Blood Collection Guidelines


All non-terminal blood collection without replacement fluids is limited to 10% of the total circulating blood volume of a healthy animal during a 2 week period. On average, the total circulating blood volume is equal to 5.5 -7.0 % (~66 ml/kg) of the animal’s body weight. If larger amounts are needed, then up to 15 % of the total circulating blood volume may be withdrawn if replacement fluids are given at the time of blood withdrawal. Example: a 4 kg rabbit is calculated to have a total blood volume of 264 ml (66 ml/kg x 4.0 kg). Thus, 26 ml (10% of 264 ml) may be collected without giving replacement fluids and 40 ml (15% of 264 ml) may be collected if replacement fluids are given once every two weeks. Removal of 15% of total blood volume must be justified in the IACUC protocol and approved by the IACUC.


The limitations for blood collection preserve the health status of the animal and maintain the validity of experimental results based on blood samples. 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 respiratory disease may not be able to tolerate this amount of blood loss.

Guidelines for rodents

Table 1: Approximate Blood Sample Volumes for a Range of Body Weights

Body weight (g)


Circulating Blood Volume

1% CBV (ml)

every 24 hrs†

7.5% CBV (ml)

every 7 days†

10% CBV (ml)

every 2 wks†


1.10 – 1.40

.011 – .014

.082 – .105

.11 – .14


1.37 – 1.75

.014 – .018

.10 – .13

.14 – .18


1.65 – 2.10

.017 – .021

.12 – .16

.17 – .21


1.93 – 2.45

.019 – .025

.14 – .18

.19 – .25


2.20 – 2.80

.022 – .028

.16 – .21

.22 – .28


6.88 – 8.75

.069 – .088

.52 – .66

.69 – .88


8.25 – 10.50

.082 – .105

.62 – .79

.82 – 1.0


11.00 – 14.00

.11 – .14

.82 – 1.05

1.1 – 1.4


13.75 – 17.50

.14 – .18

1.0 – 1.3

1.4 – 1.8


16.50 – 21.00

.17 – .21

1.2 – 1.6

1.7 – 2.1


19.25 – 24.50

.19 – .25

1.4 – 1.8

1.9 – 2.5

*Circulating blood volume

†maximum sample volume for that sampling frequency


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.

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





















Guinea Pig






Blood collection sites in mice and rats

The following guidelines refer to the most frequently used survival sampling sites: a) submandibular veins; b) tail veins; c) saphenous veins; d)jugular veins; and e) retroorbital. Blood withdrawal by cardiac puncture is considered a euthanasia procedure and should be performed only after ensuring that the animal is under deep anesthesia, as evidenced by lack of response to a painful stimulus (e.g., toe or tail pinch). A list of the issues that should guide the choice of survival blood collection route(s) is noted below, and an abbreviated summary is provided in Table 2.

Facial vein as a method of blood collection in mice

  • Blood collection from the submandibular facial vein is a safe and fast technique in mice. It requires momentary restraint and approximately 200ul of blood can be obtained easily from a healthy adult mouse. The vessel is located just beneath the skin immediately caudal to the facial vibrissae (whiskers) at the corner of the jaw. Repeated sampling is possible by alternating sides of the face. Materials needed include a 20 or 22 g hypodermic needle, blood collection tubes and sterile gauze. Training is necessary before this procedure is performed.

Lateral Tail Vein or Ventral Tail Artery Sampling:

  • 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.
  • Repeated collections possible. With tail vein nicking, the clot/scab can be gently removed for repeated small samples if serial testing is required (e.g., glucose measures, etc.)
  • Tail artery sampling yields larger volumes but requires the animal to be anesthetized and placed in dorsal recumbency. Good hemostasis is also required, as always whenever an artery is incised.

Saphenous Vein Sampling (medial or lateral approach):

  • Can be used in both rats and mice by piercing the saphenous vein with a needle.

Jugular Vein Sampling (limited to the rat):

  • Obtainable blood volumes: medium to large.
  • Results in high quality sample.
  • Jugular sampling can be conducted without anesthesia, although the use of anesthesia greatly facilitates the procedure.
  • Does not easily lend itself to repeated serial sampling.

Retro-orbital Sinus/Plexus Sampling:

  • Retro-orbital sampling can be used in both mice and rats (though usually not a method of choice in the rat) by penetrating the retro-orbital sinus in mice or plexus in rats with a glass capillary tube or Pasteur pipette.
  • Repeat sampling from the same orbit may be difficult (10 days to 2 weeks recommended between successive bleeds). However, alternating orbits should not be attempted until the phlebotomist is proficient with the technique in the same orbit.


Animals will need to be physically restrained to prevent any movement that would result in lacerating the blood vessel or other potentially serious complications. Blood may be collected from awake animals that are appropriately restrained provided that persons performing the procedure are skilled.


Anesthesia is required if blood collection is being performed either via the retro-orbital sinus or by cardiac puncture due to the distress and pain which can be caused and for the serious complications (injury to the eye, cardiac tamponade and death) associated with these routes. For survival procedures requiring anesthesia isoflurane is recommended as it is short-acting and allows replacing the rodent in its cage within minutes.

Common sites for blood collection in large animals


Sites of collection and permitted conditions


Cardiac (under anesthesia as a terminal procedure only), jugular vein, marginal ear vein (for small volume only), ear artery (requires good hemostasis)

Dog and cat

Cephalic, saphenous veins, femoral and jugular veins


Jugular vein


Jugular vein, anterior vena cava, ear veins

Nonhuman Primates

Femoral, cephalic veins, saphenous vein

Fluid replacement

Lactated Ringer’s Solution (LRS) is recommended as the best for fluid replacement. For mice administered 1 ml of warmed LRS IP or SC. For rats administer 5 -10 ml warmed LRS ½ via IP and ½ via SC administration.

Nutritional supplementation

When larger volumes are withdrawn, especially when there are repeated sampling, it is recommended that the animal receives Nutrical, a dietary supplement. For rats and mice, this can easily be done by smearing Nutrical on a few pellets and placing those on the cage floor.


Training is required for blood collection in any species and by any route. Please contact the LACF/LASC Training Coordinator to schedule training.


1) NIH Guidelines for Survival Bleeding in Mice and Rats, 2007.

2) Boston University IACUC Policy for Blood Collection Guidelines

Table 2: Summary of Rodent Blood Sampling Techniques