来源: 凯斯图尔特, RVT, RLATG, CMAR;瓦莱丽. 施罗德, RVT, RLATG。圣母大学, 在
实验室动物的护理和使用指南 (和 #34; 指南和 #34;) 指出疼痛评估和减轻是实验动物兽医护理的组成部分. 1 麻醉的定义是感觉或感受的损失。这是一个动态的事件, 涉及麻醉深度的变化与动物和 #39 的新陈代谢, 外科刺激, 或在外部环境的变化.
需要精确和持续的麻醉监测, 以安全地保持程序所需的深度。需要监测的参数包括心率、呼吸速率、体温和血氧水平。对于小鼠和大鼠, 这些参数都不容易监测, 因为这些动物和 #39; 小体型。由于啮齿类动物的心率如此之快, 通常用于听诊的听诊器不足以获取准确的心率。听诊器只能用于检测心跳的存在或缺失。一只老鼠的正常心率是每分钟328-780 拍, 而老鼠的常规心率是每分钟250-600 次。啮齿类动物的呼吸率也高于可以用视觉方法或听诊时准确计算的。一只老鼠的正常呼吸率是每分钟90-220 次呼吸, 对于老鼠来说, 这个值是每分钟66-144 呼吸。为了准确地确定心率和呼吸率, 需要专门的电子监测设备。传感器要么在手术中植入动物体内, 要么置于外部, 并与动物所放置的监控平台进行交互。 34
在啮齿类动物中, 与麻醉有关的死亡最常见的原因是体温过低。啮齿目动物有高表面面积到身体质量比率。此外, 麻醉动物失去了颤抖的能力, 以保持体温。因此, 体温监测和补充热量, 如加热垫, 是必不可少的在生存手术过程中。鼠标的正常体温为 96.6-99.7 和 #176; f (35.8-37.4 和 #176; c) 5 , 对于老鼠是 96.6-99.5 和 #176; f (35.9-37.5 和 #176; c)。 5 大多数温度计是为较大的动物设计的, 并且是为人类使用的。水银温度计已在很大程度上取代了数字和电子版本。虽然数字和电子温度计已被证明是准确的, 当使用直肠, 口服, 并在耳朵, 它们的大小是不合适的小啮齿动物。专门为小鼠和大鼠设计的直肠探针在商业上可用, 并鼓励使用它们.
血液氧合水平用于评估肺部是否有足够的吸氧量, 从而在啮齿动物和 #39 的动脉血中适当集中氧气。监测吸氧量也间接监测呼吸和通气, 因为它揭示了是否有足够的氧气的启发和废气的失效。心率也与血液的氧合有关, 因为心率的降低会导致氧水平的降低, 这可能导致血液灌注不足。 6
麻醉的目标是充分地固定和缓解所有的疼痛感觉为一个动物的最低剂量或浓度的麻醉。正确评估麻醉深度需要达到这一目标。麻醉的手术阶段有四阶段的麻醉和四架飞机。在第一阶段, 动物变得迷失方向。在第二阶段, 有一个不规则的呼吸速率兴奋阶段, 包括呼吸在一些小鼠和大鼠品系。矫正反射-这是能力回滚时, 放置在背部位置-也失去了.
阶段三是麻醉的外科阶段。在平面 I, 眼睑和吞咽反应是缺席。喉和角膜反射在平面 II. 中丢失与飞机 I 和 II, 没有健忘症或镇痛作用;因此, 在外科手术开始之前, 动物必须到达第三面。平面 III 造成的肋间肌肉麻痹, 导致膈呼吸。虽然最初在平面 III 只有部分镇痛, 但随着麻醉水平的加深, 它的进展完全失忆和镇痛。在这个层次上, 动物被完全麻醉的外科手术。在飞机 IV, 该动物已过量, 并可以继续迅速进入阶段 IV.
随着麻醉水平的进一步加深, 有可能导致动物死亡的并发症。在第四阶段, 肋间肌肉和膈肌都完全瘫痪, 导致严重的窒息。这导致呼吸停止, 髓质麻痹, 血管崩溃, 和最后死亡。瞳孔扩张, 在肌肉松弛时保持固定的膨胀.
正确选择手术麻醉药和其他可能的疼痛程序必须由兽医确定。这是基于许多方面, 包括程序的范围和持续时间, 物种和应变, 年龄, 和生理状况的动物.
麻醉剂可作为剂或注射。手术麻醉可以使用注射和吸入麻醉剂的组合来完成。 2
1。吸入麻醉诱导
吸入麻醉包括异氟醚、七氟醚和地氟醚, 其中最常用的是异氟醚。这些麻醉药使用更频繁, 因为, 与他们, 它是更容易控制麻醉深度。使用吸入麻醉剂进行麻醉的诱导可以用装有精密汽化器的钟罩或感应室来完成.
2。注射用麻醉药诱导麻醉
注射麻醉剂主要是氯胺酮和镇静剂或肌肉剂的混合物.
常见的组合为: 1) 啮齿动物鸡尾酒, 由氯胺酮 (100 毫克/毫升)、嗪 (20 毫克/毫升)、乙酰 (10 毫克/毫升) 和无菌盐水 (0.9% 氯化钠) 组成;2) 氯胺酮/嗪 2:1, 由氯胺酮 (100 毫克/毫升)、嗪 (20 毫克/毫升) 和无菌盐水 (0.9% 氯化钠) 组成;和 3) 氯胺酮/嗪鼠混合, 其中包括氯胺酮 (100 毫克/毫升), 嗪 (20 毫克/毫升), 和无菌生理盐水 (0.9% 氯化钠). 和 #160; 当使用氯胺酮/嗪组合, 促进只应该做与氯胺酮, 而不是嗪, 由于半衰期这些药. 和 #160; #160;
将氯胺酮与镇静剂和/或肌肉松弛的结合, 需要准备作为一个股票的解决方案, 从中可以抽取个人剂量。试剂必须精确测量, 用无菌生理盐水稀释, 以确保适当的剂量被管理的动物。由于氯胺酮是一种受控物质, 因此, 从瓶子中所使用的数量必须在 a 和 #34 上注明; 受控的药物记录和 #34; 和混合物必须有个人和 #34; 受控物质日志. #34; 当准备混合物时, 慢慢地加入氯胺酮瓶子, 因为它倾向于泡沫, 如果注入的力量。一种无菌塞20毫升瓶用于混合物。瓶子必须正确标明化合物的名称, 日期混合, 到期日期, 氯胺酮批号 (因为它是一个受控物质), 和建议的剂量。到期日可由配料的日期确定, 以最快的时间届满 (取决于设施/状态的规则/准则)。为了准确记录氯胺酮, 必须权衡空瓶和填充瓶。然后, 重量必须记录在标签上的混合物和单独的控制物质的记录表, 为每个瓶子准备。将氯胺酮混合物储存在黑暗、温度控制的区域以保持效力.
3。麻醉评估
麻醉深度可以通过测试对各种刺激的反应来评估。自愿运动将由身体的身体刺激产生。有关用于麻醉深度评估的物理方法的列表, 请参见表 1.
方法 | 过程 | 响应 |
脚趾夹 | 扩展腿并隔离脚趾.这一地区被牢牢地捏使用指甲或创伤钳. | 正面反射通过腿的缩回或撤退表明脚。如果有腿部或身体运动, 发声, 或明显增加的呼吸, 动物不是在外科手术的麻醉平面上. |
尾部捏 | 尾尖用手指或创伤钳夹住. | 阳性反应通过尾巴的抽动或移动来表示。如果有尾巴的运动, 发声, 或有明显增加的呼吸, 动物就不是在手术的麻醉平面上. |
耳掐 | 使用手指或创伤钳, 捏耳廓的尖端. | 正面的反应是摇动头部或向前的胡须运动。如果有头部的运动, 胡须, 发声, 或明显增加的呼吸, 该动物不是在一个外科手术平面的麻醉. |
眼睑反射 | 使用指尖, 触摸眼睛的内侧眼角 (内部角)。 | 一个积极的反射是由眨眼表示的反应, 以触摸眼睑。如果有眼睑的运动, 胡须, 或明显增加的呼吸, 该动物不是在一个外科手术平面的麻醉. |
角膜反射 | 使用棉签, 轻轻触摸角膜 (眼球)。 | 闪烁表示正响应。如果有眼睑的运动, 胡须, 或明显增加的呼吸, 该动物不是在一个足够深的手术麻醉的平面. |
表 1. 用于评估麻醉深度的物理刺激方法。 2
生理指标, 如心率、呼吸率、血压、粘膜颜色、毛细血管充盈时间也应使用。虽然一般的观察可以用来检测动物的呼吸率的变化, 利用心率, 或血压进行深度评估, 需要专门的设备。如果有心电图, 心率和心跳的强度可以测量。为了测量血压, 有各种各样的装置安装在尾部甚至整个身体。如表1所述的物理刺激将导致所有这些参数中的三增加.
粘膜, 眼睛, 耳朵, 嘴巴, 鼻子, 肛门的颜色, 和-在较小程度上-观察爪子和尾巴的变化。区域应该是粉红色的, 表示充分的呼吸作用和心脏功能。当动物移动到阶段 IV 麻醉, 呼吸停止, 导致青紫-由蓝色或灰色表明-对黏膜和周围的皮肤.
毛细管重填时间定义为颜色返回到外部毛细管床的时间量, 因为在该区域施加压力后, 它已经被漂白。在麻醉动物的牙龈、耳廓或指甲床上按压涂抹棒或手指。脱皮区域返回粉红色的秒数不应该更多。1-2 秒延长的再填充时间表明心脏收缩的心率或强度降低, 表明动物可能是太深麻醉和接近死亡.
重要的是要利用几个不同的参数来评估麻醉深度。使用相同的脚趾或耳朵反复捏将敏的区域, 和反应将被压抑, 而不是给一个准确的评估麻醉深度。使用备用站点进行脚趾和耳箍评估。麻醉深度应每10-30 分钟在整个手术中重新评估。 2
研究表明, 在麻醉动物中有心肺的变化。当麻醉与注射药物, 动物体验稳定的呼吸率;然而, 它们显示出心脏输出的变异性。对注射麻醉剂的反应在不同菌株之间有很大的差异, 因此很难对其剂量进行标准化。 7 吸入剂倾向于降低呼吸速率, 但对心血管系统的影响较小。由于吸入麻醉的剂量很容易在整个过程中进行调整, 因此通常是首选方法.
麻醉诱导和维护是接受任何形式的外科手术的实验动物的兽医护理的一个组成部分。麻醉的目的是充分地固定动物和减轻所有的疼痛感觉。除了感应外, 还需要精确和恒定的监测, 以便在整个过程中安全地保持正确的麻醉深度.
在本视频中, 我们将首先简要讨论啮齿类动物麻醉的水平, 以及一个应该达到的阶段。接下来, 我们将审查不同的归纳和维护方法, 各种方法, 以确保动物总是在理想的麻醉阶段, 最后一些现实世界的实验, 涉及使用不同的麻醉剂, 为各种目的.
让与 #39; s 从讨论级别开始。有四阶段的麻醉和四架飞机在三阶段或手术阶段.
在第一阶段, 动物就会迷失方向。第二阶段是以不规则的呼吸速率和失扶正反射为标志。在三阶段的第一个平面上, 眼睑和吞咽反应是缺席的。在第二个平面中, 喉和角膜反射失去了。直到这一点, 麻醉剂并没有导致健忘或镇痛.
在平面三中, 健忘症和镇痛的进展从部分到完整, 动物完全麻醉为外科手术。平面三也是由肋间肌肉麻痹的标志, 这导致膈呼吸是浅呼吸。在飞机四, 该动物已经过量, 并可以迅速进入四阶段, 那里有完全瘫痪的肋间肌肉和隔膜, 这可能导致呼吸骤停, 并最终导致死亡.
麻醉剂可作为吸入或注射剂, 兽医必须决定使用什么程序才能执行。这个选择是基于许多方面, 包括: 程序的范围和持续时间, 物种和应变, 年龄和动物的生理状况.
通常使用的吸入麻醉剂类包括异氟醚、七氟醚和地氟醚等化合物。这些化合物可以方便地控制麻醉深度。在设备上有几个选项可以选择来管理吸入麻醉剂.
其中一个选择是响铃罐子, 应该使用在敞篷之下-和不在长凳-避免人员接触到麻醉剂气体。用陶瓷或塑料穿孔平台组装罐子, 在罐子的底部和平台之间创造一个空间。接着, 戴上防渗手套, 用麻醉剂浸透棉球, 放在平台下, 使其停留在罐子的底部。然后立即保护盖子, 以防止麻醉蒸气逃逸。将动物放在一边, 将盖子滑向一侧, 介绍动物并立即将其固定。随后, 观察活动和呼吸, 以确定麻醉深度, 并将动物暴露于吸入 效果 。请注意, 该平台作为屏障, 防止动物与液体麻醉剂直接接触.
响铃罐的另一种替代方法是与与氧气罐连接的精密汽化器机一起使用的感应室。第一步是确保汽化器充满适量的液体麻醉剂。下一步, 检查废气净化系统。如果它是通常使用的被动系统, 然后称量罐, 以确定它是否仍然有效。一般来说, 增加五十克以上的起始重量是一个点, 在这个容器的花费。下一步是组装感应室。确保输入是从蒸发器和输出的废气清除系统.
启动时, 将动物放入感应室并固定盖子。一旦动物是在会议厅, 首先开始的氧气流量以每分钟1公升的速度, 然后调整精确度汽化器设置到感应水平 3-4% 的异氟醚。象响铃罐子, 暴露动物到麻醉剂 起作用. 一旦动物完全麻醉, 将异氟醚关闭, 在轻轻地移除动物之前, 用氧气冲洗房间。这是为了防止人员接触麻醉剂气体.
麻醉诱导的另一种方法是通过鼻锥或口罩也连接到精密汽化器。然而, 因为麻醉气体有难闻的气味, 动物可能反对被掩没为归纳。此外, 由于抓得太紧, 还有导致窒息的危险。因此, 首选的方法是使用感应箱或钟罩来诱导麻醉, 然后用鼻锥进行维护。通常情况下, 装配是这样的, 锥和感应室都是连接到同一个蒸发器与一个开关之间的麻醉蒸汽传递从感应室到鼻锥, 反之亦然。麻醉后, 在房间里的动物, 安全它的脸在锥, 并切换到油管的开关, 以重定向气体流到鼻子锥。监测呼吸和确认动物放松后, 将麻醉剂降低到维持 0.5-1.5% 的水平。同时, 将眼部软膏应用于眼部, 防止角膜干燥.
用于注射麻醉剂, 混合氯胺酮和其他镇静剂或肌肉剂包括嗪和/或乙酰。使用这些化合物可以制备不同的组合。请参阅下面的文本以了解常用比率。请注意, 氯胺酮是一种受控物质, 因此所使用的量必须在受控的药物记录上注明, 而混合物必须有各自的受控物质记录。根据动物的种类、年龄和健康状况, 选择麻醉药的混合物和剂量, 并腹腔或注。通常, 注射和吸入麻醉剂用于联合手术麻醉.
现在您知道如何诱导 anesthe让 & #39 了解麻醉深度评估, 这是很重要的, 每10-30 分钟监测, 以确保在程序中的动物不会受到伤害。在啮齿目动物中有几种方法.
通常使用的方法是脚趾捏。伸展动物和 #39 的腿, 把脚趾间的带子隔离开来。然后用指甲或创伤钳牢牢地捏住区域。正面反射由腿的缩回或撤退表示。另一种方法是在尾部的尖端进行的尾部捏。阳性反应表现为抽搐或尾部运动。你也可以捏耳廓的尖端, 如果有晃动的头部或向前的胡须运动, 那么动物不是在外科手术平面的麻醉.
检查麻醉深度, 也可以触摸内侧眼角或眼内眼角, 以引起眼睑反射–由眨眼表示, 以响应眼睑的接触。即使有眼睑、胡须或明显增加的呼吸, 动物也不在麻醉的手术平面上.
最后, 你可以通过用戴手套的手指或棉花尖的涂抹器触摸角膜来检查角膜反射。闪烁表示正响应.
在评估麻醉深度时, 必须在站点间交替进行。使用相同的脚趾或耳朵反复捏将敏的区域和反应将被压抑, 而不是给出一个准确的麻醉深度评估.
除了这些物理刺激的评估方法外, 还应监测生理指标, 包括心率、呼吸速率、血压、粘膜颜色和毛细血管充盈时间。虽然一般的观察可以用来检测呼吸频率的变化, 利用心率进行深度评估, 但可以使用像心电图这样的专业设备。为了测量血压, 有各种各样的设备可以安装在尾部甚至整个身体。粘膜、眼睛、耳朵、嘴巴、鼻子、肛门、爪子和尾巴的颜色也可以显示麻醉深度。这些区域应该是粉红色的, 建议足够的呼吸和心率.
检查毛细管重填时间, 按下麻醉动物的耳廓, 并计算脱皮区域返回到粉红色颜色所需的秒数。这不应该超过1到2秒。延长的再填充时间表明心脏收缩的心率或强度降低, 表明动物可能是太深麻醉和接近死亡。将动物从麻醉中取出后, 除非在住房区持续监测, 否则不应将其送回住房设施, 直到从麻醉中恢复.
现在我们和 #39; 我了解了啮齿类麻醉诱导和维护的原理和程序, 让我们看看当今生物医学研究中常用的麻醉剂的一些应用.
可能最常用的啮齿动物麻醉是在手术之前和。例如, 在这里, 研究人员希望开发一种由脑内血栓形成引起的中风模型。为了达到这一目的, 他们在小鼠体内进行麻醉, 然后钻到头盖骨上, 形成一个薄薄的窗户。当这只动物还在服用镇静剂时, 这些科学家在循环中注入了光敏染料。接着, 他们借助激光通过钻孔的颅骨诱导活化, 导致颅内血管形成血块.
需要进行啮齿动物麻醉的另一个实例是进行生理分析。例如, 科学家经常在麻醉动物身上使用心电图电极来监测心脏活动。或者他们使用超声波探头来确定膈肌运动的速度, 以更准确地量化呼吸速率.
最后, 使用麻醉是强制性的, 当预生存 宫内 实验。例如, 在子宫内 电穿孔–一种方法, 其中怀孕的女性被麻醉, 一个切口, 以揭露发育中的胚胎, 和电极被用来诱导胚胎细胞摄取注射的遗传物质.
您刚刚观看了朱庇特和 #39; 关于麻醉管理和维护的视频。由于啮齿类麻醉促进了如此广泛的生物实验的执行, 因此, 每一位科学家都必须具备在整个实验中诱导和维持正确的麻醉深度的技巧。一如既往, 感谢收看!
正确使用麻醉药进行手术或其他潜在的痛苦的程序, 不仅对动物和 #39 的福祉至关重要, 而且对在过程中收集的科学数据的完整性也很重要。选择合适的麻醉团有很多变数。麻醉深度必须密切监测, 因为每个个体动物对药物的反应都不同。使用适当的麻醉和仔细的监测, 痛苦的程序可以完成, 没有疼痛和最小的生理变化的动物.
Anesthesia induction and maintenance forms an integral component of veterinary care of laboratory animals undergoing any form of surgical procedure. The goal of anesthesia is to adequately immobilize the animal and alleviate all pain sensations. In addition to induction, precise and constant monitoring is required to safely maintain the correct anesthetic depth throughout the procedure.
In this video, we will first briefly discuss the levels of rodent anesthesia and what stage one should aim to achieve. Next, we will review the different induction and maintenance methods, various ways to ensure that the animal is always in the desired anesthetic stage, and finally a few real-world experiments involving use of different anesthetics for varied purposes.
Let’s start by discussing the levels. There are four stages of anesthesia and four planes within the stage three or the surgical stage.
During stage one, the animal becomes disoriented. Stage two is marked by an irregular respiratory rate and loss of the righting reflex. In plane one of stage three, the palpebral and swallowing reflexes are absent. During the plane two, the laryngeal and corneal reflexes are lost.Up until this point, the anesthetic has not induced amnesia or analgesia.
It is in plane three that amnesia and analgesia progresses from partial to complete, and the animal is fully anesthetized for a surgical procedure. Plane three is also signified by paralysis of the intercostal muscles, which results in diaphragmatic respiration that is shallow breathing. In plane four, the animal has been overdosed and can proceed quickly into stage four, where there is complete paralysis of both intercostal muscles and diaphragm, which can cause respiratory arrest, and ultimately lead to death.
Anesthetics are available as an inhalant or injectable, and a veterinarian must decide what to use for the procedure to be performed. This choice is based on numerous aspects including: the extent and duration of the procedure, the species and strain, the age, and the physiological status of the animal.
The class of commonly used inhalant anesthetics includes compounds like Isoflurane, Sevoflurane, and Desflurane. These compounds allow for easy control of the anesthesia depth. There are a few options in equipment that one can choose from to administer inhalant anesthetics.
One of the choices is a bell jar, which should be used under the hood — and not on the bench — to avoid personnel exposure to the anesthetic gases. Assemble the jar with a ceramic or plastic perforated platform creating a space between the bottom of the jar and the platform. Next, while wearing impervious gloves, saturate a cotton ball with anesthetic and place it under the platform so that it rests at the bottom of the jar. Then immediately secure the lid to prevent escape of the anesthetic vapor. To place the animal, slide the lid to one side, introduce the animal and secure it immediately. Following that, observe the activity and respirations to determine the depth of anesthesia, and expose the animal to the inhalant to effect. Note that the platform serves as a barrier and prevents the animal from coming into direct contact with the liquid anesthetic.
An alternative to the bell jar is an induction chamber used in conjunction with a precision vaporizer machine connected to an oxygen tank. The first step is to ensure that the vaporizer is filled with appropriate amount of the liquid anesthetic. Next, check the waste gas scavenging system. If it is the commonly used passive system, then weigh the canister to determine if it is still effective. Generally an increase of fifty grams above the starting weight is the point at which the canister is spent. The next step is to assemble the induction chamber. Ensure that the input is from the vaporizer and output is to the waste gas scavenging system.
To start, place the animal into the induction chamber and secure the lid. Once the animal is in the chamber, first start the oxygen flow at the rate of 1 liter per min, and then adjust the precision vaporizer setting to an induction level of 3-4 % for isoflurane. Like bell jar, expose the animal to the anesthetic to effect. Once the animal is fully anesthetized, flush the chamber with oxygen by turning the isoflurane off before gently removing the animal. This is to prevent personnel exposure to anesthetic gases.
Another method for anesthesia induction is via nose cone or facemask also connected to the precision vaporizer. However, because anesthetic gases have an unpleasant smell, animals may object to being masked for induction. In addition, there is also a risk of causing asphyxiation because of grasping too firmly. Therefore, the preferred method is to use the induction box or the bell jar to induce anesthesia followed by maintenance with the nose cone. More often than not, the assembly is such that the cone and the induction chamber both are connected to the same vaporizer with a toggle in between to switch the anesthetic vapor delivery from the induction chamber to the nose cone and vice versa. After anesthetizing the animal in the chamber, secure its face in the cone, and switch the toggle on the tubing to redirect the gas flow to the nose cone. Monitor the respiration and after confirming that the animal is relaxed, reduce the anesthetic to a maintenance level of 0.5 – 1.5 %. Also, apply ophthalmic ointment to the eyes to prevent corneal drying.
For injectable anesthetics, a mixture of Ketamine and other sedatives or muscle relaxers including Xylazine and/or Acepromazine. Different combinations can be prepared using these compounds. See text below for commonly used ratios. Note that Ketamine is a controlled substance and therefore the amount used must be noted on the Controlled Drug Log and the mixtures must have their individual Controlled Substance Logs. Depending on the species, age and health status of the animal, the mixture and the dose of the anesthetic are selected and the solution may be injected intraperitoneally or intramuscularly. Usually, injection and inhalation anesthetics are used in combination to achieve surgical anesthesia.
Now that you know how to induce anesthesia, let’s learn about anesthetic depth assessment, which is important to monitor every 10-30 minutes to ensure that the animal is not harmed during the procedure. There are several methods for doing so in rodents.
A commonly used method is the toe pinch. Extend the animal’s leg and isolate the webbing between the toes. Then firmly pinch the area using either the fingernails or atraumatic forceps. A positive reflex is indicated by the retraction of the leg or withdrawing of the foot. Another method is the tail pinch performed at the tip of the tail. A positive reaction is demonstrated by twitching or tail movement. You can also pinch the tip of the pinna, and if there is shaking of the head or the movement of the whiskers forward, then the animal is not in the surgical plane of anesthesia.
To check the anesthesia depth, one can also touch the medial canthus or the inner corner of the eye to elicit the palpebral reflex — indicated by a blink in response to touching of the eyelids. Even if there is movement of the eyelids, whiskers, or marked increase in respirations the animal is not in the surgical plane of anesthesia.
Lastly, one can check the corneal reflex by touching the cornea with gloved finger or a cotton-tipped applicator. A positive response is indicated by a blink.
It is important to alternate between sites to assess the anesthetic depth. Using the same toe or ear for repeated pinches will desensitize the area and the response will be repressed and not give an accurate assessment of anesthetic depth.
In addition to these physical stimuli methods of assessment, one should also monitor the physiological indicators including the heart rate, respiratory rate, blood pressure, mucous membrane color, and capillary refill time. While general observations can be useful to detect changes in the respiratory rate, to utilize the heart rate for depth assessment, specialized equipment like electrocardiograph may be used. For measuring the blood pressure, there are a variety of devices that can be fitted over the tail or even over the entire body. The color of mucous membranes, eyes, ears, mouth, nose, anus, paws, and tail can also indicate anesthetic depth. These areas should be pink, suggesting adequate respiration and heart rate.
To check the capillary refill time, press on the pinna of the anesthetized animals, and count the number of seconds that it takes for the blanched area to return to a pink color. This should not be more than 1 to 2 seconds. An extended refill time suggests a reduction in heart rate or strength of cardiac contraction, indicating the animal may be too deeply anesthetized and near death. After removing the animal from anesthesia, they should not be returned to the housing facility until recovered from anesthesia, unless they are continuously monitored in the housing area.
Now that we’ve learned the principles and procedures of rodent anesthesia induction and maintenance, lets look at some of the frequent applications of anesthetics in biomedical research today.
Probably the most common use for rodent anesthesia is prior to and during surgery. For example, here researchers wanted to develop a model of stroke caused by clot formation in brain. In order to achieve that, they induced anesthesia in mice and then drilled the cranium to create a thin window. And while the animal was still sedated, these scientists injected a photosensitive dye into the circulation. Next, they induced photoactivation with the help of a laser through the drilled cranium to cause formation of a clot in the cranial vasculature.
Another instance in which rodent anesthesia is required is for performing physiological analysis. For example, scientists often use ECG electrodes on anesthetized animals to monitor heart activity. Or they use ultrasound probes to determine the rate of diaphragm movement to more accurately quantify the respiratory rate.
Lastly, use of anesthesia is mandatory when preforming survival in utero experiments. For example, in utero electroporation — a method in which a pregnant female is anesthetized, an incision is made to expose the developing embryos, and electrodes are used to induce embryonic cellular uptake of the injected genetic material.
You have just watched JoVE’s video on anesthesia administration and maintenance. Since rodent anesthesia facilitates the execution of such a wide range of biological experiments, it is imperative that every scientist possesses the skill of inducing and maintaining the correct anesthetic depth throughout an experiment. As always, thanks for watching!
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