这项工作由Shriners Children's Boston（B. E. U.， K.U.， C.L.C.）资助。L.C.由“La Bourse des Gueules Cassées”，“La Bourse Année Recherche”和“La Bourse de l'Amicale des Anciens Internes des Hôpitaux de Paris”资助。Y.B.由“La Bourse des Gueules Cassées”资助。Y.B.和I.F.v.R.由Shriners儿童医院资助（奖学金ID分别为#84308-BOS-22 #84302-BOS-21）。本材料部分基于美国国家科学基金会根据资助号支持的工作。欧洲经济共同体1941543。非常感谢美国国立卫生研究院（R01EB028782、R56AI171958 和 R01DK114506）的部分支持。图 1 是使用 BioRender.com 创建的。

该程序在3个月大的雄性刘易斯大鼠上进行，每只重300-400克。共纳入24只动物，具有3种穿刺条件：12只大鼠在无麻醉的情况下进行尾静脉穿刺（组TV无麻醉），另外12只大鼠被麻醉进行尾静脉穿刺（组TV麻醉）和静脉穿刺（组PV麻醉）。所有程序均已获得批准，并遵守机构动物护理和使用委员会（IACUC）的指导方针。所有动物在研究结束时（随访1个月后）通过二氧化碳过量被安乐死。有关与本协议中使用的所有材料和仪器有关的详细信息，请参阅 材料表 。
1. 一般准则
<ol><li>根据 IACUC 指南，确保每 2 周抽取的最大血容量不超过总血容量的 10%9。例如，一只 300 g 的大鼠的总血容量应约为 19.2 mL。对于仅在第一周（第 0 天、第 1 天、第 3 天、第 7 天）需要抽血四次的方案，请将每个样品的采集限制为最多 250 μL。</li>
	<li>对于在麻醉下进行的手术，通过精密蒸发器 施 用异氟醚以麻醉动物。在剂量为3%-5%异氟醚的腔室中诱导麻醉5分钟，并在手术过程中通过鼻锥使用1%-3%异氟烷的剂量维持。根据呼吸频率的连续监测调整异氟烷水平。在开始手术之前，通过捏住脚趾来验证镇静是否足够。</li>
	<li>在手术过程中或直到动物恢复足够的意识以维持胸骨卧位之前，不要让动物无人看管。</li>
	<li>采血后，监测动物直到完全康复，然后再将其放回笼子，并且在完全恢复之前不要将其引入其他动物的陪伴下。 注意：与兽医服务达成协议，尾静脉或阴茎静脉穿刺后不需要术后止痛药。</li>
</ol>2. 从静脉抽血
<ol><li>制备<ol><li>准备以下设备：无菌纱布，手套，酒精湿巾，微毛细血管采血EDTA管（紫色帽）和30 G胰岛素注射器（30 U或50 U）。</li>
		<li>将大鼠从笼子中取出，并将其放入室中，通过精密蒸发器（剂量：3%-5%） 用 异氟醚诱导。一旦动物被镇静，将其转移到手术台上，然后将动物仰卧，鼻子放在鼻锥中以维持麻醉。监测呼吸频率，并相应地调整异氟醚水平（维持剂量：1%-3%）。在开始手术之前，通过捏脚趾来验证动物是否有足够的镇静剂。</li>
	</ol></li>
	<li>血液采样<ol><li>在注射器中来回移动柱塞几次以平滑退出。通过拉动柱塞去除几微升，在注射器中产生负压。</li>
		<li>在非惯用手的帮助下，将包皮从阴茎末端收回，并在食指和拇指之间握住龟头，轻轻拉动。背静脉将显示为浅表蓝带。请参见图 1 和 图 2。</li>
		<li>针眼朝上，将胰岛素注射器以35°角插入静脉。一旦针头进入静脉，血液就会流入注射器。</li>
		<li>以缓慢而稳定的速率缓慢抽出注射器的柱塞以收集所需的体积。 注意：不要太快地抽出柱塞，因为这会导致静脉塌陷并停止血流。</li>
		<li>如果血流量减少，请顺时针或逆时针稍微旋转。</li>
		<li>取出注射器。穿刺部位会形成血滴，在非无菌手术的情况下，抽吸将允许收集更多的微升血液。</li>
		<li>如果第一次穿刺失败，请将针头重新插入静脉近端。 注意：与尾静脉取样不同，背静脉的迭代穿刺通常不成功。</li>
		<li>对穿刺部位施加轻微压力以止血，并用新的酒精湿巾擦拭该区域。</li>
		<li>将阴茎放回中性位置。</li>
		<li>关闭异氟醚，并监测大鼠直到完全恢复。将老鼠放回笼子里。</li>
	</ol></li>
</ol>3.尾静脉穿刺
<ol><li>制备<ol><li>准备以下设备：塑料约束架，无菌纱布，手套，酒精湿巾，微毛细血管采血EDTA管（紫色帽）和28 G 1/2胰岛素注射器（30 U或50 U）。</li>
		<li>将老鼠从笼子里拿出来，并迅速将其固定在塑料约束锥中。关闭尾巴底部周围的锥体大端。确保动物舒适，并且在整个过程中呼吸不受限制。</li>
		<li>将尾巴浸入温水（37°C）中约1分钟以扩张静脉。用纸巾擦干尾巴。将动物（在其约束器中）面朝下放置，尾巴放在加热垫上。</li>
		<li>通过将整个动物旋转到任一侧来选择右尾静脉或左尾静脉（蓝线）进行采样（这样可以避免尾巴扭曲）。使用尾巴的末端三分之一进行血管穿刺，因为血管在这个区域变得更加浅表。动脉为腹侧，两条静脉为外侧10。</li>
		<li>在穿刺部位用70%乙醇湿巾擦拭尾巴。</li>
		<li>将尾巴放在加热垫的边缘，在尾巴的末端三分之一处形成一个角度。这将静脉带到表面，并为取样创造更多空间。</li>
	</ol></li>
	<li>血液采样<ol><li>在注射器中来回移动柱塞几次以平滑退出。通过拉动柱塞去除几微升，在注射器中产生负压。</li>
		<li>在非惯用食指和中指的帮助下，将尾巴平放在加热垫上。将中指放在近端，食指放在远端，穿刺部位在这两根手指之间。对中指施加比食指更大的压力以固定尾巴，仅在近端阻塞血管并让血液淤积。请参见图 1 和 图 3。</li>
		<li>针眼朝上，将胰岛素注射器滑过食指，直到将其插入静脉（这在针头和尾部之间产生35°的角度）。一旦针头进入静脉，血液就会流入注射器。此时，释放食指和中指上的压力，以确保血流不被阻塞。</li>
		<li>以稳定的速率缓慢抽出注射器的柱塞以收集所需的体积。 注意：不要太快地拔出柱塞;这将导致静脉塌陷并停止血液流动。</li>
		<li>如果血流量减少，请向任一方向稍微旋转针头。</li>
		<li>从尾部取出注射器。尾巴的穿刺部位会形成血滴。在非无菌手术的情况下，抽吸这种血液允许收集更多的微升血液。</li>
		<li>如果第一次穿刺失败，请将针头重新插入静脉近端。 注意：静脉在接近尾基时逐渐变得更深。如果注射器中没有血流，请增加注射器和尾部之间的角度，或旋转针头。</li>
		<li>对穿刺部位施加压力以止血，并用新的酒精湿巾擦拭该区域。将大鼠从塑料锥中取出，然后将其放回笼子中。</li>
	</ol></li>
	<li>麻醉下尾静脉穿刺<ol><li>执行步骤2.1.1和步骤2.1.2以诱导和维持麻醉。</li>
		<li>执行步骤3.1.3-3.2.7进行采血;参见 图1。</li>
		<li>执行步骤 2.2.10 以进行动物恢复。</li>
	</ol></li>
</ol><img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65513/65513fig01.jpg"> 图1：本协议中不同穿刺方法的示意图。 （A）对有意识的、克制的动物进行改良的尾静脉穿刺;（二）麻醉下改良尾静脉穿刺和静脉穿刺。 <a href="https://www.jove.com/files/ftp_upload/65513/65513fig01large.jpg" target="_blank">请点击此处查看此图的大图。</a>

大鼠采血技术的比较研究

<ol>
	<li>Lee, G., Goosens, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sampling+blood+from+the+lateral+tail+vein+of+the+rat.">Sampling blood from the lateral tail vein of the rat.</a> Journal of Visualized Experiments. (99), e52766 (2015).</li><li>Beeton, C., Garcia, A., Chandy, K. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drawing+blood+from+rats+through+the+saphenous+vein+and+by+cardiac+puncture.">Drawing blood from rats through the saphenous vein and by cardiac puncture.</a> Journal of Visualized Experiments. (7), 266 (2007).</li><li>Luzzi, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Collecting+blood+from+rodents:+A+discussion+by+the+Laboratory+Animal+Refinement+and+Enrichment+Forum.">Collecting blood from rodents: A discussion by the Laboratory Animal Refinement and Enrichment Forum.</a> Animal Technology and Welfare. 4 (2), 99-102 (2005).</li><li>Wang, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Repetitive+blood+sampling+from+the+subclavian+vein+of+conscious+rat.">Repetitive blood sampling from the subclavian vein of conscious rat.</a> Journal of Visualized Experiments. (180), e63439 (2022).</li><li>Van Herck, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blood+sampling+from+the+retro-orbital+plexus,+the+saphenous+vein+and+the+tail+vein+in+rats:+Comparative+effects+on+selected+behavioural+and+blood+variables.">Blood sampling from the retro-orbital plexus, the saphenous vein and the tail vein in rats: Comparative effects on selected behavioural and blood variables.</a> Laboratory Animals. 35 (2), 131-139 (2001).</li><li>Harikrishnan, V. S., Hansen, A. K., Abelson, K. S., S&#248;rensen, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparison+of+various+methods+of+blood+sampling+in+mice+and+rats:+Effects+on+animal+welfare.">A comparison of various methods of blood sampling in mice and rats: Effects on animal welfare.</a> Laboratory Animals. 52 (3), 253-264 (2018).</li><li>Zeller, W., Weber, H., Panoussis, B., B&#252;rge, T., Bergmann, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Refinement+of+blood+sampling+from+the+sublingual+vein+of+rats.">Refinement of blood sampling from the sublingual vein of rats.</a> Laboratory Animals. 32 (4), 369-376 (1998).</li><li>Nightingale, C. H., Mouravieff, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reliable+and+simple+method+of+intravenous+injection+into+the+laboratory+rat.">Reliable and simple method of intravenous injection into the laboratory rat.</a> Journal of Pharmaceutical Sciences. 62 (5), 860-861 (1973).</li><li>Blood collection: The rat. IACUC Guideline. UCSF Office of Research Institutional Animal Care and Use Program Available from: <a target="_blank" href="https://iacuc.ucsf.edu/sites/g/files/tkssra751/f/wysiwyg/guidelines%20-%20Blood%20Collection%20-%20Rat.pdf">https://iacuc.ucsf.edu/sites/g/files/tkssra751/f/wysiwyg/guidelines%20-%20Blood%20Collection%20-%20Rat.pdf</a> (2022)</li><li>Staszyk, C., Bohnet, W., Gasse, H., Hackbarth, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blood+vessels+of+the+rat+tail:+A+histological+re-examination+with+respect+to+blood+vessel+puncture+methods.">Blood vessels of the rat tail: A histological re-examination with respect to blood vessel puncture methods.</a> Laboratory Animals. 37 (2), 121-125 (2003).</li><li>Constantinides, C., Mean, R., Janssen, B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+isoflurane+anesthesia+on+the+cardiovascular+function+of+the+C57BL/6+mouse.">Effects of isoflurane anesthesia on the cardiovascular function of the C57BL/6 mouse.</a> ILAR Journal. 52 (3), e21-e31 (2011).</li><li>Hernaningsih, Y., Akualing, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effects+of+hemolysis+on+plasma+prothrombin+time+and+activated+partial+thromboplastin+time+tests+using+photo-optical+method.">The effects of hemolysis on plasma prothrombin time and activated partial thromboplastin time tests using photo-optical method.</a> Medicine. 96 (38), 7976 (2017).</li><li>Powles-Glover, N., Kirk, S., Jardine, L., Clubb, S., Stewart, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessment+of+haematological+and+clinical+pathology+effects+of+blood+microsampling+in+suckling+and+weaned+juvenile+rats.">Assessment of haematological and clinical pathology effects of blood microsampling in suckling and weaned juvenile rats.</a> Regulatory Toxicology and Pharmacology. 69 (3), 425-433 (2014).</li></ol>

作者均无任何利益冲突需要声明。

尾静脉穿刺是从有意识的大鼠身上获取血液的有效方法。然而，当动物处于麻醉状态时，异氟醚的作用会导致血管痉挛，并使尾静脉穿刺不合适11。如本研究所示，在这种情况下的另一种选择是从静脉收集血液，这更成功，并且在更短的时间内产生显着更大的血液量。请务必记住，如果此方法在第一次尝试时失败，则后续尝试可能会失败。相比之下，尾静脉穿刺允许在第一次尝?...

血液采样对于大多数动物研究都是必要的，无论是 体内 还是 体外。在大鼠中，由于血液采样的频率和数量可能很大，因此有不同的收集选择是有帮助的。在以前的研究中已经描述了各种方法。
最常用的技术是尾静脉穿刺和大隐静脉抽血。尾静脉取样适用于所有大鼠品系。通过适当的训练，该程序易于执行，并且对动物造成的痛苦最小1.同样，大隐静脉抽血，只要操作得当，也是一种快速简单的采集方法。这两种方法都不需要麻醉，并且都允许重复抽取少量血液。然而，大隐静脉穿刺通常产生较低的血容量1 ，并且需要两个人在场，留下一个后肢暴露进行穿刺2。
如果需要从单个动物收集大量血液，可以使用心脏穿刺或腔静脉穿刺（可以从心脏穿刺的 150 g 大鼠中抽取多达 10 mL 的血液2）。这些技术需要麻醉，并且是终末程序。在这两种技术中的任何一种之后，动物都必须被安乐死2.颈静脉棒是一种替代方案，如果在尚未达到终点的研究中需要收集大量血液，可以使用。然而，这种技术还需要大量的技术技能来避免对动物造成伤害;因此，应限制其使用3。
其他技术，如锁骨下静脉抽血，在采血前不需要使用麻醉剂，并允许对少量血液进行重复采样。然而，这种技术需要克制的处理和适当的针切口。操作不当可能导致动物疼痛甚至死亡，这种方法的培训可能很挑剔4. 
其他轶事程序包括眼眶穿刺和舌下静脉穿刺，两者都需要麻醉剂，既不推荐也不广泛使用。尽管先前的研究表明，通过眼眶穿刺比尾静脉穿刺更快地采血，但发现在乙醚麻醉下进行眼眶穿刺的耐受性不如后一种方法（基于动物的兴奋评分和尿液产生）5。此外，这种方法在很大程度上受到执行该程序的人的技能的影响，并且主要由经验丰富的兽医执行。相比之下，舌下静脉穿刺的痛苦较小，建议重复采血6。然而，这种技术存在严重的不良反应，例如食物和水摄入量减少，这可能导致动物死亡7。 
本研究描述了我们实验室用于重复血液采样的两种方法。尾静脉穿刺可以对有意识的动物进行，组织损伤和不良反应很小。在这项研究中，这项技术的修改包括用食指和中指稳定尾巴，这允许单个操作员进行采血。背静脉穿刺已被描述为简单的静脉注射。该技术在麻醉下进行，并在其他方法遇到困难的情况下（例如，在术后立即，与小动物一起，在麻醉下进行围手术期抽血时）允许可靠的血源。与尾静脉取样类似，与上述技术相比，穿刺部位的损伤对动物的整体影响较小8.该方法论文的目的是根据上下文为没有经验的研究人员提供简单可靠的血液采样替代方案（例如，对于在麻醉下进行的手术，对于包括雄性大鼠的研究，对于具有细尾静脉的动物）。

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.5 mL | 28 G &#189;&#160; Insulin Syringes</td>
			<td>BD</td>
			<td>329424</td>
			<td>for tail vein puncture</td>
		</tr>
		<tr>
			<td>0.5 mL | 30 G x 5/16 Insulin Syringes</td>
			<td>BD</td>
			<td>320468</td>
			<td>for penile vein puncture</td>
		</tr>
		<tr>
			<td>250 L&#160; Microtainer blood collection tubes with K2EDTA</td>
			<td>BD</td>
			<td>365974</td>
			<td></td>
		</tr>
		<tr>
			<td>Gauze Sponges</td>
			<td>Curity</td>
			<td>6939</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane Auto-Flow Anesthesia Machine</td>
			<td>E-Z Systems</td>
			<td>EZ-190F</td>
			<td>for penile vein puncture</td>
		</tr>
		<tr>
			<td>Isoflurane, USP</td>
			<td>Patterson Veterinary</td>
			<td>1403-704-06</td>
			<td>for penile vein puncture</td>
		</tr>
		<tr>
			<td>Nosecone for Anesthesia</td>
			<td>World Precision Instruments</td>
			<td>EZ-112</td>
			<td>for penile vein puncture</td>
		</tr>
		<tr>
			<td>Rodent Restraint Cone</td>
			<td>Harvard Apparatus</td>
			<td>ST2 52-95-86</td>
			<td>for tail vein puncture</td>
		</tr>
		<tr>
			<td>Small Animal Heated Operating Table (Adjustable)</td>
			<td>Peco Services Ltd</td>
			<td>69023</td>
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modified tail vein and penile vein puncture for blood sampling in the rat model

大多数实验动物设计都需要血液样本来评估各种血液学参数。本文介绍了两种大鼠采血程序：侧尾静脉穿刺和背静脉穿刺，与其他先前描述的技术相比具有显着优势。该研究表明，这两种程序允许快速采样（10 分钟内）并为大多数测定（202 μL ± 67.7 μL）产生足够的血容量。背静脉穿刺必须在麻醉下进行，而侧尾静脉穿刺可以在有意识、约束的动物身上进行。
因此，交替使用这两种技术可以在任何情况下抽血。虽然始终建议在手术过程中协助操作员以确保动物福利，但这些技术只需要一名操作员，这与大多数需要两名血液采样方法不同。此外，虽然这些先前描述的方法（例如颈静脉棒，锁骨下静脉抽血）需要广泛的事先训练以避免动物的伤害或死亡，但尾静脉和背静脉穿刺很少是致命的。由于所有这些原因，并根据具体情况（例如，对于包括雄性大鼠的研究，在围手术期或术后立即，对于尾静脉细的动物），这两种技术可以交替使用以实现重复抽血。

Blood sampling is necessary for most animal studies, both in vivo and in vitro. In rats, as the frequency and amount of blood sampling can be significant, it is helpful to have different alternatives for collection. Various methods have been described in previous studies.
The most commonly used techniques are tail vein puncture and saphenous vein blood draw. Tail vein sampling is suitable for all rat strains. With proper training, the procedure is simple to perform and causes minimal distress to the animal1. Similarly, the saphenous vein blood draw, provided it is done properly, is also a quick and simple collection method. Neither method requires anesthesia, and both allow for repeated draws of small amounts of blood. However, the saphenous vein puncture usually yields a lower blood volume1 and requires the presence of two people to leave one hindlimb exposed for puncture2.
If large amounts of blood need to be collected from a single animal, cardiac puncture or puncture of the vena cava may be used (up to 10 mL of blood can be drawn from a 150 g rat with cardiac puncture2). These techniques require anesthesia and are terminal procedures. The animal must be euthanized after any of these two techniques2. The jugular stick is an alternative that can be used if large amounts of blood need to be collected in a study that has not yet reached its endpoint. However, this technique also requires significant technical skills to avoid harm to the animal; hence, its use should be limited3.
Other techniques, such as the subclavian vein blood draw, do not need the use of anesthetics before blood collection and allow for repeated sampling of small volumes of blood. However, restrained handling and appropriate needle incision are required for this technique. An improper operation may result in animal pain or even mortality, and the training for this method may be fastidious4. 
Other anecdotal procedures include the orbital puncture and the sublingual vein puncture, both of which require an anesthetic, and neither are recommended nor widely used. Though previous studies have shown faster blood collection by orbital puncture than by tail vein puncture, it was found that orbital puncture under diethyl-ether anesthesia was less well tolerated than the latter method (based on the animals' excitation scores and urine production)5. Moreover, this method is highly influenced by the skill of the person who performs the procedure and is mainly performed by experienced veterinarians. Comparably, the sublingual vein puncture is less distressing and is recommended for repeated blood sampling6. However, this technique presents severe adverse effects such as reduced food and water intake, which can lead to the animal's death7. 
This study describes two methods used in our laboratory for repeated blood sampling. Tail vein puncture can be performed on a conscious animal, and the tissue damage and adverse effects are minimal. The modification of this technique in this study includes stabilizing the tail with the index and middle finger, which allows a single operator to perform the blood collection. The dorsal penile vein puncture has already been described for simple intravenous injections. This technique is performed under anesthesia and allows for a reliable blood source in case of difficulties with other methods (e.g., during the immediate postoperative period, with a small animal, when performing perioperative blood draw under anesthesia). Similar to tail vein sampling, the injury at the puncture site will have a minor overall effect on the animal compared to the techniques mentioned above8. The aim of this methods paper is to offer inexperienced researchers simple and reliable blood sampling alternatives according to the context (e.g., for procedures done under anesthesia, for studies including male rats, for animals with thin tail veins).

作者聚焦：大鼠的上下文依赖性采血方法 

改良尾静脉和静脉穿刺用于大鼠模型中的血液采样

The issue of repeated blood collection methods needs to be addressed since the majority of studies on rats require them. It's important to take into account that a sampling method can be very efficient in a certain context, but become unsuitable for another study. This protocol aims to offer simple and reliable blood sampling alternatives to researchers.Both methods can be done by a single operator and require no extensive prior training, which allows for standardized procedure. When an animal is under anesthesia, the effect of isoflurane can lead to vessel spasm and make the tail vein puncture unsuitable for blood collection. Our protocol offers an alternative in this situation, the penile vein puncture, which presents a better success rate and yields significantly more amounts of blood in less time.

成功定义为抽血在 10 分钟内（从穿刺时间到采血结束）产生至少 100 μL 的血液，失败定义为抽血产生少于 100 μL 的血液或需要超过 10 分钟才能取回所需的血容量。每个样品最多允许 250 μL 血液。统计分析采用单因素方差分析检验进行多重比较和卡方检验。数据以平均值±标准差表示， p < 0.05 作为确定统计显著性的临界值。
比较成功率显示，清醒大鼠尾静脉穿刺（92%）...

Watch this Scientific Journal Video about Context Dependent Blood Collection Methods in Rats at JoVE.com

在这里，我们提出了一种协议，为大鼠模型提供快速，简单和可靠的血液采集替代方案。我们根据上下文描述了三种不同的血液采样方法：麻醉下或有意识的动物的尾静脉穿刺，以及麻醉下的背静脉穿刺。

Blood samples are required in most experimental animal designs to assess various hematological parameters. This paper presents two procedures for blood ...

重复采血方法的问题需要解决，因为大多数关于大鼠的研究都需要它们。重要的是要考虑到抽样方法在特定情况下可能非常有效，但不适合另一项研究。该协议旨在为研究人员提供简单可靠的血液采样替代方案。这两种方法都可以由一名操作员完成，并且不需要大量的事先培训，从而实现标准化程序。当动物处于麻醉状态时，异氟醚的作用会导致血管痉挛，使尾静脉穿刺不适合采血。在这种情况下，我们的方案提供了一种替代方案，即静脉穿刺，它提供了更好的成功率，并在更短的时间内产生更多的血液。

modified-tail-vein-penile-vein-puncture-for-blood-sampling-rat

Research

JoVE Journal

Medicine

Modified Tail Vein and Penile Vein Puncture for Blood Sampling in the Rat Model

2.6K 次观看.  Massachusetts General Hospital. 重复采血方法的问题需要解决，因为大多数关于大鼠的研究都需要它们。重要的是要考虑到抽样方法在特定情况下可能非常有效，但不适合另一项研究。该协议旨在为研究人员提供简单可靠的血液采样替代方案。这两种方法都可以由一名操作员完成，并且不需要大量的事先培训，从而实现标准化程序。当动物处于麻醉状态时，异氟醚的作用会导致血管痉挛，使尾静脉穿?...

视频: 作者聚焦：大鼠的上下文依赖性采血方法 

Blood samples are required in most experimental animal designs to assess various hematological parameters. This paper presents two procedures for blood collection in rats: the lateral tail vein puncture and the dorsal penile vein puncture, which offer significant advantages over other previously described techniques. This study shows that these two procedures allow for fast sampling (under 10 min) and yield sufficient blood volumes for most assays (202 &#956;L &#177; 67.7 &#956;L). The dorsal penile vein puncture must be done under anesthesia, whereas the lateral tail vein puncture can be done on a conscious, restrained animal.
Alternating these two techniques, therefore, enables blood draw in any situation. While it is always recommended for an operator to be assisted during a procedure to ensure animal welfare, these techniques require only a single operator, unlike most blood sampling methods that require two. Moreover, whereas these previously described methods (e.g., jugular stick, subclavian vein blood draw) require extensive prior training to avoid harm to or death of the animal, tail vein and dorsal penile vein puncture are rarely fatal. For all these reasons, and according to the context (e.g., for studies including male rats, during the perioperative or immediate postoperative period, for animals with thin tail veins), both techniques can be used alternately to enable repeated blood draws.

Here, we present a protocol to offer rapid, easy, and reliable blood collection alternatives for the rat model. We describe three different blood sampling methods according to the context: tail vein puncture under anesthesia or on a conscious animal, and dorsal penile vein puncture under anesthesia.

科研

医学

Tail Vein Puncture Without Anesthesia

To begin, secure the rat in a plastic restraining cone, enclose the large end of the cone around the base of the tail. Ensure that the animal is comfortable and breathing is unrestricted throughout the procedure. Dip the tail in warm water for about one minute to dilate the vein, and dry the tail with a paper towel.In the restrainer, position the animal facing down with the tail lying on a heating pad. To select the appropriate tail vein for sampling, rotate the entire animal to the right or left side. Use the terminal third of the tail for blood vessel puncturing since the vessels become more superficial in this zone.Wipe the tail with 70%ethanol wipes at the puncture site and place it on the edge of the heating pad, creating an angle in the terminal third of the tail. This brings the vein to the surface and creates more space for taking the sample. For blood sampling, move the plunger back and forth in the syringe several times to smoothen the withdrawal and then pull on the plunger to create negative pressure in the syringe.Secure the tail flat on the heating pad using the non-dominant index and middle finger. Place the middle finger proximally and the index finger distally with the puncture site between these two fingers. Slide the insulin syringe against the index finger with the eye of the needle pointing upward until it is inserted into the vein.This creates an angle of 35 degrees between the needle and the tail. Once the needle has entered the vein, blood will flow into the syringe. Slowly withdraw the plunger of the syringe at a steady rate to collect the desired volume.Remove the syringe from the tail and allow a blood drop to form on the puncture side of the tail. Aspirate this blood to collect a few more microliters of blood, in case of a non-sterile procedure. Apply pressure to the puncture site to stop the bleeding and wipe the area with a new alcohol wipe.After removing the rat from the cone, return it to its cage. The success rates with the tail vein puncture methods were compared with the success rate of the penile vein puncture method under anesthesia. Interestingly, under anesthesia, the tail vein became very unreliable and had a success rate of only 25%The success rates showed almost similar results for the tail vein puncture in conscious rats and the penile vein puncture under anesthesia.The blood sampling was performed in less than six minutes in the tail vein puncture without anesthesia, and the penile vein puncture method under anesthesia, whereas the tail vein puncture method under anesthesia took more than eight minutes due to multiple failures.

无麻醉的尾静脉穿刺

Drawing Blood from the Penile Vein

Move the plunger back and forth in the syringe several times to smoothen the withdrawal and then pull on the plunger to create negative pressure in the syringe. Retract the foreskin of the anesthetized rat from the end of the penis and hold the glands between the index and thumb pulling gently. The dorsal penile vein will appear as a superficial blue cord.With the eye of the needle pointing upward, insert the insulin syringe into the vein at an angle of 35 degrees. Once the needle has entered the vein, blood will flow into the syringe. Slowly withdraw the plunger of the syringe at a slow and steady rate to collect the desired volume.Remove the syringe. A blood drop will form on the puncture site. Aspirate the drop to collect a few more microliters of blood in case of a non-sterile procedure.Apply light pressure to the puncture site to stop the bleeding and wipe the area with a new alcohol wipe. Place the penis back in its neutral position. After turning off the isoflurane, monitor the rat until complete recovery and return it to its cage.The blood volume collected in the dorsal penile vein puncture method under anesthesia was compared with the blood collected in the tail vein puncture method with and without anesthesia. The result shows that the penile vein puncture method under anesthesia and the tail vein puncture method without anesthesia yielded comparable quantities of blood.