Methods
Blood sample collection in small laboratory animals
www.jpharmacol.com
Parasuraman S, Raveendran R, Kesavan R
Department of the Pharmacology, Jawaharlal Institute of Postgraduate Medical Education and Research, Pondicherry, India
Collection of blood from small laboratory animals is necessary any method in any species.
for a wide range of scientific research and there are a number •• In general, blood sample is withdrawn from venous,
of efficient methods available for that. It is important that blood arterial blood vessels or heart chambers.
sample collection from experimental animals should be least •• Frequency of blood collection is important. Once in two
stressful because stress will affect the outcome of the study. weeks is ideal for nonrodents. If the study needs multiple
Various regulatory agencies and guidelines have restricted the blood samples, lagomorphs (e.g., hares and rabbit) can
use of animals and the techniques used for blood collection in be used.
laboratory animals. This article deals with the approved blood •• All nonterminal blood collection without replacement of
collection techniques for laboratory animals like rodents, fluids is limited up to 10% of total circulating blood volume
lagomorphs and nonrodents. Permission of the Institute Animal in healthy, normal, adult animals on a single occasion and
Ethics Committee has been obtained for the use of animals for collection may be repeated after 3 to 4 weeks. In case
demonstrating the techniques. repeated blood samples are required at short intervals, a
maximum of 0.6 ml/kg/day or 1.0% of an animal’s total
blood volume can be removed every 24 hour.[2,3]
GENERAL PRINCIPLES OF BLOOD •• If the study involves repeated blood sample collection, the
COLLECTION IN ANIMALS samples can be withdrawn through a temporary cannula.
This may reduce pain and stress in the experimental animals.
•• The method of blood collection should be described in the •• The estimated blood volume in adult animals is 55
protocol approved by the Institute animal ethics committee. to 70 ml/kg body weight. Care should be taken for
•• It should be least painful and stressful. Blood sample may be older and obese animals.[4] If blood collection volume
collected under anesthesia [Table 1] or without anesthesia.[1] exceeds more than 10% of total blood volume, fluid
•• Adequate training is required for blood collection using replacement may be required. Lactated Ringer’s solution
Table 1: Commonly recommended anesthetic agents for laboratory animal experiments
Animal species Short anesthesia Medium anesthesia Long anesthesia
Mice Isoflurane (inhalation) Xylazine + ketamine Xylazine + ketamine
Halothane (inhalation) (5 mg + 100 mg i.m.) (16 mg+60 mg i.m./i.p.)
Rat Xylazine + ketamine Xylazine + ketamine
(5 mg + 100 mg i.m.) (16 mg +60 mg i.m./ i.p.) or
Urethane (1200 mg/kg i.p.)
Guinea pig Isoflurane (inhalation) Xylazine + ketamine Xylazine + ketamine
(2 mg + 80 mg i.m.) (4 mg + 100 mg i.m.)
Rabbits Isoflurane (inhalation) Xylazine + ketamine Xylazine + ketamine
(5 mg + 15 – 30 mg i.m.) (5 mg + 100 mg i.m.)
Atropine (0.02 mg/kg s.c./i.m.) is used as a preanesthetic medication for all the species to reduce salivation, bronchial secretion and protect heart from vagal
inhibition.[6-8]
Address for correspondence:
Parasuraman S, Department of the Pharmacology, Jawaharlal Institute of Postgraduate Medical Education and Research, Pondicherry, India.
E-mail: parasuphd@gmail.com
DOI: 10.4103/0976-500X.72350
Journal of Pharmacology & Pharmacotherapeutics | July-December 2010 | Vol 1 | Issue 2 87
Parasuraman, et al.: Blood sample collection in small laboratory animals
(LRS) is recommended as the best fluid replacement by PROCEDURE FOR DORSAL PEDAL VEIN
National Institutes of Health (NIH). If the volume of blood BLOOD SAMPLE COLLECTION
collection exceeds more than 30% of the total circulatory
blood volume, adequate care should be taken so that the Requirements include animal (rat or mice), rodent handling
animal does not suffer from hypovolemia.[5] gloves, cotton, capillary tube, 23G/27G needle and blood
sample collection tubes.
GENERAL METHODS FOR BLOOD •• The animal is kept in a restrainer.
COLLECTION •• The hind foot around ankle is held and medial dorsal pedal
vessel is located on top of the foot.
•• The foot is cleaned with absolute alcohol and dorsal pedal
Blood samples are collected using the following techniques:[1]
vein is punctured with 23G/27G needle.
•• Blood collection not requiring anesthesia
•• Drops of blood that would appear on the skin surface are
Saphenous vein (rat, mice, guinea pig)
collected in a capillary tube and a little pressure is applied
Dorsal pedal vein (rat, mice)
to stop the bleeding [Figure 1].
•• Blood collection requiring anesthesia (local/general
anesthesia)
Tail vein (rat, mice) PROCEDURE FOR TAIL VEIN BLOOD SAMPLE
Tail snip (mice)
COLLECTION
Orbital sinus (rat, mice)
Jugular vein (rat, mice) Requirements include animal, rodent handling gloves, towel,
Temporary cannula (rat, mice) cotton, sample collection tube and animal warming chamber.
Blood vessel cannulation (rat, guinea pig, ferret) •• This method is recommended for collecting a large volume
Tarsal vein (guinea pig) of blood sample (up to 2ml /withdrawal)
Marginal ear vein/artery (rabbit) •• The animal is made comfortable in a restrainer while
•• Terminal procedure maintaining the temperature around at 24 to 27°C.
Cardiac puncture (rat, mice, guinea pig, rabbit, ferret) •• The tail should not be rubbed from the base to the tip as
Orbital sinus (rat, mice) it will result in leukocytosis. If the vein is not visible, the
Posterior vena cava (rat, mice) tail is dipped into warm water (40°C).
•• Local aesthetic cream must be applied on the surface of
PROCEDURE FOR SAPHENOUS VEIN BLOOD the tail 30 min before the experiment.
•• A 23G needle is inserted into the blood vessel and blood is
SAMPLE COLLECTION[9] collected using a capillary tube or a syringe with a needle.
In case of difficulties, 0.5 to 1 cm of surface of the skin
Requirements include animal, rodent handling gloves, towel,
is cut open and the vein is pricked with bleeding lancet
cotton, sample collection tubes and 20G needle.
or needle and blood is collected with a capillary tube or a
•• Lateral saphenous vein is used for sampling while taking syringe with a needle.
aseptic precautions. •• Having completed blood collection, pressure/silver nitrate
•• The back of the hind leg is shaved with electric trimmer ointment/solution is applied to stop the bleeding.
until saphenous vein is visible. Hair removal cream can •• If multiple samples are needed, temporary surgical cannula
also be used. may be used.
•• The animal is restrained manually or using a suitable •• Restrainer is washed frequently to avoid/prevent
animal restrainer. pheromonally induced stress or cross infection [Figure 2].
•• Hind leg is immobilized and slight pressure may be applied
gently above the knee joint.
•• The vein is punctured using a 20G needle and enough PROCEDURE FOR TAIL SNIP BLOOD SAMPLE
volume of blood is collected with a capillary tube or a COLLECTION
syringe with a needle. The punctured site is compressed
to stop the bleeding. While collecting blood: Requirements include animal, anesthetic agent, cotton, surgical
the local anesthetic cream may be applied on the blade and blood sample collection tubes.
collection site •• This method is recommended for blood collection only
no more than three attempts are made in mice.
continuous sampling should be avoided and •• This method should be avoided as far as possible because
collecting more than four samples in a day (24-hour it can cause potential permanent damage on the animal tail.
period) is not advisable. If needed, it should be done under terminal anesthesia only.
88 Journal of Pharmacology & Pharmacotherapeutics | July-December 2010 | Vol 1 | Issue 2
Parasuraman, et al.: Blood sample collection in small laboratory animals
Figure 1: Blood sample collection from rat dorsal pedal vein
Figure 2: Blood sample collection from mouse tail vein
•• Before collecting the blood, local anesthesia is applied capillary tube and blood sample collection tubes.
on the tail and a cut is made 1 mm from the tip of the tail •• This technique is used with recovery in experimental
using scalpel blade. circumstances and this method is also called periorbital,
•• Blood flow is stopped by dabbing the tail tip. posterior-orbital and orbital venous plexus bleeding.
•• Blood sample is collected under general anesthesia.
PROCEDURE FOR ORBITAL SINUS BLOOD •• Topical ophthalmic anesthetic agent is applied to the eye
SAMPLE COLLECTION before bleeding.
•• The animal is scruffed with thumb and forefinger of the
Requirements include animal, anesthetic agent, cotton, nondominant hand and the skin around the eye is pulled taut.
Journal of Pharmacology & Pharmacotherapeutics | July-December 2010 | Vol 1 | Issue 2 89
Parasuraman, et al.: Blood sample collection in small laboratory animals
Figure 3: Blood sample collection from rat orbital sinus
with sterile cotton. Bleeding can be stopped by applying
gentle finger pressure.
•• Thirty minutes after blood collection, animal is checked
for postoperative and periorbital lesions [Figures 3 and 4].
•• Caution:
Repeated blood sampling is not recommended.
Skill is required to collect blood.
Even a minor mistake will cause damage to the eyes.
Two weeks should be allowed between two bleedings.
•• Adverse effects reported from this method is around 1 to
2% which includes hematoma, corneal ulceration, keratitis,
pannus formation, rupture of the globe, damage of the
optic nerve and other intraorbital structures and necrotic
dacryoadenitis of the harderian gland.
PROCEDURE FOR JUGULAR VEIN BLOOD
SAMPLE COLLECTION
Figure 4: Blood sample collection from mouse orbital sinus Requirements include animal, anesthetic agent, cotton, 25G
needle and blood sample collection tubes.
•• A capillary is inserted into the medial canthus of the eye •• In this method, warming of the animals is not required and
(30 degree angle to the nose). is used to collect micro volumes to one ml of blood sample.
•• Slight thumb pressure is enough to puncture the tissue and •• This method has to be carried out under general/inhalation
enter the plexus/sinus. anesthesia and two persons are needed to collect blood
•• Once the plexus/sinus is punctured, blood will come sample.
through the capillary tube. •• One person has to restrain the animal and monitor the
•• Once the required volume of blood is collected from animal. Another person is required to collect the blood
plexus, the capillary tube is gently removed and wiped sample from the animal.
90 Journal of Pharmacology & Pharmacotherapeutics | July-December 2010 | Vol 1 | Issue 2
Parasuraman, et al.: Blood sample collection in small laboratory animals
•• The neck region of the animal is shaved and kept in
hyperextended position. The jugular veins appear blue in
color and is found 2 to 4 mm lateral to sternoclavicular
junction. A 25G needle is inserted in the caudocephalic
direction (back to front) and blood is withdrawn slowly
to avoid collapse of these small blood vessels. Animal has
to be handled carefully and not more than 3 to 4 mm of
needle is to be inserted into the blood vessel.
•• If the attempt to collect blood fails, the needle is slowly
removed and the site is monitored for bleeding. If there
is no bleeding, one more attempt can be made. Further
attempts should be avoided in case of bleeding as it may
collapse the vein.
•• Finger pressure is applied to stop bleeding.
•• Caution:
Number of attempts is limited to three.
Apply local anesthetic cream 30 minutes prior to
sampling. Figure 5: Blood vessel cannulation of rat femoral vein
PROCEDURE FOR BLOOD SAMPLE
COLLECTION WITH TEMPORARY CANNULA Table 2: Needle size used for blood vessel
cannulation in different species
Requirements include animal, anesthetic agent, cotton, 25G Species Needle to be used Maximum collection
volume
needle, animal warming chamber and blood sample collection
Mice 23 – 25G 1 ml
tubes.
Rat 19 – 21G 10 – 15 ml
•• Usually a temporary cannulation is made in the tail vein Rabbit 19 – 21G 60 – 200 ml
and used for a few hours. Guinea pig 20 – 21G 1 – 25 ml
•• The animal is restrained and local anesthetic cream is
applied on the tail (1 – 2 cm above the tail tip).
•• The tail is either cannulated or a 25G needle is used. •• Blood sample may be collected over 24 hour at the volume
•• Tail bleeding normally requires the animal to be warmed in of 0.1 to 0.2 ml/sample.
order to dilate the blood vessels (37 – 39°C for 5 – 15 min). •• After withdrawing the blood, the cannula is flushed with an
•• After cannulation, animal has to be housed individually anticoagulant and the withdrawn volume may be replaced
in large cages. (if required) with LRS and cannula should be closed tightly
[Figure 5].
PROTOCOL FOR BLOOD VESSEL •• Caution: The experiment has to be conducted fully under
aseptic precautions. Infection, hemorrhage, blockage of
CANNULATION
cannula and swelling around the cannulation site should
Requirements include animal, anesthetic agent, cotton, be looked for. The needle size and maximum blood volume
25G needle, i.v. cannula, surgical blade, heparin (or any to be collected are given in Table 2.
anticoagulant) and blood sample collection tubes.
•• This method involves continuous and multiple sampling PROTOCOL FOR TARSAL VEIN BLOOD
in the experimental animal. SAMPLE COLLECTION
•• This method requires close and continuous monitoring
of the animal. Requirements include animal, anesthetic agent, cotton, 22G
•• Usually blood vessel cannulation is done in the femoral needle, hair remover and blood sample collection tubes.
artery, femoral vein, carotid artery, jugular vein, vena cava •• Tarsal vein is identified in one of the hind legs of large
and dorsal aorta. animals. This method is commonly recommended for
•• Surgery is required for this method and appropriate guinea pig.
anesthesia and analgesia should be used to minimize the •• One person has to restrain the animal properly. Tarsal vein
pain. may be visible in blue color.
•• After surgical cannulation, animal should be housed singly •• The surface hairs are removed by applying a suitable
in a large and spacious cage. hair remover. A local anesthetic cream is applied on the
Journal of Pharmacology & Pharmacotherapeutics | July-December 2010 | Vol 1 | Issue 2 91
Parasuraman, et al.: Blood sample collection in small laboratory animals
Figure 6: Blood sample collection from guinea pig tarsal vein
Figure 7: Blood sample collection from rabbit marginal ear vein using Figure 8: Blood sample collection from rabbit marginal ear vein using
26 G needle incision method.
•• Caution:
Not more than six samples from both hind legs are taken.
The number of attempts is three or less.
PROTOCOL FOR MARGINAL EAR VEIN/
ARTERY BLOOD SAMPLE COLLECTION
Requirements include animal, anesthetic agent, cotton, 26G
needle, 95% v/v alcohol, o-Xylene, surgical blade and blood
sample collection tube.
•• This method is commonly adopted for rabbits.
•• The animal should be placed in a restrainer.
•• Ear is cleaned with 95% v/v alcohol and local anesthetic
cream is applied on the collection site 10 min prior to
sampling. (If required, the o-Xylene/topical vasodilator
Figure 9: Blood sample collection through cardiac puncture in rat
may be applied topically on the collection site to dilate
blood vessels).
collection site. •• Size 11 surgical blade is used to cut the marginal ear vein
•• After 20 to 30 minutes, blood sample is collected slowly and blood is collected in a collecting tube. Otherwise, a
by using 22G needle. 26G needle may be used to collect blood from animal
•• Maximum three samples can be taken per leg and 0.1 to marginal vein.
0.3 ml of blood can be collected per sample. •• After collecting blood, clean sterile cotton is kept on the
•• After the sample collection, gentle pressure is applied with collection site and finger pressure is applied to stop the
finger for 2 minutes to stop bleeding [Figure 6]. bleeding [Figures 7 and 8].
92 Journal of Pharmacology & Pharmacotherapeutics | July-December 2010 | Vol 1 | Issue 2
Parasuraman, et al.: Blood sample collection in small laboratory animals
PROTOCOL FOR CARDIAC PUNCTURE[10,11] error in the collection procedure may lead to a lot of variation
in the results.
Requirements include animal, anesthetic agent, towel, cotton,
19 to 25G needle with 1 to 5 ml syringe, surgical blade, tube Points to be remembered
(internal diameter of 0.1 to 0.3 mm) for thoracotomy, plastic •• Before starting any kind of blood sample collection, it must
disposable bag and blood sample collection tubes. be ensured that all chemical, surgical, fluid requirements
•• In general, cardiac puncture is recommended for terminal are available in the working site.
stage of the study to collect a single, good quality and large •• Not more than two to three attempts should be made to
volume of blood from the experimental animals. collect any kind of in vitro biological sample (excluding
•• During blood sample collection, animal will be in terminal biological secretion).
anesthesia. •• The blood collection tube must be labeled before starting
•• Appropriate needle is used for blood sample collection the experiment and blood sample collected in the
with or without thoracotomy. Blood sample will be taken appropriately labeled collection tube.
from the heart, preferably from the ventricle slowly to
avoid collapsing of heart [Figure 9].
•• Caution: If animal has dextrocardia, sampling may fail. REFERENCES
1. Hoff J, Rlagt LV. Methods of blood collection in the mouse. Lab animals
PROTOCOL FOR BLOOD SAMPLE 2.
2000;29:47-53.
Blood sampling online. 2009; [12 screens]. Available from:http://www.nc3rs.
COLLECTION THROUGH POSTERIOR VENA org.uk/bloodsamplingmicrosite/page.asp?id=313[Last cited on 2010 Feb 24].
3. Guidelines for Survival Bleeding of Mice and Rats. NIH-ARAC Guidelines
CAVA [online]. 2005 Jan 12. Available from: http://oacu.od.nih.gov/ARAC/
Bleeding.pdf [accessed on 2010 Mar 09].
Requirements include animal, anesthetic agent, surgical blade, 4. McGuill MW, Rowan AN. Biological Effects of Blood Loss: Implications
small glass rods, surgical scissor, 21 to 25G needle with 1 to for Sampling Volumes and Techniques. ILAR J 1989. p. 31.
5. Procedure for rabbit blood collection [online]. Available from:http://
5 ml syringe and blood sample collection tube. www.research.uky.edu/ori/univet/resources/sop/Procedure_rabbit_blood_
•• In general, posterior vena cava blood sample is collection.pdf [Last cited on 2010 Feb 23].
recommended for terminal stage of the study. 6. CPCSEA guidelines for laboratory animal facility. Indian J Pharmacol
2003;35:257-74.
•• Animal have to be anesthetized and ‘Y’- or ‘V’-shaped 7. Vogel HG, editor. Drug discovery and evaluation: Pharmacological assays.
cut in the abdomen is made and the intestines are gently 2nd ed. Berlin: Springer; 2002.
removed. 8. Anaesthesia and Analgesia in Laboratory Animals at UCSF [online].
Available from :http://www.iacuc.ucsf.edu/Index.asp[Last cited on 2010
•• The liver is pushed forward and the posterior vena cava Feb 24]. and Available from:http://www.iacuc.ucsf.edu/Proc/awRatFrm.asp.
(between the kidneys) is identified. [Last cited on 2010 Feb 24].
•• 21 to 25G needle is inserted to collect blood from the 9. Hem A, Smith AJ, Solberg P. Saphenous vein puncture for blood sampling
of the mouse, rat, hamster, gerbil, guinea pig, ferret and mink. Lab Anim
posterior vena cava. 1998;32:364-8.
•• This procedure will be repeated three to four times to 10. Paulose CS, Dakshinamurti K. Chronic catheterization using vascular-access-
collect more volume of blood sample. port in rats: Blood sampling with minimal stress for plasma catecholamine
determination. J Neurosci Methods 1987;22:141-6.
11. Yoburn BC, Morales R, Inturrisi CE. Chronic vascular catheterization in the
rat: Comparison of three techniques. Physiol Behav 1984;33:89-94.
DISCUSSION
Blood collection from the experimental animals is one of the
Source of Support: Nil, Conflict of Interest: None declared
important procedures in biomedical research. Even a small
Journal of Pharmacology & Pharmacotherapeutics | July-December 2010 | Vol 1 | Issue 2 93
