Guidelines for Blood Collection in Laboratory Animals


This document is to provide guidance to investigators regarding safe blood collection volumes in common laboratory animals. All procedures must be approved by the Institutional Animal Care and Use Committee (IACUC). The method of blood collection to be used, the intervals between blood collection procedures, and the volume of blood to be removed, should be listed in the approved protocol specific to each study.


The method and volume of blood to be collected will depend on the animal species, frequency of collection, and experimental needs.  Researchers should plan and perform each sampling protocol with an appreciation for the stresses associated with blood loss to the animal and do whatever they can to minimize the animal's reaction to the stress. Careful planning and control of blood sampling and all experimental variables associated with it should not only benefit the welfare of the animal, but also minimize confounding influences on research data. (7)

Training and experience of the individual in the chosen procedure are of critical importance.  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.

DLAR has a number of training resources available.

Please contact our Training Coordinator for details or to schedule a session.


Recommended volumes for blood collection are intended to preserve the health status of the animal and maintain the validity of experimental results. The guidelines provided are for healthy, normal adult animals. Animals that are young, aged, stressed, have undergone experimental manipulations, or are suffering from cardiac or other disease conditions may not be able to tolerate these recommended blood volume withdrawals .


On average, the total circulating blood volume is equal to 5.5 -8.0 % of the animal’s body weight.  Non-terminal blood collection without additional monitoring (see below) should be limited to 10% of the total circulating blood volume on a single collection or every 2 week period for serial collections.

Example (Using mean blood volume table below): a 4 kg rabbit is calculated to have a total blood volume of 224 ml (56 ml/kg x 4.0 kg). Thus, 22.4 ml (10% of 224 ml) may be collected without giving replacement fluids once every two weeks.

Estimated Total Blood Volume and Safe Bleeding Volume of Selected Species:

Species Blood volume (ml/kg) One Bleeding -
max - 10% of blood volume (ml/kg)
Mean Range
Cat 55 55 5.5
Cattle 55 55 5.5
Chicken 60 60 6.0
Dog 86 86 (79-90) 8.6
Ferret 75 75 7.5
Frog 95 95 9.5
Gerbil 67 67 6.7
Goat 66 -- 6.6
Guinea Pig 75 75 (67-92) 7.5
Hamster 78 78 7.8
Minipig 65 65 (61-68) 6.5
(Cynomolgous Macaque)
65 65 (55-75) 6.5
(Rhesus Macaque)
54 54 (44-67) 5.4
Mouse 79 79 (63-80) 7.9
Pig 65 65 (61-68) 6.5
Rabbit 56 56 (44-70) 5.6
Rat 64 64 (58-70) 6.4
Sheep 66 66 (60-74) 6.6

Source: Adapted from Formulary for Laboratory Animals, Hawk, Leary, and Morris 2005

Note:  If the amount of blood volume removed is 7.5% of total blood volume, allow 1 week recovery; if amount removed is 10%, allow 2 weeks recovery; if amount removed is 15%, allow 4 weeks recovery.

Common Sites for Blood Collection:

Species Recommended Sites & Conditions
Mouse Superficial temporal vein (a.k.a., "submandibular" or "facial"), saphenous vein, tail vein, retro-orbital (anesthetized), cardiac (anesthetized, terminal)
Rat Tail vein, saphenous vein, superficial temporal vein (a.k.a., "submandibular" or "facial"), cardiac (anesthetized, terminal), sublingual, jugular
Dog, Cat, Non-human Primate Cephalic, saphenous veins, femoral and jugular viens
Guinea pig, Hamster Saphenous, cardiac (anesthetized, terminal)
Rabbit Marginal ear vein, cardiac (anesthetized, terminal)
Swine Jugular, ear vein
Pigeon, Quail Brachial/wing vein
Chicken Brachial/wing vein, jugular
Ruminant, Equine Jugular

Guidelines for Rodents:

Similarly, of the circulating blood volume, approximately 10% of the total volume can be safely removed every 2 to 4 weeks, 7.5% every 7 days, and 1% every 24 hours. Volumes greater than recommended should be scientifically justified and appropriate fluid and/or cellular replacement provided. Approximate blood sample volumes for a range of body weights are included in the table below:

Body Weight (g) *CBV (ml) 1% CBV (ml)
every 24 hours**
7.5% CBV (ml)
every 7 days**
10% CBV (ml)
every 2-4 weeks**
20 1.10-1.40 0.011-0.014 0.082-0.105 0.11-0.14
25 1.37-1.75 0.014-0.018 0.10-0.13 0.14-0.18
30 1.65-2.10 0.017-0.021 0.12-0.16 0.17-0.21
35 1.93-2.45 0.019-0.025 0.14-0.18 0.19-0.25
40 2.20-2.80 0.022-0.28 0.16-0.21 0.22-0.28
125 6.88-8.75 0.069-0.088 0.52-0.66 0.69-0.88
150 8.25-10.50 0.082-0.105 0.62-0.79 0.82-1.00
200 11.00-14.00 0.11-0.14 0.82-1.05 1.10-1.40
250 13.75-17.50 0.14-0.18 1.00-1.30 1.40-1.80
300 16.50-21.00 0.17-0.21 1.20-1.60 1.70-2.10
350 19.25-24.50 0.19-0.25 1.40-1.80 1.90-2.50
*Circulating Blood Volume ** Maximum sample volume for that sampling frequency


unanesthetized collection methods

Blood Collection methods in Mice and Rats (unanesthetized):

Superficial temporal (a.k.a. “Submandibular” or “Facial” vein/artery) Sampling:

  • Obtainable blood volumes: medium to large.
  • Repeated sampling is possible by alternating sides of the face.
  • Sample may be a mixture of venous and arterial blood.
  • Requires less hands-on training than tail or retro-orbital sampling to reliably withdraw a reasonable quantity of blood.
  • Perform on awake animals to achieve proper restraint, which in turn results in proper site alignment and venous compression for good blood flow.
  • Can be performed rapidly and with a minimal amount of equipment, allowing for rapid completion.
  • Sample volume can be partially controlled with the size of needle (20 gauge or smaller) or lancet (4 mm) used to puncture the site.
  • Use of a lancet is recommended to control depth of puncture and reduce potential for complications.  These can be significant if puncture is too deep or homeostasis is not assured prior to returning the mouse to its cage (5).

Saphenous Sampling (medial or lateral approach):

  • Obtainable blood volumes: small to medium.
  • Can be used in both rats and mice by piercing the saphenous vein with a needle or lancet.
  • Variable sample quality.
  • The procedure is customarily done on an awake animal but effective restraint is required.
  • Requires more hands-on training than tail or retro-orbital sampling to reliably withdraw more than a minimal amount of blood.
  • Although more aesthetically acceptable than retro-orbital sampling, prolonged restraint and site preparation time can result in increased animal distress when handling an awake animal.
  • Temporary favoring of limb may be noted following the procedure.
  • Application of sterile petroleum jelly to the site facilitates blood droplet formation, which can enhance the total blood volume captured.
  • The clot/scab can be gently removed for repeated small samples if serial collection is required

Lateral Tail Vein:

  • Obtainable blood volumes: small to medium.
  • Can be used in both rats and mice by cannulating the blood vessel or by superficially nicking the vessel perpendicular to the tail.
  • Sample collection by nicking the vessel is easily performed in both species, but produces a sample of variable quality that may be contaminated with tissue and skin products. Sample quality decreases with prolonged bleeding times and “milking” of the tail.
  • Sample collection using a needle (cannulation) minimizes contamination of the sample, but is more difficult to perform in the mouse.
  • Repeated collections possible. With tail nicking, the clot/scab can be gently removed for repeated small samples if serial testing is required (e.g., glucose measures, etc.)
  • In most cases warming the tail with the aid of a heat lamp or warm compresses will increase obtainable blood volume.
  • Cannulation and tail nicking are routinely done without anesthesia, although effective restraint is required.

Jugular Sampling (typically limited to the rat, but can be performed in mice as well (6):

  • Obtainable blood volumes: medium to large.
  • May be needed when a larger blood volume withdrawal and survival are needed.
  • Results in high quality sample.
  • Jugular sampling can be conducted without anesthesia in rats, although the use of anesthesia greatly facilitates the procedure and reduces the potential for injury to the animal as well as the individual (e.g., bites, needle stick).
  • Does not easily lend itself to repeated serial sampling

anesthetized collection methods

Blood Collection methods in Mice and Rats (anesthetized):

NB:  The following methods require anesthesia (local and/or general depending on species and technique) to relieve pain and distress associated with the technique or for restraint. Use of these methods may require scientific justification for why less painful techniques cannot be used in the animal care and use protocol.

Retro-orbital Sinus/Plexus Sampling: 

  • Obtainable volume: medium to large.
  • Retro-orbital sampling can be used in both mice and rats (though not a preferred method in the rat) by penetrating the retro-orbital sinus in mice or plexus in rats with a capillary tube or Pasteur pipette.
  • Rapid – large number of animals can be bled within a short period of time.
  • Good sample quality. Potential contamination with topical anesthetic, if used, should be taken into account.
  • A minimum of 10 days should be allowed for tissue repair before repeat sampling from the same orbit. Otherwise the healing process may interfere with blood flow.
  • Alternating orbits should not be attempted until the phlebotomist is proficient in obtaining samples from the orbit accessed most readily by the dominant hand ie a right handed individual should gain proficiency withdrawing samples from the right orbit before attempting to obtain samples from the left orbit.
  • In the hands of an unskilled phlebotomist, retro-orbital sampling has a greater potential than other blood collection routes to result in complications.
  • General anesthesia must be used unless scientific justification is provided and approved by the IACUC. In addition, a topical ophthalmic anesthetic, e.g. proparacaine or tetracaine drops, is recommended prior to the procedure.
  • In both mice and rats, care must be taken to ensure adequate hemostasis following the procedure.
  • Use of sterile capillary tubes and pipettes are recommended for use to help avoid periorbital infection and potential long-term damage to the eye. The edges of the tubes should be checked for smoothness to also decrease likeliness of eye damage.
  • Animals must be monitored following collection for damage to the orbit or globe of the eye.  If damage is noted, veterinary staff should be contacted for treatment options up to and including euthanasia.

Tail Clip Sampling:

  • Obtainable volume: small.
  • Can be used in both rats and mice by clipping (e.g. amputating) no more than 1mm of the distal tail in mice or 2 mm in rats
  • Produces a sample of variable quality that may be contaminated with tissue and skin products. Sample quality decreases with prolonged bleeding times and “milking” of the tail.
  • Obtainable volume: small.
  • Repeated collections possible. The clot/scab can be gently removed for repeated small samples if serial testing is required (e.g., glucose measures, etc.)
  • In most cases warming the tail with the aid of a heat lamp or warm compresses will increase obtainable blood volume.
  • When performing tail clipping, consideration should be given to anesthesia/analgesia, particularly if the tail has been previously clipped for genotyping. If a topical hypothermic anesthetic is used, blood will flow as the tail re-warms. If a local anesthetic is applied, adequate contact time should be allowed for it to take effect.

anesthetized terminal

Blood Collection methods in Mice and Rats (anesthetized and terminal):

Cardiac Puncture:

  • Obtainable Volume:  medium – large.
  • Cardiac puncture is done only as a terminal procedure and always under anesthesia as evidenced by lack of response to a painful stimulus (e.g., toe pinch)
  • The use of a 1cc syringe with a 25 gauge needle is recommended. Find the xiphoid process as a reference point. Insert the needle at a 35-40 degree angle just under and to the left of the xiphoid process. As the needle is inserted into the chest, gently aspirate until blood begins to flow.
  • Ensure death of the animal after collection. This determination may be made by auscultation for cessation of both heartbeat and respiration by a qualified individual in larger animals or by utilizing an unequivocal secondary means of ensuring death (decapitation, opening thoracic cavity, etc.) following euthanasia with an inhalant agent (anesthetic overdose or CO2)



If the animal is being bled routinely, the red blood cell packed volume (PCV) should be checked weekly to determine when blood collection should be suspended in order for the animal to recover from potential anemia. While healthy adult animals can recover their blood volume within 24 hours, it may take up to 2 weeks for all the other blood constituents (i.e. cells, proteins) to be replaced.

By monitoring the hematocrit (Hct or packed cell volume- PCV) and/or hemoglobin of the animal, it is possible to evaluate whether the animal has sufficiently recovered from a single or multiple blood draws. After a sudden or acute blood loss, it takes up to 24 hours for the hematocrit and hemoglobin to reflect this loss. In general, if the animal’s hematocrit is less than 35% or the hemoglobin concentration is less than 10 g/dl, it is not safe to remove blood.  Please contact a DLAR Veterinarian if you need assistance with monitoring PCVs in animals. 

Normal Packed Cell Volume (PCV) for some lab animals (%)

Mouse 39-49 Dog 29-55
Rat 36-54 Cat 25-41
Gerbil 43-60 Rhesus 26-48
Hamster 40-61 Baboon 33-43
Rabbit 30-50 Swine 32-50
Guinea Pig 37-48 Cow 24-48
Sheep 24-45 Avian 35-55


Fluid replacement

Lactated Ringer’s Solution (LRS) is the recommended balanced crystalloid solution for fluid replacement. Alternatively, 0.9% sterile isotonic saline may be used. For adult mice, 1.0 ml of warmed LRS or isotonic saline can be given by IP or SC administration. For adult rats, administer 5 -10 ml warmed LRS or 0.9% saline (½ of the total volume via IP and ½ via SC routes).


1. Hawk TC, Leary, ST, and Morris, TH. 2005.  Formulary for Laboratory Animals, 3rd ed.  Ames, IA  Blackwell Publishing

2. BVA/FRAME/RSPCA/UFAW Joint Working Group on Refinement. Removal of blood from laboratory mammals and birds. Laboratory Animals (1993) Jan; 27(1):1-22

3. NIH/OACU Guidelines for Survival Bleeding in Mice and Rats, 2010 Available at:  Accessed 7/11/11

4. Boston University IACUC Policy for Blood Collection Guidelines.  Available at:  Accessed 7/11/11

5. Forbes N, et. al. Morbidity and mortality rates associated with serial bleeding from the superficial temporal vein in mice. Lab Animal (2010) Aug; 39(8):236-40.

6. Hoff, J. Methods of Blood Collection in the Mouse. Lab Animal (2000) Nov; 29(10):47-53.

7. McGuill M.W. and Rowan A.N. Biological Effects of Blood Loss: Implications for Sampling Volumes and Techniques. ILAR News (1989), 31(4): 5-18.

8. Nahas K, Provost J-P, Baneux PH, and Rabemampianina Y. Effects of acute blood removal via the sublingual vein on haematological and clinical parameters in Sprague-Dawley rats.  Laboratory Animals (2000) Oct; 34(4):362-71.

9. NC3R’s (National Centre for the Replacement, Refinement and Reduction of Animals in Research) Blood Sampling Microsite.  Available at: Accessed 7/11/11