1. Preparación
NOTA: Un solo experimentador puede llevar a cabo todos....
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Resultados
La Figura 1 ilustra el diseño experimental utilizado en un modelo de animal pequeño. La figura 2 muestra el aparato de perfusión de Langendorff modificado, que incluye un oxigenador para animales pequeños. El orden de la anastomosis para el implante abdominal heterotópico se presenta en la Figura 3.
En la figura 4 se muestran los parámetros utilizados para evaluar la viabilid.......
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Discusión
Nuestro objetivo al establecer este modelo fue replicar el trasplante de corazón humano normotérmico. Los modelos sin eyección son la técnica comúnmente preferida para preservar el corazón del donante en un ambiente ex situ 16. Si bien los modelos de eyección ofrecen muchas ventajas en la evaluación de la función cardíaca durante la perfusión ex situ 17, no son adecuados para los modelos de trasplante heterotópico. En el trasplante heterotópico, el corazón.......
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Divulgaciones
Los autores no tienen conflictos de intereses.
Agradecimientos
Este trabajo contó con el apoyo de una subvención B2021-0991 del Instituto de Investigación Biomédica del Hospital Universitario Nacional de Chonnam y NRF-2020R1F1A1073921 de la Fundación Nacional de Investigación de Corea
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Materiales
| Nombre | Empresa | Número de catálogo | Comentarios |
|---|---|---|---|
| Sistema de evacuación activa AES | Smiths medical | PC-6769-51A | Utilice CO2 y exceso de isoflurano |
| Máquina de anestesia | Smiths medical | PC-8801-01A | Mezcla isoflurano y oxyegn y entrega al animal |
| B20 monitor | de paciente GE medical systems | B20 | para observar la presión aórtica media y la temperatura |
| Sistema de monitoreo homeotérmico | Aparato de Harvard | 55-7020 | Para controlar y mantener la temperatura de los animales |
| Micro-1 Oxigenador de ratas | Dongguan Kewei Instrumentos médicos | Micro-MO | Para el intercambio de gases en el circuito de Langendorff |
| Introductor de micropuntura Set | COOK medical | G48007 | para administrar solución cardiopléjica al arco a través de la aorta abdominal |
| Microscopio | Amscope | MU1403 | Para zoom quirúrgico campo (Receptor) |
| Lupa quirúrgica | SurgiTel | L2S09 | Para hacer zoom campo quirúrgico (Donante) |
| Bomba de jeringa | AMP all | SP-8800 | Para administrar solución cardiopléjica |
| Sensor de flujo transónico | Transonic | ME3PXL-M5 | Sensor de flujo de circuito de perfusión |
| Módulo de flujo de tubo transónico Transonic | TS410 | sistema de adquisición de flujo | |
| Watson - Bombas Marlow | Aparato Harvard | 010.6131.DAO | Bomba peristáltica utilizada para la recirculación de perfusión |
| WBC-1510A | JEIO TECH | Baño calefactor | |
| Ratas Sprague-Dawley | Samtako Bio Korea Co., Ltd., Osan City Korea | ||
| Medicamentos | |||
| BioHAnce Gel Eye Drops | SENTRIX Cuidado | Ungüentos húmedos para los ojos | |
| Cefazolina | JW farmacéutica | Para la profilaxis | |
| Custodiol | DR, FRANZ KOHLER CHEMIE GMBH | Para la extracción del corazón | |
| Diclofenaco | Myungmoon Pharm. Co. Ltd | Para el control del dolor | |
| Heparina | JW farmacéutica | Anticoagulante | |
| Insulina | JW farmacéutica | hormonal | |
| Salina | JW pharmaceutical | Para la terapia de hidratación |
Referencias
- Langmuur, S. J. J., et al. Normothermic ex-situ heart perfusion with the organ care system for cardiac transplantation: A meta-analysis. Transplantation. 106 (9), 1745-1753 (2022).
- Ardehali, A., et al.
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Reimpresiones y permisos
Erratum
Formal Correction: Erratum: Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation
Posted by JoVE Editors on 8/28/2023. Citeable Link.
An erratum was issued for: Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation. The Protocol section was updated.
Section 4 of the Protocol was updated from:
4. Implantation
- Preparation of recipient
- Begin the recipient preparation 30 min before the cessation of ex situ perfusion.
- Anesthetize the recipient animal using the same method as mentioned in step 2.2.
- Place the rat in a supine position on the heating pad and insert the temperature probe into the rectum to maintain the body temperature at 37 °C.
- Apply eye lubricant, shave the pubic to the epigastric area, and cleanse the area with an iodine-based scrub and 70% alcohol.
- Medications
- Inject 2 mL of warm saline subcutaneously to compensate for the fluid lost during the surgery. Inject 200 IU of heparin subcutaneously.
- Administer antibiotic prophylaxis by injecting 10 mg/kg cefazolin dissolved in 0.3 mL of saline subcutaneously or intramuscularly.
- Administer pain control by injecting 20 mg/kg of diclofenac subcutaneously.
- Perform the mid-line laparotomy and insert a retractor to widen the abdominal cavity. Mobilize the abdominal organs to the left side of the recipient using cotton swabs to make space for the procedure.
- Prevent dehydration by wrapping the abdominal organs with warm and wet gauze. Intermittingly spread warm saline with a 50 mL syringe during the surgery.
- Utilizing a surgical microscope with a 10x magnification, mobilize the duodenum and proximal jejunum by blunt dissection with cotton swabs to expose the Abd. A. and IVC. Prepare the Abd. A and IVC for anastomosis and systematically implant the donor heart, in accordance with Figure 3 or previously documented methods15.
NOTE: Do not separate the Abd. A. and IVC.- Assuming vascular anastomosis to be placed infrarenal, prepare a sufficient portion of the aorta and IVC for clamping.
- Perform blunt preparation using cotton swabs or sharp-serrated forceps to remove the fats and fascia around the vessels.
- Place 5-0 silk ligatures to the mesenteric branches and both the cranial and caudal sides of the major vessels. Elevate the abdominal vessels and coagulate or ligate the lumbar branches with 5-0 silk sutures. Remember to spare the testicular arteries and veins and do not clamp them.
- Use ligatures to lift the vessels and position the micro-clamps to the mesenteric branches, caudal, and cranial sides of the major vessels to stop the blood flow at the anastomosis site. Be sure to switch off the heating pad before placing the clamps, as excess heating can exacerbate limb ischemia.
- Puncture the aorta using a 27 G needle and elongate the incision with micro scissors to a length equal to or slightly larger than the opening of the donor ascending aorta (Asc. A), which is approximately 5 mm.
- Make a longitudinal incision on the IVC in the same way as the aortotomy, but make it 3 mm closer to the caudal side compared to the aorta incision.
- Starting the anastomoses, placed the donor heart on the right side of the recipient's abdomen and attach the donor Asc. A to the recipient's Abd. A with one simple interrupted stitch (9-0 polypropylene) at the cranial corner of the longitudinal incision.
- Move the heart to the left side of the recipient abdomen and perform anastomosis of the donor's Asc. A with the recipient's Abd. A using a running 9-0 polypropylene suture.
- Fixate the donor pulmonary artery to the IVC with two interrupted sutures (9-0 polypropylene) at the caudal and cranial corners of the longitudinal incision.
- Perform the first half of the venous anastomosis from the intraluminal side of the vessel and complete the second half from the extraluminal side of the vessel. Before tightening the knots, flush the field with saline to prevent air embolism.
- De-airing and de-clamping
- Remove the mesenteric vein clamp first after completing the anastomosis to allow the right side of the heart to fill with venous blood.
- Remove the air in the coronary circuit and Asc. A. by applying retrograde coronary perfusion for several seconds.
- Place a piece of gauze on both sides of the vessels and remove the caudal clamp and the cranial clamp.
- Apply gentle compression with cotton swabs for 1-2 min. After ensuring adequate hemostasis, remove the swabs and wash the anastomoses with warm saline.
NOTE: The heart should begin beating within the first minute of reperfusion. If the recipient rat's body temperature is below 35 °C, the heart rhythm will normalize after the temperature reaches 36 °C.
- Replace the abdominal organs in a meander-like manner and close the layers of the abdominal incision using continuous 5-0 polypropylene sutures.
to:
4. Implantation
- Preparation of recipient
- Begin the recipient preparation 30 min before the cessation of ex situ perfusion.
- Anesthetize the recipient animal using the same method as mentioned in step 2.2.
- Place the rat in a supine position on the heating pad and insert the temperature probe into the rectum to maintain the body temperature at 37 °C.
- Apply eye lubricant, shave the pubic to the epigastric area, and cleanse the area with an iodine-based scrub and 70% alcohol.
- Medications
- Inject 2 mL of warm saline subcutaneously to compensate for the fluid lost during the surgery. Inject 200 IU of heparin subcutaneously.
- Administer antibiotic prophylaxis by injecting 10 mg/kg cefazolin dissolved in 0.3 mL of saline subcutaneously or intramuscularly.
- Administer pain control by injecting 20 mg/kg of diclofenac subcutaneously.
- Perform the mid-line laparotomy and insert a retractor to widen the abdominal cavity. Mobilize the abdominal organs to the left side of the recipient using cotton swabs to make space for the procedure.
- Prevent dehydration by wrapping the abdominal organs with warm and wet gauze. Intermittingly spread warm saline with a 50 mL syringe during the surgery.
- Utilizing a surgical microscope with a 10x magnification, mobilize the duodenum and proximal jejunum by blunt dissection with cotton swabs to expose the Abd. A. and IVC. Prepare the Abd. A and IVC for anastomosis and systematically implant the donor heart, in accordance with Figure 3 or previously documented methods15.
NOTE: Do not separate the Abd. A. and IVC.- Assuming vascular anastomosis to be placed infrarenal, prepare a sufficient portion of the aorta and IVC for clamping.
- Perform blunt preparation using cotton swabs or sharp-serrated forceps to remove the fats and fascia around the vessels.
- Place 5-0 silk ligatures to the mesenteric branches and both the cranial and caudal sides of the major vessels. Elevate the abdominal vessels and coagulate or ligate the lumbar branches with 5-0 silk sutures. Remember to spare the testicular arteries and veins and do not clamp them.
- Use ligatures to lift the vessels and position the micro-clamps to the mesenteric branches, caudal, and cranial sides of the major vessels to stop the blood flow at the anastomosis site. Switch off the heating pad before placing the clamps, as excess heating can exacerbate limb ischemia. Ensure to switch on the heating pad after de-clamping the vessels to avoid hypothermia.
- Puncture the aorta using a 27 G needle and elongate the incision with micro scissors to a length equal to or slightly larger than the opening of the donor ascending aorta (Asc. A), which is approximately 5 mm.
- Make a longitudinal incision on the IVC in the same way as the aortotomy, but make it 3 mm closer to the caudal side compared to the aorta incision.
- Starting the anastomoses, placed the donor heart on the right side of the recipient's abdomen and attach the donor Asc. A to the recipient's Abd. A with one simple interrupted stitch (9-0 polypropylene) at the cranial corner of the longitudinal incision.
- Move the heart to the left side of the recipient abdomen and perform anastomosis of the donor's Asc. A with the recipient's Abd. A using a running 9-0 polypropylene suture.
- Fixate the donor pulmonary artery to the IVC with two interrupted sutures (9-0 polypropylene) at the caudal and cranial corners of the longitudinal incision.
- Perform the first half of the venous anastomosis from the intraluminal side of the vessel and complete the second half from the extraluminal side of the vessel. Before tightening the knots, flush the field with saline to prevent air embolism.
- De-airing and de-clamping
- Remove the mesenteric vein clamp first after completing the anastomosis to allow the right side of the heart to fill with venous blood.
- Remove the air in the coronary circuit and Asc. A. by applying retrograde coronary perfusion for several seconds.
- Place a piece of gauze on both sides of the vessels and remove the caudal clamp and the cranial clamp.
- Apply gentle compression with cotton swabs for 1-2 min. After ensuring adequate hemostasis, remove the swabs and wash the anastomoses with warm saline.
NOTE: The heart should begin beating within the first minute of reperfusion. If the recipient rat's body temperature is below 35 °C, the heart rhythm will normalize after the temperature reaches 36 °C.
- Replace the abdominal organs in a meander-like manner and close the layers of the abdominal incision using continuous 5-0 polypropylene sutures.
- After the surgery, place the anesthetized animal on a clean area over a heating pad until the body temperature reaches 37°C.
NOTE: Do not initiate the postoperative examinations till the body temperature reaches 37°C. Maintain anesthesia at 2-2.5% isoflurane until the end of the experiments. - Monitor ECG of the transplanted donor heart for 3 h. Then, excise the heart under deep anesthesia for histological studies.
NOTE: Confirm anesthesia depth via lack of pedal reflex before excising the heart. The surgical procedure and the ECG monitoring take less than 6 h. Diclofenac, administered perioperatively (step 4.2.3.), enables pain management for the entire duration of this procedure. The analgesia regimen can be adjusted per the institutional animal use guidelines.
