General principles

Blood sampling can be stressful for laboratory animals because of the handling, restraint, anaesthesia or discomfort associated with a particular technique. It is in the interests of good science, as well as of animal welfare, that stress should be kept to a minimum. Adherence to the following general principles will help to ensure that this is the case.

A comprehensive table of blood sample volumes is also available, which includes volume ranges for laboratory animals, domestic species and non-human primates.

Additional information can be found in the JWGR report on the removal of blood.

Licence authority, training and competency

Project and personal licence authority

  • In the UK, blood sampling is considered a scientific procedure when the sample is taken for a scientific purpose. Although removal of blood can itself have adverse consequences, it is the potential for pain from the insertion of the needle into the skin that is deemed to exceed the level above which work needs to be regulated by the ASPA.
  • A project licence should detail the site, volume and frequency of blood sampling. These should be justified in terms of science and animal welfare.
  • A personal licence is also required for the person taking the blood sample. Blood collection techniques can only be observed and not practised before applying for the personal licence. Once the personal licence is obtained, blood collection can only be carried out under supervision until the new licensee is deemed competent in the particular technique/s.

Training and competency

  • Criteria for competency should be defined for each technique. For example, competency in blood sampling from the canine cephalic vein would involve demonstrating:
    • Knowledge of the technique (e.g. suitability for purpose, appropriate needle size, permissible collection volumes, potential adverse effects and how to cope with them, how to handle the blood sample once taken)
    • The ability to restrain the dog calmly
    • The ability to suitably prepare the sampling site - including hair removal and skin preparation in a way that does not distress the dog or damage the skin
    • Accurate localisation and good dilation of the vessel
    • Insertion of the needle without causing distress to the dog
    • Removal of a non-haemolysed blood sample at an appropriate speed without causing bruising
  • The amount of training and practice required to achieve a given level of competence in a particular technique varies from individual to individual depending on, for example, manual dexterity, prior experience, attitude, and the skills of the instructor. Retraining or additional supervision is necessary if a technique is not conducted routinely.
  • Inexperienced persons should first examine dead animals (euthanised for another purpose) to learn the relevant anatomy and thereby avoid having to make repeated unsuccessful entries when trying to locate a blood vessel. Use should be made of demonstration and instruction videos, as well as inanimate objects (e.g. veterinary simulators, oranges) to gain familiarity in handling and using needles and syringes, before carrying out any work on live animals. Observing experienced personnel will also help in learning the technique.


Handling and restraint

  • Firm, empathetic handling is very important, as is the time required to withdraw the sample. Both these parameters can affect the degree of stress for the animal and consequently the quality of the sample and research data.
  • The animal should be restrained by an experienced person (preferably one known to the animal, especially for larger laboratory animal species). The correct level of restraint is that which allows a satisfactory sample to be taken at the first attempt but which does not cause the animal to become unnecessarily distressed.
  • Inanimate restrainers can be used, although these may not always be the best method for individual animals. Manual restraint facilitates recognition of distress more effectively. 
  • A vein will collapse if a sample is taken too quickly, so care should be taken to ensure that blood is withdrawn at an appropriate speed.
  • Depending on the species consideration should be given to offering a reward after each bleed.

Needle size

  • A sterile needle (or lancet) should be used to puncture the skin and underlying blood vessel.
  • The size of the needle (length and bore) is very important.
  • It is recommended to use as large a bore as possible to ensure rapid blood withdrawal without collapsing the vein, within the constraint of avoiding haematoma (i.e. the bore should be just less than the diameter of the vessel).
  • Recommendations on appropriate needle size are given for specific techniques featured within these pages.

Site and localisation of the vein

  • Choose a site that is fit for purpose and one which causes minimal stress to the animal.
  • Note that samples taken from different sites may have differences in biochemical / haematological values.
  • Time should be spent accurately locating and dilating  the vein before puncturing the vessel.

Dilation of the vein

  • In conscious rodents, blood can be more easily obtained if the animal (or part of the animal the sample is taken from, e.g. tail) is warmed first. Animals may be placed in a thermostatically controlled warming box for a brief period during which they should be kept under constant observation in order to prevent hyperthermia (indicated by breathing more rapidly, panting or salivating).
  • In anaesthetised animals, vasodilation can occur as a result of the anaesthetic. Anaesthetised animals may not, therefore, need warming so consideration should be given to bleeding when animals are anaesthetised for another purpose.
  • Anaesthesia is not normally necessary for venous access, since the associated stress would probably be greater than the discomfort of a needle prick or of a puncture with a lance.
  • Local anaesthetic creams can be beneficial in reducing discomfort in some species if applied at least 30 minutes before taking the sample.
  • Veterinary advice should be taken on the most appropriate anaesthetic. Some anaesthetics or components of anaesthetic mixtures (e.g. medetomidine or xylazine) cause vasoconstriction and so should be avoided.

Potential adverse effects

Potential adverse effects (e.g. stress, haemorrhage, bruising, thrombosis, infection at the site of needle entry, phlebitis, scarring, nerve damage) should be avoided. Advice on treatment for adverse effects should be sought from the Named Veterinary Surgeon.

  • Haemorrhage due to poor haemostasis is not a common problem, unless the animal has a clotting defect, and in some cases gentle continuous pressure applied for several minutes is all that is needed to stop the bleeding. Longer compression of the puncture site may be needed to stop bleeding following arterial sampling.
  • 'Bruising' is due to subcutaneous bleeding at the time of venpuncture or after the animal has been placed in its cage or pen, when the site might be aggravated by the animal itself through licking or rubbing. The animal should be checked after approximately 30 minutes and, if necessary, appropriate action taken (e.g. consult the Named Veterinary Surgeon).
  • Thrombosis (clotting) and phlebitis (inflammation of the vein) are usually caused by failure to employ aseptic technique or leaking of an irritant substance (e.g. alcohol-based chemicals) around the vein. Occasionally they can result from self-mutilation.

Arterial puncture

The main reason for collecting blood from arteries is that large samples can be obtained rapidly and relatively easily. Many of the principles described above for venepuncture also apply to arterial puncture.


Cannulation is an important technique for removal of blood because it reduces the stress of multiple sampling associated with, for example, repeated restraint and needle sticks.

  • Cannulation should be considered when repeated samples are required, especially over relatively short time periods.
  • In some species, it may be necessary to restrain the animal in some way to stop it removing the cannula. For example, rats are often restrained by a harness, swivel and tether system, which restricts normal movement. Animals should be acclimatised to any restraint system before cannulation.
  • Tethered animals are often housed singly, thus adding to the stress and severity of the procedure. When dealing with social animals, every effort should be made to keep them in social groups. Cannulated pigs, cats and marmosets can be group-housed successfully with appropriate bandaging and protection for the cannula.
  • Cannulation has the potential to cause discomfort to the animal and therefore warrants post-operative administration of analgesics and careful post-operative care and monitoring for the duration of time the cannula is in place.
  • Cannula associated infections can be avoided through the use of sterile equipment and solutions, and by employing an aseptic technique.

We have a number of pages dedicated to vascular catheters:

Click here for an introduction to implanting catheters in laboratory animals Click here for information on preventing thrombosis when implanting catheters in laboratory animalsClick here for information on the design of catheters for use in laboratory animalsClick here for information on preventing infection when implanting catheters in laboratory animalsClick here for information on planning and designing experiments which will involve implanting catheters into laboratory animalsClick here for a glossary of terms used in our pages on implaning catheters into laboratory animals

Cardiac puncture

Cardiac puncture should only be carried out under deep terminal anaesthesia or on euthanised animals.

It is common to exsanguinate animals under terminal anaesthesia using this method. Where the procedure is intended as terminal, death after exsanguination should be ensured via anaesthetic overdose or by incising the heart or major blood vessels.

Cardiac puncture can be used for blood sampling in the following species:

Click here for information on cardiac puncture blood sampling in the mouseClick here for information on cardiac puncture blood sampling in the rat Click here for information on cardiac puncture blood sampling in the hamster Click here for information on cardiac puncture blood sampling in the guinea pig Click here for information on cardiac puncture blood sampling in the rabbit Click here for information on cardiac puncture blood sampling in the ferret

Volume of blood to be removed

The volume of blood removed and the frequency of sampling should be based on the purpose of the scientific procedure and the total blood volume of the animal. It is essential to take account of the combined effect of sample volume and the frequency of sampling. If too much blood is withdrawn too rapidly, or too frequently without replacement, an animal can go into short-term hypovolaemic shock and/or in the longer term suffer anaemia. Data interpretation and scientific validity may be confounded if excessive sampling is employed.

  • As a general principle, sample volumes and number of samples should be kept to a minimum.
  • As a general guide, up to 10% of the total blood volume can be taken on a single occasion from a normal, healthy animal on an adequate plane of nutrition with minimal adverse effects; this volume may be repeated after three to four weeks. For repeat bleeds at shorter intervals, a maximum of 1.0% of an animal's total blood volume can be removed every 24 hours; the effects of stress, site chosen and anaesthetic used, should be taken into account.
  • If frequent samples are necessary, the use of cannulation as a less stressful alternative to repeated venepuncture should be considered.
  • See the decision trees for sampling from the mouse and sampling from the rat.
  • As a general rule, total blood volume can generally be estimated as 55 - 70 ml/kg body weight. However, care should be taken in these calculations as the percentage of total blood will be lower (-15%) in obese and older animals.
  • Information on total blood volumes and safe blood sample volumes for laboratory animals, domestic species and non-human primates is given here. The EFPIA/ECVAM good practice guide to the administration of substances and removal of blood (Diehl et al. 2001) also contains recommended mean total blood volumes and maximum blood sample volumes for species of a given bodyweight.

A comprehensive table of blood sample volumes is available, which includes volume ranges for laboratory animals, domestic species and non-human primates.

Signs of shock and anaemia

It is essential to be able to recognise the clinical signs of shock and anaemia and be able to take appropriate action.

  • Signs of hypovolaemic shock include a fast and thready pulse, pale dry mucous membranes, cold skin and extremities, restlessness, hyperventilation, and a sub-normal body temperature. The Named Veterinary Surgeon should be consulted immediately if shock occurs. If more than 10% of the total blood volume has been removed, a routine replacement with the same volume of warm (30-39oC) normal buffered saline constitutes good animal care.
  • Signs of anaemia include pale mucous membranes of the conjunctiva or inside the mouth, pale tongue, gums, ears or footpads (non-pigmented animals), intolerance of exercise and, at the more extreme level, an increased respiratory rate when at rest. Where there is concern about the development of anaemia, packed cell volume, haemoglobin level, red blood cell and reticulocyte counts should be monitored throughout the series of bleeds using the results from the first sample from each animal as the baseline for the animal.

Resources and references