MCGILL UNIVERSITY
UNIVERSITY ANIMAL CARE COMMITTEE
UACC Standard Operating Procedure # 1
October 2005 version
BLOOD COLLECTION - RODENTS (Rats, mice and guinea pigs)
1. INTRODUCTION
Standard Operating Procedures (SOPs) provide a detailed description of commonly used procedures. SOPs offer
investigators an alternative to writing detailed procedures on their protocol forms. Any deviation from the approved
procedures must be clearly described and justified in the Animal Use Protocol form (AUP). Approval of the protocol
indicates approval of the deviation from the SOP for that project only. A signed SOP cover page must be attached
to the Animal Use Protocol form. The relevant SOP number must be referred to in the Procedures section.
2. INFORMATION REQUIRED
2.1 Species:
Age:
Approximate weight:
2.2 Route: Put an X next to the one to be employed:
X
Route
Survival procedures:
1. Lateral saphenous vein - preferred
2. Tail artery
3. Jugular vein (percutaneous)
4. Tail tip
Terminal procedures:
7. Cardiac puncture – preferred
8. Trunk blood
9. Retro-orbital sinus
10. Other, specify:
2.3 Volume per collection:
2.4 Frequency of collection:
if repeated how frequent (interval)?
Replacement volume: No
Yes
specify:
2.5 Are there changes to this SOP indicated in the AUP form?
If yes, specify changes:
2.6 PI signature: __________________________________
YES
NO
Date: ________________________
PLEASE ATTACH ONLY THIS SIGNED COVER SHEET TO THE BACK OF EACH
RELEVANT AUP FOR ANIMAL CARE COMMITTEE APPROVAL.
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3. ACCEPTABLE COLLECTION VOLUMES FOR ALL SPECIES
3.1 Single collection:
 Up to 10% of circulating blood volume with fluid replacement in normal, healthy animals.
 Blood withdrawal and fluid replacement must be performed slowly (over 1-2 minutes) and at a
steady rate.
 Fluid replacement: Sterile, warm physiologic saline equal to the volume of blood collected
administered intravenously or intraperitoneally.
 Can be repeated only in 3-4 weeks (time required for erythrocyte and blood volume
regeneration).
3.2 Maximal acceptable volume in a single bleed (ml) :

= 10% X circulating blood volume (ml/kg) X weight (kg)
3.3 Circulating blood volume :
 Rat - 60 ml/kg (e.g. 15 ml for 250 gram rat)
 Mouse – 80 ml/kg (e.g. 2 ml for 25 gram mouse)
 Guinea pig – 80 ml/kg
3.4 Repetitive blood collections at short intervals :
 Up to total of 1% of circulating blood volume can be removed every 24 hours in normal, healthy
animals.
 Blood withdrawal and fluid replacement must be performed slowly (over 1-2 minutes) and at a
steady rate.
 Fluid replacement: Sterile, warm physiologic saline equal to the volume of blood collected
administered intravenously or intraperitoneally.
BLOOD COLLECTION: IMPORTANT CLINICAL SIGNS
 Hypovolemic shock: fast and thready pulse, pale and dry mucous membranes, cold skin and extremities,
restlessness, hyperventilation and subnormal body temperature.
 Anemia: pale mucous membranes, pale tongue, gums and footpads, exercise intolerance, increased
respiratory rate at rest.
4. ACCEPTABLE BLOOD COLLECTION PROCEDURES
PREANAESTHETIC FASTING:
 Rats and mice do not vomit, and as they also possess a high metabolic rate preanaesthetic fasting is not
recommended. Guinea pigs must be fasted 2-4 hours prior to anaesthesia to reduce the volume of
gastric contents. Do not withhold water.
4.1 Volumes less than 100μl :
Method 1. Tail tip – Rats and mice
 Perform on unanaesthetized animal if mouse is less than three (3) weeks of age and rat less
than two (2) weeks of age. Anesthesia is required if mouse is older than 3 weeks, rat older than 2
weeks.
 Place animal in approved animal restrainer. In order to vasodilate the tail blood vessels the entire
animal can be carefully warmed with a carefully placed heat lamp for 5-10 minutes or the tail can
be immersed in warm water.
 Remove any bedding material or faeces from the tail, wash with a surgical skin disinfectant and
rinse with water.
 Place the animal on a clean work surface.
 Using a fresh scalpel blade, cut 1-2mm of the distal tail at an angle perpendicular to the work
surface. Change scalpel blades between animals.
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Apply gentle pressure proximal to the collection site to occlude venous return and ease
collection. Collect the blood in a capillary tube or other suitable collection device.
Apply gentle digital pressure to the wound for 30 - 45 seconds with a clean gauze pad to stop any
haemorrhaging. For persistent bleeding, apply a silver nitrate stick to the wound.
Return the animal to its cage.
Serial samples can be obtained over a short period of time by gently removing the scab.
Maximum tail length harvested over a single individual’s lifetime is 1-2 mm. If excessive amounts
of tail are harvested, it can lead to locomotory problems in rats and the development of
granulomas.
Method 2. Lateral saphenous vein – Rats, mice and guinea pigs
 Performed on unanaesthetized animal as anaesthesia causes hypotension and makes blood
collection difficult.
 Place animal in approved restrainer.
 Extend the hind leg and fix the limb by holding fold of skin between tail and thigh.
 Shave or clip hair on outer surface of lower hind leg to visualize the vein. Swab skin with alcohol
and allow it to dry completely.
 Apply vaseline on skin prior to nick to prevent wicking of blood onto skin and to ease collection.
 Apply pressure to vein upstream from nick to raise vein.
 Perpendicular to vein, nick it with a 25 gauge needle and collect blood with microhematocrit
(capillary) tube.
 Apply gentle digital pressure with a clean gauze pad for 30-45 seconds to site to stop blood flow
and prevent hematoma formation.
 Serial samples can be obtained over a short period of time by gently removing the scab.
Method 3. Jugular vein (Percutaneous) - Rats and guinea pigs
 The animal is anaesthetized (UACC SOP # Rodent Aneasthesia ) and at an appropriate depth of
anaesthesia prior to commencement. Signs that indicate a satisfactory plane of anaesthesia
includes a lack of response to a toe pinch and respirations that are relaxed and regular.
 The anaesthetized animal is placed with its back with its head towards the operator.
 Extend the neck and clip or shave the hair lateral to the midline on the side where the blood will
be collected. Disinfect the skin with 70% alcohol or povidone iodine.
 Using the angle of the jaw as a guide, place a 3 ml syringe with a 21 gauge needle alongside the
jaw. Angle the syringe at a 30 angle and gently push through the skin, following the line between
the angle of the jaw and the shoulder.
 Slowly withdraw the plunger until blood is seen indicating puncture of the vein. Collect blood.
 Apply gentle digital pressure with a clean gauze pad for 30-45 seconds to site to stop blood flow
and prevent hematoma formation.
 Jugular vein bleeding can be repeated over a short period of time by gently removing the scab.
 Serial samples can be obtained over a short period of time by gently removing the scab. Interval
between samples should be at least 24 hours and preferably 48 hours.
CHRONIC INTRAVENOUS CATHETERIZATION: IMPORTANT POINTS
 The vein will vasoconstrict when handled so this must be kept to a minimum. Leaving the vein
untouched for 1-2 minutes will allow it to regain some of its original size.
 Daily aseptic catheter maintenance is mandatory in order to ensure its patency and decrease the
likelihood of infection at the catheter entry site. The heparin solution in the catheter must be
removed daily and sterile saline used to flush any residual blood in the catheter is flushed back
into the circulation. Omitting this step might lead to a steady decline in the animal’s hematocrit.
The catheter is then refilled with a fresh heparinized saline solution.
 Daily inspection of the incision site any signs of inflammation (redness, swelling) or infection
(purulent discharge, unusual odor).
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Method 4. Jugular vein (Indwelling intravenous catheter) - Rats and guinea pigs
 Recommended technique for repetitive, chronic blood sampling studies.
a) Catheter preparation
 Common catheter materials include polyethylene, silicone rubber and polyurethane.
 A catheter guide provides rigidity to the catheter, facilitating introduction and placement in a
blood vessel.
 The end of the catheter must be fitted with a standard connector that is compatible with the hub of
standard syringe sizes.
 The catheter and all accompanying materials must be sterilized prior to use.
b) Surgical procedure
 The animal is anaesthetized (UACC SOP # 2 Rodent Aneasthesia ) and at an appropriate depth
of anaesthesia prior to commencement. Signs that indicate a satisfactory plane of anaesthesia
ncludes a lack of response to a toe pinch and respirations that is relaxed and regular.
 The anaesthetized rat is laid on its back with the head toward the surgeon.
 Measure the distance the catheter must be advanced at this time. For example, for rats weighing
between 300-500 grams, 3-4 cm of the catheter needs to be inserted to ensure proper seating of
the catheter.
 Shave or clip hair on the ventral neck.from the midline and extending laterally 1 cm past the
indented jugular “groove”, cranial to the angle of the jaw and caudal to the scapula. Swab the
skin with alcohol or povidone iodine.
 Using aseptic technique, a 1.5-2mm incision is made in the ventral neck to one side of the
midline. Clear the fat and connective tissue gently by blunt dissection. Avoid disturbing the vein
as it will vasoconstrict.
 A pair of mosquito hemostats or a needle driver is passed under the vein and two pieces of
suture (i.e. silk) are placed around the vein. The anterior ligature is tied tightly to completely
occlude the vein. The posterior ligature is placed loosely around the vein.
 The vein is placed under light tension by raising the posterior ligature vertically and the vein is
semi transected with ophthalmological scissors at a point between the two ligatures.
 A sterile heparinized saline filled catheter (20 units heparin / ml) is introduced into the vein and
advanced towards the heart. A rigid catheter introducer will greatly aid its passage down the vein.
The posterior ligature is now tied snugly around the catheter and vein.
 Catheter position can be verified by attaching a sterile saline filled syringe to the metal connector
of the catheter. If blood withdrawal is satisfactory then it should be flushed back into the
circulation and the catheter filled with heparinized saline (20 units heparin / ml) in order to provide
a “heparin lock”.
 A third ligature is placed and between the first and second ligatures, where the catheter exits the
jugular vein, tying the catheter to the vein.
 To exteriorize the catheter the dorsal neck is shaved and the incision site cleaned using alcohol.
 A 0.5-1.0 cm incision is made and a straight pair of hemostats is used to create a subcutaneous
tunnel extending from the dorsal neck incision along the side of the neck to the entry point of the
catheter.
 The catheter is grasped and a generous loop “stress loop” must be created at this time in the
area of the subcutaneous pocket near the catheter exit point in order to provide slack to minimize
disruption of the catheter and to accommodate an animal’s normal movements.
 The catheter is cut with 2.5-3.0 cm exteriorized and the end capped.
 The skin incisions are sutured and a sterile topical antibiotic applied.
 The neck is wrapped with a light bandage to incorporate the catheter. To avoid applying the
bandage too tightly a finger should be able to fit comfortably between the bandage and the neck.
Method 5: Femoral vein (Indwelling intravenous catheter) - Rats and guinea pigs
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 Recommended technique for repetitive, chronic blood sampling studies.
a) Catheter preparation
 Common catheter materials include polyethylene, silicone rubber and polyurethane.
 A catheter guide provides rigidity to the catheter, facilitating introduction and placement in a
blood vessel.
 The catheter and all accompanying materials must be sterilized prior to use.
b) Surgical procedure
 The animal is anaesthetized (UACC SOP # 2 Rodent Aneasthesia) and at an appropriate depth
of anaesthesia prior to commencement. Signs that indicate a satisfactory plane of anaesthesia
includes a lack of response to a toe pinch and respirations that are relaxed and regular.
 The anaesthetized rat is laid on its back with the head toward the surgeon.

Measure the length of catheter required. For example, for rats weighing 200-400 grams,
catheter lengths must be 3.5-4.5 cm in length.
 Shave or clip hair on the medial surface of the leg, extending from the groin to the mid thigh.
Swab the skin with alcohol or povidone iodine.
 Using aseptic technique, a 1.0-1.5mm midline incision is made on the inner surface of the leg,
exposing the femoral artery and vein.
 The large blue vein is covered with fine connective tissue which is gently cleared by blunt
dissection.
 A pair of mosquito hemostats or a needle driver is passed under the vein and two pieces of
suture (i.e. silk) are placed around the vein. The anterior ligature is tied loosely around the vein.
The posterior ligature is tied tightly around the vein.
 The vein is very mobile and can be steadied by the thumb. The vein is transected with
ophthalmological scissors at a point between the two ligatures.
 A sterile heparinized saline (20 units heparin / ml) filled catheter is introduced into the vein and
advanced towards the posterior vena cava.
 The posterior ligature is now tied snugly around the catheter and vein.
 Catheter position can be verified by attaching a sterile saline filled syringe to the metal connector
of the catheter. If blood withdrawal is satisfactory then it should be flushed back into the
circulation and the catheter filled with heparinized saline (20 units heparin / ml) in order to provide
a “heparin lock”.
 A third ligature is placed and between the first and second ligatures, where the catheter exits the
femoral vein, tying the catheter to the vein.
 To exteriorize the catheter the dorsal neck is shaved and the incision site swabbed with
alcohol.or povidone iodine.
 A 0.5-1.0 cm incision is made and a straight pair of hemostats is used to create a subcutaneous
tunnel extending from the back of the leg, along the back and to the nape of the neck.
 The catheter is cut with 2.5-3.0 cm exteriorized and the end capped.
 The skin incisions are sutured and a sterile topical antibiotic applied.
 The neck is wrapped with a light bandage to incorporate the catheter. To avoid applying the
bandage too tightly a finger should be able to fit comfortably between the bandage and the neck.
4.2 Volumes greater than 100l
Survival methods:
 Jugular vein - percutaneous (Section 4.1, Method 3).
 Jugular vein - indwelling intravenous catheter (Section 4.1, Method 4)
 Femoral vein- indwelling intravenous catheter (Section 4.1, Method 5)
Terminal methods:
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Method 1. Retro-orbital - Rats, mice and guinea pigs
 The animal is anaesthetized (UACC SOP # 2 Rodent Aneasthesia ) and at an appropriate depth
of anaesthesia prior to commencement. Signs that indicate a satisfactory plane of anaesthesia
includes a lack of response to a toe pinch and respirations that are relaxed and regular.
 The anaesthetized rat is laid on its back with the head toward the surgeon.
 Fix the head with your thumb and forefinger, tightening the skin over the sides of the face. This
retracts the eyelids and protracts the globe.
 Insert a microhematocrit (capillary) tube into the medial canthus and twist to break through the
bulbar conjunctiva. Direct the tube toward the medial aspect of the bony orbit.
 Collect blood into tubes and seal. Continue until unable to collect. If difficulty arises change to
other eye.
 Euthanize animal when blood collection is completed.
Method 2. Cardiac puncture - Rats, mice and guinea pigs
 The animal is anaesthetized (UACC SOP # 2 Rodent Aneasthesia) and at an appropriate depth
of anaesthesia prior to commencement. Signs that indicate a satisfactory plane of anaesthesia
include a lack of response to a toe pinch and respirations that are relaxed and regular.
 Attach a 21gauge needle to10 ml syringe (rats, guinea pigs) or a 23 gauge needle to a 3ml
syringe (mice).
 There are 2 approaches:
a) Sternal
 Place animal in dorsal recumbency.
 Clean chest area of dirt and debris.
 Palpate heart beat and xiphoid process.
 Introduce a 3 ml syringe with a 21gauge needle (rats, guinea pigs) at the angle where the xiphoid
process meets the last ribs.
 Insert needle towards the strongest palpable heartbeat, slightly left of midline.
 Apply light negative pressure and withdraw blood.
 Euthanize animal when blood collection is completed.
b) Lateral
 Place animal with its right side down
 Clean chest area of dirt and debris
 Fully flex the forelimb and note the point where the elbow meets the chest.
 Introduce a 3 ml syringe with a 21gauge needle (rats, guinea pigs) at the angle where the xiphoid
process meets the last ribs.
 Apply light negative pressure and withdraw blood.
 Euthanize animal when blood collection is completed.
Method 3. Trunk blood - Rats, mice and guinea pigs
 Optional: anaesthetize the animal (UACC SOP # 2 Rodent Anaesthesia).
 Decapitate using approved apparatus.
 Collect blood into an appropriate collection device.
UACC SOP # 1
Approved April 1999
Revised October 2005
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