Blood Collection Guidelines

Find general information on blood collection methods and recommended blood collection sites for common laboratory animals.

All procedures must have prior approval from the Institutional Animal Care and Use Committee (IACUC). The blood collection method, interval between blood collection procedures, and volume of blood to remove must be listed in the approved protocol specific to each study.

For training on specific blood collection methods and techniques, please contact Research Animal Resources training staff at rartrain@umn.edu. View all training sessions.

Restraint

For some species, blood collection may be adequately performed while the animal is awake using the appropriate restraint.

Restraint is necessary to prevent movement that may result in laceration of a blood vessel or other organ and serious complications. Restraint can be physical or chemical in nature, and should be chosen based on the species and the site being used for blood collection:

  • Dogs, cats, sheep, and calves: Usually require only physical restraint to collect blood. 
  • Swine: Will likely require physical or chemical restraint. 
  • Small mammals, birds, and amphibians: Use manual restraint or place animals in appropriate restraining devices. 

Chemical restraint may be required to collect blood from some animals/species. View anesthesia guidelines.

Please consult with Research Animal Resources (RAR) veterinary staff for specific instructions, assistance selecting a restraint device, or guidance on the appropriate anesthesia of individual species.

Collection guidelines: Volumes

RAR veterinary staff provide the following criteria to determine the maximum, safe amount of blood to withdraw.

Keep in mind the serum fraction represents about half of the total blood volume. When calculating blood volumes based on body weights (see below), remember that body weights in kilograms (kgs) will convert to blood volumes in liters, and weight in grams will convert to volumes in milliliters (mls). 

Appropriate blood volumes for collection

Appropriate blood volumes for collection

5-10% of body weight = total blood volume

The circulating blood volume can generally be estimated as 55-70 ml/kg of total body weight. However, care should be taken in these calculations as the percent of total blood volume will be lower (-15%) in sick, obese, and older animals. 

Approximate blood volumes by species

The following notes the mean blood volume (ml/kg), and is adopted from Heinz-Diehl, 2001 and Hawk et al., 2005:

  • Mouse: 72 ml/kg
  • Rat: 64
  • Hamster: 78
  • Gerbil: 67
  • Guinea pig: 75
  • Rabbit: 65
  • Ferret: 75
  • Cat: 55
  • Dog (beagle): 85
  • Sheep: 66
  • Minipig: 65
  • Macaque (rhesus): 56
  • Macaque (cyno): 65

Single blood draw

Single blood draw

A maximum of 1% of body weight can be removed (e.g., 0.15 ml from a 15 gram mouse, 50 ml from a 5 kg cat, 400 ml from a 40 kg dog) as a single blood draw every 14 days without requiring supplemental replacement fluids. Withdrawing the minimum amount of blood necessary is strongly recommended.

If the total volume is over 1% (not to exceed 2%), fluid volume replacement must be considered. Please consult with  RAR veterinary staff if you have questions about fluid replacement.

Multiple blood draws

Multiple blood draws

If the total volume withdrawn during several blood draws during a 14-day period is:

  • Less than 1% body weight: No additional action needs to be taken.
  • 1-2% body weight (or over): Perform fluid replacement. Please contact the RAR veterinary staff with questions.

If frequent blood draws will be necessary, it may be prudent to place a port or purchase animals with vascular catheters. Always follow vendor instructions regarding blood sampling, flushing, and animal housing.

Fluid replacement

Fluid replacement

If the blood volume removed from an animal exceeds the maximum recommended blood collection volumes listed above, replace the removed volume of blood with warm (30-35ºC) isotonic solution (e.g., 0.9% saline, lactated Ringer’s solution). 

Exsanguination

Exsanguination

Approximately 50-75% of total blood volume (4-5% of body weight) can be obtained by terminal exsanguination.

The animal must be deeply anesthetized, or recently euthanized, prior to exsanguination. Since the amount of blood obtained is substantially increased if the heart is beating during the bleeding procedure, using a surgical plane of anesthesia is recommended.

The procedure for anesthesia and/or euthanasia must be described and approved in the IACUC protocol.

Monitoring

Monitoring

An animal may go into hypovolemic shock if too much blood is withdrawn too rapidly or too frequently without replacement (approximately 2% of the animal’s body weight at one time).  

If signs of shock are observed (e.g., increased pulse, pale mucus membranes, cold skin/extremities, hyperventilation), immediately contact RAR veterinary staff.

By monitoring hematocrit (Hct or PCV) and/or hemoglobin (Hb), it is possible to evaluate if the animal has sufficiently recovered from single or multiple blood draws. Remember it may take up to 24 hours for hematocrit or hemoglobin to reflect sudden or acute blood loss.

In general, it is not safe to remove the volumes of blood listed above if:

  • The animal is anemic (i.e., below a species’ normal PCV range; ranges are listed below).
  • The hemoglobin concentration is less than 10 gm/dL.

Normal Packed Cell Volume (PCV) for common laboratory animals:

  • Mouse: 39-49%
  • Rat: 36-54%
  • Hamster: 40-61%
  • Gerbil: 43-60%
  • Guinea pig: 37-48%
  • Rabbit: 30-50%
  • Dog: 29-55%
  • Cat: 25-41%
  • Sheep: 24-45%
  • Cow: 24-48%
  • Swine: 32-50%
  • Rhesus: 26-48%

Blood collection sites and methods

Below you’ll find blood collection sites for common laboratory animal species. Sites are listed from most common/desirable to least common/desirable based on ease of collection.

For uncommon laboratory animal species, please contact the RAR veterinarians for more  information. 

Key considerations

  • For smaller species, the approximate volume of blood attainable for each site is listed. Volumes are an estimate and will also depend on the size, health, and hydration status of the animal as well as the experience and skill level of the person collecting the sample. 
  • Certain sites may be preferable based on study goals and requirements. Additionally, publications indicate that blood analysis results (especially cellular indices) may vary based on blood collection site; consult the literature for more information. 
  • Cardiac puncture may be used to obtain a single, large volume of blood from heavily anesthetized (terminal procedure only) or euthanized animals.
  • Collecting blood by lacerating an ear or tail vessels requires review and approval by the IACUC. There is potential that an artery will be lacerated rather than a vein, resulting in severe hemorrhage. In addition, these procedures are more painful than needle punctures because of the prolonged time for wound healing. Also, the procedure site is very susceptible to infection, hemorrhage, and other complications. To receive training in other methods listed below, contact RAR training staff at rartrain@umn.edu. View training calendar.
  • An animal may not be returned to its cage until complete hemostasis has been achieved (i.e., no more blood is coming from the collection site), regardless of the collection method. Achieve hemostasis using gauze and direct pressure. Up to several minutes of pressure may be required following arterial puncture.

Recommended blood collection sites by species

Mice

Mice

Saphenous vein
Anesthesia: No
Repeat bleeds: Yes
Expected volume: 100-200 ul

Submandibular vein
Anesthesia: No
Repeat bleeds: Yes
Expected volume: 200-500 ul

Distal tail transection (1-3 mm)
Anesthesia: No
Repeat bleeds: Yes – limited
Expected volume: <100 ul

Retro-orbital sinus
Anesthesia: Required
Repeat bleeds: Yes – limited
Expected volume: 200 ul

Lateral tail vein
Anesthesia: No
Repeat bleeds: Yes
Expected volume: 50-100 ul

Sublingual vein
Anesthesia: Required
Repeat bleeds: Yes
Expected volume: 500 ul

Jugular vein
Anesthesia: Recommended
Repeat bleeds: Yes

Cardiac puncture (terminal only)
Anesthesia: Required
Repeat bleeds: Terminal
Expected volume: ~1 ml

Rats

Rats

Saphenous vein
Anesthesia: No
Repeat bleeds: Yes
Expected volume: 300-400 ul

Lateral tail vein
Anesthesia: No
Repeat bleeds: Yes
Expected volume: 200-400 ul

Jugular vein
Anesthesia: Required
Repeat bleeds: Yes
Expected volume: 0.5- 2.0 ml

Sublingual vein
Anesthesia: Required
Repeat bleeds: Yes
Expected volume: 0.5-1.0 ml

Retro-orbital sinus
Anesthesia: Required
Repeat bleeds: Yes – limited
Expected volume: 0.5-1.0 ml

Cardiac puncture (terminal only)
Anesthesia: Required
Repeat bleeds: Terminal
Expected volume: ~3 ml

Guinea pigs

Guinea pigs

Saphenous vein
Anesthesia: No
Repeat bleeds: Yes
Expected volume: 400-500 ul

Jugular vein
Anesthesia: Recommended
Repeat bleeds: Yes
Expected volume: 2-3 ml

Cranial vena cava
Anesthesia: Recommended
Repeat bleeds: Yes
Expected volume: 2-3 ml

Cardiac puncture (terminal only)
Repeat bleeds: Terminal

Rabbits

Rabbits

Marginal ear vein/central ear artery
Anesthesia: Local anesthesia recommended
Repeat bleeds: Yes
Expected volume: 1-3 ml

Lateral saphenous vein
Anesthesia: No
Repeat bleeds: Yes

Cephalic vein
Anesthesia: No
Repeat bleeds: Yes

Jugular vein
Anesthesia: Recommended
Repeat bleeds: Yes

Cardiac puncture (terminal only)
Repeat bleeds: Terminal

Ferrets

Ferrets

Cephalic vein
Anesthesia: No
Repeat bleeds: Yes

Jugular vein
Anesthesia: Recommended
Repeat bleeds: Yes

Anterior vena cava vein
Anesthesia: Recommended
Repeat bleeds: Yes

Cardiac puncture (terminal only)
Repeat bleeds: Terminal

Cats

Cats

Medial saphenous vein
Anesthesia: No
Repeat bleeds: Yes

Cephalic vein
Anesthesia: No
Repeat bleeds: Yes

Jugular
Anesthesia: No
Repeat bleeds: Yes

Dogs

Dogs

Lateral saphenous vein
Anesthesia: No
Repeat bleeds: Yes

Cephalic vein
Anesthesia: No
Repeat bleeds: Yes

Jugular vein
Anesthesia: No
Repeat bleeds: Yes

Sheep/ruminants

Sheep/ruminants

Jugular vein
Anesthesia: No
Repeat bleeds: Yes

Cephalic vein
Anesthesia: No
Repeat bleeds: Yes

Saphenous vein
Anesthesia: No
Repeat bleeds: Yes

Pigs

Pigs

Marginal ear vein
Anesthesia: No
Repeat bleeds: Yes

Cephalic vein
Anesthesia: No
Repeat bleeds: Yes

Right jugular vein
Anesthesia: No
Repeat bleeds: Yes

Anterior vena cava
Anesthesia: Recommended
Repeat bleeds: Yes

Non-human primates

Non-human primates

Femoral vein
Anesthesia: Recommended
Repeat bleeds: Yes

Saphenous vein
Anesthesia: Required
Repeat bleeds: Yes

Cephalic vein
Anesthesia: Required
Repeat bleeds: Yes

Brachial vein
Anesthesia: Required
Repeat bleeds: Yes

Rodents: Common blood collection procedures and techniques 

Generally, one person can perform mouse procedures. Some techniques for rats may require two people: One to restrain the animal and one to collect blood.

Other procedures will require a restraint device or light anesthesia. Terminal blood collection by cardiac puncture can only be performed under anesthesia.

Lateral tarsal or saphenous vein

Lateral tarsal or saphenous vein

This procedure can be used to collect blood from mice or rats. The lateral tarsal vein on the foot is commonly used for hamsters, while the lateral saphenous vein on the lower leg is commonly used for mice and rats.

This procedure may require two people: One to restrain the animal and one to collect blood.  Alternatively, light anesthesia can be used if one person is working alone.

Technique

  1. The restrainer manually restrains the animal and extends one hind leg. Alternatively, lightly anesthetize the animal with isoflurane.
  2. The bleeder gently grasps the foot and shaves the leg if necessary. Apply alcohol to the skin and allow it to dry.
  3. The bleeder applies gentle pressure to the leg or foot above the site where blood will be collected to occlude the vein and cause it to dilate.
  4. Using a 25G needle, pierce the vein and allow blood to well out of the nick. The leg may be gently stroked to encourage blood flow.
  5. Either allow blood to drip into a collection tube or collect it with a hematocrit tube.
  6. Once blood collection is complete, apply gentle pressure with gauze for 15-30 seconds to stop the flow of blood.
  7. Release the animal from the restraint device and return it to its cage. If the animal was anesthetized, monitor the animal until it is fully awake and able to walk normally.

Facial vein

Facial vein

The following technique can be used in mice:

  1. Restrain (scruff) the mouse using the one-handed technique. It is critical to grasp a lot of skin. Your fingertips should almost be touching the mouse’s elbows, the mouse’s eyes should bulge slightly, the mouth should be slightly held open, and the forelimbs should stick out to the sides.
  2. The puncture point is at the back of the jaw, slightly behind the hinge of the jawbone and toward the ear on the hairless freckle. Avoiding the jawbone, puncture just behind the point where the upper and lower jawbones meet.
  3. Prick the freckle with a lancet or 21G needle. The puncture should be done with enough force to quickly insert the lancet point to the hilt.
  4. Quickly position the blood collection vessel. Collect 4-7 drops of blood. The blood may flow rapidly then slow in a very short time period.
  5. After collecting the sample, apply gentle pressure to the site with a gauze sponge to stop the bleeding.
  6. Return the animal to its cage.
  7. Observe the animal for bleeding or any signs of pain or distress.

Tail poke

Tail poke

The following procedure can be used to collect blood from mice:

  1. Place a heat lamp over the animal’s cage for no more than five minutes to warm mice and dilate vessels.
  2. Once the animals are warmed, place them in an appropriately sized restraint device with the tail extended. An alternative to the heat lamp is placing the animal in a restraint device, then rubbing the tail for several seconds with a gauze pad soaked in very warm, but not hot, water to dilate the vessels.
  3. Wipe the tail with 70% alcohol and allow it to dry.
  4. Holding the end of the tail, gently twist the tail to expose the lateral tail vein on either side.
  5. Approximately 2-3 cm from the tip of the tail, make a needle poke. When the poke is made deep enough, blood should immediately start welling up from the nick.
  6. Collect blood either by touching a capillary tube to the bead of blood, or by allowing the blood to drop into a collection tube. The tail can be gently stroked from the base of the tail toward the tail to encourage blood flow.
  7. After collecting a sufficient amount of blood, apply pressure to the nick with clean gauze for 15-30 seconds to stop the flow of blood.
  8. Once the bleeding is controlled, apply a drop of tissue glue to the nick for hemostasis.
  9. Release the animal from the restraint device and return it to its cage.

Tail vein

Tail vein

The following procedure can be used to collect blood from rats:

  1. Place the animal in a restraint device with the tail extended. Alternatively, the rat can be lightly anesthetized with isoflurane.
  2. Place a heat lamp over the tail for 2-5 minutes to warm the tail and dilate vessels.
  3. Wipe the tail with 70% ethanol and allow it to dry.
  4. Using a 23G needle attached to a syringe, insert the needle into one of the lateral veins, approximately 2-3 cm from the tip of the tail.
  5. When blood appears in the hub, slowly retract the plunger to collect the desired amount of blood.
  6. Once blood collection is complete, remove the needle and apply gentle pressure with gauze for 15- 30 seconds to stop the flow of blood.
  7. Release the animal from the restraint device and return it to its cage. If the animal was anesthetized, monitor it until it is fully awake and able to walk normally.

Retro-orbital sinus

Retro-orbital sinus

This procedure can be used to collect blood from mice, rats, and hamsters. Anesthesia is required for the following procedure:

  1. Manually restrain the animal and place one drop of topical ophthalmic anesthetic in the eye that will be bled.
  2. Anesthetize the animal following the method described in the IACUC-approved protocol.
  3. Gently pull the skin around the head and neck taut to cause the eyeball to protrude slightly. Be sure not to obstruct breathing.
  4. Carefully insert a sterile Pasteur pipet or hematocrit tube into the inner corner of the eye and direct it toward midline, angling toward the back of the head. Be careful not to scratch the eye when inserting the tube or break the tip of the tube.
  5. Apply gentle pressure to the hematocrit tube to pierce the retro-orbital sinus. Once the blood begins flowing, tip the tube down so gravity can assist the flow of blood into the tube.
  6. Once blood collection is complete, remove the hematocrit tube or Pasteur pipet and apply gentle pressure to the eye with gauze for 15-30 seconds to stop the flow of blood.
  7. After achieving hemostasis, apply antibiotic ophthalmic ointment to the eye to minimize the risk of infection.
  8. Place the animal in the recovery cage and continuously monitor it until it is fully awake and able to walk normally.

Cardiac puncture

Cardiac puncture

This is a terminal procedure to collect a large volume of blood from mice and rats. Anesthesia is required for the following procedure:

  1. Anesthetize or euthanize the animal following the method approved in the individual protocol.
  2. Place the animal on its back and wipe the chest and abdomen with ethanol.
  3. Attach an appropriately sized needle to a syringe and insert it at a 30º angle just below the xyphoid process, angling the needle slightly toward the left shoulder. A 25G 5/8 inch needle is sufficient for mice. hamsters and rats may require a 23G 1 inch or longer needle.
  4. Slightly retract the plunger to create a vacuum inside the syringe, then advance the needle until blood appears in the hub of the needle.
  5. Slowly retract the plunger to collect the desired amount of blood.
  6. Once blood collection is complete, withdraw the needle and euthanize the animal. Cervical dislocation or thoracotomy are recommended euthanasia procedures.

References for these guidelines are available by request.

Research Animal Resources