Rodent Blood Collection techniques
Guidelines: The procedures described in this SOP should be followed when collecting blood
samples from mice, rats, or hamsters. Blood volume of an individual animal is approximately 68% of body weight in adults. Up to 10% of the total blood volume can safely be removed from a
healthy animal without fluid replacement at two week intervals. If a larger sample is needed at
one time, up to 15% can be slowly removed if IV fluids are given to replace the lost volume.
Maximum withdrawal volumes must be followed to minimize the risk of anemia and
cardiovascular collapse.
Collection
Interval
One time
Every 2 weeks
Weekly
Terminal
Adult body
weight
Adult Blood
volume
Weekly sample
volume (7.5%)
Every two week
sampling (10%)
One time with
fluid replacement
(15%)
Terminal
Maximum Blood collection volumes
Percent of
Percent of body
Blood volume
weight
1.5%
1.0%
0.75
3-5%
Fluid
replacement
required
Yes
No
No
N/A
Mouse
25-35g
Rat
250-500g
Hamster
90-120g
2-2.75mL
15-30mL
5.4-7.2mL
0.15-0.21
1.125-2.25mL
0.4-0.54mL
0.2-0.28mL
1.5-3.0mL
0.54-0.72mL
0.3-0.42mL
2.25-4.5mL
0.81-1.08mL
1-1.5mL
8-12mL
3-5mL
15%
10%
7.5%
30-50%
Body weights can vary with age, strain, and gender. Therefore, it is recommended that animals
are weighed to calculate maximum blood collection volumes. For experimental groups of the
same age, strain, and gender the average group weight can be used to calculate blood collection
limits.
KMJ/ Drafted March 2010/ Updated September, 2012 KG
Rodent Blood Collection techniques
Materials:
Sterile Hematocrit tubes
Sterile Pasteur pipets
Sterile needles- 25G and 23G
Sterile syringes-1CC, 3CC, 5CC and 10CC
Sterile scalpel blades- size 15
Clippers
70% ethanol
Gauze
Blood collection tubes
Heat lamp
Tissue glue
Topical ophthalmic anesthetic, such as proparacaine
Ophthalmic antibiotic ointment
Rodent restrainer
Rat restraint board for jugular blood collection with attached leg restraint ropes
Plastic nose cone for head restraint
Procedures in the mouse can generally be performed by one person. For rats and hamsters some
techniques may require two people, one to restrain the animal and one to collect the blood. Other
procedures will require a restraint device or light anesthesia. Terminal blood collection by
cardiac puncture can only be performed under anesthesia.
Methods:
Tail nick:
This procedure can be used to collect blood from mice.
1. Place a heat lamp over the cage of animals for no more than 5 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 method to using the heat lamp is to place the animal in a
restraint device, then rub 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 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 small nick over the lateral tail vein
using a sterile scalpel blade. When the nick is made deep enough, blood should start welling
up from the nick immediately.
6. Blood can be collected 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. Once a sufficient amount of blood is collected, apply pressure to the nick with clean gauze
for 15-30 seconds to stop the flow of blood.
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Rodent Blood Collection techniques
8. Once the bleeding is controlled, a drop of tissue glue should be applied to the nick for
hemostasis.
9. Release the animal from the restraint device and return it to its cage.
Tail vein:
This 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 using the rodent EZ anesthesia machine following the
anesthesia SOP.
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 retract the plunger slowly 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 the animal until it is fully awake and able to walk normally.
Jugular vein:
This procedure can be performed on rats and requires two people, one to restrain the rat and one
to collect the blood.
1. The restrainer grasps the rat in a viper grip: Using one hand the rat’s head is placed between
the forefinger and middle finger, which are curled over the shoulders. The thumb and ring
finger are wrapped around the lower chest. With the other hand, the abdomen is grasped from
the ventral side, hooking the forefinger and middle finger over the hips and wrapping the
remaining fingers around the back. This grip gently restrains the animal and minimizes
struggling.
2. The restrainer lays the animal, back down, on the restraint board with the head toward the
bleeder with the xyphoid process approximately at the edge of the slanted surface of the
board.
3. The bleeder grasps the restraint ropes, twists them into slip knots then places the loops on the
forelimbs of the rat. The loops are cinched just behind the metacarpals so the hold is firm, but
does not significantly impede circulation to the paws.
4. When the loops are secured to the rats forelimbs, the restrainer releases the upper body of the
rat and the bleeder hooks the restraint ropes to the pegs on the sides of the board to
immobilize the forelimbs. The forelimbs should be in a straight line perpendicular to midline
for proper positioning
5. The bleeder places the nose cone over the head and folds the edges over to prevent the head
from slipping out. The restrainer then rotates the head 90 degrees toward the side where
KMJ/ Drafted March 2010/ Updated September, 2012 KG
Rodent Blood Collection techniques
blood will be collected and tilts the head back gently to pull the skin over the jugular furrow
taut.
6. Once the rat is properly positioned, a 23G needle attached to a 3CC syringe is inserted into
the center of the jugular furrow-approximately 1cm lateral to midline. This distance is
estimated for an adult 200gram rat. The actual distance must be adjusted depending on the
size of the rat.
7. Once the needle has broken the skin, the plunger of the syringe is gently retracted to create
negative pressure and the needle is slowly advanced.
8. As soon as there is a flash of blood in the hub the needle is held stationary and the plunger of
the syringe is gently retracted until the desired volume of blood is collected.
9. Once the blood collection is complete, the needle is removed and manual pressure is applied
to the site for approximately 10 seconds for hemostasis.
10. The restrainer releases the restraint ropes from the restraint board and grasps the rat in a viper
grip. Then the loops are removed from the forelegs of the rat and the paws are gently
massaged for about 15 seconds to ensure proper blood flow to the paws. The rat is then
returned to its cage.
Lateral tarsal or saphenous vein:
This procedure can be used to collect blood from mice, rats, or hamsters. 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 requires two people, one to restrain the animal
and one to collect the blood. Alternatively, light anesthesia can be used if one person is working
alone.
1. The restrainer manually restrains the animal and extends one hind leg. Alternatively, the
animal can be lightly anesthetized with isoflurane using the EZ anesthesia machine following
the anesthesia SOP.
2. The bleeder gently grasps the foot an shaves the leg if necessary. Alcohol is applied to the
skin and allowed to dry.
3. The bleeder applies gently 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, the vein is pierced and blood is allowed to well out of the nick. The leg
can be gently stroked to encourage blood flow.
5. Blood can either be allowed to drip into a collection tube or collected 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.
Retro-orbital sinus:
This procedure can be used to collect blood from mice, rats, and hamsters. Anesthesia is
required for this procedure.
KMJ/ Drafted March 2010/ Updated September, 2012 KG
Rodent Blood Collection techniques
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 anesthesia SOP.
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 inner corner of the eye and
direct it towards midline, angling toward the back of the head. For hamsters, the tube
should be inserted into the outer corner of the eye. Be careful not to scratch the eye when
inserting the tube or to 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. Once hemostasis is achieved, apply antibiotic ophthalmic ointment to the eye to minimize the
risk of infection.
8. Place the animal in the recovery cage and monitor it continuously until it is fully awake and
able to walk normally.
Sublingual vein:
The sublingual technique can be used for rats, mice, and hamsters and must be performed under
anesthesia. The technique requires one person to restrain the animal and the other to collect the
blood.
1. Anesthetize the animal with isoflurane in the rodent EZ anesthesia chamber or Bell Jar
following the appropriate anesthesia SOP.
2. Gently pull the skin around the head and neck taut to partially constrict the sublingual
and jugular veins. Be sure not to obstruct breathing. Hold the animal supine so the person
collecting blood can access the tongue.
3. The person collecting blood grasps the tongue and gently extends it. The sublingual vein
runs down the midline of the tongue, and the caudal portion is punctured with a 23G
(Hamsters and rats) or 25G needle (mice).
4. Blood is either collected with a hematocrit tube, or allowed to drip into a collection tube
while the animal is held ventrally.
5. Once the desired amount of blood is collected, the restrainer releases tension on the skin
around the head and neck and direct pressure is applied to the tongue with gauze or a
cotton tipped swab for hemostasis.
Once hemostasis is achieved, the animal is placed in the recovery cage and monitored
continuously until it is fully awake and able to walk normally.
Cardiac puncture:
This is a terminal procedure to collect a large volume of blood from mice, rats, and hamsters.
Anesthesia is required for this procedure.
KMJ/ Drafted March 2010/ Updated September, 2012 KG
Rodent Blood Collection techniques
1. Anesthetize the animal following the anesthesia SOP.
2. Place the animal on its back and wipe the chest and abdomen wiped with ethanol.
3. Attach an appropriately sized needle to a syringe and insert it at a 30 º angle just below the
xyphoid process, angling the needle slightly toward the left shoulder. A 25G 5/8 inch needle
is sufficient for mice. For hamsters and rats, a 23G 1 inch or longer needle may be required.
4. Retract the plunger slightly 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 the blood collection is complete, withdraw the needle and euthanize the animal.
Cervical dislocation or thoracotomy are recommended euthanasia procedures.
7. Place the carcass in a biohazard bag and store in a waste freezer in ARF.
Post-procedural monitoring:
1. Anesthetized animals must be monitored continuously until they are awake and able to walk
normally.
2. Non-anesthetized animals should be monitored at a minimum 1 hour and 24 hours after the
procedure.
3. Observe the animals for the following signs:
a. Bleeding from the collection site
b. Labored breathing
c. Discharge from the nose or mouth
d. Ruffled fur
e. Hunched posture
f. Lethargy
g. Minimal responsiveness
4. Lethargy, labored breathing, minimal responsiveness and uncontrolled bleeding or discharge
from the blood collection site is criteria for immediate euthanasia.
5. Document all observations on the experimental health monitoring sheet. Alert the Attending
Veterinarian of any ill or injured animals.
KMJ/ Drafted March 2010/ Updated September, 2012 KG