- Resources and references
- Tail vessel sampling in other animals
- All blood sampling techniques in the rat
Tail vein sampling is suitable for all strains and for competent individuals, is quick and simple to perform. However, this technique requires the rats to be warmed in order to dilate the blood vessel prior to taking the sample. This can be stressful and can cause dehydration due to salivation, in addition to increasing metabolic rate, which may affect the experimental data. As a result, other routes such as saphenous vein sampling should be considered where possible. View the tail vein sampling technique below.
The lateral tail vein is usually used and 0.1 - 2 ml of blood can be obtained per sample depending on the size of the rat, sampling frequency and scientific justification. The tail may need to be washed with diluted Hibiscrub (1%) in order to see the blood vessel. Finger pressure 5 cm from the tail tip can enhance visibility of the tail vessels.
To avoid bruising and damage to the tail, normally no more than eight blood samples should be taken per session and in any one 24-hour period. Where it is necessary and justifiable to take more, the use of temporary cannulation or surgical cannulation should be considered. The number of attempts to take a blood sample should be minimised (no more than three needle sticks in any one attempt) and sufficient time should be given for the tail to recover between blood sampling sessions. Alternate sides of the tail should be used and needle punctures moved towards the tail base.
If it is necessary to warm the animal, a warming cabinet should be used (39oC for 10 to 15 minutes). Rats should be carefully monitored, including checking for signs of hyperthemia and dehydration. The time the rat is in the warming cabinet should be recorded and the cabinet should be calibrated regularly to avoid hyperthermia; digital displays should not be relied upon. It is important to ensure the temperature in the cabinet is uniform and that there are no 'hot spots'. Alternatively, a warm bath at a maximum of 40 o C can be used to warm just the tail of the rat. The temperature of the bath should be monitored to prevent the tail being scalded.
The lateral tail vein is usually accessed approximately one-third along the length of the tail from the tail tip, moving towards the base of the tail for multiple samples. Blood samples should only be taken from the base of the tail if no vein is visible elsewhere. Taking the first sample/s from the proximal end of the tail can result in a perivascular clot and inflammation that significantly reduces blood flow to the distal portion of the vessel.
An aseptic technique should be used. It is recommended that a local anaesthetic cream (e.g. EMLA cream) be applied to the site 30 minutes prior to blood sampling.
Rats need to be restrained, which can cause stress and therefore the duration of restraint should be minimised. Restraint can either be manual (e.g. wrapping the rat in a towel) or using a restraint tube. Anecdotal evidence suggests that holding the rat is less stressful than using a restraint tube. Where a restraint tube is used, it should be appropriate to the size of the rat in order to avoid damage to the tail, testes, limbs and back. All forms of restraining equipment should be frequently washed to prevent pheromonally-induced stress or cross-infection.
Blood flow should be stopped by applying finger pressure to the soft tissue. A finger should be placed at the blood sampling site for approximately 30 seconds before the animal is returned to its cage.
|Number of samples||No more than eight blood samples should be taken per session and in any 24-hour period.|
|Sample volume||0.1 - 2 ml|
|Equipment||21G - 23G needle or butterfly needle or lance|
|Staff resource||One person is required to take the blood sample if a tube restrainer is used. Two people are required if the rat is held for sampling.|
|Other||Rats may be warmed to dilate the blood vessel. Care should be taken to avoid hyperthermia and dehydration.|
- A good practice guide to the administration of substances and removal of bloog, including routes and volumes.
- Fluttert M, Dalm S, Oitzl MS (2000), A refined method for sequential blood sampling by tail incision in rats. Laboratory Animals. 34(4), 372-378
- Brown C (2006), Blood collection from the tail of a rat. Lab Animal Europe. 6(8), 35-36
- van Herck H et al. (2001), Blood sampling from the retro-orbital plexus, the saphenous vein and the tail vein in rats: comparative effects on selected behavioural and blood variables . Laboratory Animals. 35(2), 131-139
- Staszyk C, Bohnet W, Gasse H, Hackbarth H (2003), Blood vessels of the rat tail: a histological re-examination with respect to blood vessel puncture methods. Laboratory Animals. 37(2), 121-125
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- Mahl A, Heining P, Ulrich P, Jakubowski J, Bobadilla M, Zeller W, Bergmann R, Singer T, Meister L (2000), Comparison of clinical pathology parameters with two different blood sampling techniques in rats: retrobulbar plexus versus sublingual vein. Laboratory Animals. 34(4), 351-361
- Verbaeys A , Ringoir S, van Maele G, Lameire N (1995), Influence of feeding, blood sampling method and type of anaesthesia on renal function parameters in the normal laboratory rat. Urological Research. 22(6), 377-382
- Timmerman A (1992), Puncture of the tail vein as a possible alternative for orbital puncture in the rat. Animal Technology. 43(3), 167-172
- Removal of blood from laboratory animals and birds.
- Saphenous vein puncture for blood sampling of the mouse, rat, hamster, gerbil, guinea pig, ferret and mink.
- Liu JY, Diaz TG 3rd, Vadgama JV, Henry JP (1996), Tail sectioning: a rapid and simple method for repeated blood sampling of the rat for corticosterone determination. Laboratory Animal Science. 46(2), 243-245
- Toft MF, Petersen MH, Dragstead N, Hansen AK (2005), The impact of different blood sampling methods on laboratory rats under different types of anaesthesia. Laboratory Animals. 40, 261-274