UNIVERSITY of WISCONSIN–MADISON
COMPLIANCE MY UW MAP
RARC link
IACUC Overview
CALS IACUC
LSVC IACUC
SMPH IACUC
SVM IACUC
Meeting Calendar
Semiannual Inspections
Training
Pathology
Pharmacy
Program Assessment
Rodent Analgesia
Rodent Quarantine
Veterinary Care
Equipment Lending
Protocol Overview
What All PIs Need to Know
ARROW
Request Grant Congruence
Submit
Comply
Amend
Renew, Update
Terminate
RARC Documents and Forms
Animal Transfer
Animal Transport
Facilities
Inspections
Species

Policies Overview
Animal User Requirements
Policies by Number
Policies by Authority
Policies by Topic
My Profile
Login Contacts

Rodent Blood Collection

Contact an RARC trainer for training or guidance on using these blood collection techniques.

Factors to consider when choosing the blood withdrawal technique:

Average Blood Volume:

Approximate Blood Sample Volume Ranges and Safe Frequency of Collection
Body Weight (g) 1% (mL) 10% (mL) 15% (mL)* 20% (mL)*
20 0.01 0.14 0.22 0.30
25 0.02 0.18 0.27 0.36
30 0.02 0.22 0.32 0.43
35 0.03 0.25 0.38 0.50
40 0.03 0.28 0.43 0.58
         
125 0.08 0.8 1.2 1.6
150 0.12 1.18 1.78 2.37
200 0.13 1.3 1.92 2.56
250 0.16 1.6 2.4 3.2
300 0.19 1.92 2.88 3.84
350 0.22 2.24 3.36 4.48
Frequency of Collection Daily Once a week Every 2 weeks Every 4 weeks
*An equal volume of warmed normal saline or LRS should be given IP to the rodent after this collection to help prevent sequelae due to blood volume loss.

Survival Blood Sampling Techniques

Mouse

Retro-Orbital Sinus

Obtainable Volume

  • Small amount (~50µL)

Sample Collection Information

  • Good sample quality.
  • Samples can be collected from both lateral and medial canthus of the eye.
    • Typically only collect from one canthus per eye/per collection.
    • The medial canthus is good for collecting a single capillary tube.
    • The lateral canthus can be used for collecting blood into a blood collection tube via a capillary tube.
  • Anesthesia is required.
    • Mice must be anesthetized unless justification is provided that anesthesia is not compatible with the experimental design.
  • Poor technique can cause damage to eye.

Example Collection Description

Blood Collection Medial Canthus

Place the anesthetized mouse on its side and the thumb and index finger of your nondominant hand around the eye to stretch the skin and cause the eye to bulge slightly. Place a capillary tube at the medial canthus of the eye and advance it into the sinus. Twist the capillary tube slightly to break the vessels, and the tube fills by capillary action. Once the appropriate volume is collected, remove the tube and hold the eye closed with slight pressure to stop further bleeding.

Blood Collection Lateral Canthus

Hold the anesthetized mouse by the scruff and advance a capillary tube into the lateral canthus of the eye. Twist the capillary tube slightly to break the vessels, and the tube fills by capillary action. Allow blood to flow through the capillary tube into a blood collection tube. Once the appropriate volume is collected, remove the tube and hold the eye closed with slight pressure to stop further bleeding.

Maxillary Vein

Obtainable Volume

  • Medium to large amount (100–200µL)
  • Approximately 3 large drops of blood = 100µL

Sample Collection Information

  • Repeat sampling is possible.
  • Sample quality is typically good but can have some saliva contamination.
  • The procedure is customarily done on an unanesthetized animal; effective restraint is required.
  • Ensure that gentle pressure is applied for approximately 30 seconds postcollection to ensure hemostasis.
  • Use of needles in lieu of lancets is discouraged: they have a propensity to cause damage to underlying tissues.
  • Recommended lancet point length for mice:
    • 2–8 week old—4mm
    • 2–6 month old—5mm
    • Over 6 months old—5.5mm

Example Collection Description

Restrain a conscious mouse by the scruff and use a lancet of appropriate size to puncture the vein, slightly behind the mandible and in front of the ear canal. After the vein is punctured, collect blood in an appropriate tube. Then, if necessary, apply pressure with a gauze sponge until the bleeding stops.

Lateral Tail Vein

Obtainable Volume

  • Small to medium amount (50–100µL)

Sample Collection Information

  • Using a needle to collect a sample minimizes contamination but is difficult to perform in the mouse.
  • Nicking the vessel to collect a sample is easy but produces a sample of variable quality that may be contaminated with tissue and skin products.
  • Sample quality decreases with prolonged bleeding times and tail stroking.
  • Repeated collection is possible.
  • This method is relatively nontraumatic.
  • Routinely done without anesthesia, although effective restraint is required.
  • In most cases, warming the tail with the aid of a heat lamp or warm compresses will increase obtainable blood volume.
  • Piercing the tail vein with a needle is also a way to collect a very small blood sample (~10µL)

Example Collection Descriptions

Mouse Lateral Tail Vein Blood Collection w/Needle and Syringe

Place the mouse in a restraint tube and warm the tail using a heated gel pack, light source, or recirculating water blanket to dilate the lateral veins. Insert a 1ml syringe and 27g needle into the vein while pulling the plunger back slightly until a flash of blood appears in the hub of the syringe; continue pulling back on the plunger slowly to collect the remaining sample. Apply slight pressure after the needle is removed to ensure bleeding has stopped.

Mouse Lateral Tail Vein Blood Collection w/Sterile Blade

Place the mouse onto a wire-bar lid and make a small transverse cut to the lateral tail vein with a sterile blade. Collect blood into an appropriate tube. Prior to bleeding, warm the mouse’s tail by using a heated gel pack, light source, or recirculating water blanket to dilate the lateral veins. Apply slight pressure after the collection to ensure bleeding has stopped.

Saphenous Vein

Obtainable Volume

  • Small to medium amount (50–100µL)

Sample Collection Information:

  • Method can be used in mice by piercing the saphenous vein with a needle (22–23g) or lancet (4–5mm).
  • Repeat sampling is possible.
  • Variable sample quality.
  • The procedure is customarily done on an unanesthetized animal; effective restraint is required.
  • Can be more time-consuming than other methods due to time required for site preparation.
  • Prolonged restraint and site preparation time can result in increased animal distress when handling an unanesthetized animal.
  • The animal may temporarily favor the limb following the procedure.
  • Care must be taken to ensure adequate hemostasis following the procedure.

Example Collection Descriptions:

Place the mouse in a restraint tube (e.g., 50mL conical tube) and clip the lateral surface of the hind leg. Grasp the skin at the front of the mouse’s rear leg to both restrain the leg and cause the saphenous vein to expand. Place petroleum jelly or artificial tears on the leg to allow blood to bead up on the skin surface after the vessel is punctured. Puncture the vessel using a 22–23g needle or a 4-5mm lancet and use a capillary tube to collect the blood. Apply pressure with a gauze sponge after the sample is collected until the bleeding stops.

Pedal Vein

Obtainable Volume

  • Small amount (~50µL)

Sample Collection Information

  • Method can be used in mice by piercing the pedal vein with a needle (23g) or lancet (4–5mm).
  • Repeat sampling is possible.
  • Variable sample quality.
  • The procedure is customarily done on an unanesthetized animal; effective restraint is required.
  • Can be more time-consuming than other methods due to time required for site preparation.

Example Collection Description

Place the mouse in a restraint tube (e.g., 50mL conical tube) and place a tourniquet on the mouse’s leg. Apply petroleum jelly or artificial tears on top of the hind foot to allow blood to bead up on the skin surface after the vessel is punctured. Puncture the vessel using a 23g needle or 4-5mm lancet, and use a capillary tube to collect the blood. After the sample is collected, release the tourniquet and apply pressure with a gauze sponge until the bleeding stops.

Rat

Jugular Vein

Obtainable Volume:

  • Medium to large amount (0.5ml+)

Sample Collection Information:

  • Able to collect a larger volume of blood.
  • Two technicians are needed if the rat is not anesthetized.
  • May require special equipment (bleeding board).

Example Collection Description

Restrain the rat with the use of a bleeding board to ensure proper position. Use a head cone to rotate the rat’s head away from the bleeding site to allow a good view of the bleeding landmarks. Slowly insert the needle (20-22g on a 3ml syringe), drawing back on the plunger at the same time. A flash of blood in the hub of the syringe indicates the vein has been successfully located. Stop inserting the needle and draw the plunger back to obtain the desired sample. After the sample is obtained, release the rat from the bleeding board and observe to ensure bleeding has stopped before returning the rat to its normal housing.
Ventral Artery

Obtainable Volume:

  • Medium amount (0.2-0.4mL)

Sample Collection Information:

  • Using a needle to collect a sample minimizes contamination of the sample.
  • Nicking the vessel is easy but produces a sample of variable quality that may be contaminated with tissue and skin products.
    • Sample quality decreases with prolonged bleeding times and tail stroking.
  • Repeated collection is possible.
  • Relatively nontraumatic.
  • Routinely done without anesthesia, although effective restraint is required.
  • In most cases, warming the tail with the aid of a heat lamp or warm compresses will increase obtainable blood volume.
  • Arterial sampling produces larger volumes and is faster, but special care must be taken to ensure adequate hemostasis.
  • Piercing the tail vein with a needle is also a way to collect a very small blood sample (~10µL)

Example Collection Descriptions

Place the rat in a restraint tube and warm the tail using a gel pack, light source, or recirculating water blanket to dilate the artery. Use a 3ml syringe and a 22-23g needle. Insert the needle into the artery while pulling back slightly on the plunger until a flash of blood appears in the hub of the syringe; continue pulling the plunger back slowly to collect the remaining sample. Apply slight pressure after the needle is removed to ensure bleeding has stopped.
Saphenous Vein

Obtainable Volume

  • Small to medium amount (0.2-0.4mL)

Sample Collection Information

  • Can be used in both rats by piercing the saphenous vein with a needle (22g) or lancet (5mm).[CLARIFY MEANING OF "BOTH"]
  • Repeat sampling is possible.
  • Variable sample quality.
  • The procedure is customarily done on an unanesthetized animal; effective restraint is required.
  • Can be more time-consuming than other methods due to time required for site preparation.
  • Prolonged restraint and site preparation time can increase distress for an unanesthetized animal.
  • An animal may temporarily favor the limb following the procedure.
  • Care must be taken to ensure adequate hemostasis following the procedure.

Example Collection Description

Place the rat in a restraint tube and clip the lateral surface of the hind leg. Place a tourniquet on the rat’s leg and put petroleum jelly or artificial tears on the leg to allow blood to bead up on the skin surface after the vessel is punctured. Use a 22g needle or 5mm lancet to puncture the vessel and use a capillary tube to collect the blood. After the sample is collected, release the tourniquet and apply pressure with a gauze sponge until the bleeding stops.

Pedal Vein

Obtainable Volume

  • Small amount (0.1-0.2mL)

Sample Collection Information

  • Can be used in rats by piercing the pedal vein with a needle (25g) or lancet (5mm).
  • Repeat sampling is possible.
  • Variable sample quality.
  • The procedure is customarily done on an unanesthetized animal; effective restraint is required.
  • Can be more time-consuming than other methods due to time required for site preparation.

Example Collection Description

Place the rat in a restraint tube and place a tourniquet on the rat’s leg. Put petroleum jelly or artificial tears on top of the hind foot to allow blood to bead up on the skin surface after the vessel is punctured. Use a 22-23g needle or 5mm lancet to puncture the vessel and use a capillary tube to collect the blood. After the sample is collected, release the tourniquet and apply pressure with a gauze sponge until the bleeding stops.

Terminal Blood Sampling Techniques

Mouse

Cardiac Puncture

Obtainable Volume

  • Large amount (0.7–1.0 mL)

Sample Collection Information

  • This is a terminal collection procedure.
    • If the abdominal cavity or thoracic cavity is opened to visualize the heart, it is considered a terminal surgery procedure.
  • Animal must be anesthetized or euthanized (e.g., CO2 euthanasia) prior to the procedure.
  • Very good sample quality.
  • A secondary means of euthanasia (e.g., cervical dislocation) is strongly recommended postexsanguination.

Example Collection Description

Insert a 20–23g needle on a 1-ml syringe vertically just behind the xiphoid cartilage at an angle of 15–20 degrees and advance toward the apex of the heart for the blood withdrawal. The animal is euthanized before recovering from anesthesia.

Axillary Cut Down

Obtainable Volume

  • Large amount (0.4–0.7mL)

Sample Collection Information

  • This is a terminal collection procedure.
  • Animal must be anesthetized or euthanized (e.g., CO2 euthanasia) prior to the procedure.
    • If animal is anesthetized for procedure this is considered a terminal surgical procedure.
  • Typically contaminated sample quality.

Example Collection Description

Anesthetize the mouse and place on its back. Make an incision or cut through the skin at the side of the thorax into the angle of the forelimb to expose the axillary vessels. Cut the vessels to allow the blood to pool in the area created by the tented skin. Collect the blood using a 1–3 ml syringe or bulb pipette to aspirate the pooled sample. Euthanize the animal without recovery from anesthesia.

Dorsal Aorta

Obtainable Volume

  • Large amount (0.4–0.7mL)

Sample Collection Information

  • This is a terminal collection procedure.
  • An animal must be anesthetized or euthanized (e.g., CO2 euthanasia) prior to the procedure.
    • If the animal is anesthetized for the procedure, it is considered a terminal surgical procedure.

Example Collection Description

Open the abdomen; displace the organs to allow visualization of the dorsal aorta. Thread a 23–25g needle bevel down into the aorta and collect the blood using a 1-ml syringe. Euthanize the animal without recovery from anesthesia.

Rat

Cardiac Puncture

Obtainable Volume

  • Large amount (7-10mL)

Sample Collection Information

  • This is a terminal collection procedure.
    • If the abdominal cavity or thoracic cavity is opened to visualize the heart, it is considered a terminal surgery procedure.[IS "procedure" NECESSARY?]
  • Animal must be anesthetized or euthanized (e.g., CO2 euthanasia) prior to the procedure.
  • A secondary means of euthanasia (e.g., creation of a bilateral pneumothorax) is strongly recommended postexsanguination.
  • Very good sample quality.

Example Collection Description

Dorsal Position

Anesthetize the rat and place on its back with the nose pointing away from you and insert a 20-22-g, 1 to 1 1/2” needle attached to a 3-10 ml syringe or Vacutainer just posterior to the xiphoid cartilage and slightly right of the midline (the animal’s left). Introduce the needle at approximately 30 degrees from the horizontal axis of the sternum in order to enter the heart. Unless the rat is very small (less than 200g), insert the needle all the way to the hub. Insert a 25g needle on a 3-10 mL syringe vertically just behind the xiphoid cartilage at an angle of 15-20 degrees and advance toward the apex of the heart for the blood withdraw. If a needle and syringe is used, draw the plunger back to show a flash of blood when the heart is entered, and then draw back slowly to avoid collapsing the heart. If a Vacutainer is used, push the tube onto the needle once the needle is inserted into the chest cavity. Animals must immediately be euthanized without recovery from anesthesia.

Lateral Position

Anesthetize the rat and place with its right side down on a flat surface. Advance a 20-22g, 1 to 1-1/2” needle attached to a 3-10 ml syringe or Vacutainer between the ribs on the animal's left side, immediately posterior to the elbow at the point of maximum heartbeat during palpation. The heart is approximately 1 cm cranial to the xyphoid process. If a needle and syringe is used, draw the plunger back to show a flash of blood when the heart is entered, and then draw back slowly to avoid collapsing the heart. If a Vacutainer is used, push the tube onto the needle once the needle is inserted into the chest cavity. Animals must immediately be euthanized without recovery from anesthesia.

Axillary Cut Down

Obtainable Volume

  • Large rat (5-7mL)

Sample Collection Information

  • This is a terminal collection procedure.
  • The animal must be anesthetized or euthanized (e.g., CO2 euthanasia) before the procedure.
    • If the animal is anesthetized for the procedure, it is considered a terminal surgical procedure.
  • Typically contaminated sample quality.

Example Collection Description

Anesthetize the rat and place on its back. Make an incision or cut through the skin at the side of the thorax into the angle of the forelimb to expose the axillary vessels. Cut the vessels to allow blood to pool in the area created by the tented skin. Use a funnel and a 50 ml conical tube to collect the blood. Euthanize the animal without recovery from anesthesia.

Dorsal Aorta

Obtainable Volume

  • Large rat (5-7mL)

Sample Collection Information

  • This is a terminal collection procedure.
  • An animal must be anesthetized or euthanized (e.g., CO2 euthanasia) before the procedure.
    • If the animal is anesthetized for procedure, it is considered a terminal surgical procedure.
  • Typically contaminated sample quality.

Example Collection Description

Anesthetize the rat and place on its back. Open the abdomen and displace the organs to allow visualization of the dorsal aorta. Thread a 22-23g needle bevel down into the aorta, and collect the blood using a 5-10ml syringe. Euthanize the animal without recovery from anesthesia.

Other Collection Techniques

Rat

Sublingual Vein

Obtainable Volume

  • Medium to large amount (0.5ml+)

Sample Collection Information

  • Able to collect a larger volume of blood.
  • Less tissue damage than some techniques.
  • Anesthesia required.
  • Some contamination of blood sample with saliva.

Example Collection Description

While an assistant holds the anesthetized rat in a supine position and also gathers up the loose skin at the neck, providing a partial stasis in the venous blood flow, extend the rat's tongue and grasp it with your forefinger. The sublingual veins will be visible at the base of the tongue. Use a 23g needle to puncture the vein and then immediately turn the rat back into a prone position and allow the blood to drip into a collection tube. Once the appropriate volume of blood is collected, release the rat and return it to the supine position in order to observe that the bleeding has stopped.

jump to top button
Internet Explorer is not compatible with this website.

Please use Edge, Firefox, Safari, Chrome or Opera.