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方法文章

常温非原位灌注异位心脏移植大鼠模型

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DOI：

10.3791/64954

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2023年4月21日

 ,  ,  ,  ,  ,  ,  , 
1Department of Medical Science, Chonnam National University Graduate School, 2Department of Thoracic and Cardiovascular Surgery, Chonnam National University Hospital and Medical School, 3Cardiovascular and Respiratory Research Team, Chonnam National University Hospital, 4Department of Internal Medicine, Chonnam National University Hospital and Medical School, 5Departments of Surgery, Chonnam National University Hospital and Medical School, 6Department of Pediatrics, Chonnam National University Children’s Hospital, and Medical School

* These authors contributed equally

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Important: There has been an erratum issued for this article. View Erratum Notice

摘要

此处，我们介绍一种在常温下异位移植心脏后的评估方案 离体 大鼠模型中的保存

摘要

心脏移植是治疗终末期心力衰竭最有效的疗法。尽管治疗手段和干预措施不断进步，等待移植的心力衰竭患者数量仍在持续增加。常温 离体 保存技术已被确立为与传统静态冷保存技术相媲美的方法。该技术的主要优势在于供体心脏可在生理条件下保存长达12小时。此外，该技术可在循环死亡后对供体心脏进行复苏，并实施必要的药理学干预，以改善移植后的供体心脏功能。已有多种动物模型被建立，以优化常温机械灌注 离体 保存技术并消除与保存相关的并发症。尽管与小型动物模型相比，大型动物模型更容易操作，但其成本较高且存在挑战性。我们介绍一种大鼠的常温 离体 供体心脏保存后进行异位腹部移植。该模型成本相对较低，可由单个实验人员完成。

引言

心脏移植仍是治疗难治性心力衰竭的唯一有效疗法1,2,3,4。尽管需要心脏移植的患者数量持续增加，但供体器官的可获得性并未相应提高5。为应对这一问题，研究人员已开发出多种新型供心保存方法，旨在改善相关挑战并提高供体器官的可获得性6,7,8,9。

使用器官维护系统（OCS）设备进行常温离体心脏灌注（NESHP）已作为一种临床干预手段出现1,3。该技术被认为是传统静态冷保存（SCS）方法的一种合适替代方案2,9。NESHP能有效缩短供体器官的冷缺血时间，降低代谢需求，并在器官运输过程中提供最佳的营养供给和氧合10,11。尽管该方法在改善供体器官保存方面具有明确潜力，但其临床应用和进一步研究仍受限于高昂的成本。因此，NESHP的临床前动物模型对于识别该技术相关的关键技术挑战至关重要12,13。猪和大鼠由于其对缺血的耐受性，是临床前研究首选的动物模型9。尽管猪模型适用于基础和转化研究，但其应用受限于高昂的成本以及护理和维持所需的大量人力。相比之下，大鼠模型成本较低且更易于操作14。

本研究介绍了一种简化的NESHP大鼠模型，随后进行异位心脏移植，以评估保存技术对移植后移植物状态的影响。该模型操作简便、成本低廉，且可由单名实验人员完成。图1展示了该操作流程的示意图。

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方案

本研究的所有动物实验均经全南国立大学医院实验动物研究中心伦理委员会批准（批准号：CNU IACUC - H - 2022-36）。实验所用雄性Sprague-Dawley大鼠（350-450 g）的饲养和护理均遵循《实验动物护理与使用指南》的规定。大鼠饲养于温度可控的环境中，光照周期为12小时明/12小时暗，自由摄取标准饲料和饮水。

1. 准备

注意：单个实验人员即可完成所有实验操作。

  1. 组装Langendorff装置，包括氧合器、泵和灌流管路，在手术前完成（图2用20 mL生理盐水填充灌注回路，并循环至回路被自体血液充分预充。
    注意：此步骤的目的是预热体外循环管路。
  2. 将心脏停跳液管路连接至循环回路 通过 连接至主动脉插管的三通阀，并准备注射泵以进行最终的心肌停搏液灌注。
    注意：确保从灌注回路和心脏停搏液管路中排除所有气泡。
  3. 将温度传感器置于将要存放供体心脏的储液器内，使循环系统的温度维持在 37 °C。
  4. 手术准备
    1. 为每只供体和受体大鼠分别准备一套独立的无菌显微器械和材料。
      1. 准备供体手术器械包：手术剪一把、显微镊一把、锐利的蚊式钳一把、5-0 丝线缝合线、棉签、50 mL 注射器、停跳液（CPS）灌注管路、注射泵、18 G 留置针、一套 5 Fr 股动脉导管以及无菌纱布。
      2. 准备受体手术器械包：显微外科剪、伤口牵开器、显微镊、蚊式钳、血管显微夹、1 mL注射器、5-0和9-0聚丙烯缝线、5-0丝线、棉签及无菌纱布。

2. 供体心脏保存与血液采集

  1. 在麻醉舱内使用异氟烷（5%）对供体大鼠进行麻醉诱导，并在将其置于手术台前记录大鼠体重。
  2. 将大鼠仰卧位固定于手术台上，通过鼻罩持续给予2%-2.5%异氟烷与90%氧气混合气体以维持麻醉。
  3. 通过检查足趾捏压无反应以及呼吸频率（应为每分钟50-60次）来确认麻醉深度。
    注意：适当的麻醉水平对于避免供体大鼠遭受不必要的应激和疼痛至关重要。
  4. 涂抹眼用润滑剂，剃除从耻骨至锁骨的手术区域毛发。使用含碘消毒液和70%酒精清洁手术部位。
  5. 导管插入术
    1. 沿腹部正中线做一条7 cm的切口，并从剑突至锁骨中点向两侧各做一条3 cm的切口。去除胸部区域的皮肤。
    2. 使用棉签将腹腔器官移至腹腔左侧。从腹膜后筋膜和脂肪组织中分离腹主动脉。
    3. 通过下腔静脉（IVC）使用1 mL注射器注入1,000 IU肝素溶于0.3 mL等渗盐水的溶液。用棉签轻轻按压针孔部位以止血。
      注意：注射时需警惕空气栓塞，因其可导致心脏骤停。
    4. 将5 Fr. 股动脉导管插入腹主动脉（Abd. A），确保导管尖端到达主动脉弓。通过评估导管插入部分的近似长度来确认导管位置。
  6. 血液采集
    1. 采集约 10 mL 血液 通过 插入腹主动脉的导管
    2. 随后，用等渗盐水稀释引血，直至总体积达到12 mL。加入溶于0.3 mL生理盐水中的头孢唑林5 mg和胰岛素（20 IU）。
  7. 心脏骤停
    1. 将先前准备好的CPS灌注管路连接至腹腔导管，并使用注射泵以800 mL/h的速度开始CPS给药。
    2. 从膈肌处打开胸腔，并在靠近膈肌处剪断下腔静脉，以防止心室扩张。沿胸椎两侧剪断肋骨，直至胸廓入口。用蚊式钳将游离的腹侧胸壁向上翻起。
    3. 使用显微镊完全移除胸腺以暴露主动脉弓。若胸腺动脉出血，可施加轻柔压迫止血。
  8. 提取
    1. 注射完所有CPS后，从周围组织中分离主动脉弓。在左侧锁骨下动脉稍下方仔细解剖。
    2. 在远离分叉处切断头臂干和左颈总动脉，保留较长的主动脉弓残端，以便在主动脉插管时易于操作。尽可能靠近分叉处切断主肺动脉（MPA），注意避免损伤左心耳。
    3. 仔细用5-0丝线结扎上腔静脉（SVC）和下腔静脉（IVC），避免阻塞右心房（RA）和冠状窦。用湿纱布覆盖胸腔左侧边缘，将心脏置于其上，并轻柔牵拉SVC和IVC的结扎线，以暴露肺门。
    4. 用5-0丝线将肺静脉和奇静脉一起结扎。切断结扎线背侧的组织，取出心脏。检查心脏是否有损伤。最后，在主动脉插管前称量心脏重量。

3. 离体 灌注

  1. 主动脉插管与灌注
    1. 在主动脉插管前，将生理盐水预充的管路更换为血液预充。
    2. 将主动脉插管插入主动脉弓，并用临时微型夹子固定。确保插管尖端位于头臂干分叉处。
    3. 用显微镊轻轻夹持主动脉，确认插管位置正确。
    4. 以2–3 mL/min的流速开始灌注，使灌注液从插管部位少量渗出，以排除任何气泡。
    5. 通过连接至监测系统的传感器监测灌注压力和温度。
    6. 用拇指和食指轻轻按摩心脏，直至静脉血从主肺动脉（MPA）渗出。
    7. 用1-0丝线结扎主动脉，在确认所有设置（灌注管路、灌注压力、温度）无误后移除夹子。
    8. 一旦完成永久性结扎，应确保心脏在数秒内开始收缩，并在60秒内恢复至正常节律。在37 °C条件下，平均灌注压为55–65 mmHg，冠状动脉流量为3–4 mL，提示灌注充分。
    9. 从储液器中采集0.15 mL血液，在灌注开始时及之后每20分钟进行一次血气分析（BGA）。灌注期间监测并记录pH、pCO2、pO2、葡萄糖、血细胞比容、钾和乳酸水平。灌注120分钟后，通过注射泵以250 mL/h的速度注入3 mL Custodiol溶液，使心脏停搏。

4. 植入

  1. 受体准备
    1. 在供体手术结束前30分钟开始受体准备工作 离体 灌注
    2. 使用与步骤2.2中所述相同的方法对受体动物进行麻醉。
    3. 将大鼠仰卧置于加热垫上，并将体温探针插入直肠，以维持体温在 37 °C。
    4. 涂抹眼用润滑剂，剃除从会阴至剑突区域的毛发，并使用含碘消毒液和70%酒精清洁该区域。
  2. 药物
    1. 皮下注射2 mL温热生理盐水，以补偿手术期间丢失的体液。皮下注射200 IU肝素。
    2. 皮下或肌内注射0.3 mL生理盐水溶解的头孢唑林（10 mg/kg）进行抗生素预防。
    3. 皮下注射20 mg/kg的双氯芬酸以进行疼痛控制。
  3. 行正中腹切口，插入拉钩以扩大腹腔。使用棉签将受体腹腔器官向左侧移动，为手术操作创造空间。
  4. 用温热的湿纱布包裹腹腔器官以防止脱水。术中使用50 mL注射器间歇性地喷洒温盐水。
  5. 利用10倍放大倍率的手术显微镜，使用棉签钝性分离十二指肠及近端空肠，暴露腹主动脉（Abd. A.）和下腔静脉（IVC）。为腹主动脉和下腔静脉做好吻合准备，并按照规范系统地植入供体心脏 图3 或已有文献记载的方法15.
    注意：不要分离腹主动脉（Abd. A.）和下腔静脉（IVC）。
    1. 假设血管吻合口位于肾下，需准备足够长度的主动脉和下腔静脉用于夹闭。
    2. 使用棉签或锐齿镊进行钝性分离，以清除血管周围的脂肪和筋膜。
    3. 用5-0丝线结扎肠系膜分支以及主要血管的头侧和尾侧。抬起腹腔血管，用5-0丝线电凝或结扎腰部分支。注意保留睾丸动脉和静脉，避免夹闭。
    4. 使用结扎线提起血管，并将微血管夹放置在主要血管吻合部位的肠系膜分支、尾侧和头侧，以阻断吻合处的血流。放置血管夹前应关闭加热垫，因为过度加热可能加重肢体缺血。在移除血管夹后应及时打开加热垫，以避免低体温。
    5. 使用27 G针头穿刺主动脉，并用显微剪刀将切口延长至等于或略大于供体升主动脉（Asc. A）开口的长度，约5 mm。
    6. 以与主动脉切开相同的方式在下腔静脉上做一纵行切口，但位置较主动脉切口向尾侧偏移3 mm。
    7. 开始吻合时，将供体心脏置于受体腹部右侧，用一根间断单针（9-0聚丙烯缝线）在纵行切口的头侧角处将供体升主动脉与受体腹主动脉相连。
    8. 将心脏移至受体腹腔左侧，使用连续的9-0聚丙烯缝线将供体的升主动脉（Asc. A）与受体的腹主动脉（Abd. A）进行吻合。
    9. 用两针间断缝线（9-0聚丙烯）将供体肺动脉固定于下腔静脉，缝合位置为纵行切口的尾侧和头侧角。
    10. 从血管腔内侧开始进行静脉吻合的前半部分，然后从血管外侧完成后半部分。在收紧结之前，用生理盐水冲洗术区，以防止空气栓塞。
  6. 脱气与去夹
    1. 完成吻合后，先移除肠系膜静脉钳，以使静脉血流入右心。
    2. 通过逆行冠状动脉灌注数秒，排出冠状动脉循环和升主动脉中的空气。
    3. 在血管两侧各放置一块纱布，然后移除尾侧夹钳和头侧夹钳。
    4. 用棉签轻轻按压1-2分钟。确认止血充分后，移除棉签，并用温盐水冲洗吻合口。
      注意：再灌注后1分钟内心脏应开始搏动。若受体大鼠体温低于35 °C，待体温升至36 °C时，心律将恢复正常。
  7. 以迂回方式将腹腔器官复位，并使用连续的5-0聚丙烯缝线逐层关闭腹部切口。
  8. 手术后，将麻醉的动物置于加热垫上的清洁区域，直至体温达到37°C。 
    注意：体温未达到37°C前，不得开始术后检查。实验结束前，维持2-2.5%异氟烷麻醉。
  9. 监测移植供体心脏的心电图3小时。随后在深度麻醉下切除心脏，用于组织学研究。
    注意：在切除心脏前，需通过确认足底反射消失来判断麻醉深度。手术操作和心电图监测全过程耗时少于6小时。围手术期给予双氯芬酸（步骤4.2.3）可实现整个操作期间的有效镇痛。镇痛方案可根据各机构的动物使用指南进行调整。

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结果

图1 展示了在小型动物模型中使用的实验设计。图2 显示了改良的朗根多夫灌注装置，该装置包含一个小型动物氧合器。图3 展示了异位腹部移植的吻合顺序。

图4 显示用于评估心脏存活性的参数 离体 灌注，例如乳酸、钾和平均主动脉压。在本研究中，采用常温 原位以外 保存使六例成功病例的总缺血时间缩短至46.2 ± 4.7分钟，而总离体时间为166.2 ± 4.7分钟（图5）。供体心脏的摘取及移植前准备 原位以外 灌注和异位移植耗时5.8 ± 1.3分钟，如图所示 图5手术的总体成功率为70%，六例成功病例的平均吻合时间为38.4 ± 3.4分钟。在所有实验中，植入后心率均显著下降，但随...

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讨论

我们建立该模型的重点是复制常温下的人类心脏移植。非射血模型是离体环境中保存供体心脏的常用技术16。尽管射血模型在评估离体灌注期间的心脏功能方面具有诸多优势17，但并不适用于异位移植模型。在异位移植中，植入的供体心脏需要克服受体循环系统中宿主心脏产生的收缩期后负荷压力，从而导致供体心脏功能受限，并在评估中产生低估18。因此，在异位移植中，非射血模型更具优势。在非射血模型中，供体心脏虽被灌注，但不支持受体的循环，从而显著限制了对心脏功能的评估。当功能评估受限时，组织形态学和分子学评价（如组织学染色和印迹分析）可用于检查供体心脏的状态。此外，可利用正电子发射断层扫描（PET）或磁共振成像（MRI）等先进技术评估代谢标志物19。该模型可用于在移植前测试药物和基因干预措施的长期有效性。

许多研究团队已开发出一种常温ex situ（离体）保存模型，并已...

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致谢

本工作由全南国立大学医院生物医学研究所资助项目B2021-0991以及韩国国家研究基金会项目NRF-2020R1F1A1073921资助

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材料

本文使用的材料清单
姓名公司目录编号评论
主动排风系统（AES）Smiths medicalPC-6769-51A用于清除二氧化碳和过量异氟烷
麻醉机Smiths medicalPC-8801-01A混合异氟烷与氧气并输送至动物
B20 患者监护仪GE medical systemsB20用于监测平均主动脉压和体温
恒温监测系统Harvard apparatus55-7020用于监测并维持动物体温
Micro-1 大鼠氧合器东莞科威医疗器械有限公司Micro-MO用于Langendorff循环中的气体交换
微穿刺导入器套装COOK medicalG48007通过腹主动脉向主动脉弓输送停跳液
显微镜AmscopeMU1403用于放大手术视野（受体）
手术放大镜SurgiTelL2S09用于放大手术视野（供体）
注射泵AMP allSP-8800用于输送停跳液
Transonic 流量传感器TransonicME3PXL-M5灌注回路流量传感器
Transonic 管路流量模块TransonicTS410流量采集系统
Watson - Marlow 泵Harvard apparatus010.6131.DAO用于灌注液循环的蠕动泵
WBC-1510AJEIO TECHE03056D加热浴槽
SD大鼠（Sprague-Dawley rats）韩国三塔科生物有限公司，韩国乌山市
药物
BioHAnce 凝胶滴眼液SENTRIX Animal care眼部湿润软膏
头孢唑林JW pharmaceutical用于预防性治疗
Custodiol 器官保存液DR, FRANZ KOHLER CHEMIE GMBH用于心脏获取
双氯芬酸明文制药有限公司用于疼痛控制
肝素JW pharmaceutical抗凝剂
胰岛素JW pharmaceutical激素治疗
生理盐水JW pharmaceutical用于补液治疗

参考文献

  1. 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).
  2. Ardehali, A., et al. Ex-vivo perfusion of donor hearts for human heart transplantation (PROCEED II): a prospective, open-label, multicentre, randomized non-inferiority trial. Lancet. 385 (9987), 2577-2584 (2015).
  3. Dang Van, S., et al. Ex vivo perfusion of the donor heart: Preliminary experience in high-risk transplantations. Archives of Cardiovascular Diseases. 114 (11), 715-726 (2021).
  4. Zhou, P., et al. Donor heart preservation with hypoxic-conditioned medium-derived from bone marrow mesenchymal stem cells improves cardiac function in a heart transplantation model. Stem Cell Research and Therapy. 12 (1), 5f6(2021).
  5. Messer, S., Large, S. Resuscitating heart transplantation: the donation after circulatory determined death donor.European. Journal of Cardio-Thoracic Surgery. 49 (1), 1-4 (2016).
  6. Trahanas, J. M., et al. Achieving 12 hour normothermic ex situ heart perfusion: an experience of 40 porcine hearts. ASAIO Journal. 62 (4), 470-476 (2016).
  7. Yang, Y., et al. Keeping donor hearts in completely beating status with normothermicblood perfusion for transplants. The Annals of Thoracic Surgery. 95 (6), 2028-2034 (2013).
  8. Van Caenegem, O., et al. Hypothermic continuous machine perfusion enables preservation of energy charge and functional recovery of heart grafts in an ex vivo model of donation following circulatory death. European Journal of Cardiothoracic Surgery. 49 (5), 1348-1353 (2016).
  9. Lu, J., et al. Normothermic ex vivo heart perfusion combined with melatonin enhances myocardial protection in rat donation after circulatory death hearts via inhibiting NLRP3 inflammasome-mediated pyroptosis. Frontiers in Cell and Developmental Biology. 9, 733183(2021).
  10. Pinnelas, R., Kobashigawa, J. A. Ex vivo normothermic perfusion in heart transplantation: a review of the TransMedics Organ Care System. Future Cardiology. 18 (1), 5-15 (2022).
  11. Fuchs, M., et al. Does the heart transplant have a future. European Journal of Cardiothoracic Surgery. 55, i38-i48 (2019).
  12. Pahuja, M., Case, B. C., Molina, E. J., Waksman, R. Overview of the FDA's circulatory system devices panel virtual meeting on the TransMedics Organ Care System (OCS) Heart - portable extracorporeal heart perfusion and monitoring system. American Heart Journal. 247, 90-99 (2022).
  13. Jawitz, O. K., Devore, A. D., Patel, C. B., Bryner, B. S., Schroder, J. N. Expanding the donor pool: quantifying the potential impact of a portable organ-care system for expanded criteria heart donation. Journal of Cardiac Failure. 27 (12), 1462-1465 (2021).
  14. van Suylen, V., et al. Ex situ perfusion of hearts donated after euthanasia: a promising contribution to heart transplantation. Transplantation Direct. 7 (3), e676(2021).
  15. Westhofen, S., et al. The heterotopic heart transplantation in mice as a small animal model to study mechanical unloading - Establishment of the procedure, perioperative management and postoperative scoring. PLoS One. 14 (4), e0214513(2019).
  16. Qin, G., Jernryd, T., Sjoberg, S., Steen, S., Nilsson, J. Machine perfusion for human heart preservation: A systematic review. Transplant International. 35, 10258(2022).
  17. Dang Van, S., Brunet, D., Akamkam, A., Decante, B., Guihaire, J. Functional assessment of the donor heart during ex situ perfusion: insights from pressure-volume loops and surface echocardiography. Journal of Visual Experiments. (188), e63945(2022).
  18. Fu, X., Segiser, A., Carrel, T. P., Tevaearai Stahel, H. T., Most, H. Rat heterotopic heart transplantation model to investigate unloading-induced myocardial remodeling. Frontiers in Cardiovascular Medicine. 3, 34(2016).
  19. Niimi, M. The technique for heterotopic cardiac transplantation in mice: experience of 3000 operations by one surgeon. The Journal of Heart and Lung Transplantation. 20 (10), 1123-1128 (2001).
  20. Qi, X., et al. The evaluation of constant coronary artery flow versus constant coronary perfusion pressure during normothermic ex-situ heart perfusion. The Journal of Heart and Lung Transplantation. 41 (12), 1738-1750 (2022).
  21. Okahara, S., et al. A novel blood viscosity estimation method based on pressure-flow characteristics of an oxygenator during cardiopulmonary bypass. Artificial Organs. 41 (3), 262-266 (2017).
  22. Quader, M., Torrado, J. F., Mangino, M. J., Toldo, S. Temperature and flow rate limit the optimal ex-vivo perfusion of the heart - an experimental study. Journal of Cardiothoracic Surgery. 15 (1), 180(2020).

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重印与许可

勘误


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

  1. Preparation of recipient
    1. Begin the recipient preparation 30 min before the cessation of ex situ perfusion.
    2. Anesthetize the recipient animal using the same method as mentioned in step 2.2.
    3. 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.
    4. Apply eye lubricant, shave the pubic to the epigastric area, and cleanse the area with an iodine-based scrub and 70% alcohol.
  2. Medications
    1. Inject 2 mL of warm saline subcutaneously to compensate for the fluid lost during the surgery. Inject 200 IU of heparin subcutaneously.
    2. Administer antibiotic prophylaxis by injecting 10 mg/kg cefazolin dissolved in 0.3 mL of saline subcutaneously or intramuscularly.
    3. Administer pain control by injecting 20 mg/kg of diclofenac subcutaneously.
  3. 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.
  4. Prevent dehydration by wrapping the abdominal organs with warm and wet gauze. Intermittingly spread warm saline with a 50 mL syringe during the surgery.
  5. 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.
    1. Assuming vascular anastomosis to be placed infrarenal, prepare a sufficient portion of the aorta and IVC for clamping.
    2. Perform blunt preparation using cotton swabs or sharp-serrated forceps to remove the fats and fascia around the vessels.
    3. 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.
    4. 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.
    5. 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.
    6. 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.
    7. 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.
    8. 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.
    9. 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.
    10. 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.
  6. De-airing and de-clamping
    1. Remove the mesenteric vein clamp first after completing the anastomosis to allow the right side of the heart to fill with venous blood.
    2. Remove the air in the coronary circuit and Asc. A. by applying retrograde coronary perfusion for several seconds.
    3. Place a piece of gauze on both sides of the vessels and remove the caudal clamp and the cranial clamp.
    4. 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.
  7. 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

  1. Preparation of recipient
    1. Begin the recipient preparation 30 min before the cessation of ex situ perfusion.
    2. Anesthetize the recipient animal using the same method as mentioned in step 2.2.
    3. 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.
    4. Apply eye lubricant, shave the pubic to the epigastric area, and cleanse the area with an iodine-based scrub and 70% alcohol.
  2. Medications
    1. Inject 2 mL of warm saline subcutaneously to compensate for the fluid lost during the surgery. Inject 200 IU of heparin subcutaneously.
    2. Administer antibiotic prophylaxis by injecting 10 mg/kg cefazolin dissolved in 0.3 mL of saline subcutaneously or intramuscularly.
    3. Administer pain control by injecting 20 mg/kg of diclofenac subcutaneously.
  3. 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.
  4. Prevent dehydration by wrapping the abdominal organs with warm and wet gauze. Intermittingly spread warm saline with a 50 mL syringe during the surgery.
  5. 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.
    1. Assuming vascular anastomosis to be placed infrarenal, prepare a sufficient portion of the aorta and IVC for clamping.
    2. Perform blunt preparation using cotton swabs or sharp-serrated forceps to remove the fats and fascia around the vessels.
    3. 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.
    4. 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.
    5. 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.
    6. 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.
    7. 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.
    8. 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.
    9. 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.
    10. 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.
  6. De-airing and de-clamping
    1. Remove the mesenteric vein clamp first after completing the anastomosis to allow the right side of the heart to fill with venous blood.
    2. Remove the air in the coronary circuit and Asc. A. by applying retrograde coronary perfusion for several seconds.
    3. Place a piece of gauze on both sides of the vessels and remove the caudal clamp and the cranial clamp.
    4. 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.
  7. Replace the abdominal organs in a meander-like manner and close the layers of the abdominal incision using continuous 5-0 polypropylene sutures.
  8. 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.
  9. 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.

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常温离体灌注大鼠心脏移植供体心脏保存离体心脏灌注Langendorff 装置显微外科技术灌注回路心脏停搏液免疫反应
常温非原位灌注异位心脏移植大鼠模型
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