Guidelines for Blood Collection in Laboratory Animals (2023)


This document is to provide guidance to investigators regarding safe blood collection volumes in 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, should be listed in the approved protocol specific to each study.


The method and volume of blood to be collected will depend on the animal species, frequency of collection, and experimental needs. Researchers should plan and perform each sampling protocol with an appreciation for the stresses associated with blood loss to the animal and do whatever they can to minimize the animal's reaction to the stress. Careful planning and control of blood sampling and all experimental variables associated with it should not only benefit the welfare of the animal, but also minimize confounding influences on research data. (7)

Training and experience of the individual in the chosen procedure are of critical importance. The amount of training and practice required to achieve a given level of competence in a particular technique varies from individual to individual depending on, for example, manual dexterity, prior experience, attitude, and the skills of the instructor.

DLAR has a number of training resources available.

Please contact our Training Coordinator for details or to schedule a session.


Recommended volumes for blood collection are intended to preserve the health status of the animal and maintain the validity of experimental results. 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 other disease conditions may not be able to tolerate these recommended blood volume withdrawals .



On average, the total circulating blood volume is equal to 5.5 -8.0 % of the animal’s body weight. Non-terminal blood collection without additional monitoring (see below) should be limited to 10% of the total circulating blood volume on a single collection or every 2 week period for serial collections.

Example (Using mean blood volume table below): a 4 kg rabbit is calculated to have a total blood volume of 224 ml (56 ml/kg x 4.0 kg). Thus, 22.4 ml (10% of 224 ml) may be collected without giving replacement fluids once every two weeks.

Estimated Total Blood Volume and Safe Bleeding Volume of Selected Species:

SpeciesBlood volume (ml/kg)One Bleeding -
max - 10% of blood volume (ml/kg)
Dog8686 (79-90)8.6
Guinea Pig7575 (67-92)7.5
Minipig6565 (61-68)6.5
(Cynomolgous Macaque)
6565 (55-75)6.5
(Rhesus Macaque)
5454 (44-67)5.4
Mouse7979 (63-80)7.9
Pig6565 (61-68)6.5
Rabbit5656 (44-70)5.6
Rat6464 (58-70)6.4
Sheep6666 (60-74)6.6

Source: Adapted from Formulary for Laboratory Animals, Hawk, Leary, and Morris 2005

Note: If the amount of blood volume removed is 7.5% of total blood volume, allow 1 week recovery; if amount removed is 10%, allow 2 weeks recovery; if amount removed is 15%, allow 4 weeks recovery.

Common Sites for Blood Collection:

SpeciesRecommended Sites & Conditions
MouseSuperficial temporal vein (a.k.a., "submandibular" or "facial"), saphenous vein, tail vein, retro-orbital (anesthetized), cardiac (anesthetized, terminal)
RatTail vein, saphenous vein, superficial temporal vein (a.k.a., "submandibular" or "facial"), cardiac (anesthetized, terminal), sublingual, jugular
Dog, Cat, Non-human PrimateCephalic, saphenous veins, femoral and jugular viens
Guinea pig, HamsterSaphenous, cardiac (anesthetized, terminal)
RabbitMarginal ear vein, cardiac (anesthetized, terminal)
SwineJugular, ear vein
Pigeon, QuailBrachial/wing vein
ChickenBrachial/wing vein, jugular
Ruminant, EquineJugular

Guidelines for Rodents:

Similarly, 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. Volumes greater than recommended should be scientifically justified and appropriate fluid and/or cellular replacement provided. Approximate blood sample volumes for a range of body weights are included in the table below:

Body Weight (g)*CBV (ml)1% CBV (ml)
every 24 hours**
7.5% CBV (ml)
every 7 days**
10% CBV (ml)
every 2-4 weeks**
*Circulating Blood Volume** Maximum sample volume for that sampling frequency

unanesthetized collection methods

Blood Collection methods in Mice and Rats (unanesthetized):

Superficial temporal (a.k.a. “Submandibular” or “Facial” vein/artery) Sampling:

  • Obtainable blood volumes: medium to large.
  • Repeated sampling is possible by alternating sides of the face.
  • Sample may be a mixture of venous and arterial blood.
  • Requires less hands-on training than tail or retro-orbital sampling to reliably withdraw a reasonable quantity of blood.
  • Perform on awake animals to achieve proper restraint, which in turn results in proper site alignment and venous compression for good blood flow.
  • Can be performed rapidly and with a minimal amount of equipment, allowing for rapid completion.
  • Sample volume can be partially controlled with the size of needle (20 gauge or smaller) or lancet (4 mm) used to puncture the site.
  • Use of a lancet is recommended to control depth of puncture and reduce potential for complications. These can be significant if puncture is too deep or homeostasis is not assured prior to returning the mouse to its cage (5).

Saphenous Sampling (medial or lateral approach):

  • Obtainable blood volumes: small to medium.
  • Can be used in both rats and mice by piercing the saphenous vein with a needle or lancet.
  • Variable sample quality.
  • The procedure is customarily done on an awake animal but effective restraint is required.
  • Requires more hands-on training than tail or retro-orbital sampling to reliably withdraw more than a minimal amount of blood.
  • Although more aesthetically acceptable than retro-orbital sampling, prolonged restraint and site preparation time can result in increased animal distress when handling an awake animal.
  • Temporary favoring of limb may be noted following the procedure.
  • Application of sterile petroleum jelly to the site facilitates blood droplet formation, which can enhance the total blood volume captured.
  • The clot/scab can be gently removed for repeated small samples if serial collection is required

Lateral Tail Vein:

  • Obtainable blood volumes: small to medium.
  • 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. Sample quality decreases with prolonged bleeding times and “milking” of the tail.
  • Sample collection using a needle (cannulation) minimizes contamination of the sample, but is more difficult to perform in the mouse.
  • Repeated collections possible. With tail nicking, the clot/scab can be gently removed for repeated small samples if serial testing is required (e.g., glucose measures, etc.)
  • In most cases warming the tail with the aid of a heat lamp or warm compresses will increase obtainable blood volume.
  • Cannulation and tail nicking are routinely done without anesthesia, although effective restraint is required.

Jugular Sampling (typically limited to the rat, but can be performed in mice as well (6):

  • Obtainable blood volumes: medium to large.
  • May be needed when a larger blood volume withdrawal and survival are needed.
  • Results in high quality sample.
  • Jugular sampling can be conducted without anesthesia in rats, although the use of anesthesia greatly facilitates the procedure and reduces the potential for injury to the animal as well as the individual (e.g., bites, needle stick).
  • Does not easily lend itself to repeated serial sampling
(Video) Blood collection by MSR

anesthetized collection methods

Blood Collection methods in Mice and Rats (anesthetized):

NB: The following methods require anesthesia (local and/or general depending on species and technique) to relieve pain and distress associated with the technique or for restraint. Use of these methods may require scientific justification for why less painful techniques cannot be used in the animal care and use protocol.

Retro-orbital Sinus/Plexus Sampling:

(Video) Site for Blood Collection in Animals

  • Obtainable volume: medium to large.
  • Retro-orbital sampling can be used in both mice and rats (though not a preferred method in the rat) by penetrating the retro-orbital sinus in mice or plexus in rats with a capillary tube or Pasteur pipette.
  • Rapid – large number of animals can be bled within a short period of time.
  • Good sample quality. Potential contamination with topical anesthetic, if used, should be taken into account.
  • A minimum of 10 days should be allowed for tissue repair before repeat sampling from the same orbit. Otherwise the healing process may interfere with blood flow.
  • Alternating orbits should not be attempted until the phlebotomist is proficient in obtaining samples from the orbit accessed most readily by the dominant hand ie a right handed individual should gain proficiency withdrawing samples from the right orbit before attempting to obtain samples from the left orbit.
  • In the hands of an unskilled phlebotomist, retro-orbital sampling has a greater potential than other blood collection routes to result in complications.
  • General anesthesia must be used unless scientific justification is provided and approved by the IACUC. In addition, a topical ophthalmic anesthetic, e.g. proparacaine or tetracaine drops, is recommended prior to the procedure.
  • In both mice and rats, care must be taken to ensure adequate hemostasis following the procedure.
  • Use of sterile capillary tubes and pipettes are recommended for use to help avoid periorbital infection and potential long-term damage to the eye. The edges of the tubes should be checked for smoothness to also decrease likeliness of eye damage.
  • Animals must be monitored following collection for damage to the orbit or globe of the eye. If damage is noted, veterinary staff should be contacted for treatment options up to and including euthanasia.

Tail Clip Sampling:

  • Obtainable volume: small.
  • Can be used in both rats and mice by clipping (e.g. amputating) no more than 1mm of the distal tail in mice or 2 mm in rats
  • Produces a sample of variable quality that may be contaminated with tissue and skin products. Sample quality decreases with prolonged bleeding times and “milking” of the tail.
  • Obtainable volume: small.
  • Repeated collections possible. The clot/scab can be gently removed for repeated small samples if serial testing is required (e.g., glucose measures, etc.)
  • In most cases warming the tail with the aid of a heat lamp or warm compresses will increase obtainable blood volume.
  • When performing tail clipping, consideration should be given to anesthesia/analgesia, particularly if the tail has been previously clipped for genotyping. If a topical hypothermic anesthetic is used, blood will flow as the tail re-warms. If a local anesthetic is applied, adequate contact time should be allowed for it to take effect.



If the animal is being bled routinely, the red blood cell packed volume (PCV) should be checked weekly to determine when blood collection should be suspended in order for the animal to recover from potential anemia. While healthy adult animals can recover their blood volume within 24 hours, it may take up to 2 weeks for all the other blood constituents (i.e. cells, proteins) to be replaced.

By monitoring the hematocrit (Hct or packed cell volume- PCV) and/or hemoglobin of the animal, it is possible to evaluate whether the animal has sufficiently recovered from a single or multiple blood draws. After a sudden or acute blood loss, it takes up to 24 hours for the hematocrit and hemoglobin to reflect this loss. In general, if the animal’s hematocrit is less than 35% or the hemoglobin concentration is less than 10 g/dl, it is not safe to remove blood. Please contact a DLAR Veterinarian if you need assistance with monitoring PCVs in animals.

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

Guinea Pig37-48Cow24-48

Fluid replacement

Lactated Ringer’s Solution (LRS) is the recommended balanced crystalloid solution for fluid replacement. Alternatively, 0.9% sterile isotonic saline may be used. For adult mice, 1.0 ml of warmed LRS or isotonic saline can be given by IP or SC administration. For adult rats, administer 5 -10 ml warmed LRS or 0.9% saline (½ of the total volume via IP and ½ via SC routes).


1. Hawk TC, Leary, ST, and Morris, TH. 2005. Formulary for Laboratory Animals, 3rd ed. Ames, IA Blackwell Publishing

2. BVA/FRAME/RSPCA/UFAW Joint Working Group on Refinement. Removal of blood from laboratory mammals and birds. Laboratory Animals (1993) Jan; 27(1):1-22

3. NIH/OACU Guidelines for Survival Bleeding in Mice and Rats, 2010 Available at: Accessed 7/11/11

4. Boston University IACUC Policy for Blood Collection Guidelines. Available at: Accessed 7/11/11

5. Forbes N, et. al. Morbidity and mortality rates associated with serial bleeding from the superficial temporal vein in mice. Lab Animal (2010) Aug; 39(8):236-40.

6. Hoff, J. Methods of Blood Collection in the Mouse. Lab Animal (2000) Nov; 29(10):47-53.

7. McGuill M.W. and Rowan A.N. Biological Effects of Blood Loss: Implications for Sampling Volumes and Techniques. ILAR News (1989), 31(4): 5-18.

8. Nahas K, Provost J-P, Baneux PH, and Rabemampianina Y. Effects of acute blood removal via the sublingual vein on haematological and clinical parameters in Sprague-Dawley rats. Laboratory Animals (2000) Oct; 34(4):362-71.

9. NC3R’s (National Centre for the Replacement, Refinement and Reduction of Animals in Research) Blood Sampling Microsite. Available at: Accessed 7/11/11


(Video) Tips for Blood Collection from Lab Animals l Learn with Labmonk l Part - 1


What is the procedure for collection of blood sample in animals? ›

The animal is restrained manually or using a suitable animal restrainer. Hind leg is immobilized and slight pressure may be applied gently above the knee joint. The vein is punctured using a 20G needle and enough volume of blood is collected with a capillary tube or a syringe with a needle.

What are the standards of practice in blood collection? ›

Ask the patient to form a fist so the veins are more prominent. Enter the vein swiftly at a 30 degree angle or less, and continue to introduce the needle along the vein at the easiest angle of entry. Once sufficient blood has been collected, release the tourniquet BEFORE withdrawing the needle.

What is the maximum volume of blood that can be collected from a laboratory animal over a 24 hour period and in two 2 weeks? ›

III. Guidelines: As a general rule, 1% of an animal's body weight (measured in grams) can be collected in blood (measured in milliliters) within a 24-hour period, every 14 days. For example, 0.3 ml can be collected once every two weeks from a 30-gram mouse.

How much blood is required to collect in each animal? ›

Volume: On average, the total circulating blood volume is equal to 5.5 -8.0 % of the animal's body weight. Non-terminal blood collection without additional monitoring (see below) should be limited to 10% of the total circulating blood volume on a single collection or every 2 week period for serial collections.

What is the Iacuc policy for blood collection in laboratory animals? ›

In all cases, non-terminal blood collection without replacement fluids is limited to 10% of the total circulating blood volume of a healthy animal during a two-week period, unless otherwise approved in the IACUC protocol.

What are the eight steps to follow when collecting specimens? ›

The following are essential safeguards for your patients.
  • Avoid patient identification errors. ...
  • Draw the tubes in the proper sequence. ...
  • Use proper containers for collection. ...
  • Mix all tubes ten times by gentle inversion immediately after collection.
  • Do not decant specimens from one type of container into another.

What are OSHA standards for phlebotomy? ›

One of OSHA's primary requirements for phlebotomists is washing hands before and after working with patients, after removing personal protection equipment such as gloves, and anytime they are exposed to blood or anything else that may carry infection. In addition, phlebotomists must keep their work areas clean.

What is the most important consideration when collecting a blood culture? ›

It is therefore most important to use proper, sterile technique to decontaminate the patient's skin before collecting the blood culture. If disinfecting steps are not properly followed, skin bacteria can transfer to the bottle and cause false positive results.

What are the six steps of routine blood collection? ›

List the six steps of routine blood collection.
  1. Correctly and positively identify the patient.
  2. Choose the appropriate equipment for. obtaining the sample.
  3. Select and prepare the site for collection.
  4. Collect the sample, ensuring patient. ...
  5. Correctly label the sample.
  6. Transport the sample to the lab in a timely.

What is minimum blood collection volume? ›

collect a minimum of 1.5 ml of whole blood.

What are the 5 common sites of blood collection in dogs? ›

Common sites for blood collection in large animals
SpeciesSites of collection and permitted conditions
Dog and catCephalic, saphenous veins, femoral and jugular veins
RuminantsJugular vein
SwineJugular vein, anterior vena cava, ear veins
Nonhuman PrimatesFemoral, cephalic veins, saphenous vein
1 more row

How do you measure the blood volume of an animal? ›

Blood volume can be related to body weight using the experimentally determined equation of Lee and Blaufox; BV = 0.06 X BW+ 0.77. In which blood volume = BV in mL and BW = body weight in grams. The equation was developed for the rat, but is reasonably accurate for mice, rabbits, guinea pigs etc.

What is the maximum blood volume that can be taken during a survival collection? ›

In general, no more than 10% of the animal's blood volume should be removed at one sampling....” Per information extracted from McGuill, M.W. and Rowan, A.N., "Biological Effects of Blood Loss: Implications for Sampling Volumes and Techniques," ILAR News, Vol.

What is the maximum amount of blood in ml that can be collected from a 30 kg dog? ›

Previous studies indicate the collection of approximately 16-18 ml (15-20% of the blood volume) of blood for each kilogram of weight of the donor. According to the formula: Estimated blood volume (liters) = 0.08-0.09 X Body weight (kg), an animal weighing 30 kg has a total blood volume of approximately 2.6L.

What is the maximum blood volume that should be collected from a healthy fish? ›

The maximum volume of blood sampling is dependent on body weight of the fish. The volume for repeated blood sampling at intervals should be ≤0.4% of body weight every week or ≤1% every 2 weeks, which were evaluated by measurements of blood hemoglobin.

What are the regulations for laboratory animals for? ›

The federal law called the Animal Welfare Act (AWA) sets high standards of care for lab animals with regard to their housing, feeding, cleanliness, ventilation and medical needs. It also requires the use of anesthesia or analgesic drugs for potentially painful procedures and during post-operative care.

What are the Three Rs in the legal standards for animal research? ›

The Three Rs principle was launched in the early 1960s by two English biologists, Russel and Burch in their book “The Principle of Humane Experimental Technique”. The 3 Rs stand for Replacement, Reduction and Refinement.

What are the ethical guidelines for animal research? ›

Ethical Guidelines for the Use of Animals in Research
  • Respect for animals' dignity.
  • Responsibility for considering options (Replace)
  • The principle of proportionality: responsibility for considering and balancing suffering and benefit.
  • Responsibility for considering reducing the number of animals (Reduce)
Jul 8, 2019

What 5 pieces of information must be on every specimen collected? ›

Last, first, and middle name of patient. Medical record number (8 digits) Patient's date of birth. Last and first name of ordering physician.

What is the correct order of specimen collection? ›

The draw order for specimen tubes is as follows:

Red No Gel. Gold SST (Plain tube w/gel and clot activator additive) Green and Dark Green (Heparin, with and without gel) Lavender (EDTA)

What are the general steps for processing blood samples? ›

There are four steps involved in obtaining a good quality specimen for testing: (1) preparation of the patient, (2) collection of the specimen, (3) processing the specimen, and (4) storing and/or transporting the specimen.

What is OSHA's 6 foot rule? ›

The 6-foot rule. Subpart M requires the use of fall protection when construction workers are working at heights of 6 feet or greater above a lower level.

What are the 4 OSHA standards? ›

There are four groups of OSHA standards: General Industry, Construction, Maritime, and Agriculture. (General Industry is the set that applies to the largest number of workers and worksites). These standards are designed to protect workers from a wide range of hazards.

What are 10 of the most violated OSHA standards? ›

OSHA's Top 10 Safety Violations 2022
  • 1 Fall Protection–General Requirements – 5,260 citations. ...
  • 2 Hazard Communication – 2,424 citations. ...
  • 3 Respiratory Protection – 2,185 citations. ...
  • 4 Ladders – 2,143 citations. ...
  • 5 Scaffolding – 2,058 citations. ...
  • 6 Lockout/Tagout – 1,977 citations. ...
  • 7 Powered Industrial Trucks – 1,749 citations.
Nov 1, 2022

What are the precautions when collecting sample for blood culture? ›

  • Aseptic techniques should be used so that bacteria from the skin do not contaminate the culture bottles.
  • Ideally the skin should be washed with soap and rinsed with sterile water. ...
  • Alcoholic chlorhexidine solutions have been shown to reduce blood culture false positives compared with aqueous povidone-iodine.
Aug 17, 2015

What would be the appropriate timing and number for the collection of routine blood cultures? ›

A maximum of three sets per patient per day for three consecutive days is recommended. Obtain three blood culture sets within two hours, then begin therapy.

What is the preparation that patient needs to do before proceed with blood collection? ›

General practice is to have the patient fast twelve hours prior to blood draw, especially when testing for a lipid profile and glucose tests. There is also evidence that fasting for creatinine levels should be considered, especially when high protein/meat diets are consumed.

What are the 3 systems used in blood collection? ›

Three popular methods of blood collection or sampling are:
  • Arterial Sampling.
  • Venipuncture Sampling.
  • Fingerstick Sampling.
Mar 2, 2017

What is the single most important step of the blood collection procedure? ›

Patient ID, the process of verifying a patient's iden- tity, is the most important step in specimen collection. Obtaining a specimen from the wrong patient can have serious, even fatal, consequences, especially specimens for type and cross-match prior to blood transfusion.

What type of procedure is the process of collecting blood? ›

Venipuncture is the collection of blood from a vein. It is most often done for laboratory testing. Blood is drawn from a vein (venipuncture), usually from the inside of the elbow or the back of the hand. A needle is inserted into the vein, and the blood is collected in an air-tight vial or a syringe.

What type of collection procedure is used to obtain blood for a blood culture? ›

Blood culture specimens are usually drawn using either a needle and syringe or a vacuum-extraction collection system that draws blood into vacuum-sealed blood culture bottles. In both cases, a hollow-bore needle is inserted into the lumen of a patient's vein to obtain the blood culture specimen.

How do they collect blood samples from cattle? ›

Stabilise the vacutainer holder with your thumb, index and middle fingers. Hold the blood tube with your 4th and 5th fingers and against your palm. Push the blood tube onto the needle i.e. so that the needle pierces the bung. When all samples have been collected, remove the vacutainer holder and needle from the tail.

What are the steps in collecting blood evidence? ›

Place thread on bloodstain with a pair of clean forceps or a clean cotton swab. Roll the thread on the bloodstain, so the stain is absorbed onto the thread. Repeat until a minimum of four threads are collected. Place the threads (and swabs, if used) in a secure area and allow them to air dry.

What precautions should be taken during sample collection? ›

Personal protective equipment (apron, hand gloves, face shield, N95 Masks etc.) need to be used and all biosafety precautions should be followed while carrying out sample collection and packaging.

What happens if you draw blood without a tourniquet? ›

Complications include pain, swelling, skin sores, varicose veins, post-thrombotic syndrome, amputation, pulmonary embolism and death. Performing venipunctures without tourniquets is not an option. Constriction of the circulation causes veins to distend as they fill up with blood that can no longer circulate.

What is the most important requirement in blood culture collection? ›

A sufficient volume of blood withdrawn per culture is one of the most important features to optimize recovery of organisms from blood. Mark each bottle label before collection with the desired fill line to indicate the desired collection volume. Overfilling of bottles should be avoided.

What are the important factors to consider when collecting blood for culture? ›

Various factors affect the positivity rate of blood culture, including the timing of sample collection, skin antiseptic preparation, number of blood culture sets, and blood volume inoculated in individual culture bottles.

What is the most common blood draw site for a bovine? ›

Coccygeal (Tail) Vein

Cattle are generally more tolerant of venipuncture in the tail than in the neck.

Why necessary to maintain the aseptic environment while collecting blood from animals? ›

Following aseptic technique helps prevent the spread of pathogens that cause infection.

What are the 3 things that must be answered when examining blood? ›

The criminalist must be prepared to answer the following questions when examining dried blood: – (1) Is it blood? – (2) From what species did the blood originate? – (3) If the blood is of human origin, how closely can it be associated to a particular individual?

What is the most preferred method of evidence collection? ›

Swabbing is the most common technique used to collect various samples of DNA evidence.


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