Blood sampling: Mouse

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. Sample volumes and and the total number of samples should be kept to the minimum necessary. Further advice is given in the general principles.

How much blood does a mouse have?

On average, mice have around 58.5 ml of blood per kg of bodyweight.

A mouse weighing 25 g would therefore have a total blood volume (TBV) of approximately 58.5 ml/kg x 0.025 kg = 1.46 ml.

How to decide on the most appropriate blood sampling technique for mice?

The two tables below are designed to assist in determining the amount of blood to sample from the animal, and depending on that volume, the most appropriate techniques to use.

1. Do you require more than one blood sample from the same mouse?

2. Microsampling

Advances in bioanalytical techniques have opened up the potential to use smaller sample volumes (microsamples of ≤50µl) to assess drug and chemical exposure in blood, plasma and/or serum. Proper consideration should be given to microsampling before obtaining larger blood samples.

Information on microsampling (e.g. study designs, sampling protocols, videos) can be found on our microsampling page. 

3. How much blood do you require?

* Saphenous or tail vein sampling are recommended over these approaches. 

# Blood may be mixed with tissue fluid.

• Technique
• Summary
• Resources and references

Technique

Sampling from the lateral saphenous vein is a relatively quick method of obtaining blood samples, including microsamples, from all strains of mice. The vein is easily visualised and it does not require the animal to be warmed for sample collection, but shaving hair away from the sampling site is necessary.

Blood is collected from the lateral saphenous vein which runs dorsally and then laterally over the tarsal joint. 

Conscious mice should be restrained manually for the minimum duration necessary. Mice should be habituated to restraint; this will improve the experience of the animal and handler and will lead to obtaining a better-quality blood sample. 

Every opportunity should be taken to habituate mice for low stress procedures. This should including habituating mice to to the sound of the electric shaver to minimise additional stress. If sedation is required on welfare grounds care should be taken due to the vasodilation action of some sedatives. Where sedatives contain peripheral vasodilators, doses should be low to avoid prolonged bleeding from the puncture site.

To collect blood, the hind leg should be immobilised in the extended position by applying gentle downward pressure immediately above the knee joint. This stretches the skin over the ankle, making it easier to shave away hair and immobilise the saphenous vein. Please note that hair removal by shaving with a scalpel blade is no longer recommended as it removes the epidermal layers of the skin. Aseptic technique should be used. 

The number of attempts to take a blood sample should be minimised (no more than three needle sticks in any one attempt). Blood is collected by capillary action into a haematocrit tube or passively into a tube.

Blood flow can be stopped by the operator relaxing their grip on the animal's leg and gently pressing a swab over the puncture site. Animals should not be returned to their cage before the blood flow has stopped.

If more than one sample is required legs can be alternated. No more than four blood samples should be taken within any 24-hour period. If more samples are needed, then temporary or surgical cannulation should be considered. 

Note that single-use needles are designed to be used once. If they are reused, there is a risk that they will dull and cause animals pain, as well as potentially transferring tissue products or spreading infection between animals. 

Summary

Number of samples

No more than four blood samples should be taken within any 24-hour period. The total number of samples should be kept to the minimum necessary.

Sample volume

Up to 0.2 ml for a single sample, which can usually be repeated at 2-week intervals without disturbances to haematological status. Alternatively, multiple smaller samples (e.g. 0.01 ml daily), taking into account limits on sample volume. Proper considering should be given to microsampling to keep the total volume of samples to the minimum necessary.

Equipment

27G or 25G needle or blood lancet (not a scalpel)

Staff resource

One person is required to take the blood sample.

Adverse effects

Adverse effects are not typical when this technique is performed competently, but animals should be monitored for:

• Bruising
• Haemorrhage
• Infection
• Temporary favouring of the opposite limb

Resources and references

• Technique
• Summary
• Resources and references

Technique

Tail vein sampling is suitable for obtaining ≤50µl (microsample) to 0.2 ml of blood depending on the size and health status of the mouse. For competent individuals it is quick and simple to perform by using a needle, butterfly needle or blood lancet to pierce the skin and vein. Note that it is not appropriate to cut the tail with a scalpel to obtain a blood sample.

It is suitable for all strains but is more difficult in black or pigmented mice as their vasculature can be difficult to observe through the skin. Illumination devices, such as small LED lights, can be used to improve tail vein visualisation. Sampling should not occur if it is not possible to visualise the tail vein and saphenous vein sampling should be considered instead.

This technique may require mice to be warmed to dilate the blood vessel prior to taking the sample. This can speed up sampling but can negatively impact animal welfare and data quality. For example, warming can cause dehydration and an increase metabolic rate, which may affect experimental data depending on the parameters observed. If it is necessary to warm mice best practice should be followed, including monitoring the mice for overheating. Signs of overheating include the ears or muzzle becoming red and shaking of the head.

With suitable acclimatisation and training, restraint is not be necessary for tail vein sampling (see 21:39 of the webinar Handling and training of mice and rats for low stress procedures). Habituating mice to the blood sampling procedure will improve the experience of the animal and handler and will lead to obtaining a better-quality blood sample. 

If restraint is necessary, manual restraint should be used rather than a device, particularly if mice have been warmed. If a restraint device is used great care should be taken and best practice should be followed. 

Aseptic technique should be used. The tail should be cleansed with an antimicrobial solution such as diluted chlorohexidine, then dried, to disinfect the area and to improve visualisation of the blood vessel. This is particularly useful for black and pigmented mice. 

The lateral tail vein is usually accessed approximately one-third along the length of the tail from the tail tip. If more than one sample is required alternate sides of the tail should be used and successive needle punctures should move towards the tail base. Where it is necessary and justifiable to take repeat samples, the use of temporary or surgical cannulation methods should be considered.

To avoid bruising and damage to the tail, normally no more than two blood samples should be taken in any one 24-hour period. 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. It is not appropriate to remove a scab from a previous needle puncture and reattempt to draw blood from this site.

Note that single-use needles are designed to be used once. If they are reused, there is a risk that they will dull and cause animals pain, as well as potentially transferring tissue products or spreading infection between animals. 

Blood flow should be stopped by applying finger pressure on the soft tissue. A swab should be placed at the blood sampling site and held gently in place using the fingers for approximately 30 seconds before the animal is returned to its cage. It is particularly important to ensure that bleeding has stopped if warming (vasodilation) techniques have been used. Once returned to the cage, the mouse should be monitored for adverse effects. 

Summary

Number of samples

One or two blood samples can be taken per session and in any 24-hour period, depending on sample volume. The total number of samples should be kept to the minimum necessary. 

Sample volume

≤50µl up to 0.2 ml may be obtained. Ensure that proper consideration has been given to using microsampling to keep the total volume of samples to the minimum necessary.

Equipment

25G needle or butterfly needle or blood lancet. Note that cutting the tail with a scalpel to obtain a blood sample is an outdated approach that should not be used.

Staff resource

One person can take the blood sample when mice are well-habituated. Where restraint is necessary, two operators using careful manual restraint is preferable to one person using a restraint tube. If a restraint device is used ensure that best practice is followed. For large groups of animals, more staff members are required.

Adverse effects

Adverse effects are not typical when this technique is performed competently, but animals should be monitored for:

• Infection <1%
• Haemorrhage <1%

Other

Mice may be warmed to dilate the blood vessel. Care should be taken to avoid thermal stress and dehydration and best practice should be followed. Using microsampling techniques reduces the need for warming mice and refines the overall procedure.

Resources and references

1. Diehl KH et al. (2001). A good practice guide to the administration of substances and removal of blood, including routes and volumes. Journal of Applied Toxicology 21(1): 15-23. doi: 10.1002/jat.727
2. Hoff J (2000). Methods of Blood Collection in the Mouse. Lab Animal 29(10): 47-53.
    
3. Morton DB et al. (1993). Removal of blood from laboratory mammals and birds. Laboratory Animals 27(1): 1-22. doi: 10.1258/002367793781082412
4. Durschlag M et al. (1996). Repeated blood collection in the laboratory mouse by tail incision - modification of an old technique. Physiology and Behaviour 60(6): 1565-8. doi: 10.1016/s0031-9384(96)00307-1
5. Hem A et al. (1998). Saphenous vein puncture for blood sampling of the mouse, rat, hamster, gerbil, guinea pig, ferret and mink. Laboratory animals 32(4): 364-8. doi: 10.1258/002367798780599866
6. Sadler MA and Bailey SJ (2017). Validation of a refined technique for taking repeated blood samples from juvenile and adult mice. Laboratory Animals 47(4):316-9. doi: 10.1177/0023677213494366 
7. David JM and Chen X (2018). Evaluation of warming devices for lateral tail vein blood collection in mice (Mus musculus). Journal of Pharmacological and Toxicological Methods 94(Pt 1):87-93. doi: 10.1016/j.vascn.2018.06.002

Technique

Snipping the tail is a crude method of sampling and should be avoided as it involves the removal of soft tissue from the tip of the tail using a scalpel, resulting in permanent damage to the tail and pain to the mouse. An additional limitation of the technique includes contamination, with the sample containing tissue fluid as well as blood. 

• Technique
• Summary
• Resources and references

Technique

Blood vessel cannulation should be considered when repeated samples are required, as it avoids multiple needle entries at any one site. It is suitable for use in all strains of mice and can be used to take blood from the carotid artery, vena cava and femoral vein. 

Depending on the sample volume and scientific justification up to six samples and a maximum of 1% of the total blood volume may be taken in a 24-hour period. 

Surgery is required and appropriate anaesthesia, analgesia and aseptic technique should be used to minimise any pain caused. Mice should be allowed to fully recover from surgery and ideally regain their preoperative body weight for a period of approximately 7 days before they are enrolled in studies and blood samples are taken.

A refinement to the use of a traditional catheterisation system and exteriorised technique is the transcutaneous skin button. Skin buttons provide a closed system; they feature an external port which contains a septum, a disk which is placed subcutaneously in the interscapular area and a connector to join the surgically implanted catheter to the external port. The external port allows for quick and clean repeated access. A metal cap may be placed on the external port, protecting it and allowing mice to be group housed.

When using conventional cannulation methods, the cannula may be exteriorised at the nape of the neck through a jacket and tether system. The jacket can cause swelling and skin abrasion and mice require regular and detailed observation to identify any problems. Use of a subcutaneous access port may be more appropriate than exteriorisation because these eliminate the need for tethering and hence allow group housing. For terminal work, the cannula is not exteriorised.

Mice cannulated using the conventional approach are usually housed singly. The caging, bedding and environmental enrichment need to be appropriate to prevent the tether becoming entangled and the wound contaminated. In addition, the bedding needs to be sand free.

The cannulas used for mice are small, which can promote blood clotting (however, larger cannulae can abrade the blood vessel wall). To prevent this, the cannula requires regular maintenance (e.g. flushing with an anticoagulant). Aseptic technique should be used when accessing the catheter and the catheter should be locked accordingly. 

The following should be checked daily

• Skin in contact with the jackets should be checked for abrasion.
• The jacket should be checked for tightness.
• The site of the catheter should be checked for dehiscence /infection/irritation/bruising/swelling/haemorrhage.
• The cannula should be checked for patency (without blockage).
• The weight of the mouse (remember weight will include that of the device).

Changes in any of the above may require veterinary advice or treatment, or may indicate that a humane endpoint has been reached and appropriate action should be taken.

Summary

Number of samples

It is recommended up to six samples may be taken in a 24-hour period, depending on sample volume.

Sample volume

0.01 - 0.02 ml

Equipment

25G cannula

Staff resource

One person is required to take the blood sample. Further staff resource is required for surgery, post-operative care for as long as necessary for the individual animal, and daily animal observations post-surgery.

Adverse effects and incidence

• Infection 1-5%
• Haemorrhage 1-5%
• Poor recovery after surgery 1-5%
• Blocked cannula 1-5%
• Swelling around the jacket 1-5%
• Skin sores from the jacket 1-5%

Be sure to use our advice on vascular catheters to reduce the incidence of adverse effects.

Other

Mice should be back at their pre-operative weight before blood sampling starts.

Resources and references

• Technique
• Summary
• Resources and references

Technique

Also referred to as peri-orbital, posterior-orbital and orbital venous sinus bleeding. Blood is collected from the venous sinus.

The mouse is placed under terminal anaesthesia and restrained, the neck gently scruffed and the eye made to bulge. A capillary tube/pipette is inserted medially, laterally or dorsally. Blood is allowed to flow by capillary action into the capillary tube/pipette. The sample obtained is a mixture of venous blood and tissue fluid, and is not representative of venous blood.

Summary

Number of samples

One 

Sample volume

Up to 0.5 ml 

Equipment

A glass capillary tube or Pasteur pipette.

Staff resource

One person is required to take the blood sample.

Adverse effects

• Retro-orbital haemorrhage resulting in haematoma and excessive pressure on the eye
• Corneal ulceration, keratitis, pannus formation, rupture of the globe and micro-ophthalmia caused by proptosis of the globe
• Damage to the optic nerve and other intra-orbital structures which can lead to deficits in vision and blindness
• Fracture of the fragile bones of the orbit and neural damage by the micro-pipette
• Penetration of the eye globe itself with a loss of vitreous humour

Other

This is non-recovery procedure and should be always be carried out under terminal anaesthesia.

Resources and references

• Technique
• Summary
• Resources and references

Technique

Appropriate for all strains of mouse, this is a suitable technique to obtain a single, large, good quality blood sample from a humanely killed mouse or a mouse under terminal anaesthesia. A sample size of 0.4 -1.0 ml can be collected depending on the size of the mouse. As the heart is not punctured, this technique can be used when it is necessary to avoid cardiac damage.

Blood is collected either from the abdominal vena cava, abdominal aorta or aortic arch which can be accessed via a laparotomy or thoracotomy in larger mice. Removal of connective tissue and application of finger pressure is necessary to dilate the vessel. Blood should be withdrawn slowly to prevent the vessel collapsing. 

Summary

Number of samples

One

Sample volume

Up to 1 ml

Equipment

25G needle

Staff resource

One person is required to take the sample.

Other

This is non-recovery/terminal procedure.

Resources and references

• Technique
• Summary
• Resources and references

Technique

Cardiac puncture is a suitable technique to obtain a single, large, good quality sample from a mouse following humane killing, or from a mouse under deep terminal anaesthesia if coagulation parameters, a separate arterial or venous sample or cardiac histology are not required. It is appropriate for all strains of mouse.

0.1 - 1 ml of blood can be obtained depending on the size of the mouse and whether the heart is beating. Blood samples are taken from the heart, preferably the ventricle, which can be accessed either via the left side of the chest, through the diaphragm, from the top of the sternum or by performing a thoracotomy. Blood should be withdrawn slowly to prevent the heart collapsing.

Summary

Number of samples

One

Sample volume

Up to 1 ml

Equipment

23G-25G needle

Staff resource

One person is required to take the sample.

Other

This is non-recovery procedure and should be always be carried out under terminal anaesthesia.

Resources and references

• Technique
• Summary

Technique

The primary reason for using this technique is to obtain a large volume of blood that has not been affected by anaesthetic drugs or carbon dioxide. A large volume of blood can be collected from the trunk if necessary, but it should be noted there is a risk of contamination from other body fluids and tissues.

In order to be deemed a Schedule 1 method of humane killing, mice which have been stunned must be determined as dead before decapitation (e.g. via confirmation of cessation of circulation or exsanguination - see Section 1(4) of the amended ASPA). This method should only be carried out by people competent in this method for the species and size of the animal. Training for stunning and decapitation should be undertaken on dead animals.

Summary

Number of samples

One

Sample volume

Up to 1 ml

Equipment

Suitable sharp instrument to decapitate (e.g., sharp scissors for neonatal mice, guillotine for adult mice).

Staff resource

One person is required to take the sample.

Other

A high level of expertise is required for this technique.

• Technique
• Summary

Technique

Trunk blood is collected from the site where the animal is decapitated, under deep terminal anaesthesia. It should be noted there is a risk of contamination from other body fluids and tissues. Training for decapitation should be undertaken on dead animals.

Summary

Number of samples

One

Sample volume

Up to 1 ml

Equipment

Suitable sharp instrument to decapitate (e.g., sharp scissors for neonatal mice, guillotine for adult mice).

Staff resource

One person is required to take the sample.

Other

A high level of expertise is required for this technique.