What is the maximum percentage of blood volume collected from an experimental animal is considered safe in a day?

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. As a general principle, sample volumes and frequency of sampling should be kept to a minimum. 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. How much blood do you require?

* Not recommended as more refined methods of sampling exist. Plus a mixed arterial / venous sample is obtained.

** Not widely used [1]

*** Not widely used [2]

# Blood may be mixed with tissue fluid.

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.

Information on microsampling (e.g. study designs, sampling protocols, videos) can be found in our dedicated microsampling resource. The technique for sampling is as described for tail vein below.

1. Golde WT et al. (2005). A rapid, simple, and humane method for submandibular bleeding of mice using a lancet. Lab Animal 34(9): 39-43. doi: 10.1038/laban1005-39

2. Hiemann M et al. (2009). Blood collection from the sublingual vein in mice and hamsters: a suitable alternative to retrobulbar technique that provides large volumes and minimizes tissue damage. Laboratory Animals 43(3): 255-60. doi: 10.1258/la.2008.007073

3. 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

4. Parasuraman S et al. (2010). Blood sample collection in small laboratory animals. Journal of Pharmacology and Pharmacotherapeutics 1(2): 87-93. doi: 10.4103/0976-500X.72350

5. Teilmann AC et al. (2014). Physiological and pathological impact of blood sampling by retro-bulbar sinus puncture and facial vein phlebotomy in laboratory mice. PLoS ONE 9(11): e113225. doi: 10.1371/journal.pone.0113225 

6. Meyer N et al. (2020). Impact of three commonly used blood sampling techniques on the welfare of laboratory mice: Taking the animal’s perspective. PLoS ONE 15(9): e0238895. doi: 10.1371/journal.pone.0238895

• Technique
• Summary
• Resources and references

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. Surgery is required and appropriate anaesthesia, analgesia and aseptic technique should be used to minimise any pain caused. Mice should be allowed to regain their pre-operative body weight before blood samples are taken.

For recovery work, 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.

Cannulated mice 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 cannula used is 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).

Using a cannula 0.01 - 0.02 ml of blood can be taken and, 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. Aseptic technique should be used. Sterile saline with anticoagulant should be used to flush the cannula after blood sampling to prevent occlusion. A pin is then inserted into the exteriorised end of the cannula to stop the blood from flowing. A sterile locking solution can be used to lock the cannula after a series of samples have been taken, allowing flushing to be avoided for a number of days.

The following should be checked daily

• Skin in contact with the jackets should be checked for abrasion.
• The jacket should be checked for tightness.
• Wound sites should be checked for infection/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

Resources and references

1. Kmiotek EK et al. (2012). Methods for intravenous self administration in a mouse model. Journal of visualized experiments: JoVE (70): e3739. doi: 10.3791/3739
2. UC Davis (2015). Mouse Tail Vein Catherterization Procedure. 
3. Gunaratna PC et al. (2004). An automated blood sampler for simultaneous sampling of systemic blood and brain microdialysates for drug absorption, distribution, metabolism and elimination studies. Journal of Pharmacological and Toxicological Methods 49(1): 57-64. doi: 10.1016/S1056-8719(03)00058-3
4. Bardelmeijer HA et al. (2003). Cannulation of the jugular vein in mice: a method for serial withdrawal of blood samples. Laboratory Animals 37(3): 181-7. doi: 10.1258/002367703766453010
5. Nolan TE and Klein HJ (2002). Methods in vascular infusion biotechnology in research with rodents. Institute for Laboratory Animal Research journal 43(3): 175-82. doi: 10.1093/ilar.43.3.175

Tail snip is not recommended. The saphenous vein and tail vein are more refined and appropriate routes of sampling for most studies and strains of mice. 

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
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Tail vein sampling is suitable for obtaining a small volume of blood (less than 0.2 ml) by incision (with a needle or lance) of the tail vein. 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. For competent individuals, it is quick and simple to perform. View videos of the mouse tail vein sampling technique below (restrained and unrestrained).

This technique may require the animals 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 experimental data depending on the parameters observed. As a result, other routes such as saphenous vein sampling should considered. If it is necessary to warm the animal, a warming cabinet should be used (no more than body temperature for up to 10 minutes) following best practice. Male mice may need to be warmed singly to avoid fighting.

The lateral tail vein is usually used and ≤50µl (microsample) to 0.2 ml of blood can be obtained per sample depending on the size of the animal and its health status. The tail should be washed with an antimicrobial solution such as diluted chlorohexidine to disinfect the area and to see the blood vessel. This is particularly useful for black and pigmented mice. Illumination devices can be used to improve tail vein visualisation, if necessary. 

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. Alternate sides of the tail should be used and successive needle punctures moved towards the tail base. Where it is necessary and justifiable to take more than four samples, the use of temporary or surgical cannulation methods should be considered. If multiple microsamples are required in one day (e.g. when performing a glucose tolerance test), the scab or clot may be gently soaked and removed.

With suitable training acclimatisation and training, restraint may not be necessary. Where animals need to be restrained this can cause stress, therefore the duration of restraint should be minimised.

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. Aseptic technique should be used. 

Blood flow should be stopped by applying finger pressure on 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, and the animal monitored for adverse effects.

Summary

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
8. Meyer N et al.  (2020). Impact of three commonly used blood sampling techniques on the welfare of laboratory mice: Taking the animal’s perspective. PLoS ONE 15(9): e0238895. doi: 10.1371/journal.pone.0238895

• Technique
• Summary
• Resources and references

Sampling from the lateral saphenous vein is a relatively quick method of obtaining blood samples from all strains of mice. It does not require the animal to be warmed for sample collection.

Slides and videos of restraint and sampling for this technique are available on the Norecopa website.

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

Conscious mice should be restrained either manually or using a restraint tube. This can cause stress and therefore the duration of restraint should be minimised. Where a restraint tube is used, it should be appropriate for the size of the mouse. All forms of restraining equipment should be frequently washed to prevent pheromonally-induced stress or cross-infection.

Saphenous bleeding of an untrained (top) and trained (bottom) mouse. The untrained mouse is agitated, flipping a stiff tail, struggling and trying to get away; its ears are flipped back, and eyes half shut; it is hard for the handler to find the blood vessel as the blood flow is "turned off". In contrast, the trained mouse is calm with a relaxed tail and open eyes; the mouse's colour is normal, and it is easy for the technician to find the blood vessel and fill up the capillary tube.

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 clip 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. Anaesthesia is not necessary but may be used on welfare grounds for animals that are difficult to hold. Where sedatives contain peripheral vasodilators, doses should be low to avoid prolonged bleeding from the puncture site. 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 gentle finger pressure over the puncture site, or simple relaxation of the operator's grip on the animal's leg. Animals should not be returned to their cage before the blood flow has stopped.

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. The scab or blood clot is removed for multiple samples.

Mice may show temporary favouring of the opposite limb following sampling from the saphenous vein.

Summary

Resources and references

• Technique
• Summary
• Resources and references

Retro-orbital bleeding should only be performed under terminal anaesthesia because of the severity of adverse effects that can occur with this technique, even in skilled hands (summarised below).

Also referred to as peri-orbital, posterior-orbital and orbital venous sinus bleeding. 

Blood is collected from the venous sinus. The mouse is 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

Resources and references

1. Jo EJ et al. (2021). Comparison of murine retroorbital plexus and facial vein blood collection to mitigate animal ethics issues. Laboratory Animal Research 37(1): 12. doi: 10.1186/s42826-021-00090-4 
2. Meyer N et al. (2020). Impact of three commonly used blood sampling techniques on the welfare of laboratory mice: Taking the animal’s perspective. PLoS ONE 15(9): e0238895. doi: 10.1371/journal.pone.0238895
3. Harikrishnan VS et al. (2018). A comparison of various methods of blood sampling in mice and rats: Effects on animal welfare. Laboratory Animals 52(3): 253-64. doi: 10.1177/0023677217741332
4. Tsai PP et al. (2015). Effects of different blood collection methods on indicators of welfare in mice. Lab Animal 44(8): 301-10. doi: 10.1038/laban.432
5. Fried JH et al. (2015). Type, duration, and incidence of pathologic findings after retroorbital bleeding of mice by experienced and novice personnel. Journal of the American Association for Laboratory Animal Science 54(3): 317-27. PMCID: PMC4460946
6. Teilmann AC et al. (2014). Physiological and pathological impact of blood sampling by retro-bulbar sinus puncture and facial vein phlebotomy in laboratory mice. PLoS ONE 9(11): e113225. doi: 10.1371/journal.pone.0113225
7. Holmberg H et al. (2011). Impact of blood sampling technique on blood quality and animal welfare in haemophilic mice. Lab Animal 45(2): 114-20. doi: 10.1258/la.2010.010129
8. Forbes N et al. (2010). Morbidity and mortality rates associated with serial bleeding from the superficial temporal vein in mice. Lab Animal 10(9): 14-22. doi: 10.1038/laban0810-236
9. Heimann M et al. (2009). Blood collection from the sublingual vein in mice and hamsters: a suitable alternative to retrobulbar technique that provides large volumes and minimizes tissue damage. Lab Animal 43(3): 255-60. doi: 10.1258/la.2008.007073
10. Luzzi M et al. (2005). Collecting blood from rodents: a discussion by the laboratory animal refinement and enrichment forum. Animal Technology and Welfare 4(2): 99-102. 
11. 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
12. Hoff J (2000). Methods of Blood Collection in the Mouse. Lab Animal  29(10): 47-53.

• Technique
• Summary
• Resources and references

Appropriate for all strains of mouse, this is a suitable technique to obtain a single, large, good quality blood sample from a euthanised 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

Resources and references

1. Hedrich H (2012). The laboratory mouse. 2nd edition. Academic Press
2. Parasuraman S et al. (2010). Blood sample collection in small laboratory animals. Journal of Pharmacology and Pharmacotherapeutics 1(2): 87-93. doi: 10.4103/0976-500X.72350
3. Morton DB et al. (2001). Refining procedures for the administration of substances. Laboratory animals 35(1): 1-41. doi: 10.1258/0023677011911345

• Technique
• Summary
• Resources and references

Cardiac puncture should not be used if the peritoneum needs to be lavaged to harvest cells, as this technique can cause blood to escape into the peritoneal cavity.

Cardiac puncture is a suitable technique to obtain a single, large, good quality sample from a euthanised mouse or 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

Resources and references

Although suitable for all strains, this technique should only be used in rare circumstances and where there is exceptional scientific justification.

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 euthanasia, 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

This technique should only be used in rare circumstances and with exceptional scientific justification. In the UK this technique is not a Schedule 1 method of euthanasia, therefore personal and project licence authority is required.

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