介绍了一种用于自由活动小鼠的市售遥测发射器植入手术技术，该发射器可连续测量生物电位（单导联心电图）、心率、核心体温和运动活动。同时提供了术后护理和镇痛的建议与操作规程，以促进动物恢复、改善健康状况并提高存活率。
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摘要
摘要
在学术界和制药行业中，实验室小鼠是大多数生物医学研究的首选动物物种。小鼠体型适中，相对容易饲养。这些特点，加上存在大量自发突变和实验诱导突变品系，使实验室小鼠非常适合用于多种研究领域。
在心血管、药理学和毒理学研究中，通常需要准确测量实验动物循环系统相关的参数。心率、心率变异性以及PQ间期和QT间期持续时间的测定均基于心电图（ECG）记录。然而，使用传统测量技术在小鼠中获取可靠的心电图曲线以及核心体温等生理数据可能较为困难，因为这些方法需要将传感器和导线连接到被固定、束缚甚至麻醉的动物身上。对此类方法获得的数据必须谨慎解读，众所周知，固定和麻醉会对生理参数产生显著的人为干扰影响1, 2。
遥测技术能够从清醒且无束缚的动物中采集数据。即使在自由活动的动物中也可进行测量，且无需研究人员靠近动物。因此，可避免已知的人为干扰因素，确保测量结果的准确性和可靠性。该方法还可减少个体间差异，从而降低所需使用的动物数量，使该技术成为监测实验动物生理参数最人道的方法3, 4。随着数据采集技术和植入设备微型化的持续进步，如今已可连续、实时地长时间记录生理参数和运动活动，持续时间可达数小时、数天甚至数周3, 5。
本文介绍了一种用于植入商业化遥测发射器的外科手术技术，该发射器可用于自由活动、无束缚小鼠的核心体温、运动活动以及生物电势（即单导联心电图）的连续测量，由此可确定心率、心率变异性以及PQ间期和QT间期。我们还提供了术前准备程序以及术后重症监护和镇痛治疗方案，以提高植入小鼠的恢复质量、福利状况和存活率5, 6。
方案
动物实验已获得瑞士苏黎世州兽医办公室的批准。饲养和实验程序均符合瑞士动物保护法，并遵循欧洲议会和理事会关于用于科学目的动物保护的指令（2010年9月22日第2010/63/EU号指令）。
1. 术前注意事项
1.1 小鼠：饲养条件、一般状况及健康监测
建议从供应商处获得或从外部啮齿类动物种群转移的小鼠，应在手术前至少两周到达饲养设施。此期间可使动物适应新环境及设施特定的饲养条件。由于小鼠是群居动物，在适应期间应以相容的群体进行合笼饲养。为监测个体的食物和饮水摄入量，每只小鼠需从术前3天起至手术植入遥测发射器后10天期间单独饲养。建立携带遥测发射器小鼠的时间线如图1所示。动物必须以良好的健康状况接受手术，因此在术前应每天观察2-3天，评估其总体状况（外观、姿势、自发行为）以及体重、食物和饮水摄入情况。这些数据应记录在医疗档案中（总体状况与健康监测数据表，表1），以建立个体在总体状况、整体健康和福利方面的基线水平。任何在手术前出现疾病症状或总体状况不良的动物均应排除在实验之外。
在最后一次健康检查后或手术前，为了剃除动物手术部位的毛发，将小鼠置于一个小型（8×8×8 cm）有机玻璃舱内，使用8%的七氟烷或5%的异氟烷在纯氧（600 mL/min）中进行短暂麻醉。在手术前一天剃毛可避免手术区域出现毛茬。当小鼠失去翻正反射后，将其从舱中取出，并让其仰卧，用剃毛器去除颈部前侧和腹部的毛发；通过面罩以600 mL/min的流速给予3–4%七氟烷或1.5–3%异氟烷维持纯氧麻醉，持续约5分钟。剃毛完成后，让动物苏醒，并放回其原有饲养笼中。
2. 植入
2.1 操作环境、遥测发射器的准备
在植入当天，所有关于发射器准备和手术的操作均在配备手术显微镜的层流罩工作台中进行。通过使用经高压灭菌的器械和灭菌材料，并对工作台面进行消毒，以确保无菌条件7。在植入前，首先准备遥测发射器（ETA-F10，Data Sciences International，St. Paul，MN，USA）。从无菌包装中取出后，根据待植入小鼠的体型，将发射器的导线剪短至适当长度。在大多数成年远交系或近交系小鼠中，红色电极可剪短至约42 mm，白色/无色电极则剪短至约55 mm。去除导线远端（感知端）的绝缘套管：从红色电极上去除约20 mm套管，从白色/无色电极上去除约10 mm套管。将每个电极的远端（此时已无套管）用细丝线（PERMA-Handseide，6-0，Ethicon，Norderstedt，德国）固定成环状。电极准备完成后，将发射器置于温热的无菌生理盐水中，待动物麻醉并完成外科准备后立即植入。
2.2 麻醉
在吸入麻醉诱导前5-10分钟，皮下注射咪达唑仑（4 mg/kg）和芬太尼（0.04 mg/kg）混合液作为预麻醉用药，以提供镇静作用及预先镇痛。通过将动物置于诱导舱内，并通入挥发性麻醉剂（8%七氟烷或5%异氟烷，纯氧流速600 ml/min）来诱导全身吸入麻醉。当动物出现翻正反射消失时，将其转移至超净台下的操作台面上，仰卧位放置于特制的金属板上，该金属板配备有鼻罩及连接麻醉设备的管路。麻醉维持通过自主呼吸进行（3-4%七氟烷或1.5-3%异氟烷，纯氧流速600 mL/min）。麻醉期间，使用眼膏（维生素A，Baush & Lomb，Steinhausen，瑞士）保护动物的眼睛。动物仰卧于金属板上时，由操作台面内置水浴加热系统维持体温（39°C ±1）。
2.3 手术
使用浸有70%乙醇、氯己定或碘溶液的棉签对前颈部和腹部区域的皮肤进行5分钟消毒。沿中线从胸部下部至腹部做一条长1至1.5厘米的皮肤切口。将阴性（白色/无色）电极导线经皮下隧道从胸部引至颈部，并在该处纵向做一小切口（≤0.5厘米）。分离皮肤及皮下组织，为电极金属环的固定预留空间。将金属环使用两根细丝线（PERMA-Handseide, 6-0, Ethicon, Norderstedt, Germany）固定于气管右侧的肌肉之间。随后分层使用可吸收缝线（VICRYL 6-0, Ethicon, Norderstedt, Germany）缝合颈部切口。在腹白线处切开腹壁，将遥测发射器本体置入小鼠腹腔内。将阳性（红色）电极的金属环用丝线缝合固定于剑突处，使其位于左上腹部区域，并置于肝脏与膈肌之间（图2）。随后使用可吸收缝线（VICRYL 6-0, Ethicon, Norderstedt, Germany）逐层缝合腹壁肌肉层。在最终关闭腹壁前，向腹腔内注射磺胺多辛与甲氧苄啶混合液［分别为30 mg/kg和6 mg/kg，溶于1 mL生理盐水（0.9%），并加热至接近体温（38–39°C）］，以达到抗感染预防和维持体液稳态的目的。最后，使用皮肤钉合器（Precise, 3 M Health Care, St. Paul, MN, USA）或皮内连续可吸收缝线（VICRYL 6-0, Ethicon, Norderstedt, Germany）闭合腹部皮肤。
3. 术后护理
手术和麻醉结束后，皮下注射0.1 mg/kg的丁丙诺啡（Temgesic, Essex Chemie AG, Lucerne, Switzerland）和5 mg/kg的美洛昔康（Metacam, Boehringer Ingelheim, Basel, Switzerland）用于镇痛，动物置于工作台温暖（39°C ±1）的表面上恢复约2小时。除镇痛处理（每日两次：丁丙诺啡0.1 mg/kg，美洛昔康5 mg/kg）外，支持性治疗包括每日两次皮下注射300 μL葡萄糖溶液（5%）和300 μL生理盐水（0.9%），均预热至体温。为促进进一步恢复，建议为动物额外提供一瓶含15%葡萄糖溶液的饮水瓶。在4至10天的恢复期间，建议保持动物体温适宜。因此，在本实验中，小鼠被饲养于恒温箱（30°C ±1）内。术后10天内每日根据一般状况与健康监测记录表（表1）监测动物的一般状况、体重变化以及食物和水的消耗情况。人道终点，即在恢复进展不理想时为避免动物遭受不必要的痛苦和疼痛而实施安乐死，适用于以下情况：
- 若动物总体状况较差，即受到触碰或推动后仍明显表现出冷漠（无活动），且尽管已采取保温措施，其体表仍感觉冰冷，则应立即实施安乐死。
- 在植入发射器后的第4天，若动物表现出明显的冷漠症状、极度攻击性或完全不进食，应立即实施安乐死。
- 在植入发射器后的第8天，动物体重必须较术后前期有明显增加，并且其食物摄入量必须达到术前每日摄入量的至少80%。若上述任一条件未满足，应立即实施安乐死。
植入后第10天，将动物转移回标准饲养条件的动物房。如果使用了皮肤钉合钉，应在手术后7-10天将其拆除；可吸收缝线无需拆除。小鼠应以相容的群体进行饲养，以允许社会互动，并防止长期单独饲养可能对后续实验结果产生的不利影响8, 9。在进行首次实验并开始数据采集之前，小鼠应在发射器植入术后至少经过4周的恢复期。
4. 数据采集
通过使用磁铁触碰动物来启动数据采集，此时发射器将被开启。Dataquest A.R.T. 软件（Data Sciences International，美国明尼苏达州圣保罗）负责协调来自一个或多个动物信号的检测、采集、分析和图形化显示（以波形形式呈现）。采集程序通过数据交换矩阵（Data Sciences International）从转换器和接收器收集发送至计算机的数据信号。该程序可按设定时间间隔采集特定时长的数据，也可进行连续采样并将数据保存在计算机硬盘中。由于发射信号的范围和质量在很大程度上取决于笼具及周围设备的材料组成（例如金属与塑料），建议将接收板尽可能靠近动物放置，例如置于动物笼具下方或实验区域上方（如实验台或跑步机上方）。建议通过在连续采样模式下进行短时间的实时测量检查，以确认记录和数据传输系统的正确配置。数据采集并存储后，可使用分析程序对多种不同参数进行绘图、列表和分析。记录系统的配置细节（例如定义采样模式）以及分析软件的使用方法（例如基于生物电位/心电图曲线计算心率变异性参数、PQ间期和QT间期）可在制造商手册中找到。有关适用于遥测数据采集与解释的生物测量实验设计和统计方法的宝贵建议，已在其他文献中发表3。
5. 代表性结果：
图1展示了所述实验流程的整体示意图。图2显示了植入式发射器的位置，包括用于从心脏获取生物电位信号（单导联心电图）的电极位置。图3和图4分别给出了单只小鼠短期生物电位曲线（单导联心电图）以及长期心率、核心体温和运动活动记录的原始数据示例。图5展示了实验后对小鼠群体进行长期测量所得数据的已发表结果示例。此外，还可从生物电位曲线中提取多个其他参数。心率变异性参数5、QT间期和PQ间期10, 11的分析示例已在其他文献中发表。
表1. 小鼠一般状况与健康监测数据表。点击此处下载该表格。此模板用于监测单只小鼠的一般状况与健康情况。在植入手术前，需连续3天、每天1次对动物的外观、姿势和自发行为进行基础检查，并记录体重、食物和水的消耗量。术后连续10天每日监测上述指标，并与术前基线值进行比较，以评估术后恢复进程。此外，术后护理和镇痛治疗措施也以医疗记录形式详细记录。表格中提供了人道终点的操作指导，以便在动物未能达到快速恢复标准时，及时决定是否实施安乐死，避免不必要的疼痛与痛苦。
图1. 建立携带遥测发射器小鼠的时间安排。与发射器植入相关的各项操作的时间顺序，显示小鼠可用于实验和数据采集的时间点。
图 2. X射线影像/示意图显示植入式遥测发射器的位置。发射器本体置于腹腔内。正极导线弯成环状，并用缝线固定于剑突处。负极导线从胸部皮下穿至颈部，并在气管旁肌肉之间弯成环状固定。该X射线影像引自作者先前发表于《实验动物》（Laboratory Animals）9 的研究。
图3. 生物电位曲线。来自清醒小鼠及同一小鼠在七氟烷吸入麻醉状态下的单导联ECG原始打印曲线。心率由遥测系统自动计算。麻醉状态下记录的3秒序列显示心率为440次/分钟（bpm）。清醒小鼠记录的曲线显示心率为660 bpm，处于中等体力活动（如理毛或进食）期间预期的心率范围内。通过生物电位/单导联ECG曲线，可使用制造商提供的软件确定心率变异性参数、心动周期间期以及PQ间期和QT间期。
图 4. 健康与患病小鼠长期测量的原始数据。在小鼠单独饲养于家中笼具且不受人为或实验操作干扰的条件下，持续监测心率（bpm）、核心体温（°C）和运动活动（计数）。心率每5分钟记录30秒（采样频率1000 Hz）；核心体温每5分钟采样10秒；运动活动连续记录，并以5分钟为间隔存储数据。数据连续追踪6.5天，共采集三个不同生理状态小鼠的遥测数据。健康小鼠在黑暗（夜间）阶段表现出清晰的昼夜节律，生理指标值及运动活动行为均呈现正常升高。相比之下，接受重大手术后的小鼠心率升高，尤其在光照阶段更为明显，且运动活动受到抑制。第三只小鼠患有慢性肿瘤疾病，其心率与核心体温的昼夜节律趋于平缓，运动活动显著减少。代表性的心率测量数据（正常值及重大手术后）取自作者此前在Altex12发表的文献。
图 5. 实验后长期遥测测量结果展示示例。 该图取自作者先前发表于《实验动物》（Laboratory Animals）的论文1。以一项示例实验为例，对动物实施了持续 50 分钟的异氟烷或七氟烷麻醉。比较了麻醉剂在动物苏醒后对心率、核心体温和运动活动的长期影响。使用 16 只植入遥测发射器的小鼠，在每种麻醉条件下各记录 8 只小鼠的数据；实验期间小鼠单笼饲养，可在笼内自由活动。在分析麻醉后的长期效应时，考虑到小鼠主要在夜间活动，其各项指标在 24 小时周期内存在显著波动，因此分别计算每只动物在夜间（12 小时黑暗期）和白天（12 小时光照期）的遥测数据均值。通过计算麻醉前连续三天的均值，确定每只动物的正常基准值。对于麻醉后的每一天，将黑暗期和光照期的均值分别与该动物的正常基准值进行比较，得到差值（delta 值）。因此，delta 值表示在相应 12 小时白天或夜间时段相对于正常基准值（麻醉前建立）的偏离程度。柱状图表示 8 只小鼠的均值；误差线表示标准差。星号表示 P ≤0.05 的显著性差异（采用单因素方差分析，比较麻醉后四天内各组均值与正常值）。
讨论
遥测技术是生物医学研究中生理参数测量传统方法的一种强大替代方案。目前市场上已有高质量的遥测系统，包括可植入式发射器、接收器以及数据采集与分析的硬件和软件，甚至适用于小至小鼠的动物。遥测是目前唯一可用于从无束缚、自由活动的小鼠中采集数据的技术。利用该方法，现在可以在动物处于其熟悉环境中的情况下，持续和/或长时间地收集数据，从而最大限度地减少动物的应激反应以及由此产生的实验干扰。导联的形状和位置经过优化，可在快速运动（如挣扎、奔跑、打斗）或直立姿势时仍能获取信号9。因此，可在多种实验过程中获得准确的测量结果，例如在麻醉期间、应激诱导期间、跑台运动时、行为学实验期间、感染实验期间以及其他多种实验情境中。
然而，为了获得可靠、可重复且无伪迹的数据，排除环境干扰至关重要，我们特别强调标准化条件的重要性。建议将实验空间与电子噪声和声学噪声（包括小鼠尤为敏感的超声波）隔离开来。此外，在进行测量时，应禁止任何干扰，例如访客进入或不相关的实验操作。为了避免干扰因素（特别是在进行笼内测量时），所有必要的动物饲养操作应在每次测量开始前于实验室内完成。另外，小鼠的饲养方式——尤其是使用雄性小鼠时——无论是群养还是单养，都可能影响测量结果，在实验设计时必须予以考虑9。同时，小鼠必须健康且无鼠源性病原体，因为潜在或明显的感染、疾病或其他任何健康问题均可能对生理参数和活动行为产生显著影响。因此，小鼠在植入手术后应充分恢复，并在开始任何实验前给予足够时间以适应体内发射器的存在。
在小鼠中通过遥测技术进行数据采集，需要预先通过手术植入遥测发射器。该操作只能由具备外科技能的受过培训的人员进行，以尽量减少组织创伤以及后续的疼痛和痛苦。对于具备基本甚至高级（显微）外科技能的研究人员，建议首先使用新鲜的小鼠尸体，并采用训练用植入物（即制造商提供的模拟装置）进行初步练习，以建立操作流程并熟悉此类手术的具体细节。经过此类培训后，大多数研究人员将能够成功植入此类发射器，并在数次植入操作后达到实用的熟练程度。
手术过程中应保持无菌条件，以降低微生物负荷和感染风险。然而，如果因小鼠身上某些特定的、与无菌相冲突的情况而怀疑无菌操作已被破坏（例如大面积脱毛和消毒带来的冷却效应，以及包扎伤口在实际操作中的困难），则应在植入过程中给予抗感染预防措施。适当调整的镇痛治疗、明确的监测计划以及充分的术后护理，对于实验的顺利结果起着至关重要的作用。
总体而言，对小鼠进行遥测发射器手术植入将对其造成一定程度的应激。特别是当特定小鼠品系的基因修饰影响其表型并损害动物身体状况时，围手术期可能出现并发症，且植入后死亡率升高的风险增加。为避免不必要的痛苦，对于恢复不理想或康复期延长的个体，应在其达到濒死状态之前及时终止实验并实施安乐死。为此，已制定一份数据记录表（表1：一般状况与健康监测记录表），用于系统性地监测关键症状，并提供有关人道终点的指导建议。因此，恢复过程将以医疗记录或实验记录本的形式进行记录，从而使本方法（即植入手术及术后恢复）的实施过程对相关监管机构及负责动物实验的动物福利机构（如IACUC）保持透明。
披露
未声明任何利益冲突。
致谢
作者谨此感谢 Charles River Germany 提供 CD-1 小鼠。我们还感谢 Robin Schneider 及中央生物实验室工作人员在小鼠饲养方面提供的支持。衷心感谢 Flora Nicholls 提供出色的技术协助，以及 Kurt Burki 教授慷慨提供科研设施与资源。
参考文献
- Cesarovic, N. Isoflurane and sevoflurane provide equally effective anesthesia in laboratory mice. Lab. Anim. 44, 329-336 (2010).
- Gross, V., Luft, F. C. Exercising restraint in measuring blood pressure in conscious mice. Hypertension. 41, 879-881 (2003).
- Kramer, K., Kinter, L. B. Evaluation and applications of radiotelemetry in small laboratory animals. Physiol. Genomics. 13, 197-205 (2003).
- Kramer, K. Use of telemetry to record electrocardiogram and heart rate in freely moving mice. J. Pharmacol. Toxicol. Methods. 30, 209-215 (1993).
- Arras, M., Rettich, A., Cinelli, P., Kasermann, H. P., Burki, K. Assessment of post-laparotomy pain in laboratory mice by telemetric recording of heart rate and heart rate variability. BMC. Vet. Res. 3, 16-16 (2007).
- Schuler, B., Rettich, A., Vogel, J., Gassmann,, Arras, M. Optimized surgical techniques and postoperative care improve survival rates and permit accurate telemetric recording in exercising mice. BMC. Vet. Res. 5, 28-28 (2009).
- Pritchett-Corning, K. R., Mulder, G. B., Luo, Y., White, W. J. Principles of Rodent Surgery for the New Surgeon. J. Vis. Exp. (47), e2586-e2586 (2011).
- Rettich, A., Kasermann, H. P., Pelczar, P., Burki, K., Arras, M. The physiological and behavioral impact of sensory contact among unfamiliar adult mice in the laboratory. J. Appl. Anim. Welf. Sci. 9, 277-288 (2006).
- Spani, D., Arras, M., Konig, B., Rulicke, T. Higher heart rate of laboratory mice housed individually vs in pairs. Lab. Anim. 37, 54-62 (2003).
- Zeller, A., Arras, M., Jurd, R., Rudolph, U. Mapping the contribution of beta3-containing GABAA receptors to volatile and intravenous general anesthetic actions. BMC. Pharmacol. 7, 2-2 (2007).
- Zeller, A., Arras, M., Jurd, R., Rudolph, U. Identification of a molecular target mediating the general anesthetic actions of pentobarbital. Mol. Pharmacol. 71, 852-859 (2007).
- Arras, M. Improvement of pain therapy in laboratory mice. Altex. 24, 6-8 (2007).
重印与许可
勘误
Formal Correction: Erratum: Implantation of Radiotelemetry Transmitters Yielding Data on ECG, Heart Rate, Core Body Temperature and Activity in Free-moving Laboratory Mice
Posted by JoVE Editors on 10/09/2016. Citeable Link.
Corrections in the Protocol and Discussion sections have been made to: Implantation of Radiotelemetry Transmitters Yielding Data on ECG, Heart Rate, Core Body Temperature and Activity in Free-moving Laboratory Mice
Step 1.2 in the Protocol has been updated from:
1.2 Hair clipping at one day prior to surgery
The day prior to implantation, in order to shave the animals for surgery, mice are anesthetized briefly in a small (8x8x8cm) Perspex chamber using sevoflurane (8%) or isoflurane (5%) in pure oxygen (600 mL/min). After loss of the righting reflex, the mouse is taken out of the chamber and the anterior neck and abdominal hair is clipped with the animal lying in dorsal recumbence; anesthesia is maintained for approximately 5 minutes with a nose mask with sevoflurane 3-4% or isoflurane 1.5-3% in pure oxygen at a flow rate of 600 mL/min. After clipping the hair, the animals are allowed to awaken and are then brought back to their home cage.
to:
After the last health check or directly prior surgery, in order to shave the animals for surgery, mice are anesthetized briefly in a small (8x8x8cm) Perspex chamber using sevoflurane (8%) or isoflurane (5%) in pure oxygen (600 mL/min). Shaving the animals one day before surgery prevents hair stubbles in the operating field. After loss of the righting reflex, the mouse is taken out of the chamber and the anterior neck and abdominal hair is clipped with the animal lying in dorsal recumbence; anesthesia is maintained for approximately 5 minutes with a nose mask with sevoflurane 3-4% or isoflurane 1.5-3% in pure oxygen at a flow rate of 600 mL/min. After clipping the hair, the animals are allowed to awaken and are then brought back to their home cage.
Step 2.3 in the Protocol has been updated from:
2.3 Surgery
The skin of the anterior neck and abdominal region is disinfected with 70% ethanol. A 1- to 1.5-cm-long incision in the skin is made from the lower thorax along the midline to the abdomen. The negative (white/colourless) lead is tunnelled subcutaneously from the thorax to the neck, where a small incision (≤0.5 cm) is made in the longitudinal direction. The skin and underlying tissues are prepared to make space for the fixation of the wire loop of the electrode. The wire loop is fixed between the muscles located to the right of the trachea, using two thin silk sutures (PERMA-Handseide, 6-0, Ethicon, Norderstedt, Germany). The wound in the neck is then closed with absorbable sutures (VICRYL 6-0, Ethicon, Norderstedt, Germany) in layers. The abdominal wall is then opened at the linea alba and the body of the telemetric transmitter is placed into the abdominal cavity of the mouse. The wire loop of the positive (red) electrode is sutured to the xiphoid process with silk sutures in such a way that it lies between the liver and the diaphragm in the left upper abdominal region (Figure 2). Then, the muscle layers of the abdominal region are closed with absorbable sutures (VICRYL 6-0, Ethicon, Norderstedt, Germany). Before finally closing the abdominal wall, a mixture of Sulfadoxin and Trimethoprim [(30 mg/kg and 6 mg/kg, respectively; dissolved in 1 mL of saline (0.9%) and at approximately body temperature (38-39°C)] is injected into the abdominal cavity for the purposes of anti-infective prophylaxis and to support fluid homeostasis. Finally, the skin of the abdominal region is restored with staples (Precise, 3 M Health Care, St. Paul, MN, USA).
to:
2.3 Surgery
The skin of the anterior neck and abdominal region is disinfected for 5 minutes with 70% ethanol, chlorhexidine or iodine using a soaked cotton swab. A 1- to 1.5-cm-long incision in the skin is made from the lower thorax along the midline to the abdomen. The negative (white/colourless) lead is tunnelled subcutaneously from the thorax to the neck, where a small incision (≤0.5 cm) is made in the longitudinal direction. The skin and underlying tissues are prepared to make space for the fixation of the wire loop of the electrode. The wire loop is fixed between the muscles located to the right of the trachea, using two thin silk sutures (PERMA-Handseide, 6-0, Ethicon, Norderstedt, Germany). The wound in the neck is then closed with absorbable sutures (VICRYL 6-0, Ethicon, Norderstedt, Germany) in layers. The abdominal wall is then opened at the linea alba and the body of the telemetric transmitter is placed into the abdominal cavity of the mouse. The wire loop of the positive (red) electrode is sutured to the xiphoid process with silk sutures in such a way that it lies between the liver and the diaphragm in the left upper abdominal region (Figure 2). Then, the muscle layers of the abdominal region are closed with absorbable sutures (VICRYL 6-0, Ethicon, Norderstedt, Germany). Before finally closing the abdominal wall, a mixture of Sulfadoxin and Trimethoprim [(30 mg/kg and 6 mg/kg, respectively; dissolved in 1 mL of saline (0.9%) and at approximately body temperature (38-39°C)] is injected into the abdominal cavity for the purposes of anti-infective prophylaxis and to support fluid homeostasis. Finally, the skin of the abdominal region is restored with staples (Precise, 3 M Health Care, St. Paul, MN, USA) or intracutaneous, running, absorbable sutures (VICRYL 6-0, Ethicon, Norderstedt, Germany).
Step 3 in the Protocol has been updated from:
3. Post-operative care
After completion of surgery and anesthesia, 0.1 mg/kg of buprenorphine (Temgesic, Essex Chemie AG, Lucerne, Switzerland) and 5 mg/kg of meloxicam (Metacam, Boehringer Ingelheim, Basel, Switzerland) is administered subcutaneously for pain treatment, and the animals are left on the warm (39°C +/-1) surface of the work bench to recover for approximately 2h. Together with pain relief (twice daily: buprenorphine, 0.1 mg/kg and meloxicam 5 mg/kg), supportive therapy consisting of 300 μL glucose (5%) and 300 μL saline (0.9%) warmed to body temperature, is applied subcutaneously twice daily for 4 days. For further recovery support, it is worthwhile providing the animals with an additional drinking bottle containing 15% glucose solution During the recovery period of 4-10 days, it is recommended that the animals are kept warm. Therefore, in our case, the mice are housed in a warming cabinet (30°C +/- 1). Monitoring of general condition and body weight, as well as food and water consumption, is performed once daily according to the general condition and health monitoring data sheet (Table 1) for 10 days post-operatively. Humane endpoints, i.e. the sacrifice of an animal to avoid unnecessary suffering and pain if progression of recovery is unsatisfactory, are realised under the following conditions:
i. If in poor general condition, i.e. the animal is substantially apathetic (no movement after being touched/pushed) and its body surface feels cold despite warming, the animal should be euthanatized immediately
ii. If, on day 4 after transmitter implantation, the animal shows clear signs of apathy, is extremely aggressive or does not show any food intake, it should be euthanatized immediately.
iii. On day 8 after transmitter implantation, the animal has to display a clear increase in body weight in comparison to the preceding post-operative days. Moreover, it has to consume at least 80% of the pre-operative daily food intake. If one of these conditions is not met, the animal should be euthanatized immediately.
At 10 days after implantation, the animal is transferred back to the animal room under standard housing conditions. Mice should be housed in compatible groups to allow social interaction and to prevent the adverse effects of long-term individual housing, which can have substantial impacts on the read-out of subsequent experiments8, 9. Mice should have a period of at least 4 weeks convalescence after transmitter implantation before the first experiment is conducted and data acquisition begins.
to:
After completion of surgery and anesthesia, 0.1 mg/kg of buprenorphine (Temgesic, Essex Chemie AG, Lucerne, Switzerland) and 5 mg/kg of meloxicam (Metacam, Boehringer Ingelheim, Basel, Switzerland) is administered subcutaneously for pain treatment, and the animals are left on the warm (39°C +/-1) surface of the work bench to recover for approximately 2h. Together with pain relief (twice daily: buprenorphine, 0.1 mg/kg and meloxicam 5 mg/kg), supportive therapy consisting of 300 μL glucose (5%) and 300 μL saline (0.9%) warmed to body temperature, is injected subcutaneously twice daily for 4 days. For further recovery support, it is worthwhile providing the animals with an additional drinking bottle containing 15% glucose solution During the recovery period of 4-10 days, it is recommended that the animals are kept warm. Therefore, in our case, the mice are housed in a warming cabinet (30°C +/- 1). Monitoring of general condition and body weight, as well as food and water consumption, is performed once daily according to the general condition and health monitoring data sheet (Table 1) for 10 days post-operatively. Humane endpoints, i.e. the sacrifice of an animal to avoid unnecessary suffering and pain if progression of recovery is unsatisfactory, are realised under the following conditions:
i. If in poor general condition, i.e. the animal is substantially apathetic (no movement after being touched/pushed) and its body surface feels cold despite warming, the animal should be euthanatized immediately
ii. If, on day 4 after transmitter implantation, the animal shows clear signs of apathy, is extremely aggressive or does not show any food intake, it should be euthanatized immediately.
iii. On day 8 after transmitter implantation, the animal has to display a clear increase in body weight in comparison to the preceding post-operative days. Moreover, it has to consume at least 80% of the pre-operative daily food intake. If one of these conditions is not met, the animal should be euthanatized immediately.
At 10 days after implantation, the animal is transferred back to the animal room under standard housing conditions. In case staples have been used, these should be removed 7-10 days after surgery; absorbable sutures have not to be removed. Mice should be housed in compatible groups to allow social interaction and to prevent the adverse effects of long-term individual housing, which can have substantial impacts on the read-out of subsequent experiments8, 9. Mice should have a period of at least 4 weeks convalescence after transmitter implantation before the first experiment is conducted and data acquisition begins.
The 4th paragraph in the Discussion has been updated from:
Aseptic conditions should be maintained during surgery to keep the microbiological burden and the risk of infections low. However, complete sterility cannot be provided because of some specific, sterility conflicting conditions in mice (e.g., cooling effect of extensive hair clipping and disinfection, impracticality of bandages to protect the wounds). Thus, anti-infective prophylaxis is administered during the implantation. Well tailored analgesic treatment and a clearly defined monitoring plan as well as adequate post-operative care play a crucial role in the satisfactory outcome of the experiment.
to:
Aseptic conditions should be maintained during surgery to keep the microbiological burden and the risk of infections low. However, if there are doubts that asepsis was breached because of some specific, sterility conflicting conditions in mice (e.g., cooling effect of extensive hair clipping and disinfection, impracticality of bandages to protect the wounds). Anti-infective prophylaxis should be administered during the implantation. Well-tailored analgesic treatment and a clearly defined monitoring plan as well as adequate post-operative care play a crucial role in the satisfactory outcome of the experiment.
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