This document provides guidelines for blood sample collection in small laboratory animals, emphasizing the necessity of ethics approval and minimizing pain and stress during the procedure. It details methods of blood collection, common anesthetic agents, and specific techniques for various species while also specifying recommended sites, volumes, and frequencies for blood withdrawal. The document concludes with references for further reading on blood collection practices in laboratory settings.

1. Welcome to Journal
Club Presentation
Presented by,
Dnyanshree V. Tijare
Guide: Dr. Manoj Aswar
M. Pharm second year (Dept. of Pharmacology)
Rajarshi Shahu College of Pharmacy, Buldhana

2. Title and Affiliation

3. Continue…
Blood sample collection in small
laboratory animals
Author : Parasuraman S. Ravindran R. Kesavan R
DOI no. : 10.4103/0976-500X.72350
Journal name : Journal of Pharmacology & Pharmacotherapeutics
Publication year : July-December 2010

4. Content
General principles of blood collection method
Common anaesthetic agents required for lab animals
General methods for blood collection
Discussion & References

5. General principles of blood
collection method
1. Protocol approved by the Institute animal ethics committee.
2. It should be least painful and stressful, Blood sample may be collected
under anesthesia or without anesthesia.
3. Adequate training (any species)
4. In general, blood sample is withdrawn from venous, arterial blood vessels or
heart chambers.
5. Frequency (Once in two weeks is ideal for nonrodents).
6. The estimated blood volume in adult animals is 55 to 70 ml/kg body weight.

6. Common anaesthetic agents
required for lab animals

7. General methods for blood
collection
Blood samples does not required anaesthesia
Blood samples required anaesthesia
Terminal procedures
A
B
C

8. General requirement for blood
collection method
• Animal
• Rodent handling gloves
• Towel
• Cotton
• Sample collection tubes
• Surgical blade
• Capillary tube
• Anesthetic agent
• Animal warming chamber
• Needles
• 95% v/v Alcohol
• 1-5 ml syringe
• Electronic trimmer

9. No. Blood sample collection method Needle size/ Other collecting tool
1. Saphenous vein 20G
2. Dorsal pedal vein 23G/ 27G
3. Tail vein 23G
4. Orbital sinus Capillary
5. Jugular vein 25G
6. Blood vessel cannulation R- 23-25G, M- 19-21G
7. Tarsal vein 22G
8. Marginal ear vein/ artery 26G
9. Cardiac puncture 19-25G
10. Posterior vena cava 21-25G
11. Tail snip 1 mm cut

10. Blood sample collection
without anaesthesia
1. • Saphenous vein (rat, mice, guinea pig)
2. • Dorsal pedal vein (rat, mice)

11. Saphenous vein method
Click here to video

12. Dorsal pedal vein method

13. Blood sample collection with
anaesthesia
Tail vein & Tail snip
Orbital sinus
Jugular vein
Blood vessel cannulation
Tarsal vein
Marginal ear vein/ artery

14. Tail vein and Tail snip method
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15. Orbital sinus method
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16. Jugular vein method
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17. Blood vessel cannulation

18. Tarsal vein method

20. Marginal Ear vein method
Click here to play video

21. Terminal procedures
Posterior
vena cava
Cardiac
puncture
1
2

22. Cardiac Puncture
Click here to play video

23. Posterior vena cava

24. Procedure
This procedure is recommended for terminal stage of study
Give V/ Y shape cut in rat abdomen, and remove intestine gently
Anesthetized the animal
The liver is pushed forward, and identified the inferior/ posterior vena cava
Insert 21 to 25G needle and collect the blood

25. 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).

26. Estimated total blood volume and
safe bleeding volume of selected
species (once every two week)
No. Animal Total blood volume
(ml/kg)
Max. bleeding (ml/kg)
(10% of BW)
1. Mice 79 7.9
2. Rat 64 6.4
3. Guinea pig 75 7.5
4. Mini pig 65 6.5
5. Hamster 78 7.8
6. Frog 95 9.5
7. Rabbit 56 5.6
8. Chicken 60 6.0
9. Cat 55 5.5

27. Recommended site for blood
collection
No. Animals Recommended sites and conditions
1. Mice "submandibular" or "facial", saphenous vein,
retro-orbital (anesthetized), cardiac
(anesthetized, terminal)
2. Rat Tail vein, saphenous vein, "submandibular" or
"facial", cardiac (anesthetized, terminal),
sublingual, jugular vein
3. Guinea pig, Hamster Saphenous, cardiac (anesthetized, terminal)
4. Rabbit Marginal ear vein, cardiac (anesthetized,
terminal)
5. Chicken Brachial/wing vein, jugular

28. References
• Hoff J, Rlagt LV. Methods of blood collection in the mouse. Lab
animals 2000;29:47-53.
• Blood sampling online. 2009; [12 screens]. Available
from:http://www.nc3rs.org.uk/bloodsamplingmicrosite/page.asp?id=3
13[Last cited on 2010 Fe24].
• Guidelines for Survival Bleeding of Mice and Rats. NIH-ARAC
Guidelines [online]. 2005 Jan 12. Available from
http://oacu.od.nih.gov/ARAC/Bleeding.pdf [accessed on 2010 Mar
09].
• McGuill MW, Rowan AN. Biological Effects of Blood Loss:
Implications for Sampling Volumes and Techniques. ILAR J 1989. p.
31

29. Continue…
• Procedure for rabbit blood collection [online]. Available from:http://
www.research.uky.edu/ori/univet/resources/sop/Procedure_rabbit_bl
ood_ collection.pdf [Last cited on 2010 Feb 23].
• CPCSEA guidelines for laboratory animal facility. Indian J
Pharmacol 2003;35:257-74.
• Vogel HG, editor. Drug discovery and evaluation: Pharmacological
assays. 2nd ed. Berlin: Springer; 2002.
• Anaesthesia and Analgesia in Laboratory Animals at UCSF [online].
Available from :http://www.iacuc.ucsf.edu/Index.asp[Last cited on
2010 Feb 24]. and Availabl
from:http://www.iacuc.ucsf.edu/Proc/awRatFrm.asp. [Last cited on
2010 Feb 24].

30. Thank You