Guidelines on Blood Collection
This document is designed to provide general information on blood collection methods for 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, must be listed in the approved protocol specific to each study.
The ULAM veterinary staff provides the following criteria to determine the maximum safe amount of blood to be withdrawn. It is recommended to take no more blood than is absolutely necessary. Remember to calculate beforehand the minimum amount of blood necessary to perform all tests and assays, and that the serum fraction is about ½ of the total blood volume. When calculating blood volumes based on body weights, remember that body weights in kilograms (kgs) will convert to blood volumes in liters, and weights in grams will convert to volumes in milliliters (mls).
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Procedures
1. Approximate Blood Volume
- 5-10% of body weight = total blood volume
- In healthy animals the circulating blood volume can generally be estimated as 55-70 ml/kg of the total body weight.
- In sick, obese and older animals the circulating blood volume can be up to 15% lower than in healthy animals, therefore, calculations should be adjusted accordingly.
- See Table 13 at end of this document for average blood volumes in certain species.
2. Blood Collection Volumes
- 1% of body weight (BW) = maximum volume that can be collected every 14 days, without requiring supplemental fluids. Examples:
- 0.15 ml from a 15 g mouse
- 3 ml from a 300 g rat
- 50 ml from a 5 kg cat
- 100 ml from a 10 kg monkey
- 400 ml from a 40 kg dog
- If blood collections are needed multiple times within a 14-day period, calculate the total amount needed over the 14-day span and divide 1% by that number. Examples:
- 0.07% of body weight maximum per collection if 14 samples are needed within a 14-day period.
- 0.1% of body weight maximum per collection if 10 samples are needed within a 14-day period.
- 0.14% of body weight maximum per collection if 7 samples are needed within a 14-day period.
- 4-5% of body weight = amount expected at exsanguinations (see section 4)
3. Fluid Replacement Recommendations
- If the total volume withdrawn over a 14-day period is greater than 1% BW, fluid volume replacement must be considered.
- Use of a warm (30-35ºC) isotonic solution (e.g., 0.9% saline, lactated Ringer's solution) constitutes accepted veterinary practice. Replacement fluids should be infused at a slow and steady rate.
- Examples of Fluid Replacement
- For a 15 g mouse:
- Up to 0.15 ml (1% withdrawn) of blood collected over 14 days does not require fluid replacement.
- Up to 0.3 ml (2% withdrawn) of blood collected over 14 days, requires fluid replacement with 0.3 ml appropriate fluids SQ.
- For a 300 g rat:
- Up to 3 ml (1% withdrawn) of blood collected over 14 days does not require fluid replacement.
- Up to 6 ml (2% withdrawn) of blood collected over 14 days, requires fluid replacement with 0.3 ml appropriate fluids SQ.
- For a 20 kg dog:
- Up to 200 ml (1% withdrawn) of blood collected over 14 days does not require fluid replacement.
- Up to 400 ml (2% withdrawn) of blood collected over 14 days, requires fluid replacement with 0.3 ml appropriate fluids SQ or IV.
- For a 15 g mouse:
4. Exsanguination
- Approximately 50-75% of total blood volume (4-5% of body weight) can be obtained by terminal exsanguination. The animal must be deeply anesthetized, or recently euthanized, prior to exsanguination. Since the amount of blood obtained is substantially increased if the heart is beating during the bleeding procedure, use of a surgical plane of anesthesia is recommended. The procedure for anesthesia and/or euthanasia must be described fully in the approved IACUC protocol. Examples:
- 0.60-0.75 ml from a 15 g mouse
- 12-15 ml from a 300 g rat
- 200-250 ml from a 5 kg cat
- 400-500 ml from a 10 kg monkey
- 1600-2000 ml from a 40 kg dog
5. Monitoring
- If too much blood is withdrawn too rapidly or too frequently without replacement (approximately 2% of the animal's body weight at one time), the animal may go into hypovolemic shock. If signs of shock are observed, contact the ULAM veterinary staff immediately. Signs of shock include:
- Fast and thready pulse
- Pale dry mucous membranes
- Cold skin and extremities
- Restlessness
- Hyperventilation
- Sub-normal body temperature
- If 15-20% of total blood volume is removed, cardiac output and blood pressure will be reduced.
- If 30-40% of total blood volume is removed, death will result in at least 50% of animals.
- If > 40% of total blood volume is removed, death of the animal is expected.
- Stressed, sick, or otherwise compromised animals may not tolerate the blood collection criteria noted above, which is for healthy animals.
- By monitoring hematocrit (Hct or PCV) and/or hemoglobin (Hb) it is possible to evaluate if the animal has sufficiently recovered from single or multiple blood draws. Remember it may take up to 24 hours for hematocrit or hemoglobin to reflect a sudden or acute blood loss. In general, if the animal is anemic (below the normal PCV range for the species), or if the hemoglobin concentration is less than 10 gm/dL, it is not safe to remove the volumes of blood listed above.
6. Blood Collection Sites & Methods
The tables below list the most frequently used blood collection sites for common laboratory animal species. They are listed from most common/desirable to least common/desirable based on ease of collection from the site. For uncommon laboratory animal species, please contact the ULAM veterinarians at [email protected]. For smaller species, the volume of blood attainable for each site is listed based; however, this is an estimation and will also depend on the size, heath, and hydration status of the animal as well as the experience level of the person collecting the sample. Based on the goals and requirements of the study, certain sites may be preferable). Additionally, publications have indicated that the results from blood analysis (especially cellular indices) may vary based on the site of blood withdrawal; consult the literature for more information. In all cases, cardiac puncture may be used to obtain a single, large volume of blood from heavily anesthetized (terminal procedure only) or euthanized animals.
- Table 1: Mouse
Site
Anesthesia
Repeat Bleeds
Expected Volume
Lateral Tail Vein
No
Yes
50 - 100 ul
Saphenous Vein
No
Yes
100 - 200 ul
Facial Vein (submandibular & submental veins)
No
Yes
200 - 500 ul
Distal Tail Transection (less than 3mm)
Required a
Yes - limited
< 100 ul
Retro-Orbital Sinus
Required
Yes - limited
200 ul
Sublingual Vein
Required
Yes
500 ul
Jugular Vein
Recommended
Yes
Cardiac Puncture (Terminal Only)
Required
Terminal
~ 1 ml
- Table 2: Rat
Site
Anesthesia
Repeat Bleeds
Expected Volume
Saphenous Vein
No
Yes
300 - 400 ul
Lateral Tail Vein
No
Yes
200 - 400 ul
Distal Tail Transection (less than 3mm)
Required a
Yes
200 - 400 ul
Dorsal Metatarsal
No
Yes
100 - 200 ul
Submandibular / Facial Vein
No
Yes
200 - 500 ul
Jugular Vein
Required
Yes
0.5 - 2.0 ml
Sublingual Vein
Required
Yes
0.5 - 1.0 ml
Retro-Orbital Plexus
Required
Yes - limited
0.5 - 1.0 ml
Cardiac Puncture (Terminal Only)
Required
Terminal
~3 ml
- Table 3: Hamster
Site
Anesthesia
Repeat Bleeds
Expected Volume
Lateral Tarsal Vein
No
Yes
100 - 200 ul
Toenail Clipping
No
Yes
100 - 200 ul
Retro-Orbital Sinus
Required
Yes - limited
100 - 200 ul
Jugular Vein
Required
Yes
0.5 - 2.0 ml
Cardiac Puncture (Terminal Only)
Required
Terminal
~3 ml
- Table 4: Gerbil
Site
Anesthesia
Repeat Bleeds
Expected Volume
Lateral Tail Vein
No
Yes
200 - 400 ul
Toenail Clipping
No
Yes
100 - 200 ul
Distal Tail Transection (less than 3mm)
Required a
Yes (1-2 times only)
100 - 200 ul
Retro-Orbital Sinus
Required
Yes - limited
100 - 200 ul
Jugular Vein
Required
Yes
0.5 - 2.0 ml
Cardiac Puncture (Terminal Only)
Required
Terminal
~3 ml
- Table 5: Guinea Pig
Site
Anesthesia
Repeat Bleeds
Expected Volume
Auricular Vein
No
Yes
50 - 100 ul
Cephalic Vein
No
Yes
50 - 100 ul
Saphenous Vein
No
Yes
400 - 500 ul
Jugular Vein
Recommended
Yes
2 - 3 ml
Cranial Vena Cava
Recommended
Yes
2 - 3 ml
Cardiac Puncture (Terminal Only)
Required
Terminal
- Table 6: Rabbit
Site
Anesthesia
Repeat Bleeds
Expected Volume
Marginal Ear Vein / Central Ear Artery
Local Anesthesia Recommended
Yes
1 - 3 ml
Lateral Saphenous Vein
No
Yes
Cephalic Vein
No
Yes
Jugular Vein
Recommended
Yes
Cardiac Puncture (Terminal Only)
Required
Terminal
- Table 7: Ferret
Site Anesthesia
Repeat Bleeds
Expected Volume
Cephalic Vein
No
Yes
Jugular Vein
Recommended
Yes
Anterior Vena Cava
Recommended
Yes
Cardiac Puncture (Terminal Only)
Required
Terminal
- Table 8: Cat
Site
Anesthesia
Repeat Bleeds
Expected Volume
Medial Saphenous Vein
No
Yes
Cephalic Vein
No
Yes
Jugular Vein
No
Yes
- Table 9: Dog
Site
Anesthesia
Repeat Bleeds
Expected Volume
Lateral Saphenous Vein
No
Yes
Cephalic Vein
No
Yes
Jugular Vein
No
Yes
- Table 10: Sheep
Site
Anesthesia
Repeat Bleeds
Expected Volume
Jugular Vein
No
Yes
Cephalic Vein
No
Yes
Saphenous Vein
No
Yes
- Table 11: Pig
Site
Anesthesia
Repeat Bleeds
Expected Volume
Marginal Ear Vein
No
Yes
Cephalic Vein
No
Yes
Right Jugular Vein
No
Yes
Anterior Vena Cava
Recommended
Yes
- Table 12: Non-Human Primate
Site
Anesthesia
Repeat Bleeds
Expected Volume
Femoral Vein
Recommended
Yes
Saphenous Vein
Required
Yes
Cephalic Vein
Required
Yes
Brachial Vein
Required
Yes
- Table 13: Circulating Blood Volumes in Common Lab Animal Species (adopted from Heinz-Diehl, 2001 and Hawk et al. 2005)
Species
Mean Blood
Volume (ml/kg)Range of Mean
Blood Volume (ml/kg)Mouse
72
63 - 80
Rat
64
58 - 70
Hamster
78
Gerbil
67
Guinea Pig
75 67 - 92 Rabbit
56 44 - 70 Ferret
75 Cat
55 47 - 66 Dog (Beagle)
85 79 - 90 Sheep
66 60 - 74 Minipig
65 61 - 68 Macaque (Rhesus)
56 44 - 67 Macaque (Cynomolgus)
65 55 - 75 Marmoset
70 58 - 82
- 5-10% of body weight = total blood volume
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References
- Clemons DJ, Seeman JL. 2011. The Laboratory Guinea Pig. Boca Raton (FL): CRC Press, p. 89-98.
- Ebert RV, Stead EA, Gibson JG. 1941. Response of normal subjects to acute blood loss. Arch Int Med; 68:578-90.
- BVA/FRAME/RSPCCA/UFAW Joint Working Group on Refinement. 1993. Removal of blood from laboratory mammals and birds (first report). Laboratory Animals; 27:1-22.
- Field KJ, Sibold AL. 1999. The Laboratory Hamster and Gerbil. Boca Raton (FL): CRC Press, p.108-112.
- Hawk CT, Leary SL, Morris TH. 2005. Formulary for Laboratory Animals. Ames (IO): Blackwell Publishing. p. 157.
- Heimann M, Käsermann HP, Pfister R, Roth DR, and Bürki K. 2009. Blood collection from the sublingual vein in mice and hamsters: a suitable alternative to retrobulbar technique that provides large volumes and minimizes tissue damage. Lab Animal; 43: 255-260.
- Heinz-Diehl K, Hull R, Morton D, Pfister R, Rabemampianina Y, Smith D, Vidal JM, Vorstenbosch C. 2001. A good practice guide to the administration of substances and removal of blood, including routes and volumes. J Appl Toxicol; 21:15-23.
- Hoff J. Methods of blood collection in the mouse. 2000. Lab Animal; 29(10):47-53.
- McGuill MW, Rowan AN. 1989. Biological effects of blood loss: Implications for sampling volumes and techniques. ILAR Journal; 31(4):5-20.
- Nerenberg ST, Zedler P, Prasad R, Biskup N, Pedersen L. 1978. Hematological response of rabbits to chronic, repetitive, severe bleedings for the production of antisera. J Immunol Meth; 24:19-24.
- Otto G, Rosenbald WD, Fox JG. 1993. Practical venipuncture techniques for the ferret. Lab Anim 27:26-29
- Scipioni RL, Diters RW, Myers WR, Hart SM. 1997. Clinical and clinicopathologic assessment of serial phlebotomy in the Sprague Dawley rat. Lab Anim Sci; 47(3):293-299.
- Scipioni RL, Guidi DA, Stehr JE, Hart SM, Diters RW. 1996. Clinical, hematologic, and clinicochemical assessment of serial blood sample collection in Sprague-Dawley rats. Contemp Top Lab Anim Sci; 35(6):90. [Abstract]
- Skavlen PA, Baron SJ, Stevens JO. 1992. Effect of blood collection volumes on the hemograms of rabbits. Contemp Top Lab Anim Sci; 31(4):23. [Abstract]
- Villano JS, Sharp PE. 2013. The Laboratory Rat. Boca Raton (FL): CRC press, p. 202-212.
- Yale CE, Torhortst JB. 1972. Critical bleeding and plasma volumes of the adult germfree rat. Lab Anim Sci; 22(4):497-502.
Questions?
Questions or concerns about the content of this document should be directed to the Unit for Laboratory Animal Medicine (ULAM) at (734) 764-0277 or [email protected].
For training on specific blood collection methods and techniques, contact the ULAM Training Core via email at [email protected] or call 734-763-8039.