来源: 凯斯图尔特, RVT, RLATG, CMAR;瓦莱丽. 施罗德, RVT, RLATG。圣母大学
血液收集是一个共同的要求, 研究研究, 涉及老鼠和老鼠。小鼠和大鼠的血液提取方法取决于所需的血液体积、取样的频率、被放血的动物的健康状况以及技术员的技术水平。1所讨论的所有方法–向后眶窦出血、初始尾剪出血和心内出血–需要使用全身麻醉。
在出血过程之前, 必须确定所需样品的类型。实验程序可能需要全血、血浆或血清。对于全血, 必须在样品中加入抗凝剂。血浆, 其中含有纤维蛋白原和其他凝血因子, 当与红细胞分离, 可以从一个抗凝样本提取。血清是通过血液采集而获得的, 没有抗凝血。一旦血块形成, 血清就会从样品的离心中产生。由于样品已凝结, 血清不会含有纤维蛋白原或其他凝血因子。血浆和血清都是通过使用离心机运行在 2200-2500 RPM 至少15分钟。
对于必须产生全血或血浆的样品, 必须使用适当的抗凝剂。实验室动物常用的抗凝剂是肝素、柠檬酸钠和乙二胺乙酸酸 (EDTA);选择的依据是研究的需要。封存–EDTA、肝素和枸橼酸钠的液体形式–可以直接装入注射器以涂上表面。这就可以直接接触抗凝血的血液被绘制, 协助预防凝血。由于鼠血凝块比大多数哺乳动物的血液凝结得快, 因此必须将抗凝血的正确比例用于血液采集。
针的选择依据的是动物的大小和穿刺部位。一般而言, 针头的孔径越大, 样品收集起来就越迅速。对更大的针头来说, 减少对血液细胞的伤害是另一个好处。然而, 大口径针头的主要缺点是对容器的潜在损害。对小鼠和大鼠, 大小的选择范围从20-29 口径针, 是 0.5-1.5 英寸的长度。如果针头太长, 不仅使用起来很笨拙, 而且在针头上有多余的空间会导致凝固。”过程” 部分中的每个方法都列出了适当的针大小。
所需样本的大小也必须预先确定。由于小老鼠或老鼠的大小, 血液收集的最大数量必须计算为生存出血。每只体重25克的小鼠总血容量为1.8 毫升;平均体重250克的鼠总血容量为16毫升。对于没有液体置换的小鼠或大鼠的单个血液样本, 可以安全去除的最大血容量是总血量的 10%, 或 7.7-8 µl/克。 因而为一个平均老鼠, 10% 它的血液容量是193-200 µl。对于250克的平均鼠来说, 这相当于 1.9-2.0 毫升. 研究表明, 去除超过15% 的血容量会引起休克休克。12然而, 随着体液置换, 多达15% 的总血容量-或12µl/克-可以删除。对于25克鼠标, 这相当于300µl;对于一个250克大鼠, 它相当于3毫升。对于流体置换, 液体应加热, 并给予皮下注射。
如果有必要采取多样本, 血液体积绘制减少。每周可抽取的最大血量不超过总血量的 7.5%, 或6µl/克。 对于一个25克鼠标, 这相当于每星期145-150 µl。对于一个250克大鼠, 这相当于 1.45-1. 50 毫升每星期。 如果取样将每2周发生一次, 则可抽取10% 的总血容量 (8 µl/克)。这相当于200µl 每2周的平均鼠标, 和2.00 毫升每2周为250克鼠。一项对平均重量为250克的大鼠进行的一项研究显示, 当血液的15-20% 被移除时, 血液水平的正常化需要超过29天的时间。12对于重复的血液收集, 液体替换不允许更大的血液容量或更频繁的血液汇集, 因为它只替换容量。这只动物需要时间补充血液细胞。
使用回溯式眶丛是过去常见的做法。然而, 对这一程序的仁爱的许多关注已经出现。在手术过程中, 红细胞压积管的过度运动一旦放置在眼睛的内侧眼角, 就会对周围组织造成损伤, 导致眼睑和/或结膜膜肿胀。肿胀的组织可导致眼球凸出足够远, 使眼睑闭合受阻, 可能导致角膜干燥和损伤。肿胀的疼痛会引起抓挠和自残, 导致眼球摘除。不正确放置红细胞压积管在一个复古眶出血可以切断视神经, 导致失明。如果红细胞压积管是在一个不适当的角度推进, 眼睛可以被迫离开轨道, 让眼睑落在眼球后面。如果出现这种情况, 则很难将眼睛正确地替换到插槽中。其他可能出现的问题包括: 对脆弱的轨道骨骼的破裂, 眼球球体的穿透, 导致玻璃体液的丧失, 或者眼睛后面的血肿形成, 由于眼睛和周围的压力, 会导致极度疼痛。结构.尽管所有这些问题, 如果一个熟练的技术员执行的程序和动物完全麻醉与一般麻醉, 如异氟醚吸入麻醉, 复古眼眶出血已证明是一种有效的方法的血液收集啮齿目动物。
在小鼠和老鼠之间, 眶区的解剖结构是不同的。老鼠有一个复古的眼窝窦-一个收集的血管, 创造了一个鼻窦在轨道区域。在鼠眼的眼眶里, 有一丛血管在那眼后面流动;然而, 它们并不像老鼠那样形成一个窦。因此, 对小鼠进行这一过程比较容易。对于重复采样收集通过回溯轨道丛, 至少10天之间的出血是需要的, 使该地区的组织愈合。虽然一般麻醉建议, 该程序可以执行的小鼠没有全身麻醉, 如果一个局部眼科麻醉, 如丙或丁卡因, 是在程序之前应用。由于大鼠没有逆行眶窦, 而且由于其周围的薄膜的轨道更强, 因此必须麻醉它们。
使用尾夹法可以获得小体积的串口样品。尾巴的最初截肢必须限制在尾巴尖端, 大约 0.5-1.0 毫米长的小鼠和2.0 毫米的大鼠。1通过将原始切口的痂或血块阻断到尾部的尾部, 可以进行血液采集的尾剪程序。一般情况下, 附加的尾端截肢是不必要的。收集的血量范围从20-100 µL 的小鼠和75-150 µL 的大鼠。收集到的数量在动物之间是可变的, 并且可以受年龄、健康状况和体重的影响。
从尾部切片收集的样本可以同时包含动脉和静脉血, 以及组织产品的污染。如果尾巴被抚摸或 “挤奶” 以获得更多的血液, 样品质量就会降低。为了增加血液流量, 可以用热敷、加热灯或在温水中浸泡尾部来加热。应将压力应用于尾部止血, 每5-10 分钟检查一次动物, 以确保止血效果。止血常因反复取样而延迟。止血粉可用于止血。对于最初的截肢, 建议麻醉 (一般或局部)。随后的出血不应要求麻醉, 特别是当动物习惯于程序。麻醉会导致血压下降, 使血液收集与此技术困难。
尾巴剪的另一种选择是尾巴船尼克。这一过程很容易在老鼠和小鼠身上进行。然而, 与尾巴剪, 样品可能被污染与组织产品, 特别是在老鼠。对于大鼠, 皮下注射针头插入血管, 血液从针的中心收集。一项研究显示, 在穿刺部位上方放置止血带, 以帮助血液收集。3注射器不用于抽取血管中的血液, 因为注射器产生的压力会使血管坍塌。这种方法也可以用于连续取样, 因为血块可以被移除, 从而导致站点再次出血。与尾部剪, 这是当务之急, 以确保止血的应用压力, 对现场和复核的动物每5-10 分钟。
通常, 研究需要一个 nonsurvival, 大的血液样本, 通过失血通过心腔出血或尾静脉静脉收集。4心脏穿刺可以从老鼠或大鼠身上采集到总血量的大约一半。这相当于40µl/克或大约1毫升的平均25克鼠标。250克老鼠会产生大约10毫升的血液。动物必须被麻醉为失血。吸入麻醉或 CO2麻醉可由熟练技师使用;注射麻醉也可以使用。然而, 血压和血液循环可能减少, 这可能会减少血液的采集量。
尾腔静脉法要求对动物进行深度麻醉, 以手术暴露血管。CO2麻醉是不够的, 因为心脏必须跳动和动物呼吸在血液撤退。在手术过程中, 抽血速度过快会导致血管塌陷, 咬合, 防止血液收集。此外, 船只的墙壁是薄的, 因此, 必须避免手和针的运动, 以防止从针进入现场的血液破裂或泄漏。由于针不是通过皮肤, 这种方法导致收集一个无菌样品。必须采用辅助安乐死的方法, 以确保动物不会从麻醉中恢复。这种方法通常是心脏或主动脉灌注后。
心腔方法可在被麻醉 (闭合方法) 的动物中进行人工约束, 或者根据骶管静脉采血方法 (open 方法) 的规定, 对心脏进行手术暴露。对于封闭的方法, 针放置的标志是在剑过程中的肋骨笼形成的凹槽, 在动物的左侧。
Blood collection for mice and rats can be accomplished with a variety of techniques. Although many factors, such as sample size, frequency of sampling, and the size and age of the animal influence this, the most essential component is the skill level of the technician performing the sample collection. For the methods described here, the proper use of anesthetics is also crucial for quality samples and the wellbeing of the animals.
Blood collection is a common requirement for several research studies that involve mice and rats. The choice of method for blood withdrawal in these animals is dependent upon many factors like, the volume of blood needed, frequency of the sampling, health status of the animal to be bled, and the skill level of the technician.
Here, we will review these considerations and outline blood collection procedures including the retro-orbital eye bleed, tail snips and nicks, as well as intra-cardiac blood collection. For other methods, see the second video in this series.
Before delving into the blood withdrawal protocols, let’s first review some general considerations including sample type, needle selection, and the maximum blood volume that can be collected. Prior to collecting blood from a mouse or a rat, the type of blood sample required must be determined. Experimental procedures could require whole blood, plasma, or serum.
If collecting whole blood, an anticoagulant must be added to the sample to prevent clotting. Commonly used anticoagulants include heparin, sodium citrate, and ethylenediamine tetraacetic acid, abbreviated as EDTA. Anticoagulants can be loaded directly into the syringe to coat the surfaces. This allows contact of the anticoagulant directly as the blood is drawn aiding in the prevention of clotting. Because rodent blood clots rapidly, it is essential that the correct ratio of anticoagulant to blood be used. Plasma collection requires centrifuging the whole blood WITH anticoagulant. Following the spin, the translucent liquid above the WBC and platelet layer is plasma. It contains fibrinogen and other clotting factors. On the other hand, serum is collected from whole blood sample WITHOUT anticoagulants. And because the sample has clotted, the serum, which is the top player, does not contain fibrinogen or other clotting factors.
Needle selection is based on the size of the animal and the site of the venipuncture. In general, large bore needles cause less damage to blood cells and enable more rapid blood collection; but are more likely to cause vessel damage. Needle length should also be considered. If a needle is too long, it could be awkward to use, or blood could begin to clot while still inside the needle. The choices of size ranges from 18 to 29 gauge and 0.5 to 1.5 inches in length. The appropriate needle size for each method will be discussed in the procedures section.
Lastly, because of the small size of rodents, there is a maximum amount of blood that can be collected from a single blood draw, which will not cause serious harm to the organism. Blood withdrawal could be without or with fluid replacement – usually done using 0.9% physiological saline. The upper limit in each case is listed in the text protocol below. Furthermore, some experiments require multiple sample collection and in such cases along with fluid replacement animal will need time in between to replenish blood cells as well. Again, there is a maximum amount that can be collected during serial collection, and the upper limits are listed in the protocol below.
After reviewing some general considerations, let’s jump into the specific blood withdrawal techniques, starting with retro-orbital bleeding – a technique used by scientists to collect small volumes from the vessels near the eye. Note that the anatomical structure of the orbital area is different between the mouse and rat. The rats have a plexus of vessels that flow behind the eye, whereas the mouse has a collection of vessels that create a retro orbital sinus, which makes it is easier to perform this procedure in mice.
Begin by grabbing a tube for blood collection. Micro hematocrit tubes that hold 50-75 microliters are preferred. Lay down several paper towels or other insulating materials on the work surface. This is to maintain the animal’s body heat during the procedure. Now anesthetize the animal using an inhalation anesthetic such as isoflurane. Once the animal is fully anesthetized, remove it from the chamber and place it down on its side that is in in lateral recumbency position. Next, place a finger on the top of the head and along the jaw line and pull the skin back and down to induce eye protrusion. Avoid applying pressure to the trachea as that may cause death by asphyxia. Subsequently, place the micro-hematocrit tube in the medial canthus of the eye and direct it caudally at a 30 to 45 degree angle from the plane of the nose. Apply pressure while gently rotating the tube. This will cut through the conjunctival membranes and rupture the ocular plexus or sinus. The blood will flow into the hematocrit tube by capillary action. Avoid pushing the tube so deep that you hit the bone at the back of the ocular cavity. Once blood begins to flow, maintain pressure to keep the eye protruded. To stop bleeding, release the skin and allow the eye to return to the normal position. Apply pressure to promote hemostasis. For repeated sample collection, allow a minimum of 10 days between the bleeds. This provides tissues some time to heal.
Although retro-orbital bleeding is a common procedure, there are many concerns about its humaneness. These include swelling due to excessive movement of the hematocrit tube. This in turn can cause the eyeball protrusion and impede closure of the eyelid resulting in corneal drying, damage, and pain, which can trigger scratching and self-mutilation. Improper placement of the hematocrit tube can sever the optic nerve resulting in blindness. Another possible complication is that the eye can be forced out of the orbit, allowing the eyelids to fall behind the eyeball. Furthermore, issues can arise from the fracturing of the fragile orbit bones, penetration of the eye globe resulting in the loss of vitreous humor, or the formation of a hematoma behind the eye that can result in extreme pain. Despite all of these concerns, if a skilled technician performs the procedure and the animal is fully anesthetized, retro-orbital bleeding is an effective method of blood collection in rodents.
Now let’s review the considerations and procedures for tail bleeding, which allows collection of a serial samples of small volumes. The equipment needed for this procedure include a sterile number 11 scalpel. Scissors should not be used because the cut made by scissors is crushing, which can promote clotting and reduce blood flow. Other instruments are a restraint tube that allows access to the animal’s tail; absorbent paper towels; collection or hematocrit tubes and styptic powder – to aid in hemostasis.
Start by securing the animal into the restraint tube. Then, wipe the tail with warm water to remove debris and to cause slight vasodilation. DO NOT use hot water.Extend the tail and with the scalpel blade snip the very end of the tail to collect the blood using hematocrit or collection tubes. The tail can be stroked or “milked” from rump to tip to encourage blood flow. This will, however, decrease the quality of the sample.
To stop bleeding, apply pressure to the tail tip with a gauze pad. The styptic powder may be used to achieve hemostasis. Check the animals every 5 to 10 minutes to ensure hemostasis has been achieved, which might be delayed after repeated sampling. The sample collected from a tail snip can contain both arterial and venous blood, along with tissue product contamination. However, this procedure for blood collection allows for serial collections by disrupting the scab or clot of the original cut at the end of the tail.
An alternative blood collection method to a tail snip is the tail vessel nick, which is relatively less invasive. For this, using the same scalpel blade, make a small cut directly over the lateral tail vein, approximately two-third the distance from the rump. As with tail snips, blood can be collected in collection or hematocrit tubes. And it is imperative to assure hemostasis by applying pressure to the site and rechecking the animal every 5-10 minutes. However, as with the tail snip, the samples may be contaminated with tissue products.
Often studies that require a non-survival large blood sample, which is accomplished through exsanguination via an intra-cardiac bleed or the caudal vena cava.
For intra-cardiac method in mice, you need a 3 cc syringe with a 22 -25 gauge 1 inch needle. And for rats, a 10-12 cc syringe with an 18 gauge 1.5 inches needle is preferred. See the protocol below to understand the why these needs and syringes are ideal.
Start by euthanizing the animal using carbon dioxide. Following euthanasia, hold the rodent by the scruff with the body hanging vertically. This restrain is critical as the body should be straight to prevent deflection of the heart or a twisting of the chest. Note that the heart is located approximately at the level of the elbow. The insertion side is in the notch just to the left of the xiphoid, parallel to the spine and under the ribs.
Insert the needle, bevel up, into the chest and puncture the heart. Apply slight backpressure with the syringe. If the needle is in the heart, blood will flow into the syringe. Wait until the blood has filled the barrel before adding additional backpressure. Approximately half of the total blood volume can be collected from a mouse or rat by cardiac puncture. This is equivalent to approximately 1 mL of blood from an average mouse and approximately 10 mL of blood from an average rat
An alternative position is dorsal recumbency when using the lateral approach. In this case, place the needle between the ribs on the animal’s left side. The point of entry is measured against the point of the elbow on the chest wall. Insert the needle, bevel up, perpendicular to the plane of the table at a point midway on the chest wall. Apply slight back pressure with the syringe. If the needle is in the heart blood will flow into the syringe. Again, wait until the blood has filled the barrel before adding additional backpressure. Note that in either position, excessive backpressure may collapse the heart occluding the needle bevel and stopping blood flow into the syringe.
Another method to collect cardiac blood is through the caudal vena cava. The equipment needed for this procedure are an appropriate syringe with a correct size needle attached; scissors for opening the abdominal cavity, small atraumatic thumb forceps and gauze sponge. This technique requires that the animal be deeply anesthetized and maintained under anesthesia throughout the procedure. CO2 narcosis is not an option, as the animal heart must be beating for this procedure. Place the animal in dorsal recumbency position, and secure the limbs to the platform. The limbs should be extended away from the body.
Now lift the skin with forceps and use scissors to make a small transverse cut through the skin just above the pelvis in females or prepuce in males. Next, place the point of the scissors into the cut and make a midline incision through the skin from the pelvis or prepuce to the xiphoid. With the skin laterally reflected, lift the muscle and make a small transverse cut through the muscle, just above the skin cut.
Place the point of the scissor into the abdomen and make a midline incision through the muscle to the xiphoid. Be sure to angle the scissor point upward to prevent cutting any organs. Cut transversely along the curve of the ribs on each side. Be careful not to puncture the liver. Gently move the intestines to the animal’s left to expose the posterior vena cava. Place a gauze pad on the liver and rest your index and middle fingers on it. With your other hand, insert the needle, bevel up into the vena cava, midway between the junction of the renal vessels and iliac bifurcation. Slowly withdraw the blood while applying pressure on the liver.
Avoid hand movement, as that might cause the vessel rupture. Also, too rapid blood withdrawal can cause the vessel to collapse onto the bevel occluding the opening and preventing blood collection. The main advantage of this technique is the ability to collect a sterile sample because the needle does not pass through the skin.
Lastly, let’s look at some applications of these blood withdrawal techniques. Immuno-oncology is an emerging field, and researchers in this area often perform blood collection to study the immune cells at different stages of cancer development. For example, here researchers collected cardiac blood from cancer-bearing mice to isolate and quantify neutrophils at ten, twenty and thirty days following tumor engraftment.
On the other hand, blood composition is also frequently studied by physiologists. Like in this study, researchers were interested in evaluating kidney function in diabetic animals. In order to do that, these scientists first injected a dye into a diabetes animal model. Next, they then used tail snip method to collect blood at several time-points to evaluate dye concentration in blood, which was ultimately used to calculate glomerular filtration rate that highlighted the difference in kidney function following diabetes induction.
Lastly, stem cells researchers use blood samples to evaluate the success of incorporation of donor cells into the recipient’s system. Here, the investigators first transplanted bone marrow cells from a male mouse into a wild type and genetically modified female animal via the tail vein injection. Next, they collected blood from the retro orbital sinus of the recipient mouse to study the genomic DNA of blood cells using polymerase chain reaction. This provided the percentage of donor cells engraftment in the two types of animals.
You’ve just watched JoVE’s first installment on blood withdrawal techniques. Please see the next video in series to review how to perform other commonly employed techniques of blood collection in lab animals. As always, thanks for watching!