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Guidelines on Blood Collection

Last Updated 15 April 2016

  1. Purpose

    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. For training on specific blood collection methods and techniques, contact the ULAM Training Core via email at ulam-trainingcore@umich.edu or call 734-763-8039.

    The ULAM veterinary staff provides the following criteria to determine the maximum safe amount of blood to be withdrawn.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 (see below), 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).

  2. Procedures

    1. Approximate Blood Volume
      1. 5-10% of body weight = total blood volume
        1. The circulating blood volume can generally be estimated as 55-70 ml/kg of total body weight. However, care should be taken in these calculations as the % of total blood volume will be lower (-15%) in sick, obese and older animals.
        2. See Table 2 at end of document for some specific blood volumes.
    2. Blood Collection Volumes
      1. 1% of body weight = maximum volume per collection every 14 days, without requiring supplemental fluids. This applies for single blood collections as well as repeated collections. For irregular sampling schedules, calculate the total amount needed over a 14 day span.
      2. 0.07% of body weight = amount that can be taken daily without requiring supplemental fluids
      3. 4-5% of body weight = amount expected at exsanguinations
    3. Single Blood Draw
      1. Maximum of 1% of body weight can be removed as a single blood draw every 14 days, without requiring administration of supplemental replacement fluids. Withdrawing the minimum amount of blood necessary is strongly recommended. Examples:
        1. 0.15 ml from a 15 gm mouse
        2. 3 ml from a 300 gm 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
    4. Multiple Blood Draws
      1. If the total volume withdrawn over a 14-day period is less than 1% BW, then no additional action needs to be taken.
      2. If the total volume withdrawn over a 14 day period is up to 2% BW (or over), fluid volume replacement must be considered. Withdrawing the minimum amount of blood necessary is strongly recommended. Examples:
        1. Up to 0.15 ml withdrawn from a 15 g mouse over 2 weeks is OK
        2. Up to 0.3 ml withdrawn from a 15 g mouse over 2 weeks, replace volume with 0.3 ml saline SQ
        3. Up to 3 ml from a 300 gm rat over 2 weeks is OK
        4. Up to 6ml from a 300 gm rat over 2 weeks, replace volume with 6 ml saline SQ
        5. Up to 200 ml from a 20 kg dog over 2 weeks is OK
        6. Up to 400 ml from a 20 kg dog over 2 weeks, replace volume with 400 ml saline SQ or IV
      3. As a helpful guideline, daily blood draws under 0.07% BW will keep the total 2-week withdrawal under 1% BW.
    5. Exsanguination
      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 UCUCA protocol. Examples:
        1. 0.60-0.75 ml from a 15 gm mouse
        2. 12-15 ml from a 300 gm 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
    6. Fluid Replacement
      1. If the volume of blood removed from an animal exceeds the maximum recommended blood collection volumes (i.e., > 1% body weight every 14 days), replacement of the removed volume of blood with warm (30-35ºC) isotonic solution (e.g., 0.9% saline, lactated Ringer's solution) constitutes accepted veterinary practice. When this volume of blood is collected, it should be withdrawn at a slow, steady rate, and the volume of solution to be infused should be administered similarly.
    7. Monitoring
      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.
    8. Blood Collection Sites and Methods

      Table 1 lists the 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 ulam-vets@umich.edu. 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 – Frequently Used Sites For Blood Draws for Common Lab Animal Species

      Species Site Anesthesia Repeat Bleeds Expected Volume
      Mouse Lateral Tail Vein No Yes 50-100 ul
        Saphenous Vein No Yes 100-200 ul
        Submandibular Vein No Yes 200-500 ul
        Distal Tail Transection (1-3 mm) Required* 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
      Species Site Anesthesia Repeat Bleeds Expected Volume
      Rat Saphenous Vein No Yes 300-400 ul
        Lateral Tail Vein No Yes 200-400 ul
        Distal Tail Transection (1-3 mm) Required* 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
      Species Site Anesthesia Repeat Bleeds Expected Volume
      Hamster 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
      Species Site Anesthesia Repeat Bleeds Expected Volume
      Gerbils Lateral Tail Vein No Yes 200-400 ul
        Toenail Clipping No Yes 100-200 ul
        Distal Tail Transection (1-3 mm) Required* 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
      Species Site Anesthesia Repeat Bleeds Expected Volume
      Guinea pig 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  
      Species Site Anesthesia Repeat Bleeds Expected Volume
      Rabbit 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  
      Species Site Anesthesia Repeat Bleeds Expected Volume
      Ferret Cephalic Vein No Yes  
        Jugular Vein Recommended Yes  
        Anterior Vena Cava Recommended Yes  
        Cardiac Puncture (Terminal Only) Required Terminal  
      Species Site Anesthesia Repeat Bleeds Expected Volume
      Cat Medial saphenous Vein No Yes  
        Cephalic Vein No Yes  
        Jugular Vein No Yes  
      Species Site Anesthesia Repeat Bleeds Expected Volume
      Dog Lateral Saphenous Vein No Yes  
        Cephalic Vein No Yes  
        Jugular Vein No Yes  
      Species Site Anesthesia Repeat Bleeds Expected Volume
      Sheep Jugular Vein No Yes  
        Cephalic Vein No Yes  
        Saphenous Vein No Yes  
      Species Site Anesthesia Repeat Bleeds Expected Volume
      Pig Marginal Ear Vein No Yes  
        Cephalic Vein No Yes  
        Right Jugular Vein No Yes  
        Anterior Vena Cava Recommended Yes  
      Species Site Anesthesia Repeat Bleeds Expected Volume
      Non-Human Primate Femoral Vein Recommended Yes  
        Saphenous Vein Required Yes  
        Cephalic Vein Required Yes  
        Brachial Vein Required Yes  

      *Distal tail transection in gerbils, and adult and 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.

      Table 2 – 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


  3. References

    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.