Collecting blood from mice is necessaryfor a wide variety of scientific studies,and there are a number of efficientmethods available. It is important toremember that blood collection, becauseit can stress the animals, may have animpact on the outcome of research data.In addition, it is extremely importantthat those who collect blood becomeskilled in the techniques they employ,and seek to stress the mice as little aspossible. Drawing from experience, per-sonal communication, and publishedresources1-4, the following techniques aredescribed:• Blood collection not requiring anes-

thesia:–Saphenous vein–Dorsal pedal vein

• Blood collection requiring anesthesia:–Tail vein–Orbital sinus–Jugular vein

• Terminal procedures:–Cardiac puncture–Posterior vena cava–Axillary vessels–Orbital sinus

The Guide for the Care and Use ofLaboratory Animals mandates that ani-mal care personnel be appropriatelytrained5. Training is available throughaccredited programs in veterinary tech-nology; certification programs throughorganizations such as the AmericanAssociation for Laboratory AnimalScience; or commercially available train-ing materials appropriate for self-study.To minimize pain and stress in the ani-mal, it is recommended that all bloodcollection techniques be practiced onanesthetized or freshly euthanized mice

until the handler feels comfortable andis able to perform the procedure withoutcausing harm to the animal.

Protocol ApprovalThe method of blood collection you

choose must be described in theapproved protocol in which the animalsare used. Some IACUCs require justifi-cation for collecting blood from theorbital sinus in mice that are intended torecover, since there are other morehumane methods available. When notperformed correctly, blood collectionfrom the orbital sinus can result in irre-versible problems such as blindness orocular ulcerations. One study showedthat the competence of the person per-forming retro-orbital blood collectionin rats had an appreciable impact on theincidence of induced abnormalities6.

In addition, the Public Health ServicePolicy requires that, in order to obtainapproval to use animals (includingmice) in research, personnel must bequalified to perform the techniquesdescribed in the protocol.

When deciding which technique isbest suited for your study, you will needto consider a number of parameters.

Frequency of CollectionBlood should not be collected from

the orbital sinus more frequently thanonce every two weeks. The tail, dorsalpedal, saphenous, and jugular veins canbe used for serial blood collection asoften as needed, according to theapproved protocol.

Use of AnestheticsTo minimize discomfort to the ani-

mal, anesthesia is recommended when

November 2000 Lab Animal Volume 29, No. 10 TECHNIQUE

Methods of Blood Collection in theMouseJanet Hoff, LVT, RLATG

The author outlines various meth-

ods of blood collection from mice.

Hoff is a training technician at theUniversity of Michigan, 1150 W. MedicalCenter Dr., 018 ARF, Ann Arbor, MI48109-0614. Please send reprint requeststo Hoff at the above address.

47

vein compared to blood obtained fromother sites7.

Venous vs. Arterial BloodCertain methods of blood collection

yield arterial blood, others yield venous

blood, while still others yield a mixtureof both. The tail yields a mixture ofvenous and arterial blood. Terminalblood collection from the axillary vesselsresults in a mixture of venous and arter-ial blood and cardiac puncture can yieldvenous or arterial blood, or a mixture ofboth. The posterior vena cava and theorbital sinus yield venous blood.

Aseptic Blood CollectionThe optimal blood collection method

will also depend on whether the samplemust be taken aseptically. Collectionfrom the orbital sinus and jugular veinare good nonterminal choices when anaseptic sample is needed. Care must betaken to make sure that the blood doesnot come in contact with the mouse’sskin or fur. Cardiac puncture or bloodcollection from the posterior vena cavaare good methods of terminal blood collection when an aseptic sample isneeded.

Volume RequiredOn average, the total blood volume in

the mouse is 6-8% of its body weight, or6-8 ml of blood per 100 g of body

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TECHNIQUE Volume 29, No. 10 Lab Animal November 2000

collecting blood from the tail in miceover 28 days old and from the orbitalsinus in mice of all ages. Anesthesia isalso recommended when collectingblood from the jugular vein to ensurecorrect positioning of the mouse. Whencollecting blood from the saphenous ordorsal pedal veins, anesthesia is not nec-essary, since the procedure causes mini-mal pain and employs a restrainingdevice that holds the animal in place.

Effect on Blood ParametersBoth the source and method of blood

collection can affect blood parameters.Increases in blood hormone and glucoselevels are directly related to stressfulmethods of blood collection. Mice maybe stressed by restraining procedures orby sensing impending danger. Todecrease stress, acclimate the mice to therestraining device and prevent themfrom seeing procedures being conducted(move them to the other side of the roomor cover cages until they are needed).Packed cell volume (PCV) and hemoglo-bin measurements have been reported tobe higher in blood collected from the tail

FIGURE 1. Blood collection from the saphenous vein.

FIGURE 2. Application of petroleum jelly to the blood source.

weight. Thinner animals will have rela-tively larger blood volumes per bodymass, due to greater surface area.Younger animals, especially newborns,have proportionately larger blood vol-umes than older animals8.

If nonterminal blood collection isplanned, the orbital sinus, jugular vein,or saphenous vein will typically yieldmore blood (0.2-0.3 ml in the averagesize adult mouse) than the tail or dorsal pedal vein. Terminal blood collection from the orbital sinus or bycardiac puncture will yield 0.8-1.0 ml.The posterior vena cava and axillary ves-sels typically provide a smaller bloodvolume.

The following guidelines9 should beconsidered when determining a safe vol-ume of blood to withdraw:• 10-15% of total blood volume or 1% of

body weight is the maximum amountof blood that should be collected atone time;

• Blood volume is restored in 24 hours,but erythrocytes and reticulocytes maynot return to normal levels for up totwo weeks. Therefore, it is recommend-ed that the maximum amount of bloodbe withdrawn only once every twoweeks. Monitoring the PCV (normalrange for mice is 39-49%) or hemoglo-bin can help evaluate whether themouse has recovered from blood with-drawal;

• Removal of up to 1% of total bloodvolume daily over time is permissible,however, the effects of stress, site cho-sen, and anesthetic used must be care-fully considered;

• Removal of 2% of the total blood vol-ume is permissible if replacement flu-ids are given at the time blood is col-lected. The blood volume removedshould be replaced intravenously withtwice the volume of warm sterile fluidsat a slow, steady rate. If fluids cannotbe administered intravenously,

intraperitoneal or subcutaneous routesare other alternatives;

• 15-25% blood loss results in elevatedplasma epinephrine, norepinephrine,and corticosterone concentrations tocompensate for a decreased level ofplasma glucose concentration;

• 20-25% blood loss decreases the arter-ial blood pressure, cardiac output andoxygen delivery to vital organs leadingto hypovolemia and cardiac failure(shock). Muscular weakness, depressedmentation, and cold extremities mayalso be observed.

Immediately following blood collec-tion, always observe the mouse for signsof distress or anemia (e.g., rapid breath-ing, pale color of mucous membranes,depressed mentation, or muscle weak-ness). Observe mice daily for otherproblems, such as local trauma, infec-tion, or irritation at the blood collectionsite.

Methods of Blood CollectionNot Requiring AnesthesiaBlood Collection from theSaphenous Vein

Warming the mouse immediatelyprior to blood collection will increaseblood flow considerably. Place a lampover the cage for five minutes or placethe cage on a heating pad, on the lowestsetting. Place the mouse in a restrainingtube so its head is covered and its hindlegs are free. Grasp the fold of skinbetween the tail and thigh (Fig. 1). Thesaphenous vein is found on the caudalsurface of the thigh. Remove hair fromthe area with clippers. Apply petroleumjelly (Fig. 2) or eye lubricant to preventmigration of blood into the surroundinghair and place a tourniquet around theleg, above the knee (Fig. 3). Puncture thevein with a 25 gauge needle. Collectdrops of blood as they appear. The use ofcollection tubes with capillary action willfacilitate blood collection. Apply pres-sure or use a cauterizing agent such as astyptic pencil (silver nitrate) to stop thebleeding.

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49

FIGURE 3. Instructions for making a mouse-sized tourniquet.

1. Disassemble a 3 cc syringe. Thread aloop of OO gut suture material throughthe syringe.

3. Place a needle into the loop and pullthe loop tight against the tip of thesyringe. Tie the long ends of the suturematerial together.

2. Sew one end of the suture material intothe hub of the syringe.

4. Cut the long ends of the suturematerial and reassemble the syringe.

Methods of Blood CollectionRequiring Anesthesia

Mice can be anesthetized for a shortperiod of time using a drop jar with atight-fitting lid or a Ziploc™ bag con-taining an appropriate amount of anes-thetic10. When placed in the container,the mouse should become sufficientlyanesthetized to perform procedures,without overdosing from the anesthetic.This method of anesthetizing mice isused when the procedure may causemomentary pain or is difficult to per-form when the mouse is able to moveabout freely, and has been used for manyyears with anesthetics such as methoxy-fluorane (Metofane ) and ether. SinceMetofane is no longer available andether presents problems with handlingand storage, isoflurane is the currentanesthetic of choice. Table 1 lists the sup-plies needed; Table 2 and Table 3 outlinetwo possible procedures for anesthetiz-ing mice using isoflurane.

Blood Collection from the TailWarm the mouse and place it in a

restraining tube as described above. Do

not attempt to increase blood flow byrubbing the tail from the base to the tip,as this will result in leukocytosis(increased white blood cell count). Usinga scalpel, straight edge razor, or sharpscissors, quickly remove up to 1 cm ofthe tail. Collect blood in a capillary tubeas drops appear. Apply pressure or use acauterizing agent such as a styptic pencil(silver nitrate) to stop the bleeding.When several samples are needed withina short time period, the original woundcan be reopened by removing the clot.When additional samples are needed at alater date, blood samples can be obtainedby removing just 2-3 mm of additionaltail. Cutting the tail too short may resultin trauma to the cartilage and ultimatelyto the coccygeal vertebrae.

Blood Collection from the OrbitalSinus

Lay the anesthetized mouse on itsside on a table or hold it in your handwith its head pointing down (Fig. 5).With your first finger and thumb (fingerabove and thumb below the eye) pull theskin away from the eyeball, above andbelow the eye, so that the eyeball is pro-truding out of the socket as much aspossible. Take care not to occlude thetrachea with your thumb. Insert the tipof a fine-walled Pasteur pipette (o.d. of

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TECHNIQUE Volume 29, No. 10 Lab Animal November 2000

Blood Collection from the Top ofthe Foot

Warm the mouse and place it in arestraining tube as described above. Withyour thumb and first finger, hold a hindfoot around the ankle (Fig. 4). Yourthumb should be on top of the foot. Themedial dorsal pedal vessel is found on thetop of the foot. Apply petroleum jelly oreye lubricant to the foot. Puncture thevein with a 23–27 gauge needle. As dropsof blood appear collect them in a capil-lary tube. Apply pressure or use a cauter-izing agent such as a styptic pencil (silvernitrate) to stop the bleeding.

TABLE 2. Procedure for anesthetizingmice using isoflurane in a drop jar.

• Fill tissue cassettes with cotton or gauze and place in the jar or place cotton or gauze in the bottom of the jar and cover with wire or plastic mesh;

• Add 300 µl isoflurane to the cotton or gauze;

• Quickly close the lid;• Open the lid, place one mouse in the jar,

and quickly close the lid;• WATCH CLOSELY;• When the mouse is lying on its side and its

breathing has slowed:−Open the lid;−Remove the mouse from the jar;−Quickly close the lid;

• The mouse will be at a surgical plane of anesthesia for 30 seconds to one minute;

• Add another 300 µl of isoflurane to the jar after every third or fourth mouse.

FIGURE 4. Blood collection from the dorsal pedal vein.

TABLE 1. Supplies and equipmentneeded to anesthetize mice using adrop jar or Ziploc™ bag.

• 500 ml jar with tightly fitting lid or a Ziploc™bag

• Cotton or gauze• Tool to separate cotton or gauze from the

animal (e.g., plastic or wire mesh or tissue cassettes)

• Adequate local exhaust ventilation to pre-vent overexposure of lab personnel (e.g., scavenging devices, fume or snorkel hoods).

1-2 mm) or a microhematocrit bloodtube into the corner of the eye socketunderneath the eyeball, directing the tipat a 45-degree angle toward the middleof the eye socket. Rotate the pipettebetween your fingers during forwardpassage; do not move it from side to sideor front to back. Apply gentle downwardpressure and then release until the veinis broken and blood is visualized enter-ing the pipette. When a small amount of

November 2000 Lab Animal Volume 29, No. 10 TECHNIQUE

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blood begins filling the pipette, withdrawslightly and allow the pipette to fill. Donot let the pipette come out of the eyesocket. If the pipette is not withdrawnslightly, it may occlude the vein andblood will not flow freely.

Cover the open end of the pipette withthe tip of your finger before removing it

from the orbital sinus to prevent bloodfrom spilling out of the tube. Bleedingusually stops immediately and completelywhen the pipette is removed. It may benecessary to apply gentle pressure on theeyeball for a brief moment by closing theskin above and below the eye using yourfirst finger and thumb. It is recommend-ed that sample collection not be repeatedon the same eye for at least two weeks.

Caution: Blindness can occur if theoptic nerve is damaged as a result of theblood collection tube coming into con-tact with the nerve, which attaches to themiddle of the ventral surface of the eye.Ocular ulcerations, puncture wounds,loss of vitreous humor, infection, or ker-atitis may occur as a result of poor tech-nique or uncontrolled movement of theanimal.

Blood Collection from theJugular Vein

Restrain the anesthetized mouse byattaching a loop of string to a gauzesquare and looping it around the upperincisors. Pull the head up and back whilepulling the gauze across the back of thehand and lock the gauze between two fin-gers (Fig. 6). It may be helpful to placethe mouse in your lap. Wet the fur withalcohol or shave the neck area. In this

FIGURE 5. Retro-orbital blood collection.

TABLE 3. Procedure for anesthetizingmice using isoflurane in a Ziploc™bag9.

• Fill tissue cassette with cotton or gauze and place in the Ziploc™ bag;

• Add 100-200 ml isoflurane to the cotton orgauze;

• Quickly close the bag;• Open the bag, place one mouse in the

bag, and quickly close the bag;• WATCH CLOSELY;• When the mouse is lying on its side and

its breathing has slowed:-Open the bag;-Remove the mouse from the bag;-Quickly close the bag.

FIGURE 6. Proper positioning of mouse for blood collection from the jugular vein.

will stop breathing before the heartstops. Either perform a bilateral pneu-mothorax (puncturing both sides of thethorax) or wait for the animal to becomerigid.

Blood Collection by CardiacPuncture

Three possible approaches:• Hold the mouse by the scruff of skin

above the shoulders so that its head isup and its rear legs are down. Use a 1 mlsyringe and a 22 gauge needle. Insertneedle 5 mm from the center of thethorax towards the animal’s chin, 5-10mm deep, holding the syringe 25-30degrees away from the chest (Fig. 9);

• Lay animal on back and push syringevertically through sternum;

• Lay animal on side and insert the nee-dle perpendicular to chest wall.

If blood doesn’t appear immediately,withdraw 0.5 cc of air to create a vacuumin the syringe. Withdraw the needlewithout removing it from under the skinand try a slightly different angle or direc-tion. When blood appears in the syringe,hold it still and gently pull back on theplunger to obtain the maximum amountof blood available. Pulling back on the

plunger too much will cause the heart tocollapse. If blood stops flowing, rotatethe needle or pull it out slightly.

Blood Collection from thePosterior Vena Cava

Open the abdominal cavity of anes-thetized mouse by making a V-cutthrough the skin and abdominal wall 1cm caudal to the rib cage. Shift theintestines over to the left and push theliver forward. Locate the widest part ofthe posterior vena cava (between the kid-neys). Use a 23-25 gauge needle and a 1ml syringe. Carefully insert the needleinto the vein and draw blood slowly untilthe vessel wall collapses. Pause to allowthe vein to refill and then repeat three orfour times or until no more blood isavailable.

Blood Collection from theAxillary Vessels

Lay the anesthetized mouse on itsback. Stretch out a forelimb and pin thefront foot. Make a deep incision in theaxilla (armpit) at the side of the thorax.Hold the skin at the posterior part of theincision using forceps to create a cuppedarea. Incise the blood vessels in the area

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TECHNIQUE Volume 29, No. 10 Lab Animal November 2000

hyperextended position, the jugular veinsappear blue and are found 2-4 mm later-al to the sternoclavicular junction (Fig.7). Using a 1 ml syringe and 25 gaugeneedle, approach the vessel in a caudo-cephalic direction (from back to front).Insert the needle 1-3 mm deep, 2-4 mmlateral of the sternoclavicular junction,over the sternum; include a small amountof muscle from the sternum to stabilizethe needle. Hold the needle very stillwhen blood enters the syringe (Fig. 8).Withdraw blood slowly to avoid collapseof these small vessels.

If the first attempt to draw blood isunsuccessful, withdraw the needle slight-ly; it may have been placed too deeply. Ifblood stops flowing, do not continue todraw back on the syringe. The vein mayhave collapsed or the needle may haveattached to the vessel wall. Rotate theneedle slightly or apply slight pressure onthe needle (either above, below, or to theside of puncture site).

Terminal Methods of BloodCollection Requiring DeepAnesthesia

After performing a terminal bloodcollection, always be certain that the ani-mal is dead before placing the carcass inthe freezer. Remember that the animal

FIGURE 7. Location of the jugular vein inthe mouse.

FIGURE 8. Blood collection from the jugular vein.

Jugular Vein

with a scalpel or straight edge razor andcollect blood as it pools. It may beimportant to consider that tissue fluidswill contaminate the blood sample.

Blood Collection from theOrbital Sinus

Quickly remove the eyeball from thesocket with a pair of tissue forceps. Holdthe mouse in the palm of your hand overa collection tube. Massage the body ofthe mouse with your hand by squeezingthe rear half, then the middle, then thehead of the mouse so that you are “milk-ing” the blood toward the eye. Whendone properly, large drops of blood willflow from the orbital sinus.

Received 1/27/99; accepted 9/19/00.

References

1. Popesko, P., Rajtova, V., and Horak, J. A.Colour Atlas of the Anatomy of SmallLaboratory Animals, Volume 2, Rat,Mouse, Golden Hamster. WolfePublishing Ltd, London, England, 1992.

2. Cunliffe-Beamer, T. Biomethodology andSurgical Techniques. In: Foster, H.,

Small, D., and Fox, J.G., eds. TheMouse in Biomedical Research VolumeIII. Academic Press, New York, NY,1983.

3. Hem, A., et al. Saphenous vein puncturefor blood sampling of the mouse, rat,hamster, gerbil, guinea pig, ferret, andmink. Laboratory Animals; 32(4):364-367, 1998.

4. Kassel, R. and Levitan, S. A jugulartechnique for the repeated bleeding ofsmall animals. Science; 118:563-564,1953.

5. National Research Council, Institute forLaboratory Animal Research. Guide forthe Care and Use of Laboratory Animals.National Academy Press, Washington,DC, 1996.

6. Van herck, H., et al. Orbital sinus bloodsampling in cats as performed by differ-ent animal technician: the influence oftechnique and expertise. LaboratoryAnimals; 32: 377–386, 1998.

7. Sakaki, K. Hematological comparison ofthe mouse blood taken from the eye andthe tail. Exp. Anim.; 10:14-19, 1961.

8. Bannerman, R. Hematology. In: Foster,H., Small, D., and Fox, J.G., eds. TheMouse in Biomedical Research VolumeIII. Academic Press, New York, NY, 1983.

9. Waynforth, H.B. and Flecknell, P.A.Experimental and Surgical Technique inthe Rat, 2nd edition. Academic PressLtd., San Diego, CA, 1992.

10. Huerkamp, M. It’s in the bag: easy andmedically sound rodent gas anesthesiainduction. In press.

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FIGURE 9. Relative locations of the heart and diaphragm in the mouse thorax.

Diaphragm

Diaphragm

Last ribHeart