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I. Purpose

The purpose of this guideline is to provide recommended blood sampling volumes and guidance on a variety of acceptable blood collection techniques in the anesthetized and conscious rodent.

II. Scope

This policy applies to all personnel collecting blood samples from laboratory rodents.

II. Guidance

  1. General Information 
    1. Factors to consider when selecting the appropriate blood collection technique for research purposes include, but are not limited to:
      1. The species to be bled
      2. The size and age of the animal to be bled and the estimated total blood volume
      3. The type of the sample required (e.g. serum, whole blood cells, etc.)
      4. The quality of the sample required (sterility, tissue fluid contamination, etc.)
      5. The quantity of blood required (taking into account extraneous blood loss due to a selected method)
      6. The frequency of sampling
      7. The health status of the animal being bled
      8. The training and experience of the phlebotomist
      9. The size and type of capillary tube is appropriate
      10. The effect of the site, restraint or anesthesia on the blood parameter measured.
    2. The acceptable quantity and frequency of blood sampling is dependent on the circulating blood volume of the animal and the red blood cell (RBC) turnover rate. The approximate circulating blood volume of adult rodents varies with species and body weight. For purposes of calculating the maximum blood volume that may be sampled, the following reference values for total blood volume (TBV) are to be used:
      1. Mouse            72 ml/kg
      2. Rat                  64 ml/kg
      3. Hamster         78 ml/kg
      4. Guinea pig     75 ml/kg
    3. Of the circulating blood volume, approximately 10% of the total volume can be safely removed every 2 to 4 weeks, 7.5% every 7 days, and 1% every 24 hours.
    4. The guidance provided below is for healthy, normal adult animals.  Animals that are young, aged, stressed, have cardiac or respiratory disease, or are otherwise compromised may not be able to tolerate recommended amounts of blood removal.
    5. If the experimental design requires blood volumes and/or frequency of collection that fall outside the recommendations within this guideline, consult the AV and include justification in your protocol for IACUC consideration and approval.
  2. Table 1: Calculated Blood Sample Volumes for Species and Range of Body Weights




    Species Body Weight (g) *CBV (ml) ~1% CBV every 24 hrs.† ~7.5% CBV every 7 days† ~10% CBV every 2 - 4wks†
    Mouse 20 1.10 - 1.40  11 - 14 µl
     
     90 - 105 µl
     
     110 - 140 µl
     
      25 1.37 - 1.75  14 - 18 µl
     
     102 - 131 µl
     
     140 - 180 µl
     
      30 1.65 - 2.10  17 - 21 µl
     
     124 - 158 µl
     
     170 - 210 µl
     
      35 1.93 - 2.45  19 - 25 µl
     
     145 - 184 µl  190 - 250 µl
     
      40 2.20 - 2.80  22 - 28 µl
     
     165 - 210 µl
     
     220 - 280 µl
     
    Rat 125 6.88 - 8.75  69 - 88 µl
     
     516 - 656µl
     
     690 - 880 µl
     
      150 8.25 - 10.50  82 - 105 µl
     
     619 - 788 µl
     
     820 - 1000 µl
     
      200 11.00 - 14.00 110 - 140 µl
     
     825 – 1050 µl
     
     1.1 - 1.4 ml
     
      250 13.75 - 17.50  138 - 175 µl
     
     1.0 – 1.3 ml
     
     1.4 - 1.8 ml
     
      300 16.50 - 21.00  165 - 210 µl
     
     1.2 – 1.6 ml
     
     1.7 - 2.1 ml
     
      350 19.25 - 24.50  193 - 245 µl
     
     1.4 – 1.8 ml
     
     1.9 - 2.5 ml
     
      *Circulating blood volume (1ml = 1000µl) †Maximum sample volume for that sampling frequency
  3. Survival blood sampling
    1. Procedures that may be performed without anesthesia:
      1. Dorsal Pedal/Metatarsal vein
      2. Jugular vein
      3.  Lateral tail vein
      4.  Saphenous/Medial vein
      5.  Submandibular vein
      6. Tail nick
      7. Sublingual vein
    2. Procedures requiring anesthesia:
      1. Retro-orbital This procedure is strongly discouraged by the IACUC as a primary method of blood collection. Scientific justification and IACUC approval are required for use of this collection technique for primary survival sampling.
        1. Only one eye may be sampled at any time. If repeated sampling within an 8 hour period is necessary, the retro-orbital sinus may be resampled by disrupting the blood clot without repeated damage to the sinus.
        2. Alternate between left and right eyes per session.
        3. No more than one collection performed per 5 days.
        4. A maximum of 3 procedures may be performed per eye.
  4. Non-survival (terminal) blood sampling
    1. Terminal blood collection is only to be performed on animals maintained under a surgical plane of anesthesia. Death of the animal must be verified at the completion of the bleed.
    2. An unlimited amount of blood may be withdrawn (i.e., exsanguination) regardless of route. Animal must be immediately euthanized following blood collection.
    3. Terminal blood collection may be performed from the following:
      1. Abdominal vena cava
      2. Renal Vein
      3. Brachial plexus
      4. Cardiac puncture
  5. References
    1. Diehl KH, Hull R, Morton D et al: A Good Practice Guide to the Administration of Substances and Removal of Blood, Including Routes and Volumes.  J Appl Toxicology 21: 15-23, 2001.
    2. Montani, DJ, Cooper, DM: Management of Animal Welfare Issues following Retroorbital Blood Collection in Rats. Techtalk Vol.14/No.3, 2009.
    3. Hawk TR, Leary SL and TH Harris (eds).: Formulary for Laboratory Animals, 3rd edition. Blackwell Publishing, Ames, Iowa, 2005.
    4. McGill MW and AN Rowan. Biological Effects of Blood Loss: Implications for Sampling Volume and Techniques. ILAR News 31(4): 5-18, 1989.
    5. Removal of Blood from Laboratory Mammals and Birds: First Report of the BVA/FRAME/RSPCA/UFAW Joint Working Group on Refinement. Laboratory Animals 27: 1-22, 1993.
    6. The UFAW Handbook on the Care and Management of Laboratory Animals, Volume 1. CRC Press, New York, 2003. “Blood Sampling: pp 379-386.
    7. Raabe BM, Artwohl JE, et al:  Effects of Weekly Blood Collection in C57BL/6 Mice. JAALAS 50(5):680-685, 2011.
    8. Hoff, Janet: Methods of Blood Collection in the Mouse.  Lab Animal 29(10): 47-53, 2000
    9. Scipioni, RL; Diters, RW; et al:  Clinical and Clinicopathological Assessment of Serial Phlebotomy in the Sprague Dawley Rat.  47(3):293-299. 1997
    10. Hui, Yu-hua, Huang NH, et al: Pharmacokinetic comparison of tail-bleeding with cannula- or retro-orbital bleeding techniques in rats using six marketed drugs, Journal of Pharmacological and Toxicological Methods 56:256-264, 2007.
    11. Shirasaki, Y; Ito Y, et al:  Validation Studies on Blood Collection from the Jugular Vein of Conscious Mice: JAALAS, 51(3): 345-351, 2012.
    12. NIH Rodent Blood Collection Guidelines: http://oacu.od.nih.gov/ARAC/documents/rodent bleeding.pdf
    13. Joslin JO. Blood Collection Techniques in Exotic Small Mammals. J. of Exotic Pet Medicine. 18:117-139, 2009.
    14. Gold WT, Gollobin P. and Rodriquez LL. A rapid, simple, and humane method for submandibular bleeding of mice using a lancet. Lab Animal. 34:39-43, 2005.
    15. Beeton C, Garcia A, and Chandy KG. Drawing Blood from Rats through the Saphenous Vein and by cardiac Puncture. J Vis Exp. 7:266, 2007

IACUC Approval Date: 02/19/2020

Review Date: 02/19/2020

Issue Date: 6/22/2020