Blood samples

(Blood sample volumes)

Blood samples

Puncture sample from Vena saphena

Vena saphena runs laterally across the tarsus. The point of puncture, proximally from the tarsal joint, should be shaved. A non-sedated animal should be held still by an assistant. The hind leg should be straightened by pressing with your finger at the top of the knee, and the vein is compressed with another finger proximally to the sampling site. The vein is punctured with the tip of the injection needle. The blood coming from the puncture point is collected into a capillary tube or cuvette. After sampling the puncture point should be pressed to stop the bleeding.

Puncture sample from the tip of the tail

The vein close to the tip of the tail is punctured with a 25–23 G needle. The blood sample should be collected in a capillary or cuvette.

Blood sample from the tail vein

Usually the sample is taken from the lateral tail vein. A non-sedated animal must be immobilized so that the tail can be handled, for example in a transparent plastic tube with a hole for the tail. Sampling is greatly facilitated by warming the tail or the whole animal before taking the sample, as this dilates the tail veins. However, warming of the whole animal, especially of mice, involves a risk of excessive heating or drying, which may also affect the samples.

The sample is usually taken from the apical part of the tail. If the sampling fails or more samples are needed, you can move towards the base. A needle of a suitable thickness or a butterfly cannula should be used. An assistant presses the veins with fingers at the base of the tail. The needle is inserted into the vein, as in the i.v. injection. The blood is allowed to drop from an open needle or cannula into the sample tube, or the sample can be aspirated into a syringe attached to the needle. Blood drips more easily from a vacuum needle, after removing the rubber cap at its other end. In a normal injection needle, slowly dripping blood can easily stop at the needle hub.

Tail vein cannulation

It is possible to place a normal IV cannula in the rat’s tail vein, in which case it is e.g. easy to add anaesthetic according to the animal’s response during surgery. In any case it is more secure to administer i.v. injections through a cannula than a needle; a substance accidentally injected next to the vein may cause a local reaction and even ulceration of the tail skin.

Warming the whole animal or just the tail dilates the veins and significantly facilitates the procedure. The rat must be completely immobilised or preferably sedated. The veins should be pressed at the base of the tail. The tail should be pulled straight. A sufficiently thin cannula (e.g. 23 or 24 G) shall be inserted in the vein, as in the i.v. injection. The blood rising to the cannula shows that it is in the correct place. The needle tip must be pushed a few millimetres inside the vein, in order for the cannula tip on top of the needle to enter the vein properly. Then the cannula is all the way in while holding the needle still. Pressing the veins is stopped and the needle pulled out. The correct positioning of the cannula should be confirmed by injecting a small amount of physiological saline; it is then closed it with a plug. The saline also prevents the cannula from clogging with coagulated blood. If necessary, the cannula may be taped to the tail.

Blood sample from the retro-orbital venous plexus (orbital puncture)

Taking a blood sample from the retro-orbital venous plexus (orbital puncture) is nowadays only allowed for taking a terminal sample during anaesthesia. The anaesthetised mouse lies on a surface on its stomach or side. With one hand, you should hold the head against the surface so that your thumb presses on the jugular vein right behind the lower jawbone and your index finger is above the eye, pulling the lid up so that the eye bulges out slightly. The tip of a micro-hematocrit capillary is pressed from either corner of the eye, past the eyeball, towards the bottom of the orbit, rolling it at the same time between your fingers. When the venous plexus bursts, the blood rises to the capillary. The technique is nearly the same for a rat, but the recommended point for the capillary introduction is beneath the upper lid in the middle.


Closed cardiac puncture

A cardiac puncture should only be performed in terminal anaesthesia.


The animal lies on a surface on its back. A needle of 26–23 G and 1 ml (or 2 ml) syringe should be used. The puncture may be done next to the sternum (1), through the cranial opening of the thoracic cavity (2) or from the abdominal cavity (3).

1) The needle is inserted vertically right next to the sternum on the left side, approximately midway (between the 5th and 6th rib). When injecting, aspirate lightly by pulling on the plunger. When blood enters the syringe, the needle must be held still and slow aspiration shall continue for as long as blood enters the syringe.

2) The head and neck must be pulled completely straight. The needle is inserted through the cranial opening of the thoracic cavity, under the sternum, in parallel with the sternum or in a slight downward angle, towards the base of the tail.

3) The needle is inserted right next to the xiphoid cartilage (caudal tip of the sternum) from the left side, directing cranially towards the median line, at a 10–30° downwards angle relative to the sternum.


The puncture may be performed above the heart (1), next to or through the sternum (2) or on the side of the abdominal cavity (3).

1) The rat is on its right side. Locate with your thumb the point of the strongest pulse at the left side of the chest, while supporting the rat’s back with the rest of your fingers. Insert the needle (23–21 G) tip there between the ribs, at a right angle to the chest wall, push it slowly deeper and stop when blood enters the syringe. Aspirate the blood into the syringe slowly, trying to hold the needle and the syringe completely still. If the blood flow stops too early, you can search for the right place by slowly pushing the needle deeper or pulling it back. You can also try inserting the needle in another direction, but do not move the tip sideways inside the chest, as it could incise the big vessels.

2) The rat is on its back, and with the fingers of one hand you should hold on to both sides of the thorax. The needle is inserted vertically next to (or through) the sternum approximately midway. When blood enters the syringe, continue as described above.

3) The rat is on its back. With the thumb and other fingers of one hand, hold on to both sides of the thorax and use your index finger to lift up the xiphoid cartilage. The needle is inserted right beneath the xiphoid cartilage. It should be directed horizontally under the sternum in a cranial direction, until the heartbeat can be felt in the syringe. Then the syringe should be turned upwards to an angle of approximately 30° in relation to the vertical position, the needle inserted into the heart (at most 3–4 mm), and the blood aspirated into the syringe. Alternatively, the needle may be inserted from the start at a 30° angle, holding a slight negative pressure in the syringe, and when blood enters the syringe, you must stop the needle and continue aspiration.


Open cardiac puncture

Under surgical anaesthesia, the ventral thorax is cut off with scissors so that the organs in the thoracic cavity can be seen. A 23–21 G needle attached to a syringe is inserted in the right ventricle of the heart, and aspirated slowly.

Blood sample from the caudal vena cava (Vena cava caudalis) or abdominal aorta

A blood sample from the caudal vena cava is only possible in terminal anaesthesia. The animal is laid on its back. The abdominal wall is cut open and moved aside. The intestines are taken out of the abdominal cavity and put down on the left side of the animal (right side of the operator). The caudal vena cava and the abdominal aorta are next to each other at the roof of the abdominal cavity, midline. The fat tissue and other connective tissue that may be on top of the veins is carefully prepared away if needed, in order to improve visibility. The sample is taken with a needle and a syringe: caudal vena cava sample near the kidneys, and aorta sample cranially of the point where the iliac arteries (Arteria iliaca) branch, at the back of the abdominal cavity.

Last updated: 5.1.2017