Guidelines on Blood Collection

1. 5-10% of body weight = total blood volume
   1. In healthy animals the circulating blood volume can generally be estimated as 55-70 ml/kg of the total body weight.
   2. 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.
   3. See Table 13 at end of this document for average blood volumes in certain species.

1. 1% of body weight (BW) = maximum volume that can be collected every 14 days, without requiring supplemental fluids. Examples:
   1. 0.15 ml from a 15 g mouse
   2. 3 ml from a 300 g rat
   3. 50 ml from a 5 kg cat
   4. 100 ml from a 10 kg monkey
   5. 400 ml from a 40 kg dog
2. 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:
   1. 0.07% of body weight maximum per collection if 14 samples are needed within a 14-day period.
   2. 0.1% of body weight maximum per collection if 10 samples are needed within a 14-day period.
   3. 0.14% of body weight maximum per collection if 7 samples are needed within a 14-day period.
   4. 4-5% of body weight = amount expected at exsanguinations (see Section 4)

1. If the total volume withdrawn over a 14-day period is greater than 1% BW, fluid volume replacement must be considered.
2. 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.
3. Examples of Fluid Replacement
   1. For a 15 g mouse:
      1. Up to 0.15 ml (1% withdrawn) of blood collected over 14 days does not require fluid replacement.
      2. Up to 0.3 ml (2% withdrawn) of blood collected over 14 days, requires fluid replacement with 0.3 ml appropriate fluids SQ.
   2. For a 300 g rat:
      1. Up to 3 ml (1% withdrawn) of blood collected over 14 days does not require fluid replacement.
      2. Up to 6 ml (2% withdrawn) of blood collected over 14 days, requires fluid replacement with 0.3 ml appropriate fluids SQ.
   3. For a 20 kg dog:
      1. Up to 200 ml (1% withdrawn) of blood collected over 14 days does not require fluid replacement.
      2. Up to 400 ml (2% withdrawn) of blood collected over 14 days, requires fluid replacement with 0.3 ml appropriate fluids SQ or IV.

1. 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:
   1. 0.60-0.75 ml from a 15 g mouse
   2. 12-15 ml from a 300 g rat
   3. 200-250 ml from a 5 kg cat
   4. 400-500 ml from a 10 kg monkey
   5. 1600-2000 ml from a 40 kg dog

1. 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:
   1. Fast and thready pulse
   2. Pale dry mucous membranes
   3. Cold skin and extremities
   4. Restlessness
   5. Hyperventilation
   6. Sub-normal body temperature
2. If 15-20% of total blood volume is removed, cardiac output and blood pressure will be reduced.
3. If 30-40% of total blood volume is removed, death will result in at least 50% of animals.
4. If > 40% of total blood volume is removed, death of the animal is expected.
5. Stressed, sick, or otherwise compromised animals may not tolerate the blood collection criteria noted above, which is for healthy animals.
6. 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.

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.

1. ​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)	Requireda	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

   ^aDistal tail transection in gerbils, and adult rats and mice (>21 days) requires the use of general anesthesia and preemptive analgesia (e.g., NSAIDs, opioids) unless scientifically justified and approved in the IACUC protocol.

2. 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)	Requireda	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

   ^aDistal tail transection in gerbils, and adult rats and mice (>21 days) requires the use of general anesthesia and preemptive analgesia (e.g., NSAIDs, opioids) unless scientifically justified and approved in the IACUC protocol.

3. 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

4. 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)	Requireda	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

   ^aDistal tail transection in gerbils, and adult rats and mice (>21 days) requires the use of general anesthesia and preemptive analgesia (e.g., NSAIDs, opioids) unless scientifically justified and approved in the IACUC protocol.

5. 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

6. 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

7. 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

8. Table 8: Cat
   

   Site	Anesthesia	Repeat Bleeds	Expected Volume
   Medial Saphenous Vein	No	Yes
   Cephalic Vein	No	Yes
   Jugular Vein	No	Yes

9. Table 9: Dog
   

   Site	Anesthesia	Repeat Bleeds	Expected Volume
   Lateral Saphenous Vein	No	Yes
   Cephalic Vein	No	Yes
   Jugular Vein	No	Yes

10. Table 10: Sheep
    

    Site	Anesthesia	Repeat Bleeds	Expected Volume
    Jugular Vein	No	Yes
    Cephalic Vein	No	Yes
    Saphenous Vein	No	Yes

11. 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

12. 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

13. 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

1. Clemons DJ, Seeman JL. 2011. The Laboratory Guinea Pig. Boca Raton (FL): CRC Press, p. 89-98.
2. Ebert RV, Stead EA, Gibson JG. 1941. Response of normal subjects to acute blood loss. Arch Int Med; 68:578-90.
3. 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.
4. Field KJ, Sibold AL. 1999. The Laboratory Hamster and Gerbil. Boca Raton (FL): CRC Press, p.108-112.
5. Hawk CT, Leary SL, Morris TH. 2005. Formulary for Laboratory Animals. Ames (IO): Blackwell Publishing. p. 157.
6. 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.
7. 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.
8. Hoff J. Methods of blood collection in the mouse. 2000. Lab Animal; 29(10):47-53.
9. McGuill MW, Rowan AN. 1989. Biological effects of blood loss: Implications for sampling volumes and techniques. ILAR Journal; 31(4):5-20.
10. 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.
11. Otto G, Rosenbald WD, Fox JG. 1993. Practical venipuncture techniques for the ferret. Lab Anim 27:26-29
12. 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.
13. 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]
14. 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]
15. Villano JS, Sharp PE. 2013. The Laboratory Rat. Boca Raton (FL): CRC press, p. 202-212.
16. Yale CE, Torhortst JB. 1972. Critical bleeding and plasma volumes of the adult germfree rat. Lab Anim Sci; 22(4):497-502.