Blood Sampling in Mice and Rats

Blood Sampling in Mice and Rats

Approved September 18, 2003 Revised: November 15, 2012   (Download) 

These guidelines have been developed to assist investigators and the Institutional Animal Care and Use Committee (IACUC) in their choice and application of rodent bleeding procedures. The guidelines are based on peer-reviewed publications as well as on data and experience accumulated at the UW.

Background: Blood samples can be obtained from various sites of the body, using a variety of methods: from the veins, from the arteries, the retro-orbital sinus (mice) or plexus (rats), the jugular vein, by cardiac puncture, or by decapitation. For repeated sampling surgical placement of indwelling catheters is recommended. The choice of the method depends on several factors, including the purpose of the blood collection, need for arterial or venous sample, duration and frequency of sampling, the health status of the animal being bled, whether the experiment is terminal for the animal, and the training and experience of the phlebotomist. When selecting a method for blood sampling in the conscious animal, consideration must be given to the potential for stress-induced effects on biochemical parameters. Local anesthesia, sedation, and general anesthesia are refinements that assist in immobilization and may improve success and decrease stress for blood sampling. Noteworthy are reports of significant differences in values, e.g. hematological and plasma, depending on the site of blood collection (see references below for some examples).

Although there are no precise data to support amount and frequency of blood volume sampling from rodents, recommendations generally state that no more than 1% of the body weight should be removed at a single bleeding. For example, this would equate to removal of 0.2 ml from a 20 gram mouse or 3.0 ml from a 300 gram rat. Removal of larger volumes necessitates longer recovery periods for the animal, or may cause distress or death. This 1% body weight volume may be taken once every 2 weeks. Scientific objectives of the study may require more frequent sampling that does not exceed this total volume. If there is a scientific necessity for removal of more than 1% of the body weight within a two week period, a rationale should be given for the increased volume and a plan provided for monitoring of the animals and possibly replacement of blood products (e.g., fluid administration, red blood cell administration).

Orbital bleeding is a procedure used in rats and mice (though usually not a good choice in the rat) to obtain small blood samples. A glass capillary tube is inserted at the medial canthus to gain access to the mouse orbital venous sinus or rat orbital venous plexus. This area offers a readily accessible site for blood collection. Retro-orbital bleeding may be conducted in awake mice by a skilled operator. Alternatively, systemic anesthesia should be considered if compatible with experimental objectives. Due to restraint issues, retro-orbital sampling in the rat should be conducted under general anesthesia. Alternate eyes should be used for sequential bleeds. In the hands of an unskilled operator, retro-orbital sampling has a greater potential than other blood collection routes to result in complications and thus requires certification. A traumatized eye will require euthanasia.

Sites for blood sampling are listed in the table below. Please consult with Animal Use Training (auts@uw.edu) for training or Veterinary Services (vsreview@uw.edu) for questions regarding which technique is appropriate for your protocol.

Mouse Policy: Sites for blood collection in the mouse include the lateral saphenous vein, the tail vein, the facial vein (submandibular), the retro-orbital sinus and the heart (see chart below). Depending on the skill of the operator, volumes of up to 200 microliters can be readily collected from the lateral or medial saphenous vein. It is also possible to obtain small amounts of blood by tail vein prick with minimal restraint. Volumes of up to 1% of the body weight are readily and rapidly obtained from the retro-orbital sinus and facial vein. Since there is a risk of corneal or ocular damage with orbital bleeding, orbital bleeding must only be performed in mice by persons certified to perform it by the Attending Veterinarian or an individual designated by the Attending Veterinarian. Although there may be several individuals listed in an approved protocol, orbital bleeding must be restricted to specific individual(s) that have been certified. Cardiac blood withdrawal is performed when the mouse is in a deep plane of anesthesia and is generally a terminal (non-recovery) procedure. Tail tip removal is not a recommended procedure in the mouse. If the technique is proposed, it must be justified scientifically with an explanation of why other techniques are not satisfactory. For repeated blood sampling, indwelling catheter placement is recommended.

Rat Policy: Sites for blood collection in the rat include the lateral tail vein, the ventral tail artery, the dorsal metatarsal or lateral saphenous vein, tail tip removal, the retro-orbital plexus, the jugular vein and the heart (see chart below). Other sites used more rarely include the penile vein and lingual vein. Typically, the lateral tail vein is used for small volumes of blood (up to about 1 ml). However, the lateral saphenous or the dorsal metatarsal vein can be used as well. Very small volumes of blood can be obtained by tail tip removal (0.1 – 0.2 ml). Tail tip removal is only used when multiple short-interval, small- volume samples are required in awake rats. Volumes of up to 1% of the body weight may be obtained from retro-orbital plexus (0.5 to 3 ml depending on the size of the rat) in anesthetized rats only. Since there is a risk of corneal or ocular damage with this technique, orbital bleeding may be performed in rats by persons certified to perform it by the Attending Veterinarian or an individual designated by the Attending Veterinarian. Although there may be several individuals listed in an approved protocol, orbital bleeding must be restricted to specific individual(s) that have been certified. Volumes of up to 1% of the body weight can also be obtained from the jugular vein by a skilled individual in an anesthetized rat. Cardiac blood withdrawal is performed when the rat is in a deep plane of anesthesia and is generally a terminal (non-recovery) procedure. For repeated blood sampling, indwelling catheter placement is recommended.

Table I: Summary of Blood Sampling Techniques

MICE
Route General Anesthesia Required Certification Required Volume <µl> collected (approximately) Comments
Lateral/ medial saphenous No No 50-200 not as rapid as other techniques, low potential for tissue damage, larger volume requires skilled collector
Retro-orbital No Yes 200 rapid, potential for complications
Facial Vein (submandibular) No No 200 rapid, potential for complications if untrained, local tissue trauma seen
Prick tail vein No No 20 useful for small volume collection with minimal restraint
Tail tip removal Yes Yes 20 not recommended, must justify use of this technique
Cardiac Yes No 200-500 usually terminal, must justify use of this technique as a survival procedure
RATS
Route General Anesthesia Required Certification Required Comments
Lateral tail vein or Ventral tail artery No No repeatable, simple
Dorsal metatarsal or lateral saphenous No No not as rapid as other techniques, low potential for tissue damage
Retro-orbital Yes Yes rapid, potential for complications
Cardiac Yes No usually terminal, must justify use of this technique as a survival procedure
Jugular Recommended No limited application, poor for repeated sampling and technically difficult
Tail tip removal (distal tip, not to include bone) No No used for repeated small samples only

 

References:
Site and Anesthesia Can Change Outcome:
1. Chan YK, Davis PF, Poppitt SD, Sun X, Greenhill NS, Krishnamurthi R, Przepiorski A, McGill AT, Krissansen GW. 2012. Influence of tail versus cardiac sampling on blood glucose and lipid profiles in mice. Lab Anim. 46(2):142-7. Epub 2012 Mar 7.
2. Fernández I, Peña A, Del Teso N, Pérez V, Rodríguez-Cuesta J. 2010. Clinical biochemistry parameters in C57BL/6J mice after blood collection from the submandibular vein and retroorbital plexus. J Am Assoc Lab Anim Sci. 49(2):202-6.
3. Nyuyki KD, Maloumby R, Reber SO, Neumann ID. 2012. Comparison of corticosterone responses to acute stressors: Chronic jugular vein versus trunk blood samples in mice. Stress 15(6):618-26. doi: 10.3109/10253890.2012.655348. Epub 2012 Feb 23.
Blood Volume Removed:
1. Minabe S, Uenoyama Y, Tsukamura H, Maeda K. 2011. Analysis of pulsatile and surge-like luteinizing hormone secretion with frequent blood sampling in female mice. J Reprod Dev. 57(5):660-4. Epub 2011 Jul 30.
2. Pekow, C. and V. Baumans, V. Common Nonsurgical Techniques and Procedures In J. Hau and G.L. Van Hoosier (ed) Handbook of Laboratory animal Science, 2 ed. Essential Principles and Practices, Vol I. CRC Press, Boca Raton, Florida.
3. Raabe BM, Artwohl JE, Purcell JE, Lovaglio J, Fortman JD. 2011. Effects of weekly blood collection in C57BL/6 mice. J Am Assoc Lab Anim Sci. 50(5):680-5.
4. Weixelbaumer KM, Raeven P, Redl H, van Griensven M, Bahrami S, Osuchowski MF. 2010. Repetitive low-volume blood sampling method as a feasible monitoring tool in a mouse model of sepsis. Shock. Oct;34(4):420-6.
5. Schnell, Mam et al. 2002. Effect of Blood Collection Technique in Mice on Clinical pathology Parameters. Human Gene Therapy 13:155-162.
6. Joint working group on refinement. 1993.. Removal of Blood from laboratory mammals and birds: First report of the BVA/FRAME/RSPCA/UFAW Lab Anim 27: 1-22.
Comparision of Techniques:
1. Aasland KE, Skjerve E, Smith AJ. 2010. Quality of blood samples from the saphenous vein compared with the tail vein during multiple blood sampling of mice. Lab Anim. 44(1):25-9. Epub 2009 Jun 17.
2. Abatan OI, Welch KB, Nemzek JA. 2008. Evaluation of saphenous venipuncture and modified tail-clip blood collection in mice. J Am Assoc Lab Anim Sci. 47(3):8-15. Arnold M, Langhans W. 2010. Effects of anesthesia and blood sampling techniques on plasma metabolites and corticosterone in the rat. Physiol Behav. 99(5):592-8. Epub 2010 Feb 10.
3. Christensen SD, Mikkelsen LF, Fels JJ, Bodvarsdóttir TB, Hansen AK. 2009. Quality of plasma sampled by different methods for multiple blood sampling in mice. Lab Anim. 43(1):65-71. Epub 2008 Nov 10.
4. Heimann M, Roth DR, Ledieu D, Pfister R, Classen W. 2010. Sublingual and submandibular blood collection in mice: a comparison of effects on body weight, food consumption and tissue damage. Lab Anim. 44(4):352-8. Epub 2010 Aug 9.
5. Holmberg H, Kiersgaard MK, Mikkelsen LF, Tranholm M. 2011. Impact of blood sampling technique on blood quality and animal welfare in haemophilic mice. Lab Anim. 45(2):114-20. Epub 2011 Mar 7.
6. Van Herck H., et al. 2001. Blood Sampling from the Retro-orbital Plexus, the Saphenous Vein and the Tail in Rats: Comparative Effects on Selected Behavioral and Blood Variables. Lab. Anim. 35:131-139