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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!