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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 rodents.

II. Scope

This guideline 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 mean total blood volume (TBV) values may be used:
      1. Mouse            72 ml/kg
      2. Rat                  64 ml/kg
      3. Hamster         78 ml/kg
      4. Guinea pig     75 ml/kg
    3. Approximately 10% of the total blood volume can be safely removed every 2 to 4 weeks, 5% every 7 days, and 1% every 24 hours
    4. The guidance provided below is for healthy adult animals. Animals that are young, aged, stressed, have cardiac or respiratory disease, or are otherwise compromised may not 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, consultation with the AV and justification in the IACUC protocol is required.
       
      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
      90 - 105 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
      8.25 - 10.50
      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
      16.50 - 21.00
      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
  2. Collection site requirement and advantages / disadvantages
    Collection Sites
    ADVANTAGES
    DISADVANTAGES

    Submandibular Sampling
    • Preferred blood collection method
    • Maximum allowable sample volume with minimal trauma
    • Anesthesia recommended
    • Must be securely restrained
    • Yields a large sample so should not be used for frequent small sampling

    Tail Nick or Tail Vein  Sampling
    • Anesthesia not required
    • Multiple samples can be taken
    • Vein is easily accessed
    • Must be securely restrained
    • Yields only small quantities
    Sublingual Vein
    • Multiple samples can be taken
    • Anesthesia required
    • Must be securely restrained
    • Yields a large sample so should not be used for frequent small sampling
    Saphenous Sampling (medial or lateral approach)
    • Excellent technique for serial blood sampling
    • Moderate volume of blood can be collected
    • Multiple samples can be taken by alternating sites
    • Requires specialized training
    • Specialized equipment required
    Cardiac Puncture
    • Maximum volume of blood can be collected
    • Requires deep anesthesia.
    • Non-survival procedure only
    Retro-orbital Sinus
    • Yields a greater volume of blood
    • For multiple sampling, see IACUC standard procedure
    • Requires anesthesia
    • Involves risk of injury to the eye and surrounding structures and therefore use is discouraged.
    • Use must be justified in the protocol.
  3. 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 Blackwell Publishing, Ames, Iowa, 2005.
    4. McGill MW and AN 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 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,
    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 pdf
    13. Joslin 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 Lab Animal. 34:39-43, 2005.
    15. Beeton C, Garcia A, and Chandy Drawing Blood from Rats through the Saphenous Vein and by cardiac Puncture. J Vis Exp. 7:266, 2007
    16. Parasuraman S, Raveendran R, Kesavan R. Blood sample collection in small laboratory animals. J Pharmacol Pharmacother. 2010 Jul;1(2):87-93. doi: 10.4103/0976-500X.72350. Erratum in: J Pharmacol Pharmacother. 2017 Jul-Sep;8(3):153. PMID: 21350616; PMCID: PMC3043327.
    17. Regan RD, Fenyk-Melody JE, Tran SM, Chen G, Stocking KL. 2016: Comparison of Submental Blood Collection with the Retroorbital and Submandibular Methods in Mice (Mus musculus). J Am Assoc Lab Anim Sci. 2016; 55(5):570-6.

IACUC Approval Date: 02/19/2020

Review Date: 2/23/2023

Issue Date: 3/1/2023