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Blood Collection Guidelines (IACUC)

Last updated on January 7, 2025 9 min read Blood Collection Guidelines (IACUC)

Purpose

Boston University (BU) is committed to observing Federal policies and regulations and the Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC) International standards for the humane care and use of animals. The intent of this policy is to provide guidance on blood collection in laboratory animals. Adherence to this policy is mandatory unless a specific exception has been approved by the IACUC.

Covered Parties

This policy is applicable to all persons responsible for conducting research, teaching, training, breeding, and related activities, hereinafter referred to collectively as “activities”, involving live vertebrate animals conducted at or under the auspices of Boston University.

University Policy

All non-terminal blood collection without replacement fluids is limited to 10% of the total circulating blood volume of a healthy animal at any single draw or cumulatively over 14 days. On average, the total circulating blood volume is equal to 5.5 -7.0 % (~66 ml/kg) of the animal’s body weight. If larger amounts are needed, then up to 15% of the total circulating blood volume may be withdrawn if replacement fluids are given at the time of blood withdrawal. Where species-specific literature is available, it may be used as scientific justification for larger or more frequent volumes. 1,2 Removal of 15% of total blood volume must be justified in the IACUC protocol and approved by the IACUC.

Example: a 4 kg rabbit is calculated to have a total blood volume of 264 ml (66 ml/kg x 4.0 kg). Thus, 26 ml (10% of 264 ml) may be collected without giving replacement fluids and 40 ml (15% of 264 ml) may be collected if replacement fluids are given within a 14-day period.

The limitations for blood collection preserve the health status of the animal and maintain the validity of experimental results based on blood samples. The guidelines provided are for healthy, normal adult animals. Animals that are young, aged, stressed, have undergone experimental manipulations, or are suffering from cardiac or respiratory disease may not be able to tolerate this amount of blood loss.

Blood sample volume guidelines for rodents

Table 1: Approximate Blood Sample Volumes for a Range of Body Weights 3

Body weight (g)*CBV(ml)1% CBV (ml)

every 24 hrs†

7.5% CBV (ml)

every 7 days†

10% CBV (ml)

every 14 days†

201.10 – 1.40.011 – .014.082 – .105.11 – .14
251.37 – 1.75.014 – .018.10 – .13.14 – .18
301.65 – 2.10.017 – .021.12 – .16.17 – .21
351.93 – 2.45.019 – .025.14 – .18.19 – .25
402.20 – 2.80.022 – .028.16 – .21.22 – .28
1256.88 – 8.75.069 – .088.52 – .66.69 – .88
1508.25 – 10.50.082 – .105.62 – .79.82 – 1.0
20011.00 – 14.00.11 – .14.82 – 1.051.1 – 1.4
25013.75 – 17.50.14 – .181.0 – 1.31.4 – 1.8
30016.50 – 21.00.17 – .211.2 – 1.61.7 – 2.1
35019.25 – 24.50.19 – .251.4 – 1.81.9 – 2.5

*Circulating blood volume

†maximum sample volume for that sampling frequency

Blood sample volumes for other lab species 4

SpeciesReference weight (g)Blood volume (ml/kg)Total blood volume (TBV), normal adult (ml)10% TBV (ml)
Hamster85-15078Male 6.3 – 9.7

Female 7.1 – 11.2

Male 0.6 – 0.9

Female 0.7 – 1.1

Guinea pig400-9007028 – 632.8 – 6.3
Rabbit1000 – 600057 – 6558.5 – 5855 – 50
NHP (rhesus)55 – 80Male 420 – 770

Female 280 – 630

Male 42 – 77

Female 28 – 63

NHP (cynomolgus)50 – 96Male 280 – 560

Female 140 – 420

Male 28 – 56

Female 14 – 42

Normal Packed Cell Volume (PCV) for some lab animals (%)

Mouse39-49
Rat36-54
Hamster39-59 [5]
Chinchilla30-55 [6]
Guinea pig37-48[13]
Rabbit31-50 [7]
Rhesus26-48 [8]
Cynomolgus31-57 [9]
African Green Monkey33-54 [10]

Monitoring

If the animal is being bled routinely, the packed cell volume (PCV) should be checked frequently (e.g. weekly) to determine whether blood collection should be suspended due to anemia. While healthy adult animals can recover their blood volume within 24 hours. It may take up to 2 weeks for other blood constituents (RBCs, proteins, clotting factors) to be replaced.
By monitoring the hematocrit (Hct or PCV) and/or hemoglobin of the animal. It is possible to evaluate whether an animal has sufficiently recovered from a single or multiple blood draws. After sudden to acute blood loss, it takes up to 24 hours for hematocrit or hemoglobin to reflect this loss. For many lab species, a PCV <35% or hemoglobin concentration <10g/dL would indicate it is not safe to remove blood.

Blood collection sites in mice and rats 11

The following Guidelines refer to the most frequently used survival sampling sites: a) retroorbital b) facial vein c) saphenous vein d) tail veins e) jugular vein. Blood withdrawal by cardiac puncture is considered a terminal procedure and should only be performed once an animal is under general anesthesia.

Information to guide the choice of blood collection site or method:

Facial vein

Blood collection from the maxillary facial vein is a safe and fast technique in mice. It requires momentary restraint and approximately 200ul of blood can be obtained easily from a healthy adult mouse. The vessel is located just beneath the skin immediately caudal to the facial vibrissae (whiskers) at the corner of the jaw. Repeated sampling is possible by alternating sides of the face. Materials needed include a 20 or 22 g hypodermic needle or lancet manufactured for this purpose, blood collection tubes and sterile gauze. Training is necessary before this procedure is performed.

Submental

Must be performed in anesthetized animals (suggest isoflurane). Scruff the mouse so that the skin across the chin (submental region) is taut. Apply sterile lubricant as needed to visualize vessels. Prick the blood vessel using a sterile lancet or needle. Collect sample with a pipette or other tube. This method is normally good for obtaining a moderate sample volume (100-200uL). 12

Lateral tail vein or ventral tail artery

  • Can be used in both rats and mice by cannulating the blood vessel, or, by superficially nicking the vessel perpendicular to the tail.
  • Sample collection by nicking the vessel is easily performed in both species, but produces a sample of variable quality that may be contaminated with tissue and skin products.
  • Repeated collections possible. With tail vein nicking, the clot/scab can be gently removed for repeated small samples if serial testing is required (e.g., glucose measures, etc.)
  • Tail artery sampling yields larger volumes but requires the animal to be anesthetized and placed in dorsal recumbency. Good hemostasis is also required, as always whenever an artery is incised.

Saphenous vein (medial or lateral approach)

Can be used in both rats, mice, and other rodents by piercing the saphenous vein with a needle or combination needle/collection tube.

Jugular vein sampling (rat, hamster, larger rodents)

  • Blood volume obtained: medium to large.
  • High quality sample.
  • Jugular sampling can be conducted without anesthesia, although the use of anesthesia greatly facilitates the procedure.
  • Does not easily lend itself to repeated serial sampling.

Retro-orbital sinus/plexus sampling

  • Retro-orbital sampling can be used in mice, rats, and hamsters (though usually not a method of choice in the rat) by penetrating the retro-orbital sinus in mice or plexus in rats and hamsters with a glass capillary tube or Pasteur pipette.
  • Repeat sampling from the same orbit may be difficult (10 days to 2 weeks recommended between successive bleeds).

Restraint

Animals will need to be physically restrained to prevent any movement that would result in lacerating the blood vessel or other potentially serious complications. Blood may be collected from awake animals that are appropriately restrained provided that persons performing the procedure are skilled.

Anesthesia

Anesthesia is required if blood collection is being performed either via the retro-orbital sinus or by cardiac puncture due to the distress and pain which can be caused and for the serious complications (injury to the eye, cardiac tamponade and death) associated with these routes. For survival procedures requiring anesthesia isoflurane is recommended as it is short-acting and allows replacing the rodent in its cage within minutes. Cardiac puncture is only permitted as a terminal procedure and only after the animal is in a surgical (deep) plane of anesthesia.

Common sites for blood collection in other lab species

SpeciesSites of collection and permitted conditions
HamsterSaphenous vein, sublingual vein, retro-orbital sinus, gingival vein, jugular vein, cranial vena cava, cardiac (under anesthesia as a terminal procedure only)
ChinchillaSuperior sagittal or transverse sinus, jugular vein, cranial vena cava, saphenous vein
Guinea pigCranial vena cava/jugular vein, saphenous vein, tarsal vein, gingival sinus, cardiac (under anesthesia as a terminal procedure only)
RabbitCardiac (under anesthesia as a terminal procedure only), jugular vein, marginal ear vein (for small volume only), ear artery (requires good hemostasis), cephalic vein, saphenous vein
Nonhuman PrimatesFemoral, cephalic veins, saphenous vein

Fluid replacement

Lactated Ringer’s Solution (LRS) is recommended as the best for fluid replacement. For mice, administer 1 ml of warmed LRS IP or SC. For rats, administer 5 -10 ml warmed LRS ½ via IP and ½ via SC administration. For larger species, replacement fluid (isotonic crystalloid like LRS or 0.9% saline) may be given SC or IV under advisement of the veterinarian or in line with the study protocol. Note that phosphate-buffered saline (PBS) has been shown to be mildly inflammatory when used for peritoneal lavage in rats and should not be used for fluid replacement unless scientifically justified. 13

Nutritional supplementation

When larger volumes are withdrawn, especially when there is repeated sampling, it is recommended that rodents receive Nutrical or diet gel as a dietary supplement. For rats and mice, this can easily be done by smearing Nutrical on a few pellets or placing an opened container of diet gel on the cage floor. Larger, non-rodent species may also be offered easily-digestible and appealing food supplementation.

Training

Training is required for blood collection in any species and by any route. Please contact the BU ASC to schedule training.

Responsible Parties

Principal Investigators are responsible for: preparing and submitting IACUC applications; making modifications in applications in order to secure IACUC approval; ensuring adherence to approved protocols; ensuring humane care and use of animals; ensuring that all personnel have completed required training; and reporting any adverse events to the IACUC.

The Animal Welfare Program and the Institutional Animal Care and Use Committee are responsible for overseeing policy implementation and ensuring compliance with this policy.


[1] Hobbs, T. R., Blue, S. W., Park, B. S., Greisel, J. J., Conn, P. M., & Pau, F. K. (2015). Measurement of Blood Volume in Adult Rhesus Macaques (Macaca mulatta). Journal of the American Association for Laboratory Animal Science : JAALAS54(6), 687–693.

[2] Adams, C. R., Halliday, L. C., Nunamaker, E. A., & Fortman, J. D. (2014). Effects of weekly blood collection in male and female cynomolgus macaques (Macaca fascicularis). Journal of the American Association for Laboratory Animal Science : JAALAS53(1), 81–88.

[3] NIH (2019). Guidelines for Blood Collection in Mice and Rats. https://oacu.oir.nih.gov/system/files/media/file/2021-02/b2_blood_collection_in_mice_and_rats.pdf. Accessed November 17, 2021.

[4] NC3Rs. Blood sample volumes. Adapted from Wolfensohn & Lloyd, 2003, Handbook of Laboratory Animal Management and Welfare, 3rd Edition. https://www.nc3rs.org.uk/blood-sample-volumes. Accessed Nov 17 2021.

[5] Washington, I. M., & Van Hoosier, G. (2012). Clinical Biochemistry and Hematology. The Laboratory Rabbit, Guinea Pig, Hamster, and Other Rodents, 57–116. https://doi.org/10.1016/B978-0-12-380920-9.00003-1

[6] Washington, I. M., & Van Hoosier, G. (2012). Clinical Biochemistry and Hematology. The Laboratory Rabbit, Guinea Pig, Hamster, and Other Rodents, 57–116. https://doi.org/10.1016/B978-0-12-380920-9.00003-1

[7] Washington, I. M., & Van Hoosier, G. (2012). Clinical Biochemistry and Hematology. The Laboratory Rabbit, Guinea Pig, Hamster, and Other Rodents, 57–116. https://doi.org/10.1016/B978-0-12-380920-9.00003-1

[8] Koo, BS., Lee, DH., Kang, P. et al. Reference values of hematological and biochemical parameters in young-adult cynomolgus monkey (Macaca fascicularis) and rhesus monkey (Macaca mulatta) anesthetized with ketamine hydrochloride. Lab Anim Res 35, 7 (2019). https://doi.org/10.1186/s42826-019-0006-0

[9] Koo, BS., Lee, DH., Kang, P. et al. Reference values of hematological and biochemical parameters in young-adult cynomolgus monkey (Macaca fascicularis) and rhesus monkey (Macaca mulatta) anesthetized with ketamine hydrochloride. Lab Anim Res 35, 7 (2019). https://doi.org/10.1186/s42826-019-0006-0

[10] Liddie, S., Goody, R.J., Valles, R. and Lawrence, M.S. (2010), Clinical chemistry and hematology values in a Caribbean population of African green monkeys. Journal of Medical Primatology, 39: 389-398. https://doi.org/10.1111/j.1600-0684.2010.00422.x

[11] Parasuraman, S., Raveendran, R., & Kesavan, R. (2010). Blood sample collection in small laboratory animals. Journal of pharmacology & pharmacotherapeutics, 1(2), 87–93. https://doi.org/10.4103/0976-500X.72350

[12] Regan RD, Fenyk-Melody JE, Tran SM, Chen G, Stocking KL. 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. PMID: 27657712; PMCID: PMC5029828.

[13] Adapted from Formulary for Laboratory Animals, Hawk, Leary, and Morris 2005

History

Effective Date: 1/7/2025
Next Review Date: 1/6/2028

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