Blood Collection in Research Animals

Policy

The method of blood collection, maximum frequency of sampling, minimum interval between samples, and the maximum volume to be obtained must be included within the IACUC-approved protocol. All personnel collecting blood from animals must be appropriately trained. Some methods of collection also require certification.

The actual circulating blood volume varies based on species, age, body condition, and overall health status. For most healthy adult animals, total circulating blood volume is typically 50-80 mL of blood per kg of body weight.

Non-terminal blood collection is limited to 1% of body weight (=10 mL/kg) over any 14-day period. If larger volumes are needed, justification must be provided within the protocol and parenteral fluids should be given when practical. Depending on the species, enhanced monitoring may also be advised.

Terminal blood collections performed under anesthesia from which the animal will not recover for any length of time may exceed the 1% of body weight limit. An animal may be exsanguinated if done as a terminal procedure under a deep plane of anesthesia. Experienced staff typically obtain as much as 3% of body weight during exsanguination.

Bone marrow collection volumes must be included within the blood volume calculation. Each milliliter of bone marrow counts for the same volume of blood obtained.

Examples of Maximum Collection Volumes for Survival Procedures*

Species	Average Adult Body Weight	Total Circulating Blood Volume	Maximum Allowable Blood Draw in 14 days
Hummingbird	10 g	0.6 mL	0.1 mL / 100 µL
Mouse	25 g	1.5 mL	0.25 mL / 250 µL
Rat	300 g	18 mL	3.0 mL
Hamster/Gerbil	100 g	6 mL	1.0 mL
Rabbit	3 kg	180 mL	30 mL
Ferret	1 kg	60 mL	10 mL
Dog	12 kg	720 mL	120 mL
Pig	40 kg	2400 mL	400 mL
NHP	10 kg	600 mL	100 mL

*This table is to be used as a guide for planning purposes only; when calculating allowable volumes, you must use the actual weight of the animal

All personnel collecting blood samples must be appropriately trained or supervised by a qualified trainer. The location from which to obtain blood samples must consider the species of animal, skill of the phlebotomist, required volume to be obtained, prior blood sampling history, demeanor of the animal, and whether sampling will be done awake or sedated/anesthetized. Special consideration must be given to collection procedures likely to pose safety or injury risks to a person or animal, such as blood collection from non-human primates (NHPs). Appropriate chemical restraint may be used, or animals may be acclimated to restraint devices or undergo Positive Reinforcement Training (PRT) to allow for blood collection from conscious animals.

The blood collection method selected should be the most humane and efficient so that any pain, discomfort, or distress is kept to a minimum while adequately fulfilling the study design’s needs. Recommended and alternative sites for commonly used species can be found in the table below. In most species, venous sampling is considered lower risk than arterial sampling. When collecting blood from arterial sites, increased pressure or flow rate can delay clotting and may require that sites are held for longer to ensure hemostasis. For species not included on this table, consult with veterinary services for recommendations.

Common Sites for Blood Collection:

Species	Recommended Sites	Alternative Sites*
Mouse	Submental / Chin Bleed Superficial Temporal Vein (a.k.a., “submandibular” or “facial”) Saphenous Vein Tail Vein Tail Prick	Retro-orbital, anesthetized Retro-orbital, unanesthetized Cardiac (terminal only)
Rat	Tail Vein Tail Prick Saphenous Vein Superficial Temporal Vein (a.k.a., “submandibular” or “facial”	Sublingual Retro-orbital, anesthetized Cardiac (terminal only)
Dog, Cat, NHP	Cephalic Vein Saphenous Vein  Femoral Vein	Jugular Vein Cardiac (terminal only)
Guinea pig (GP), Hamster	Saphenous Vein Cephalic Vein (GP)	Gingival Vein (GP), anesthetized Jugular Vein (GP), anesthetized Cardiac (terminal)
Rabbit	Marginal Ear Vein Cephalic Vein Auricular Artery	Cardiac (terminal)
Swine	Saphenous Vein Ear Vein Mammary Vein Femoral Vein	Jugular Vein Vena Cava Cardiac (terminal)
Bird	Brachial/wing vein	Jugular vein Cardiac (terminal)
Ferret	Cephalic vein Saphenous vein,  Femoral vein	Cranial Vena Cava Jugular vein Cardiac (terminal only)

*In general, alternative sites (or other sites not listed) may require justification within the IACUC protocol as they may pose increased risk to the animal, have specific advanced training requirements, or may have other limitations as noted in species-specific sections below.

Animal Monitoring:

Animals undergoing frequent or large volume blood draws should have hematocrit (Hct) and/or packed cell volume (PCV) monitored regularly when practical. While healthy adult animals can recover their blood volume within 24 hours, it may take up to 2 weeks for other blood constituents (red blood cells, proteins, clotting factors, etc.) to be replenished. When assessed at least 24 hours after blood loss, the Hct or PCV allows for evaluation of recovery from blood draws over time. For most species, normal PCV ranges from 35-45%. If the PCV drops, it may be unsafe to collect additional blood and future blood draw schedules should be altered accordingly. Hemoglobin ranges are more species-specific but generally range from 10-20 g/dL. Because hemoglobin is critical for regeneration of red blood cells, decreases of hemoglobin should also be an indicator to adjust and/or cancel future blood collections.

When exceptions to volume limits within this policy are requested, enhanced monitoring is expected. Parenteral fluids are recommended in most cases. The protocol must specify monitoring parameters for development of anemia (e.g., skin pallor, weakness, apparent dizziness, hematologic values), frequency of monitoring for anemia, and a response if anemia does develop. For studies in which anemia is expected and unavoidable, blood transfusion criteria should be included. Generally, once anemia develops, future blood draws should be adjusted or cancelled until the animal has recovered. Minimal blood collection (e.g. using low volume microtainers) to monitor the clinical condition of the animal may need to continue under the direction of the clinical veterinarian.

Mouse Blood Collection

Table I: Summary of Blood Collection Techniques in Mice  

MICE
Route	Anesthesia Required?	Certification Required?	Expected Volume (µl)	Comments
Submental (chin)	No	No	200	rapid
Lateral/medial saphenous vein	No	No	50-200	not as rapid as other techniques, low potential for tissue damage, larger volumes are challenging to obtain
Facial Vein (submandibular)	No	No	200	rapid, local tissue trauma seen
Tail Prick/Tail Vein	No	No	20	useful for small volume collection with minimal restraint
Retro-orbital	Recommended	Yes	200	rapid, potential for complications, anesthesia strongly recommended
Cardiac	Yes	No	200-500	terminal use only

*In general, alternative sites (or other sites not listed) may require justification within the IACUC protocol as they may pose increased risk to the animal, have specific advanced training requirements, or may have other limitations as noted in species-specific sections below.

Generally, volumes of up to 150 microliters can be readily collected from the lateral or medial saphenous vein. It is also possible to obtain small amounts of blood by tail vein prick with minimal restraint. Volumes of up to 1% of the body weight are readily and rapidly obtained from the retro-orbital sinus, submental region (chin), and facial vein. Cardiac blood collection is performed when the mouse is in a deep plane of anesthesia and is a terminal (non-recovery) procedure. For repeated blood collection, indwelling catheter placement is recommended.

Retro-orbital blood collection is a procedure used in mice to obtain small blood samples. Retro-orbital blood collection may be conducted in awake mice, but general anesthesia is strongly encouraged if compatible with experimental objectives. A minimum healing time of 7 days between sampling from the same orbital sinus is required, and alternate eyes should be used for sequential blood collection if more frequent sampling is needed. Retro-orbital blood collection requires a high amount of technical proficiency and thus has a greater potential than other blood collection routes to result in complications such as corneal and ocular damage. Trauma to the eye may require euthanasia of the animal. Retro-orbital bleeding requires certification by CLATR and may only be performed by certified individuals.

Tail transection (also referred to as tail snip, tail tip amputation, or tail clip) is NOT considered a routine method of blood collection for mice. If necessary, this requires anesthesia, scientific justification, and should be described within the protocol as a procedure with associated monitoring and pain management where appropriate. It may only be performed a single time per animal and less than 1 millimeter of tail may be removed. If genotyping by tail snip was performed previously, this method may not be used for blood collection later in life.

Rat Blood Collection

Table II: Summary of Blood Collection Techniques in Rats

Route	Anesthesia Required?	Certification Required?	Comments
Lateral tail vein or ventral tail artery	No	No	repeatable, simple
Dorsal metatarsal or lateral saphenous	No	No	not as rapid as other techniques, low potential for tissue damage
Retro-orbital	Yes	Yes	rapid, potential for complications, anesthesia strongly recommended
Jugular	Recommended	No	limited application, poor for repeated sampling and technically difficult
Cardiac	Yes	No	Terminal use only

*In general, alternative sites (or other sites not listed) may require justification within the IACUC protocol as they may pose increased risk to the animal, have specific advanced training requirements, or may have other limitations as noted in species-specific sections below.

Rats:

Generally, volumes up to 1 mL can be obtained from the lateral tail vein with minimal restraint. The lateral saphenous or the dorsal metatarsal vein can be used as well. Volumes of up to 1% of the body weight can also be obtained from the jugular vein but require a high degree of technical proficiency and are considered higher risk. Cardiac blood collection is performed when the rat is in a deep plane of anesthesia and as a terminal (non-recovery) procedure. For repeated blood sampling, indwelling catheter placement is recommended.

Retro-orbital blood collection is a procedure used in rats to obtain small blood samples. In rats, this method requires anesthesia. Volumes of up to 1% of the body weight may be obtained from retro-orbital plexus (0.5 to 3 mL depending on the size of the rat) in anesthetized rats only. A minimum healing time of 7 days between sampling from the same orbital plexus is required, and alternate eyes should be used for sequential blood collection if more frequent sampling is needed Retro-orbital blood collection requires a high amount of technical proficiency and thus has a greater potential than other blood collection routes to result in complications such as corneal and ocular damage. Trauma to the eye may require euthanasia of the animal. Retro-orbital bleeding requires certification by CLATR and may only be performed by certified individuals.

Tail transection (also referred to as tail snip, tail tip amputation, or tail clip) may be used in rats when indicated by study design (e.g. very frequent small volume collection over a short period of time). General anesthesia is recommended for the initial transection; subsequent samples can be obtained by scab removal without anesthesia. The bones in the tail must always be avoided and only the fleshy tail tip transected (<2 mm). In most animals, this may only be done one time without risking bone involvement. If genotyping by tail snip was performed previously, it may not be possible to transect additional tail without impacting bone so alternative blood collection methods may need to be considered.