Venipuncture in Small Mammals

Key Points

  • Blood collection from small mammals can be difficult.
  • Carefully weigh the benefits of obtaining the blood sample versus the risk of the stress from the collection procedure on the patient.
  • Limit blood collection to no more than 0.5-1.0% of body weight.
  • The stress of transport and manual restraint can influence blood test results.
  • Unless the patient is extremely debilitated or extremely calm, use anesthesia or sedation to minimize the stress of handling and venipuncture, but remember that anesthesia can also affect test results.
  • Small blood samples may be collected from the lateral saphenous vein in virtually all small mammals.
  • The cranial vena cava is generally the quickest method to collect the largest amount of blood from the ferret.
  • The jugular vein and femoral vein are also popular venipuncture sites.


Hematological and serum chemistry tests are considered part of the minimum database, yet collecting blood samples from small mammals can be extremely challenging. To become proficient in small mammal venipuncture, understand the anatomic location of the vessels and their associated landmarks and then practice, practice, practice. Veterinary health professionals should also be aware of the potential risks associated with blood collection from these small species, especially those presenting in advanced diseased states.

There are a variety of published normal clinical values available, however laboratory results may be affected by a variety of factors including anesthesia, gender, age, stage of reproductive cycle, circadian rhythm, diet, season, restraint, and stress. The type of anticoagulant and the venipuncture site used can also affect blood test results. Unfortunately, published normal values rarely provide information on these parameters in exotic species. In-house reference ranges may be developed if your small mammal caseload is large enough, however this requires consistent methodology when collecting blood samples including the type of anesthetic agent and anticoagulant used.


Tips for sample collection

  1. Increase body temperature to promote vasodilation. Gently warm rodents by placing the cage on a heating pad set on low or placing the cage in an incubator at 39°C (102°F) for 5 to 10 minutes. Monitor the patient closely because overheating can lead to dehydration and hyperthermia. Dilate tail vessels by placing the tail in warm water (35-40°C or 95-104°F) for 5 to 10 seconds or placing a warm washcloth over the tail. Avoid this technique in chinchillas (Chinchilla lanigera) as they overheat very easily.
  2. With the exception of the ferret (Mustela putorius furo), most exotic pets are prey species that are easily stressed. Consider sedation (see #3 below) or isoflurane or sevoflurane anesthesia to minimize the stress of handling. An inexperienced handler and phlebotomist will also benefit from anesthetizing the patient. Many of the venipuncture sites recommended in small rodents, like rats and mice, have been developed in research settings where many animals are bled one right after another. Even with experience, few small mammal veterinarians have this opportunity. However with training, staff can become proficient in difficult venipuncture procedures and can minimize stress on conscious animals. Of course, anesthesia can also affect hematology. Ferrets anesthetized with isoflurane exhibit a rapid decrease in hematocrit, hemoglobin, and red blood cell count, and these values do not return to pre-anesthetic levels for 45 minutes.
  3. If general anesthesia is not used, consider sedating the pet with midazolam (0.25-1.0 mg/kg IM).
  4. Application of a topical anesthetic such as EMLA® cream (AstraZeneca LP, Wilmington, DE) over the vein 30 minutes prior to venipuncture can also prove helpful. Remember EMLA must be covered with a semi-occlusive dressing to work appropriately.
  5. Some small mammals, such as the rabbit (Oryctolagus cuniculus), rat (Rattus norvegicus), and Syrian hamster (Mesocricetus auratus), have a relatively short prothrombin time so whole blood clots quickly at room temperature. In some individuals, the needle and syringe can be pre-heparinized before collecting blood samples by drawing heparin into the needle and expelling the excess from the hub. Always check with your lab first, since heparin can cause changes in staining quality.
  6. The total volume of blood that can be safely collected typically ranges from 0.5-1.0% body weight (BW). Collect smaller volumes (0.5% BW) from geriatric patients or those suspected to have anemia or hypoproteinemia. Place samples into microtainers (Fig 1) (BD Microtainer Blood Collection Tubes; Becton Dickinson and Company, Franklin Lakes, NJ). For example, a maximum volume of 3.5-7.0 ml of blood may be collected from a 700-gram guinea pig (Cavia porcellus).


    Figure 1. Place small blood volumes into microtainers. Image provided by Jody Nugent-Deal. Click image to enlarge.

  7. Small veins may collapse when pierced by a needle on a syringe. It may be necessary to cannulate the blood vessel and to collect blood directly from the needle hub with a hematocrit tube (Fig 2).

    Tail vein

    Figure 2. Small vessels must sometimes be cannulated; blood is then collected directly from the needle hub. Image provided by J. Nugent-Deal. Click on image to enlarge.


Recommended sample collection sites in various species

Many of the blood collection sites and sampling techniques used in exotic mammals are described for cats and dogs. Other sites may be relatively unfamiliar, such as the cranial vena cava or femoral vein (Table 1). Click here for a PDF version of this table.

Also view the LafeberVet videos (or articles with images) illustrating Blood Collection in Rabbits and Blood Collection in Ferrets.


Table 1. Venipuncture sites for use in small mammal patients
Species Ear vessels Jugular vein Cephalic vein Femoral vessel* Saphenous vein Sub- mandibular vein Tail vessels Vena cava
Chinchilla (Chinchilla lanigera) + + + + +/-
Ferret (Mustela putorius furo) + + + +
Gerbil (Meriones unguiculatus) + +
Guinea pig (Cavia porcellus) +/- + + + +/-
Hamster + + + + +/-
Hedgehog (Atelerix albiventris) + + + + +/-

(Mus musculus)

+ + + +/- +
Rabbit (Oryctolagus cuniculus) + or +/- + + + +/-

(Rattus norvegicus)

+ + + + + +/-
Sugar glider (Petaurus breviceps) + + + + + +

Cannulation of the femoral vein is preferable, however the femoral artery may be utilized instead because of the close relationship of these vessels.
+ Vessels that are used most frequently or that may be used to collect large blood samples
+ Vessels of intermediate or moderate use
+/- Vessels that should be utilized only as a last resort


Cephalic vein

Utilize the cephalic vein for collection of small blood volumes in the chinchilla, ferret, hedgehog (Atelerix albiventris), rabbit, hamster, and sugar glider (Petaurus breviceps). Depending on patient size, use a 25- to 27-gauge needle on a 0.5-1.0-mL syringe to reduce the risk of vessel collapse.


Saphenous vein

Small blood samples may be collected from the lateral saphenous vein in virtually all small mammals (Fig 3 and Fig 4). Use of the medial saphenous vein has also been described in mice. Go to the Norwegian Reference Centre for videos of saphenous venipuncture in the mouse (Note: Use of a scalpel blade to shave the fur is likely unwise in a pet mouse).

Saphenous venipuncture

Figure 3. Saphenous venipuncture in a Syrian hamster (Mesocricetus auratus). Image provided by J. Nugent-Deal. Click image to enlarge.

Blood collection from rabbit

Figure 4. Blood collection from the saphenous vein of a rabbit (Oryctolagus cuniculus). Image provided by J. Nugent-Deal.Click image to enlarge.

  • Place the animal in ventral or lateral recumbency.
  • Shave fur or apply alcohol over the vein to improve visualization.
  • Grasp the rear leg just above or below the stifle to engorge the vessel or apply a tourniquet, such as a rubber band or penrose drain clamped with a hemostat.
  • Use a 22- to 30-gauge needle on a 0.5-1.0-mL syringe to reduce the risk of vessel collapse.
  • In a laboratory setting, petroleum jelly or silicone grease (McNett Silicone Grease 100% Pure Silicone Lubricant, McNett Corp.) is sometimes applied over the vein of small patients like mice to prevent blood from spreading into the fur. This aids in easier collection of blood into a microhematocrit tube.


Tail vessels

Small amounts of blood may be collected from lateral tail veins in the gerbil, rat, and mouse, the dorsal tail vein in the rat (Fig 2 above), or the ventral tail vein in the sugar glider. These vessels are usually superficial and can easily be seen unless the animal is very debilitated or obese. Warm the tail beforehand. Use a 22- to 27-gauge needle on a 0.3 -1.0-mL syringe. A 27 or 25-gauge needle may also be placed into the vessel. Once there is blood in the hub of the needle, a hematocrit tube can be used to collect the blood. Avoid squeezing or milking the tail during sampling, and provide appropriate hemostasis afterwards. Gerbils are prone to degloving tail injury and should be handled particularly carefully.

Lateral tail vein rat

Figure 5. Cannulation of the lateral tail vein in a rat (Rattus norvegicus). Photograph provided by J. Nugent-Deal. Click image to enlarge.

Blood collection from the ventral tail artery in the mouse and rat is similar to that described above. Place the pet in dorsal recumbency, and aseptically prepare the venipuncture site, which sits approximately one-third of the distance from the tail base. Attach a 23-30-gauge needle to a plunger-less syringe and insert the needle bevel up at a 30° angle on the ventral midline of the tail. Provide good hemostasis afterwards.


Cranial vena cava

The cranial vena cava is generally the quickest method to collect the largest amount of blood from the ferret and sugar glider. The vena cava may also be venipuncture site of last resort in the chinchilla, hamster, rabbit, rat, and particularly the guinea pig and hedgehog. In these latter two species the neck is very short and the heart lies very cranial within the thorax in these species. This increases the risk of hemorrhage if the needle punctures the thin-walled atria. General anesthesia or heavy sedation is required. The ferret heart is located caudally within the long thoracic cavity and the risk of cardiac puncture with caval sticks is minimal. Nevertheless, the vena cava can still be lacerated if the ferret moves and all but the most severely debilitated ferrets should also be sedated or anesthetized.

Caval venipuncture is a blind technique, which relies on the identification of anatomic landmarks.

  • Place the pet in dorsal recumbency with the forelegs pulled back along the sides of the chest.
  • Position the animal squarely on the table in perfect alignment.
  • Identify the thoracic notch, the angle created by the manubrium and the first rib.
  • Insert a 25-to 27-gauge needle on a 1.0-3.0 ml syringe at a 30° to 45° angle.
    (Use a more shallow angle of 20° to 35° in the guinea pig).
  • Direct the needle towards the opposite hip, while maintaining slight negative pressure.
  • Once blood starts to fill the syringe, do not move the needle.
  • If blood is not aspirated, slowly pull the needle out until it lies just beneath the skin, then redirect a little towards midline. Alternatively, pull out completely, and start over using a fresh needle.
Caval venipuncture

Figure 6. Caval venipuncture in a ferret (Mustela putorious furo). Photograph provided by J. Nugent-Deal. Click image to enlarge.


Jugular vein

Restraint for jugular venipuncture will depend on the species as well as the preference of the phlebotomist. General anesthesia or sedation is often required to collect blood from the jugular vein in the guinea pig, hedgehog, and sugar glider. Although jugular venipuncture may be performed in conscious ferrets, chinchillas, and rabbits, it may be best for both patient and phlebotomist to use chemical restraint if the patient is stressed or uncooperative.

The jugular vein tends to lie in a more lateral position in the ferret, when compared to the cat. Jugular venipuncture can be difficult in hedgehogs unless they are thin, and guinea pigs have very short necks making it difficult to draw blood from their jugular veins as well. Jugular venipuncture may also be difficult in obese rabbits or does with a large dewlap (Fig 7). Jugular venipuncture may also be attempted in the hamster, rat, and mouse but this site can be quite challenging and requires heavy sedation or general anesthesia.

Fat rabbit

Figure 7. A large dewlap may hinder jugular venipuncture in does or obese rabbits (Oryctolagus cuniculi). Image provided by J. Nugent-Deal. Click image to enlarge.

Most species may be placed in dorsal or lateral recumbency with the head extended and the legs pulled down toward the body (Fig 8). Ferrets, chinchillas, guinea pigs, and rabbits may also be held in sternal recumbency at the edge of the exam table (Fig 9). Extend the head upward and pull the front legs down as in the cat. Take care not to overextend the head and neck, and carefully monitor breathing.

Jugular venipuncture, chinchilla

Figure 8. Jugular venipuncture in a chinchilla (Chinchilla lanigera) placed in dorsal recumbency. Photo by J. Nugent-Deal. Click image to enlarge.

Jugular venipuncture in a rabbit

Figure 9. Jugular venipuncture in a rabbit (Oryctolagus cuniculus) held in ventral recumbency. Photo by J. Nugent-Deal. Click image to enlarge.


Femoral vessel

Blood sample collection from the femoral artery or vein is generally the quickest method that will yield the largest amount of blood in the guinea pig, rat, mouse, and hamster (Fig 9). This venipuncture site has been described in the literature in most small mammals except for the ferret, rabbit, and gerbil. General anesthesia is usually required to collect samples in small mammals.

Femoral venipuncture

Figure 10. Femoral venipuncture is often the quickest way to obtain a large volume of blood. Provided by J. Nugent-Deal. Click image to enlarge.

  • Place the animal in dorsal recumbency and position the lower leg so that the femur lies at a 90° angle to the long axis of the body.
  • Palpate the pulse in the femoral triangle. The pulse may be found slightly behind the femur along the upper half of the thigh. The femoral artery runs parallel and slightly in front of the vein. The optimal site for venipuncture is where the body wall meets the upper thigh. The nipple may be used as a landmark in the guinea pig.
  • Direct a 23- to 27-gauge, 3/4-inch needle on a 1- to 3-mL syringe needle perpendicular to the skin or at a 45° angle to the femur.
  • Insert the needle into the tissue using gentle negative pressure.
  • If the blood collected is bright red, the femoral artery has likely been cannulated. Apply pressure to the venipuncture site for a minimum of 2-5 minutes.


Ear vessels

Considerable variability exists in the literature and in personal opinion over use of ear vessels in the rabbit. Everyone agrees that rabbit blood vessels are fragile and prone to hematoma formation. Even development of a mild ear hematoma or bruising may be unacceptable to some owners. Opponents of ear venipuncture also point out that disruption of blood flow can potentially lead to thrombosis, ischemia, and sloughing of pinnal tissue. Therefore the ear is only used as a last resort in some private practices.

Proponents state that the central ear artery and marginal ear vein (Fig 10) are readily accessible, and that the rabbit pinna has significant collateral circulation in the form of arteriovenous anastomoses. Anecdotally, even with significant hematoma formation the incidence of problems secondary to venipuncture are rare. Some also advocate this technique in high-strung rabbits since the patient is allowed to sit normally, and only the rare rabbit seems to notice when the needle punctures the pinna.

Rabbit ear vessels

Figure 11. Central ear artery and marginal ear veins in the rabbit (Oryctolagus cuniculus). Photo provided by J. Nugent-Deal. Click image to enlarge.

  • Restrain the rabbit in sternal recumbency, wrapping the animal in a large cloth to avoid inadvertent movement.
  • Gently shave fur or apply alcohol over the vessel to improve visualization (Fig 11).
  • It may also help to warm the ear by gently stroking and tapping the ear, applying a warm towel or warm water, or directing a low-watt light bulb towards the vessel.
  • Multiple techniques are described for cannulation. Use a 21-23 gauge needle or butterfly catheter in large or medium-sized rabbits. Select a 23-26-gauge needle in small rabbits. When using a 1-3 cc syringe it is important not to apply too much negative pressure. Some individuals prefer to use a needle or butterfly catheter alone, catching blood in an appropriate collection tube. Blood often flows best if the hub is broken off of a 1-2 inch needle.
  • Applying tension to the ear will help with needle insertion. Direct the needle toward the base of the ear, with the insertion point closer to the tip of the ear.
  • To minimize the risk of hematoma formation, insert the needle parallel to the artery and then direct the needle into the vessel once it is in the subcutaneous space.
  • If the vessel collapses, gently stroke the ear until the vessel relaxes and blood begins to flow.
  • Apply digital pressure over the collection site for 5 minutes or so for hemostasis.

In a laboratory setting, ear veins are sometimes used for collection of small quantities of blood in the guinea pig. The ear veins are nicked with a 25-gauge needle so that blood may be collected into a hematocrit tube.

Rabbit ear vein venipuncture

Figure 12. Venipuncture of the marginal ear vein in a rabbit (Oryctolagus cuniculus). Photograph provided by J. Nugent-Deal. Click image to enlarge.


Submandibular vein

The submandibular vein is sometimes used in a laboratory setting, but is not recommended in the pet mouse. The orbital and submandibular veins join just behind the mandibular joint to form the jugular vein. Small amounts of blood may be collected from the anesthetized mouse or rat where these vessels meet. Potential risks include bleeding into the mouth, which can cause aspiration and potential death, or bleeding into the ear canal. See Medipoint, Inc for pictures of this blood collection site.


Although it can be difficult to collect blood from the small veins of exotic small mammals, one can take some consolation in knowing that there are similarities in the vascular anatomy between these animals and larger domestic species. Many of the venipuncture sites and sampling techniques used in small mammals are similar to those described for cats and dogs. For some sites and techniques, however, an inexperienced or infrequent handler and phlebotomist will benefit from anesthetizing the patient. General anesthesia also reduces the stress of restraint and handling on the patient.



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To cite this page:

Nugent-Deal J, Pollock C. Venipuncture in small mammals. LafeberVet website. Available at