Prior to the bleeding procedure, the type of sample required must be determined. Experimental procedures could require whole blood, plasma, or serum. For whole blood, an anticoagulant must be added to the sample. Plasma, which contains fibrinogen and other clotting factors when separated from the red blood cells, can be extracted from an anticoagulated sample. Serum is obtained through blood collection without an anticoagulant. The serum will result from centrifugation of the sample once a clot has formed. As the sample has clotted, the serum will not contain fibrinogen or other clotting factors. Both plasma and serum are obtained through the use of a centrifuge run at 2200-2500 RPM for a minimum of 15 minutes.

For a sample that must yield whole blood or plasma, an appropriate anticoagulant must be used. Commonly used anticoagulants for laboratory animals are heparin, sodium citrate, and ethylenediamine tetraacetic acid (EDTA); selection of which is based on research needs. Sequester-a liquid form of EDTA, heparin, and sodium citrate-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. As rat blood clots faster than most mammalian blood, it is essential that the correct ratio of anticoagulant to blood be used for blood collection.

Needle selection is based on the size of the animal and the site of the venipuncture. In general, the larger the bore of the needle, the more rapidly the sample can be collected. Less damage to the blood cells is another benefit to larger needles. However, the main disadvantage to large-bore needles is the potential damage to the vessel. On mice and rats, the choices of size range from 20-29 gauge needles that are 0.5-1.5 inches in length. If a needle is too long, not only is it awkward to use, but having the extra space in the needle could result in clotting. The appropriate needle size is listed for each method in the procedures section.

The size of the required sample must also be predetermined. Due to the small size of the mouse or rat, the maximum amount of blood collection must be calculated for a survival bleed. An average mouse weighing 25 grams has a total blood volume of 1.8 ml; the average rat weighing 250 grams has a total blood volume of 16 ml. For a single blood sample on a mouse or rat without fluid replacement, the maximum blood volume that can be safely removed is 10% of the total blood volume, or 7.7-8 µl/g. Thus for an average mouse, 10% of its blood volume is 193-200 µl. For an average rat of 250 grams, this is equivalent to 1.9-2.0 ml. Studies have shown that removing more than 15% of the blood volume can cause hypovolemic shock.^1,2 However, with fluid replacement, up to 15% of the total blood volume-or 12 µl/g-can be removed. For a 25 gram mouse, this is equivalent to 300 µl; for a 250 gram rat, it is equivalent to 3 ml. For fluid replacement, the fluids should be warmed and given subcutaneously.

If it is necessary to take multiple samples, the blood volume drawn is reduced. The maximum blood volume that may be drawn per week is no more than 7.5% of the total blood volume, or 6 µl/g. For a 25 gram mouse, this is equivalent to 145-150 µl per week. For a 250 gram rat, this is equivalent to 1.45-1.50 ml per week. If sampling will occur every 2 weeks, up to 10% of the total blood volume (8 µl/g) may be drawn. This is equivalent to 200 µl every 2 weeks for an average mouse, and up to 2.00 ml every 2 weeks for a 250 gram rat. One study, performed on rats with the average weight of 250 grams, revealed that when blood volumes of 15-20% were removed, it took more than 29 days for blood levels to normalize.^1,2 For repeated blood collection, fluid replacement does not allow for a larger blood volume or more frequent blood collection, as it only replaces volume. The animal will need time to replenish blood cells.

The use of the retro-orbital plexus has been a common practice in the past. However, many concerns about the humaneness of this procedure have arisen. During the procedure, excessive movement of the hematocrit tube once placed in the medial canthus of the eye can cause damage to the surrounding tissues, resulting in swelling of the eyelids and/or conjunctival membranes. The swollen tissues can cause the eyeball to protrude far enough so that closure of the eyelid is impeded, potentially resulting in corneal drying and damage. Pain from swelling can trigger scratching and self-mutilation that results in enucleation of the eye. Improper placement of the hematocrit tube during a retro-orbital bleed can sever the optic nerve, resulting in blindness. If the hematocrit tube is advanced at an improper angle, the eye can be forced out of the orbit, allowing the eyelids to fall behind the eyeball. If this occurs, it is very difficult to correctly replace the eye into the socket. Other issues that can arise include fracturing of the fragile orbit bones, penetration of the eye globe that results in the loss of vitreous humour, or the formation of a hematoma behind the eye that can result in extreme pain due to the pressure on the eye and surrounding structures. Despite all of these concerns, if a skilled technician performs the procedure and the animal is fully anesthetized with a general anesthetic, such as isoflurane inhalant anesthesia, retro-orbital bleeding has been shown to be an effective method of blood collection in rodents.

The anatomical structure of the orbital area is different between the mouse and rat. The mouse has the retro-orbital sinus-a collection of vessels that create a sinus in the orbital area. In the orbit of the rat eye, there is a plexus of vessels that flow behind that eye; however, they do not form a sinus, as in the mouse. Consequently, it is easier to perform this procedure on mice. For repeated sampling collection via the retro-orbital plexus, a minimum of 10 days between bleeds is required to allow the tissues in the area to heal. Although general anesthesia is recommended, the procedure can be performed in mice without general anesthesia if a topical ophthalmic anesthetic, such as proparacaine or tetracaine, is applied prior to the procedure. As rats do not have the retro-orbital sinus, and because their membranes around the orbit are much stronger, it is mandatory to anesthetize them for this procedure.

Serial samples of a small volume can be obtained by using a tail clip method. The initial amputation of the tail must be limited to a tail tip, approximately 0.5-1.0 mm in length in mice and 2.0 mm in rats.¹ The tail snip procedure for blood collection allows for serial collections by disrupting the scab or clot of the original cut at the end of the tail. Generally, additional amputation of the tail tip is not necessary. Volumes of blood collected range from 20-100 µL for mice and 75-150 µL for rats. The amount collected is variable between animals and can be influenced by age, health status, and weight.

The sample collected from a tail snip can contain both arterial and venous blood, along with tissue product contamination. The sample quality decreases if the tail is stroked or "milked" to obtain more blood. To increase blood flow, the tail can be heated with warm compresses, a heat lamp, or submersion in warm water. Pressure should be applied to the tail tip for hemostasis, and animals should be checked every 5-10 minutes to ensure hemostasis has been achieved. Hemostasis is often delayed with repeated sampling. A styptic powder may be used for hemostasis. For the initial amputation, anesthesia (general or local) is recommended. Subsequent bleeding should not require anesthesia, especially as the animals become habituated to the procedure. Anesthesia will cause a drop in blood pressure, making blood collection with this technique difficult.

An alternative to a tail snip is the tail vessel nick. This procedure is easily performed on both mice and rats. However, as with the tail snip, the samples may be contaminated with tissue products, especially in the mouse. For rats, a hypodermic needle is inserted into the vessel, and the blood is collected from the hub of the needle. One study demonstrated the use of a tourniquet placed above the needle puncture site to aid in blood collection.³ A syringe is not used to draw the blood out of the vessel, as the pressure created from the syringe will collapse the vessel. This method can also be used for serial sampling, as a clot can be removed to cause the site to bleed again. As with tail snips, it is imperative to ensure hemostasis by applying pressure to the site and rechecking the animal every 5-10 minutes.

Often, studies require a nonsurvival, large blood sample that is collected through exsanguination via an intracardiac bleed or the caudal vena cava.⁴ Approximately half of the total blood volume can be collected from a mouse or rat by cardiac puncture. This is equivalent to 40 µl/g or approximately 1 ml for an average 25 gram mouse. A 250 gram rat would yield approximately 10 ml of blood. The animal must be anesthetized for exsanguination. Inhalant anesthesia or CO₂ narcosis can be used by a proficient technician; injectable anesthesia can also be used. However, there may be a decrease in blood pressure and circulation, which could decrease the amount of blood collected.

The caudal vena cava method requires that the animal be deeply anesthetized to surgically expose the vessel. CO₂ narcosis is not sufficient, as the heart must be beating and the animal breathing during blood withdrawal. During the procedure, too rapid of blood withdrawal can cause the vessel to collapse onto the bevel of the syringe, occluding the opening and preventing blood collection. Also, the vessel walls are thin, and thus movement of the hand and needle must be avoided to prevent rupture or leaking of blood from the needle entry site. As the needle is not passing through the skin, this method results in the collection of a sterile sample. Adjunctive euthanasia methods must be employed to ensure that the animal does not recover from anesthesia. This method is often followed by cardiac or aortic perfusion.

The intracardiac method can be performed either with the animal restrained manually once it is anesthetized (closed method), or the heart can be surgically exposed as per the protocol for caudal vena cava blood collection method (open method). For the closed method, the landmarks for needle placement are the groove formed by the rib cage at the xiphoid process, on the animal's left side.

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.