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Guidelines SOPC201: Blood Collection – Multiple Species

SOP # TitleC201 Blood Collection by Different Routes in Different Species

1. PurposeThe purpose of this SOP is to describe the general method for blood collection by different acceptable routes in various species and recommended blood collection volume and frequency.

2. General Information/ResponsibilityThis SOP describes the various blood collection procedures approved by the AREB. Principal investigators may reference this SOP when blood collection is performed as indicated in the SOP. Any deviation from the SOP must be identified in the Animal Use Protocol and the principal investigator must provide justification for the deviation from standard procedure.

Total blood volume of the species determines the acceptable quantity and frequency of blood sampling. Volume of blood sampled must also consider the size and health status of the animal, type of sample needed (serum, whole blood), frequency of sampling.

Responsibility: Individuals performing blood collection must be adequately trained. Individuals collecting blood samples must be aware of the maximum allowable blood volume that can be collected at any one time or over a period of time to ensure that the animal is not physiologically compromised. This SOP provides only general guidance on blood volume collection.

Table of Contents for Blood Collection Techniques by SpeciesMouse (Submandibular vein, Sapheneous vein, Tail vein, Cardiac puncture)....................7Rat (Submandibular vein, Jugular vein, Saphenous vein, Cardiac puncture)....................11Rabbit (Central ear artery, Marginal ear vein, Carotid artery).........................................16Guinea Pig (Cardiac puncture, Vena Cava, Jugular vein)..................................................18Dog (Cephalic vein, Lateral saphenous vein, Jugular vein)...............................................21Cat (Cephalic vein, Medial saphenous vein, Jugular vein)................................................24Cattle (Tail vein, Jugular vein)...........................................................................................26Sheep (Jugular vein).........................................................................................................28Horse (Jugular vein, Facial sinus)......................................................................................29Pig (Cranial vena cava, Marginal ear vein.........................................................................31Chicken/Bird (Cardiac puncture, Jugular vein, Brachial vein)...........................................32

University Committee on Animal Care and Supply (UCACS)Standard Operating Procedure (SOP)

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3. Equipment/Materials (items utilized will depend upon the species and the collection method) Vacutainer tubes or Eppendorf

tubes and holders Capillary or Hematocrit tube (plain

or heparinized) Sterile disposable Vacutainer or

hypodermic needles (one inch)o Gauge 18, 20, 22, 23, 25, 27

Needle holders Syringes (e.g. 1 cc, 3 cc, 6 cc, 12 cc,

60 cc) Gauze or Cotton tipped applicators Vaseline Oil of Wintergreen Cordless hair trimmer; razor blade;

electric clippers Restraint tube (50 mL tube with

open bottom) or appropriate restraint device (e.g. towel, cat bag)

Hemostatic forceps with wide elastic band attached

Butterfly catheter with plastic tubing cut off

2% xylocaine jelly Skin disinfectant Alcohol Atropine

Ketamine Acepromazine Hydrogen peroxide Isoflurane and anaesthetic machine

– table top with face mask and induction chamber

Eye lubricant Circulating warm water blanket Halstead mosquito forceps, curved Kelly forceps – curved Allis tissue forceps Tissue forceps – toothed Microsurgery scissors, sharp points,

Vein introducer Fluids for replacement (Lactated

Ringer Solution, 0.9% Saline, 5% Dextrose)

Macrodrip administration set IV Pressure bag 2-0 silk ties 3 French or 4 French x 70 cm

urethral Catheter, open-end (Cook Urological)

Sharps container Gloves Paper clip White tape, shoelaces

4. Procedure

A. Type of Blood Collection Tube and Size of Needle

1. Blood Collection Tube: Check the relevant laboratory assay procedure for the type and method of sampling required (serum, whole blood, plasma). Obtain the correct type and size of Vacutainer/Eppendorf tube required for assays.

Table. Type of Vacutainer tubes.

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Sample Anticoagulant Color CodeSerum None (Silicone glass + SST) Red/GreySerum None (Silicone glass) RedPlasma Heparin GreenPlasma Na2EDTA LavenderPlasma Sodium Citrate BlueBlood Trace Elements None Royal Blue

Note: The specific type of anticoagulant may be critical for the study. A wide variety of specialized blood collection tubes are available to suit specific needs.

2. Size of needle: The length and bore size (gauge) are important considerations. Use the largest bore size possible to allow rapid blood withdrawal without collapsing the vein and without causing a hematoma.

B. Volume of Blood

1. Circulating Blood Volumea. The maximum allowable blood collection depends on knowledge of the circulating

blood volumes. On average the circulating blood volume is approximately 55-70 mL/kg body weight or 5-15% of body weight.

2. Blood Collection Volumea. Animal well-being and maintenance of a normal physiological response are the

primary considerations regarding the limits of blood sampling volume and repeated blood sampling.

Table. Blood volume limits and recovery periodsSingle sampling Multiple sampling

% Circulatory blood volume removed in a

single samplingApproximate

recovery period

% Circulatory blood volume

removed in 24 hApproximate

recovery period7.5% 1 week 7.5% 1 week10% 2 weeks 10-15% 2 weeks

15%* 4 weeks 20% 4 weeks*For single sampling it is not recommended to remove blood volume >15% due to risk of hypovolaemic shock.

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b. Single blood draw – a maximum of 1% body weight can be removed as a single blood draw without need to provide supplemental replacement fluids. Withdraw the minimum volume of blood necessary for the experimental needs. If >10% blood volume is required, replace collected volume by 3-4 –fold with isotonic fluids (saline, dextrose, lactated ringers).

Table. Recommended maximum blood volumes for different species*Species(weight)

Blood volume (mL) [mL/kg BW]

7.5% (mL) [mL/kg BW]

10% (mL) [mL/kg BW]

15% (mL) [mL/kg BW]

20% (mL) [mL/kg BW]

Mouse(25 g)

1.8[72]

0.1[5.4]

0.2[7.2]

0.3[10.8]

0.4[14.4]

Rat(250 g)

16[64]

1.2[4.8]

1.6[6.4]

2.4[9.6]

3.2[12.8]

Rabbit(4 Kg)

224[56]

17[4.2]

22[5.6]

34[8.4]

45 [11.2]

Hamster(100 g)

7.8[78]

0.6[5.8]

0.8[7.8]

1.2[11.7]

1.6[15.6]

Cat(3 Kg)

168[56]

12.8[4.2]

17[5.6]

25.5[8.4]

34[11.2]

Dog(10 Kg)

850[80]

64[6.4]

85[8.5]

127[12.8]

170 [17.0]

Guinea pig(200 g)

14.6[73]

1.2[5.5]

1.5[7.3]

2.3[11.7]

3.0[15.6]

Minipig(15 Kg)

975[65]

73[4.8]

98[6.5]

146[9.7]

195[13]

Pig(30 Kg)

1950[65]

146[4.8]

196[6.5]

292[9.7]

390[13]

Goat(50 Kg)

4650 350 467 700 933

Sheep(75 Kg)

4550[60]

350[4.5]

450[6]

600[9]

900[12]

Deer(85 Kg)

5100[60]

385[4.5]

510[6.0]

765[9.0]

1020[12]

Poultry(1 Kg)

60[60]

4.5[4.5]

6[6]

9[9]

12[12]

Cattle(250 Kg)

15000[60]

1000[4.5]

1500[6]

2000[9]

3000[12]

Bison(350 Kg)

21000[60]

1575[4.5]

2100[6.0]

3150[9.0]

4200[12]

*Blood volumes that can be removed that do not cause significant alterations in the animal’s normal physiology is as follows

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c. Multiple blood collection – 0.07% of body weight is the maximum volume of blood that can be taken daily without requiring supplemental fluids.

d. For exsanguination, expect blood volumes equaling 4-5% body weight when blood is collected while the heart is beating under a surgical plane of anaesthesia.

e. Monitor the animal during and immediately following the blood sampling for possible signs of hypovolemic shock. Institute emergency measures if noted (provision of oxygen, fluid replacement). Signs of hypovolemic shock include: a) fast and weak pulse; b) pale mucous membranes; c) cold skin and extremities; d) restlessness; e) hyperventilation.

Table. Normal haematological values in different species

SpeciesPCV(%)

RBC (1012/L)

Hb(g/dL)

Reticulocyte (%RBC)

WBC(109/L)

Mouse 35-45 7.7-12.5 10-20 3.3-13.3 8.0Rat 35-45 7.2-9.6 12-18 1.7-21.1 14.0Hamster 39-59 4.0-10.0 2-30 -- 7.6Guineapig 35-42 4.5-7.0 11-17 1.8-6.1 10Rabbit 30-50 4.5-7.0 8-15 2.9-8.0 9.0Cat 30-50 6.0-10.0 8-14 0-1.0 5.5-19.5Dog 38-53 4.5-8.0 11-18 0-1.5 6.0-17.0Pig 30-50 5.0-9.0 10-16 -- 7.0-20.0Chicken 23-55 1.25-4.5 7.0-18.6 -- 9-31Cattle 24-46 5-10 8-15 -- 4-12Sheep 29-38 8.0-14.0 10-12 ---- 4.0-12.0Goat 29-38 13.0-18.0 8-14 5.0-14.0

C. Collection of Blood Samples (General)

1. Manual restraint or sedation/general anaesthesia is required depending upon blood volume collection and species. General guidelines are as follows:

a. Rodents: manual restraint for small volumes; general anaesthesia for large volumesb. Rabbits: sedation with or without topical for other than small volumesc. Dog/Cat/Pig: manual restraint for small volumes, general anaesthesia for large

volumesd. Ruminants and horses: manual restraint for small volumes, general anaesthesia for

large volumese. Fish: general anaesthesia

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Table. Advantages/disadvantages of the various blood sampling routes and methods.

Vein General Anaesthesia

Tissue Trauma

Repeat sampling

Volume Species

Jugular No Low Yes +++ Rat, dog, rabbit, cat, hamster,

large animalCephalic No Low Yes +++ Dog, sheep,

catSaphenous/lateral tarsal

No Low Yes ++(+) Mouse, rat, dog, cat, hamster

Marginal ear No (local) Low Yes ++ Rabbit, pigSubmandibular No Low Yes ++ Mouse, ratLateral tail No Low Yes ++(+) Mouse, ratFemoral No Low Yes ++(+) RabbitCranial vena cava No Low Yes +++ PigTail tip amputation

Yes Moderate Limited + Mouse, rat

Cardiac Yes Moderate No +++ Mouse, rat, rabbit

Coccygeal No Low Yes +++ BovineBrachial wing vein No Low Yes ++ BirdCaudal vein Yes Fish

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D. Blood Sample Collection by Species and Route

MOUSE

I. Submandibular Vein

a. At the back of the jaw the veins draining the eye (orbital vein) and facial region (submandibular vein) meet to form the beginning of the jugular (Figure 1).

b. Assemble blood collection equipment. Open the protective covering containing the needle but do not remove the needle cover.

c. Remove the mouse from the cage by picking it up at the base of the tail.d. Transfer mouse from cage to stainless steel cage top.e. While holding onto the tail, use the other hand to manually restrain the mouse. Scruff

the mouse by grasping loose skin over the shoulders by starting low on the forelegs of the mouse and moving upwards behind the ears. Hold mouse in the air to establish a relaxed position for the mouse. (The proper restraint will result in the forelegs pulled up and backwards away from the head and the eyes will bulge out. NOTE: this restraint may cause some respiratory distress to the mouse – try to restrain the mouse for the shortest period of time possible.

f. Position a needle, bevel up at 90° to the skin at a location using the following landmarks:i. White mouse will have a grey dot located on the jawline directly below the lateral

canthus of the eye (Figure 1).ii. Colored mice do not have a landmark; the needle will be inserted on the jaw line in

a line drawn straight down from the lateral canthus of the eye.g. To determine the appropriate needle gauge use the following criteria:

i. 18 gauge is used to collect a rapid free flow sample into a blood collection tube when a maximum volume of sample is required.

ii. 20 gauge is used to collect a moderately flowing sample either into a collection tube or a capillary tube when 100-150 microliters of blood are required.

iii. 22 gauge is used to collect a controlled blood flow into a capillary tube when 50-100 microliters of blood are required.

h. When the needle is removed, blood will flow.i. Collect blood into the desired blood collection container (e.g. Eppendorf).

Grey dot

Submandibular vein

Orbital vein

Jaw line

Figure 1

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j. After collection, release the mouse from the restraint to stop the flow of blood.k. If the facial artery is struck instead of vein, blood will be bright in color and after sample

collection the artery will have to be held off to stop the bleeding using a gauze.l. Return the mouse to its cage.m. Discard the needle into the sharps container.

Figure. Mouse restraint, facial vein blood collection

Mouse restraint Facial Vein Blood Collection

II. Saphenous Vein Venipuncturea. Saphenous vein venipuncture is a convenient, rather noninvasive method for collection

of blood samples for purposes of clinical chemistry, hematology, biochemical assays, and pharmacokinetic studies. This procedure can be performed on both anaesthetized and unanaesthetized mice.

b. Warming the mouse immediately prior to blood collection by use of a heat lamp over the cage for five minutes will improve blood collection.

c. Place mouse head first into the restraint tube.d. Extend the back leg and fix by holding the fold of skin between the tail and the thigh of

the mouse.e. Shave hair from the hind leg over the saphenous vein using a cordless trimmer.f. Apply vaseline on the skin to form a thin layer as a barrier between the skin and the

blood. Vaseline will allow the blood to form a droplet for easy collection.g. Apply digital pressure at the groin to raise the femoral and saphenous veins. The vein

crosses the leg at the ankle and wraps around below the knee joint.h. Using a 22 gauge needle (one inch) make a puncture by quickly inserting the needle

perpendicular to the vessel on the inner surface of the hind leg. The blood should immediately flow and form a drop on the skin. Dispose the needle into the sharps container.

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i. Fill hematocrit tubes from the blood that pools on the surface of the skin. j. Alternatively, blood can be collected into an Eppendorf tube by collecting each droplet

of blood as it forms.k. Release leg and apply pressure with a clean gauze to the puncture site to achieve

hemostasis.l. Release mouse from restrainer and return the animal to the cage after checking that

bleeding has stopped.m. Serial samples can be obtained over a short period of time by gently removing the scab.

III. Tail Vein Microsampling (5-20 μL Blood Volumes)a. Tail vein microsampling is used for the collection of small (5-20 μL) blood samples as a

single sample or serial sampling from the same animal.b. Remove the mouse from the cage by picking it up at the base of the tail.c. Transfer mouse from cage to the workstation. d. Gently restrain the mouse at the base of the tail.e. Using a 25 gauge needle, prick the tip of the tail and transfer the mouse to the top of a

wire cage. f. Gently stroke the tail from the base of the tail to near the tip to encourage a drop of

blood to form at the site of the prick. g. Apply a hematocrit or capillary tube to the blood drop to collect the blood.h. Repeat the tail stroke until a sufficient volume of blood is collected from the mouse.i. Apply finger pressure on the tail at the blood sampling site for 10-20 seconds to stop

blood flow.j. Return mouse to its cage.k. It might be necessary to warm the mouse to dilate the blood vessel prior to sampling

when 20 μL blood volume is required.

IV. Lateral Tail Veina. Lateral tail vein sampling is used when larger volumes of blood (50 μL - 0.2 mL) are

required. Ideally, a maximum of two blood samples in a 24 h period is recommended to avoid damage to the tail vein.

b. Place mouse head first into the restraint tube.c. Extend the tail from the opening and wash the tail with diluted Hibitaine (1%), if

necessary.d. Apply Oil of Wintergreen to the sampling site and allow 30 minutes before blood

sampling to assure complete local anaesthesia.

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e. Using a 25 gauge needle, prick the lateral tail vein one-third the distance up the tail from the tail tip. This will allow additional samples to be taken from the tail vein by using sites closer to the base of the tail.

f. Apply a capillary tube to the blood drop and collect blood into the tube.g. Stop blood flow by applying finger pressure at the blood sampling site for 30 seconds.h. Return the mouse to the cage.i. It might be necessary to warm the mouse to dilate the blood vessel prior to sampling

when a larger blood volume is required.

V. Cardiac Puncturea. Cardiac puncture is used when large volumes of blood (0.1 mL – 1 mL) are required.

This procedure is normally done under general anaesthesia and is terminal (if not terminal consult the Animal Welfare Veterinarian).

b. Remove mouse from cage and place in induction chamber. c. Turn oxygen to 1 L/min and isoflurane to 5%.d. Once rodent has fallen asleep and is breathing regularly, remove from chamber and

place rodent in dorsal recumbency on circulating warm water blanket with nose in nose cone to maintain anesthesia.

e. Align animal so it is straight from nose to tail tip.f. Prepare 23 gauge needle and 1 mL syringe.g. With index finger, locate the notch between the Xiphoid process (at the end of the

sternum) and the last rib on the left side of rodent.h. Aiming toward the midline, insert the needle into the notch, at a 10 – 15 degree angle

above the body of the rodent.i. As the needle is inserted into the heart, pulsating blood will be seen in the hub of the

needle. If no blood is seen, pull the needle straight out the path it went in. Relocate the

landmarks and insert the needle again. (alternative needle placement below)

j. When blood appears in the hub, hold the needle and syringe still and withdraw sample.k. If the blood flow stops, use MINOR manipulations of the needle (rotating left or right,

inserting or withdrawing a millimeter or two) to try to restart the flow.l. When completed, withdraw the needle out of the heart and chest cavity in a straight

path.m. Place blood into collection tuben. Transfer mouse to euthanasia chamber.o. Turn off anesthetic machine.

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p. Ensure euthanasia has been accomplished by opening the chest cavity before properly disposing of mouse

q. Alternative needle placementa) Grasp the rodent’s right front foot and bend the elbow to form a 90° angle.b) At the point the of the 90° angle, insert the needle though the chest between the

ribs. The needle should be inserted closer to the sternum of the rodent as opposed

to its spine.c) Insert the needle until blood is visible in the hub of the needle.

If the needle is inserted and no blood appears, pull the needle straight back out the path it went in.

Recheck the landmarks for proper position and re-insert the needle to locate the heart.

d) Continue procedure as above.

RAT

I. Facial (Submandibular) Veina. Assemble blood collection equipment. Open the protective covering containing the

needle but do not remove the needle cover.b. Remove the rat from the cage by picking it up under its armpits.c. Restrain rat with a scruff hold so that it is lying on the palm of your hand to expose the

jaw line and masseter muscle.d. Use the following landmarks to determine where to insert the needle:

i. In a white rat located along the jaw line directly below the lateral canthus of the eye a grey dot is observed (use this as a landmark).

ii. Insert the needle (18 gauge) at a 30° angle 0.5 cm behind the grey dot.iii. For pigmented rat, insert the needle at a point straight down from the lateral

canthus of the eye to the jaw line.e. Invert the animal to collect the sample directly into the collection tube.f. After the sample is collected, apply gauze and pressure at the site of the venipuncture to

stop bleeding. g. Release restraint and return rat to the cage.h. Dispose of needle into sharps container.

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II. Jugular Vein (anesthetized)a. Put a paper towel at the bottom of the anaesthetic induction chamber and place rat in

the chamber.b. Turn the oxygen flow to 1-2 L/ min and the isoflurane to 5% on the anaesthetic machine.c. Once the rat is relaxed and has lost its righting reflex dial down isoflurane to 2-3%.d. Remove rat from chamber and put on the face mask (remember to open the valve to

the face mask and close the valve to the induction chamber) and place rat on an incontinent pad.

e. Put eye lube in each eye and then place the rat in dorsal recumbency.f. Using the pedal reflex ensure that the rat is maintained at a light surgical plane (usually

1.5-2 % isoflurane is sufficient).g. Place a 3 cc syringe underneath the neck of the rat cranial to the shoulder blades.h. Secure the rat to the pad with masking tape (extend each limb and secure to pad). It is

important that the rat is positioned with its vertebra as straight as possible and the front limbs at 90o angles to the spine.

i. Hold the skin taut in the neck region and shave the hair from midline and laterally to the right side of the rat using the small clippers (only shave a 1 inch by 1 inch area). Remove hair from the area.

j. Sterilize the area with an alcohol swab. k. Secure a needle to a syringe and unlock the plunger of the syringe. If a plasma sample is

required, first draw up the anticoagulant to allow for coating of the syringe barrel and discharge to the syringe contents to leave a small volume of anticoagulant in the hub of your needle.

l. Find the middle point between the sternum and the top of the shoulder at a point just right to the midline.

m. Advance a 23 gauge one inch needle (bevel side up) through the skin at a 30o angle in a direction towards the heart. Create a vacuum within the syringe barrel and then advance the needle slowly a few millimeters at a time.

n. Once the needle is in the trunk of the vein, a very small drop of blood will appear in the needle hub. This will indicate that the needle is properly seated into the vein.

o. Once seated, steady the syringe and slowly milk the plunger to fill the barrel. Avoid continuous pressure as this will collapse the vein.

p. Once the sample is collected, apply digital pressure with gauze to the site before withdrawing the needle.

q. Remove the needle from the syringe and dispense blood into the collection tube. Dispose needle into sharps container.

r. Remove the masking tape and syringe from under the animal’s neck.s. Turn off the isoflurane and deliver 100% oxygen for a few minutes.

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t. Place the animal in a cage on a paper towel until fully conscious. If recovery is slow, the cage can be placed on a heated circulating water blanket until complete recovery.

u. This can be conducted in an awake rat according to the following steps:i. This technique requires two people – a restrainer and a phlebotomist.ii. Remove rat from the cage and calm the rat by gently stroking it.

iii. Restrain the rat by grasping the skin over the nape of the neck and shoulders with thumb and index fingers.

iv. Turn the rat over such that it is lying in your palm and the body is straight. With the other hand restrain the head by placing the muzzle between the second and third fingers and the thumb at the thoracic inlet to hold back the forelegs. Pull back on the head until it is even with the spine.

v. The other person shaves and prepares the neck region as above.vi. Apply slight pressure in the thoracic region to raise the jugular vein.vii. With a 23 guage one inch needle insert the needle (bevel side up) through the

skin at a 30o angle at a midpoint between the sternum and shoulder in a direction towards the heart. Create a vacuum within the syringe barrel and then advance the needle slowly a few millimeters at a time.

viii. Once the needle is in the trunk of the vein, a very small drop of blood will appear in the needle hub. This will indicate that the needle is properly seated into the vein.

ix. Once seated, steady the syringe and slowly milk the plunger to fill the barrel. Avoid continuous pressure as this will collapse the vein.

x. Once the sample is collected, apply digital pressure with gauze to the site before withdrawing the needle.

xi. Remove the needle from the syringe and dispense blood into the collection tube.xii. Release animal from restrainer; give positive reinforcement.

xiii. Dispose of needle into sharps container.

III. Saphenous Veina. Use this route for collection of single or serial blood samples without anaesthesia.b. Assemble all equipment.c. Remove rat from cage and wrap the head, front legs and body in a towel leaving the

rear legs and tail free.d. Using the same hand that is holding the restrained rat, gently but firmly squeeze the leg

by placing the thumb on the upper thigh/hip area and the lower leg between the second and third fingers.

e. Shave hair using clippers over the lateral saphenous vein area and swab shaved area with 70% alcohol.

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f. The pressure applied by the thumb will result in saphenous vein engorgement. Locate the vein and apply a thin film of Vaseline® to allow for drop formation.

g. Using a 25 gauge needle puncture the vessel at a 90° angle at the most proximal (closest to the body) visible site.

h. Collect the blood drops into the appropriate collection tube.i. Apply dry gauze and pressure to the puncture site and release pressure on the upper

thigh until bleeding ceases.j. Remove the rat from the towel and return to its cage.k. Monitor the rat for an addition 5-10 minutes to ensure that bleeding has stopped.l. Repeated blood samples may simply require removal of the scab with a dry piece of

gauze or a new puncture site can be made distal to the previous site (towards the foot).

IV. Tail Veina. Assemble all equipment.b. Remove rat from cage 20 – 30 minutes before procedure – apply 2% Xylocaine Jelly to

rodent tail & return to cage.c. Restrain animal in a decapicone (see figure below) using forceps to tighten elastic band

around cone and tail.d. Clean tail with gauze, disinfectant, and warm water; apply alcohol and let dry.e. With bevel up, insert butterfly needle (21-23 gauge depending on size of rat) into the

lateral tail vein – if no blood flow, redirect needle.f. Position tail over blood collection tube to catch drops.g. After sample collection, remove needle and apply pressure with wet gauze.h. Release animal from restrainer; give positive reinforcement.i. Dispose of needle into sharps container.

Figure. Restraint, blood collection, and positive reinforcement with hemostasis

Rat restraint in decapicone and tail preparation

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Butterfly catheter inserted and collection Praising rat and holding off vein followinginto blood tube. procedure.

V. Cardiac Puncture (Terminal)a. Cardiac puncture is only used for terminal blood sampling because of the risk of

damage to the myocardium.b. Put a paper towel at the bottom of the anaesthetic induction chamber and then place

the rat in the chamber.c. Turn the oxygen flow to 1-2 L/ min and the isoflurane to 5% on the anaesthetic machine.d. Once the rat is relaxed and has lost its righting reflex, dial down the isoflurane to 3%.e. Remove the rat from the chamber, put on the face mask (remember to open the valve

to the face mask and close the valve to the induction chamber) and place rat on a incontinence pad.

f. Place the rat in dorsal recumbency. Align the rat such that it is straight from nose to tail tip.

g. Using the pedal reflex, ensure that the rat is maintained at a surgical plane of anaesthesia (usually 1.5-2% isoflurane).

h. Secure an 18 or 20 gauge needle to the syringe (10 or 20 cc) and unlock the plunger of the syringe.

If a plasma sample is required, first draw up the anticoagulant to allow for coating of the syringe barrel and discharge to the syringe contents to leave a small volume of anticoagulant in the hub of your needle.

i. Locate the notch between the xiphoid process (at the end of the sternum) and the last rib on the left side of the rat. Just below or to the side of the xyphoid cartilage advance the needle (bevel side up) through the skin at a 15-30o angle and toward the midline.

Create a vacuum within the syringe barrel and then advance the needle slowly a few millimeters at a time.

j. Once the needle is in the heart, a very small drop of blood will appear in the needle hub. If no blood is seen, pull the needle out a short distance in the same path as it went in, redirect the needle and advance it slowly again.

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k. Once seated, steady the syringe and slowly withdraw the plunger to fill the syringe barrel. Avoid continuous pressure as this will collapse the chamber of the heart.

l. When completed, withdraw the needle from the heart and chest cavity in a straight path.

m. Remove the needle from the syringe and dispense blood into the collection tube.n. Further exsanguination, cervical dislocation, decapitation, opening the chest cavity, or

removal of a vital organ should be performed to ensure death.

RABBIT

I. Central Ear Arterya. Place rabbit in a cat bag or wrap in a towel.b. Grasp ear and remove hair over the artery with a straight razor blade.c. To dilate the artery apply Oil of Wintergreen with the cotton-tipped applicator to the

skin over the artery.d. Remove caps from blood collection tubes.e. Insert a 22 gauge catheter into the artery.f. Have the rabbit restrainer lightly pinch the artery as the catheter stylette is removed.g. Place the blood collection tube under the catheter end.h. Restrainer releases pinch over the artery to allow blood to drip into the blood collection

tube.i. When filling multiple blood collection tubes, have restrainer pinch artery every time the

blood collection tube is changed.j. If blood flow stops or slows before all blood is collected, gently rub artery with finger or

with the wintergreen cotton-tipped applicator.k. Remove catheter and gently apply pressure with gauze until the blood flow stops from

the puncture site.l. Wash ear with wet gauze and then apply Hibitane ointment.m. Return rabbit to cage or pen.

II. Marginal Ear Veina. Place rabbit in a cat bag or wrap snuggly in a towel.b. Grasp ear and remove hair over the vein with a straight razor blade. c. Clean the shaved area with an alcohol swab.d. Anaesthetize the venipuncture site by application of a lidocaine containing cream (i.e.

EMLA) (this is recommended and if performed the skin requires 45 minutes for the full skin numbing effect).

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e. Remove caps from blood collection tubes.f. Massage the ear to dilate the vein.g. Insert a 23 gauge needle or catheter into the vein.h. Have the rabbit restrainer lightly pinch the artery as the catheter stylette is removed.i. Place the blood collection tube under the catheter end.j. Remove catheter and gently apply pressure with gauze until the blood flow stops from

the puncture site.k. Wash ear with wet gauze and then apply Hibitane ointment.l. Return rabbit to cage or pen.

III. Carotid Arterya. Wrap the rabbit snuggly in a towel or cat bag, and give an I.M. injection of Ketamine

25mg/kg and acepromazine 1 mg/kg.b. Once the sedation has taken effect, place the rabbit on the table and place the face

mask over the rabbit’s nose.c. Start delivering the isoflurane in small increments every minute from 1- 5% (to prevent

breath holding).d. Once the rabbit is relaxed, dial down the isoflurane to 2-3%.e. Tie mask, using gauze, around the head behind the ears.f. Place the rabbit in lateral recumbency and clip the ear over the marginal ear vein, and

then clean over the vein with chlorhexidine and alcohol.g. Place a paper clip over the proximal end of the vein and insert the catheter in the raised

vein.h. Remove the paper clip, remove the stylet of the catheter and attached the pre-flushed

extension set to the catheter.i. Give a small bolus of heparinized saline to clear the catheter.j. Using a 8-12” piece of 1/4’ white tape, make a tape butterfly over the hub of the

catheter; place a gauze roll inside the ear; wrap the long end of tape around the ear to secure the catheter in place.

k. Attach the I.V. fluids of 0.9% NaCl to the extension set and set fluid drip rate to the maintenance rate of 3 mL/kg/hr.

l. Turn the rabbit into dorsal recumbency and clip the hair on the neck from the start of the mandible to the thoracic inlet using the electric clippers with #40 blade.

m. Using the knife handle and scalpel blade, make an incision just lateral to the trachea and about 1 1/2”-2” long.

n. Using blunt dissection with a kelly forcep, separate the connective tissue between the sternohyoideus and sternocepalicus muscles.

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o. Once the carotid artery is exposed, use the mosquito forcep to gently separate the carotid artery from the vagus nerve.

p. Loop two silk ligature (2-0) lengths (3-4 inches) under the carotid artery.q. Tie the silk ligature that is most cranial so that it occludes blood flow.r. Place one throw on the ligature proximally to the heart, so that it is ready to tie down.s. Have an assistant pull up on both ligatures, so that you have good exposure of the

carotid artery.t. Make a small cut in the artery about 0.5 cm from the cranial silk tie using vascular

scissors.u. Place the tip of the vein introducer through the hole and into the artery.v. Slide the catheter under the introducer and into the artery, past the ligature.w. Insert the catheter approximately 8 cm into the carotid and then tie the proximal

ligature snuggly over the catheter.x. Tie the distal ligature over the catheter to anchor it in place.y. Pull the catheter so that it is taut (but not sliding out of the artery), and tape it to the

face mask with masking tape or white tape.z. Inflate the pressure bag over the I.V. fluids to 150 mm Hg, and open the line fully so that

it is flowing in.aa. Start removing the blood from the catheter with a 60 cc syringe.bb. Put collected blood into 50 ml centrifuge tubes. Label vials 1, 2, 3….. to identify the

order of collection.*This technique allows for the collection of a much greater percentage of the blood volume of a rabbit compared to that using cardiac puncture.*

GUINEA PIG

I. Cardiac Puncturea. NB: This technique is only used when attempts at vena cava collection are

unsuccessful.b. Day before blood sampling – fast the guinea pig overnight.c. Weigh the guinea pig and record the weight in the file.d. Administer 0.1 mg/kg atropine subcutaneously.e. Fifteen minutes after the atropine administration place the guinea pig into the paper

towel lined induction chamber.f. Turn the oxygen flow to 1-2 L/ min and the isoflurane to 5% on the anaesthetic machine.g. Once the guinea pig is relaxed and has lost its righting reflex, dial down the isoflurane to

2-3%.

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h. Remove the guinea pig from the chamber onto the incontinence pad and put on the face mask (remember to open the valve to the face mask and close the valve to the induction chamber).

i. Put eye lubricant in each eye and then place the guinea pig in dorsal recumbency.j. Locate the xyphoid process of the sternum. a. Prepare the area with an alcohol swab. k. Insert a needle (23 gauge with 6cc syringe on an extension with a T) just lateral to the

xyphoid process at a 30 degree angle directed midline below the sternum. i. Maintain a slight amount of negative pressure on the syringe while slowly

withdrawing the needle until blood readily flows. ii. Collect the desired sample volume (5-6 mL).

l. Pour the blood into the appropriate collection tube.m. Put gentle pressure over the collection site for 30-60 seconds to stop the bleeding;

clean the site with hydrogen peroxide if necessary. n. Place the animal in sternal recumbency.o. Turn the anesthetic gas off and deliver 100% oxygen.p. Give the guinea pig 10 mL of 0.9% NaCl over the dorsal back between the shoulder

blades as a subcutaneous injection.q. Maintain guinea pig on 100% oxygen for 2-3 minutes.r. Monitor the guinea pig until mobile and then return to the cage.s. Monitor the animal every two hours until the end of the work day.t. Animals that have prolonged recovery from anesthesia (>20 minutes), laboured or

raspy respiration, cyanosis or pale mucous membranes, and decreased heart rate (less than 150 bpm) will be euthanized with intracardiac Euthanyl under anesthesia.

II. Vena Cavaa. Fast the guinea pig to be bled for 24 hours prior to the procedure.b. Weigh the guinea pig and record in the file.c. Administer 0.1 mg/kg atropine subcutaneously.d. Fifteen minutes after the atropine injection place the guinea pig into the paper towel

lined induction chamber.e. Turn the oxygen flow to 1-2 L/ min and the isoflurane to 5% on the anaesthetic machine.f. Once the guinea pig is relaxed and has lost its righting reflex, dial down the isoflurane to

2-3%.g. Remove the guinea pig from the chamber and put on the face mask (remember to open

the valve to the face mask and close the valve to the induction chamber).h. Put eye lubricant in each eye and then place the guinea pig in dorsal recumbency.i. Locate the cranial portion of the sternum, the manubrium.

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i. Prepare the area with an alcohol swab. j. Insert a needle (23 gauge with 6cc syringe on an extension with a T) just lateral to the

manubrium at a 30 degree angle directed at the opposite thigh – mid-lateral abdominal region.

i. The needle should not be inserted more than ½ inch. ii. Maintain a slight amount of negative pressure on the syringe while slowly

withdrawing the needle until blood readily flows. iii. Collect the desired sample volume (5-6 mL).

k. Pour the blood into the appropriate collection tube.l. Put gentle pressure over the collection site for 30-60 seconds to stop the bleeding;

clean the site with hydrogen peroxide if necessary. m. Place the animal in sternal recumbency.n. Turn the anesthetic gas off and deliver 100% oxygen for 5 minutes.o. Give the guinea pig 10 mL of 0.9% NaCl over the dorsal back between the shoulder

blades as a subcutaneous injection.p. Monitor the guinea pig until mobile and then return to the cageq. Animals that have prolonged recovery from anesthesia (>20 minutes), laboured or

raspy respiration, cyanosis or pale mucous membranes, and decreased heart rate (less than 150 bpm) will be euthanized with intracardiac Euthanyl under anesthesia.

III. Jugular Veina. Fast guinea pig overnight (take feed away from the whole tub).b. Weigh the guinea pig and record the weight in the log.c. Give 0.1 mg/kg Atropine by subcutaneous injection 15 minutes before inducing

anesthesia.d. Put a paper towel at the bottom of the induction chamber, and then place the guinea

pig in the induction chamber.e. Turn the oxygen flow to 1-2 L/ min and the isoflurane to 5% on the anaesthetic machine.f. Once the guinea pig is relaxed and has lost its righting reflex, remove the guinea pig

from the chamber and put on the face mask (remember to open the valve to the face mask and close the valve to the induction chamber).

g. Dial down the isoflurane to 2-3%.h. Put eye lubricant in each eye. i. Give the guinea pig 10 mL of 0.9% NaCl over the dorsal back between the shoulder

blades.j. Place in dorsal recumbency on an incontinence pad resting on a stainless steel rodent

cage lid.k. Position the head into the food trough so that it is at a 30% angle to the body.

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l. With the body straight, tie the front limbs at 90o angles to the spine using shoelaces.m. Hold the skin taut in the neck region and shave the hair using small clippers. Remove

hair from the area.n. Prepare the area with an alcohol swab. o. Secure a needle (22 gauge) to the syringe and unlock the plunger of the syringe. Draw

up ~0.5 – 1 cc of anticoagulant to allow for coating of the syringe barrel and leaving a small amount in the syringe.

p. The landmark for needle insertion is a depression within the jugular furrow cranial to the clavicle and slightly off the midline.

q. Advance the needle (bevel side up) through the skin. Create a vacuum within the syringe barrel and then advance the needle slowly a few millimeters at a time.

r. Once the needle is in the trunk of the vein, a very small drop of blood will appear in the needle hub. This will indicate that the needle is properly seated into the vein.

s. Once seated, steady the syringe and slowly withdraw the plunger to fill the barrel. Avoid continuous pressure as this will collapse the vein.

t. Once the sample is collected, apply digital pressure with a gauze to the site before withdrawing the needle.

u. Remove the needle from the syringe and dispense blood into the collection tube and mix gently.

v. After putting gentle pressure over the collection site for 30-60 seconds, clean the site with hydrogen peroxide if necessary.

w. Remove the leg restraints, massage the feet to promote circulation, and then place the animal in sternal recumbency.

x. Turn the anesthetic gas off and deliver 100% oxygen.y. Give 100% oxygen for another 2-3 minutes.z. Monitor the guinea pig until in sternal recumbency and then return to the cage.

DOG

I. Cephalic Veina. Two people are needed for this procedure – one for restraint and raising the vein and

the other for taking the blood sample.b. Assemble all of the equipment. Attach appropriate sized syringe to a 21 gauge needle.

Manually restrain the dog in sternal recumbency or in a standing position. Extend the foreleg. For the right cephalic vein the restrainer is positioned on the dog’s left side. The dog is restrained close to the body of the restrainer and the restrainer’s left hand or arm is placed under the muzzle to pull the head toward the restrainer’s body. The restrainer

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extends the foreleg with the right hand positioned under the dog’s elbow while occluding the vein with the thumb positioned near the elbow of the dog.

c. If needed, clip hair around the sampling site with small clippers. Sterilize the area (skin or hair) by using a swab soaked in 70% alcohol.

d. Raise the cephalic vein with one thumb by holding pressure across the proximal end of the foreleg. This prevents blood flow and the cephalic vein will enlarge and become turgid.

e. Remove cap from needle. Grab the leg and place a thumb adjacent to the vein to pull the skin and stabilize the vein. Position the needle with bevel facing upwards parallel to the skin overlying the cephalic vein.

f. Insert the needle through the skin and into the vein at a 25° angle up to 0.5-0.75 cm and when blood appears in the syringe hub retract the syringe plunger to let blood fill the syringe. Do not apply too much suction as this may collapse the vein and disrupt blood flow into the syringe.

g. If blood is not flowing into the syringe, the needle bevel may be lodged against the vessel wall. Rotate the needle slightly and reposition. Retract the syringe plunger again to let blood collect into the syringe.

h. When blood collection complete, release pressure on the vein and remove the needle. i. Apply pressure with fingers immediately following removal of the needle until the blood

flow stops from the puncture site (about 30 seconds).j. Detach needle from syringe and add blood to the blood collection tube.k. Provide dog with a reward (e.g. food treat) and return dog to kennel.l. Subsequent venipuncture attempts can occur as necessary more proximal to the original

puncture site.

II. Lateral Sapheneous Veina. Two people are needed for this procedure – one for restraint and raising the vein and

the other for taking the blood sample.b. Assemble all of the equipment. Attach appropriate sized syringe to a 21 gauge needle.

Manually restrain the dog in lateral recumbency. For the right lateral saphenous vein the restrainer holds off the vein with the right hand positioned behind the knee.


d. Raise the vein with one thumb by holding pressure across the proximal end of the saphenous vein. This prevents blood flow and the cephalic vein will enlarge and become turgid.

e. Remove cap from needle. Grab the leg and place a thumb adjacent to the vein to pull the skin and stabilize the vein. Position the needle with bevel facing upwards parallel to the skin overlying the cephalic vein.

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flow stops from the puncture site (about 30 seconds).j. Detach needle from syringe and add blood to the blood collection tube.k. Provide dog with a reward (e.g. food treat) and return dog to kennel.l. Subsequent venipuncture attempts can occur as necessary more proximal to the original

puncture site.

III. Jugular Veina. Two people are needed for this procedure – one for restraint and raising the vein and

the other for taking the blood sample.b. Assemble all of the equipment. Attach appropriate sized syringe to a 21 gauge one inch

needle. Manually restrain the dog.c. If needed, clip hair around the sampling site with small clippers. Sterilize the area (skin

or hair) by using a swab soaked in 70% alcohol.d. Raise the jugular vein with one thumb by holding pressure across the jugular groove

near the thoracic inlet, below the venipuncture site. This prevents blood flow and the jugular vein will enlarge and become turgid.

e. Remove cap from needle and position the needle with bevel facing upwards parallel to the skin overlying the jugular vein.

f. Insert the needle through the skin and into the vein at a 25° angle and retract the syringe plunger to let blood fill the syringe. Do not apply too much suction as this may collapse the vein and disrupt blood flow into the syringe.



flow stops from the puncture site (about 30 seconds).j. Provide dog with a reward (e.g. food treat) and return dog to kennel.

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CAT

I. Cephalic Veina. Two people are needed for this procedure – one for restraint and raising the vein and

the other for taking the blood sample. b. Assemble all of the equipment. Attach appropriate sized syringe to a 21 gauge needle.

Manually restrain the cat in sternal recumbency. For the right cephalic vein the restrainer is position on the cat’s left side. The restrainer’s left hand or arm is placed under the jaw to pull the head toward the restrainer’s body and the restrainer extends the foreleg with the right hand under the elbow while occluding the vein with the right thumb positioned near the elbow of the cat.


d. Raise the cephalic vein with one thumb by holding pressure across the proximal end of the foreleg. This prevents blood flow and the cephalic vein will enlarge and become turgid.

e. Remove cap from needle and grab the leg and place a thumb lateral to the vein to pull the skin and stabilize the vein. Position the needle with bevel facing upwards parallel to the skin overlying the cephalic vein.




flow stops from the puncture site (about 30 seconds).j. Detach needle from syringe and add blood to the blood collection tube.k. Return the cat to the cage.

II. Medial Saphenous or Femoral Veina. Use this route to obtain small volumes of blood.b. Two people are needed for this procedure – one for restraint and raising the vein and

the other for taking the blood sample.

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c. Assemble all of the equipment. Attach appropriate sized syringe to a 22 gauge one inch needle. Manually restrain the cat in right lateral recumbency with the left rear leg abducted.

d. If needed, clip hair around the sampling site with small clippers. Sterilize the area (skin or hair) by using a swab soaked in 70% alcohol.

e. Grasp the trarsus and extend the right leg. Occlude the vein with pressure applied by the restrainer’s left hand in the right inguinal region. This prevents blood flow and the vein will enlarge and become turgid. To visualize the vein it may be necessary to wipe the area with alcohol and separate the hair over the vessel.

f. Remove cap from needle and position the needle with bevel facing upwards parallel to the skin overlying the jugular vein.

g. Insert the needle through the skin and into the vein at a 25° angle and retract the syringe plunger to let blood fill the syringe. Do not apply too much suction as this may collapse the vein and disrupt blood flow into the syringe.

h. If blood is not flowing into the syringe, the needle bevel may be lodged against the vessel wall. Rotate the needle slightly and reposition. Retract the syringe plunger again to let blood collect into the syringe.

i. When blood collection complete, release pressure on the vein and remove the needle. j. Apply pressure with fingers immediately following removal of the needle until the blood

flow stops from the puncture site (at least 60 seconds).

III. Jugular Veina. Two people are needed for this procedure – one for restraint and raising the vein and

the other for taking the blood sample. A cat bag can also be used for restraint.b. Assemble all of the equipment. Attach appropriate sized syringe to a 21 gauge one inch

needle. Manually restrain the dog.c. If needed, clip hair around the sampling site with small clippers. Sterilize the area (skin

or hair) by using a swab soaked in 70% alcohol.d. Raise the jugular vein with one thumb by holding pressure across the jugular groove

near the thoracic inlet, below the venipuncture site. This prevents blood flow and the jugular vein will enlarge and become turgid.

e. Remove cap from needle and position the needle with bevel facing upwards parallel to the skin overlying the jugular vein.

f. Insert the needle through the skin and into the vein at a 25° angle and retract the syringe plunger to let blood fill the syringe. Do not apply too much suction as this may collapse the vein and disrupt blood flow into the syringe.


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flow stops from the puncture site (about 30 seconds).

CATTLE AND BISON

I. Tail Veina. Use appropriate restraint to ensure the procedure will cause minimal distress and can

be conducted quickly and safely. Stand where you are protected from the hind limbs of the bison. Do not stand directly behind a bison or you will be kicked.

b. Wear gloves and disinfect gloves between animals. Use a new needle for each venipuncture. The vacutainer holder can be reused but should be cleaned between animals.

c. Assemble all of the equipment. Screw the vacutainer needle onto the vacutainer.d. Insert the collection tube into the vacutainer, using care not to puncture the stopper.

Hold the vacutainer and inserted collection tube in one hand.e. Restrain the animal.f. Remove the needle cap and store safely.g. Raise the tail vertically with one hand until it is horizontal with the ground.h. Locate the groove lying in the ventral midline of the tail approximately 15 cm from the

base of the tail.i. Clean the skin using an antiseptic cleanser prior to the venipuncture.j. Midway along the body of a tail vertebra insert a 20 gauge needle attached to a syringe

or vacutainer holder perpendicularly to the surface of the skin to a depth of a few millimeters. Once the needle is through the skin, push the collection tube onto the needle within the vacutainer. Remember that vacuum will be lost if the needle is removed from the skin, therefore use care if re-directing the needle. If vacuum is lost, a new collection tube must be used.

k. Advance the needle slowly until blood starts flowing into the collection tube. Hold the needle in this position until the tube is filled. If more than one tube is required, remove the tube from the collection needle and insert a new tube without pulling the needle from the skin.

l. Withdraw needle and apply pressure to the venipuncture site to stop bleeding.m. Discard the needle into a sharps container, using care to avid needle-stick injury.n. Label and process tubes as appropriate.

II. Jugular Vein

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a. Move animal into squeeze chute, and restrain properly. For bison, in a chute do not raise the ‘crash gate’ until you are certain that the head has be caught by the head gate.

b. When bleeding in front of the head gate, apply a halter and tie the head to one side exposing the lower neck region.

c. Identify the jugular vein by raising the vein and inspection and/or palpation. Clip the hair on the animal’s neck at the area (2 inch x 2 inch) for needle insertion.

d. Clean the clipped area using 70% alcohol and swab.e. Screw the vacutainer needle onto the vacutainer.f. Insert the collection tube into the vacutainer, using care not to puncture the stopper.

Hold the vacutainer and inserted collection tube in one hand.g. Remove the needle cap and store safely.h. Raise the jugular vein with one thumb by holding pressure across the base of the neck

within the jugular groove. This prevents blood flow and the jugular vein will enlarge and become turgid.

i. Insert the needle (20 gauge) into the jugular vein in the upper third of the neck and at approximately a 45 angle. It is helpful to rest your wrist against the animal’s neck.

j. Once the needle is through the skin, push the collection tube onto the needle within the vacutainer. Remember that vacuum will be lost if the needle is removed from the skin, therefore use care if re-directing the needle. If vacuum is lost, a new collection tube must be used.

k. Advance the needle slowly until blood starts flowing into the collection tube. Hold the needle in this position until the tube is filled. If more than one tube is required, remove the tube from the collection needle and insert a new tube without pulling the needle from the skin.

l. When collection is completed, remove the tube from the collection needle and then remove the needle from the skin. Lightly press a new swab with 70% alcohol on the site.

m. Discard the needle into a sharps container, using care to avoid needle-stick injury.n. Remove halter and release animal from chute.o. Label and process tubes as appropriate.

SHEEP AND DEER

I. Jugular Veina. Use appropriate restraint to ensure the procedure will cause minimal distress and can

be conducted quickly and safely.

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b. Wear gloves and disinfect gloves between animals. Use a new needle for each venipuncture. The vacutainer holder can be reused but should be cleaned between animals.

c. Assemble all the equipment.d. Restrain the sheep by straddling the animal, placing knees behind the shoulders of the

animal and backing the animal into a wall to control the hindquarters. Use electric shears to shave a patch over the jugular groove large enough to visual the vein (e.g. 3 inches by 5 inches).

e. White-tailed deer must be either chemically immobilized (asleep) or secured in a suitable deer chute. White-tailed deer cannot be safely held otherwise.

f. Have an assistant turn the head at a 30° angle to the side by holding the animal under the jaw to allow for easy access to the vein.

g. Identify the jugular vein by raising the vein and inspection and/or palpation. The vein lies along an imaginary line from the middle of the animal’s eye down the side of the neck. Apply alcohol to clean the skin and facilitate visualization of the vein.

h. Screw the vacutainer needle (20 gauge by one inch) onto the vacutainer holder.i. Insert the collection tube into the vacutainer holder, using care not to puncture the

stopper. Hold the vacutainer and inserted collection tube in one hand. j. Remove the needle cap and store safely. Do not remove the needle cap with your teeth.k. Raise the jugular vein with one hand by applying pressure in the jugular furrow near the

thoracic inlet while using the other hand to stroke the jugular vein until it becomes large and distended.

l. Insert the needle with a quick thrust into the jugular vein in the upper third of the neck and at approximately a 45° angle. It is helpful to rest your wrist against the horse’s neck.

m. Once the needle is through the skin, push the collection tube onto the needle within the vacutainer. Remember that vacuum will be lost if the needle is removed from the skin, therefore use care if re-directing the needle. If vacuum is lost, a new collection tube must be used.

n. Hold the needle in this position until the tube is filled. If more than one tube is required, remove the tube from the collection needle and insert a new tube without pulling the needle from the skin.

o. When collection is completed, remove the tube from the collection needle and then remove the needle from the skin. Press gently on the site of needle insertion until bleeding has ceased.

p. Discard the needle into a sharps container, using care to avoid needle stick injury. Do not recap the needle.

q. Discard disposable materials in the trash.r. Wash hands, clean area.

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HORSE

I. Jugular Veina. Use appropriate restraint to ensure the procedure will cause minimal distress and can

be conducted quickly and safely.b. Wear gloves and disinfect gloves between animals. Use a new needle for each

venipuncture. The vacutainer holder can be reused but should be cleaned between animals.

c. Assemble all the equipment.d. Restrain the horse, by having an assistant hold the horse by its halter. Do not tie the

horse. Do not perform this procedure without an assistant holding the horse. Do not proceed with the venipuncture unless the horse is adequately restrained.

e. Identify the jugular vein by raising the vein and inspection and/or palpation. Apply alcohol to clean the skin and facilitate visualization of the vein.

f. Screw the vacutainer needle (20 gauge) onto the vacutainer.g. Insert the collection tube into the vacutainer, using care not to puncture the stopper.

Hold the vacutainer and inserted collection tube in one hand. h. Remove the needle cap and store safely. Do not remove the needle cap with your teeth.i. Raise the jugular vein with one hand by applying pressure in the jugular furrow near the

thoracic inlet while using the other hand to stroke the jugular vein until it becomes large and distended.

j. Insert the needle with a quick thrust into the jugular vein in the upper third of the neck and at approximately a 45° angle. It is helpful to rest your wrist against the horse’s neck.

k. Once the needle is through the skin, push the collection tube onto the needle within the vacutainer. Remember that vacuum will be lost if the needle is removed from the skin, therefore use care if re-directing the needle. If vacuum is lost, a new collection tube must be used.

l. Hold the needle in this position until the tube is filled. If more than one tube is required, remove the tube from the collection needle and insert a new tube without pulling the needle from the skin.

m. When collection is completed, remove the tube from the collection needle and then remove the needle from the skin.

n. Discard the needle into a sharps container, using care to avoid needle stick injury. Do not recap the needle.

o. Discard disposable materials in the trash.p. Wash hands, clean area.

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II. Facial Sinusa. This procedure is used if small volumes of blood are required, e.g. for a PCV/TP

determination. Typically, only a few drops of blood will be obtained. b. This is a “blind stick”, i.e. you cannot visualize or palpate the venous sinus. Review

anatomy before performing this procedure.c. Assemble all the equipment. d. Restrain the horse, typically by having an assistant hold the horse by its halter. Do not

tie the horse. Do not perform this procedure without an assistant holding the horse. Do not proceed with venepuncture unless the horse is adequately restrained.

e. Locate the appropriate site, approximately 0.5-1.cm below the facial crest at approximately 1/3 of the way between the medial and lateral canthus of the eye.

f. Clean the skin using a moist (not wet) alcohol swab. Take care to avoid any alcohol entering the horse’s eye. Do not apply alcohol directly from the bottle.

g. Remove the needle cap from a 20 gauge, 1.5 inch needle (or 22 gauge, 1.5 inch) and store safely. Do not remove the needle cap with your teeth.

h. Rest the hand holding the needle firmly against the horse’s head. Failure to do so can lead to eye injury of the horse if the horse moves during the procedure.

i. Use your free hand to help restrain the horse by its halter. j. Rapidly insert the needle through the skin at the appropriate site, using a 90° angle to

the skin (perpendicular orientation). Most horses will flinch at this, therefore let go of the needle as soon as it is inserted. Failure to do so can make you pull out the needle inadvertently and you have to repeat the procedure.

k. Slowly advance the needle until blood is obtained. It is helpful to advance the needle in short, rapid thrusts and to rotate the needle during advancement. Let go of the needle each time after advancing it.

l. Once blood is obtained, collect blood into hematocrit tubes or blood collection tubes, or just remove the needle.

m. Remove the needle and discard in a sharps container using care to avoid needle stick injury. Do not recap the needle.

n. Discard any disposable materials in the trash. Wash hands, clean area.

PIG

I. Cranial vena cava/External jugular vein

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a. Move pig into blood collection area. Restrain the pig. For small pigs less than 25 kg one person can restrain the pig by turning the pig onto its back, holding the front legs and restraining the body with the person’s arms or by straddling the pig and using their legs. Larger animals require restraint using a snout snare and will remain standing. Allow the pig to bite the snare and then close it quickly and apply pressure. The pig will remain quiet when pressure is maintained. Ensure that the neck is stretched upwards by making sure the pig leans back against the pull of the rope.

b. A second person will take the blood sample from the jugular vein by using one hand to restrain the head of the pig by holding the snout and the other hand will take the blood sample.

c. Insert an appropriate sized needle attached to a vacuum test tube holder and tube or a syringe into the ventral side of the neck, on the right side just off the midline in the depression under the neck between the legs. The needle puncture site should be at the deepest point of the jugular groove formed between the medial sternocephalic and lateral brachiocephalic muscles. Use of the left side is not recommended due to risk of causing trauma to a vulnerable nerve. Suggested needle sizes: newborn: 21 gauge x 1 inch; <20 kg: 20 gauge x 1 inch; >20kg: 20 gauge x 1.5 inch.

d. Advance the needle caudo-dorsally towards the opposite scapula. Pull gently using a syringe or apply vacuum with the vacutainer snapped into place. If more than one vacutainer fill is required change the vacutainers without removing the needle. For large pigs the 1.5 inch needle usually must be inserted its full length.

e. Remove the vacuum or release pressure on the syringe prior to removing the needle. Remove the needle and briefly apply pressure to stop blood flow.

f. The size and type of the vacuum tube depends on the type and amount of sample required.

II. Marginal Ear Veina. Move pig into blood collection area. Restrain the pig. For small pigs less than 25 kg one

person can restrain the pig by turning the pig onto its back, holding the front legs and restraining the body with the person’s arms or by straddling the pig and using their legs. Larger animals require restraint using a snout snare and will remain standing. Allow the pig to bite the snare and then close it quickly and apply pressure. The pig will remain quiet when pressure is maintained.

b. Warm the ear to dilate the vessel – either gently stroke the ear or apply a swab soaked in warm water.

c. Occlude the vein at the base of the lateral surface of the ear.d. Insert an appropriate sized needle (21-23 gauge) attached to a vacuum test tube holder

and tube towards the base of the ear.

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e. Remove the vacuum or release pressure on the syringe prior to removing the needle. Remove the needle and briefly apply pressure to stop blood flow (about 2 minutes).

f. The size and type of the vacuum tube depends on the type and amount of sample required.

POULTRY/BIRDS

I. Cardiac Puncturea. Animal should be fasted overnight.b. Prepare vacutainers or collection vials with labels.

i. Ensure that the proper anti-coagulant is being used (if necessary).c. Have an assistant hold the rooster (hen) by the feet and legs with the left hand and the

wings over the back with the right hand.d. Pluck a few feathers from the breast area near the middle of the body. You will notice a

V-formation of thickened skin (where the larger feathers were anchored). You may use this as a guide for point of entry of the needle.

e. Wipe the area with alcohol. Try to locate a strong heart beat with the left thumb.i. Keeping the left thumb over the point of the strongest heart beat as another guide,

hold the vacutainer assembly in the right hand, insert the needle at the tip of the "V" of skin and follow through at an angle of 35-40. You may have to insert the total length of the needle before the blood will appear at the tip of the needle inside the vacutainer holder.

f. If blood is not seen, withdraw the needle and try again (avoid unnecessary jabbing). NOTE: As there is only a small space allowance on the side of the breast bone where the

needle can pass through to reach the heart, any deviation of puncture site of skin or angle of needle insertion may bring the needle against the breast bone and dull the needle tip. If this happens, start again using a fresh needle to minimize the damage to the heart.

II. Jugular Veina. The right jugular vein is the preferred vein due to its larger size relative to the left

jugular.b. Prepare equipment. Attach appropriate sized syringe to a 22-28 gauge needle

(depending upon size of bird). If an anticoagulant is necessary first rinse the syringe with the anticoagulant and discharge contents such that a small volume of the anticoagulant remains in the hub of the needle.

c. Remove the bird from the cage or holding bag.

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d. Hold the bird in one hand with the bird’s right side up (left lateral recumbency) and neck and head held between two fingers (index and middle fingers) to gently extend the bird’s neck. If the bird is too large then a towel can be used to restrain the body while the free hand is used to restrain the head and neck.

e. Gently blow on the neck to expose the unfeathered area on the lateral side of the neck. Apply a small volume of 70% alcohol to the area to wet the feathers, sterilize the area, and expose the jugular vein.

f. Apply a small amount of pressure at the base of the neck just cranial to the thoracic inlet to occlude the jugular vein and allow it to distend and become visible.

g. With the bevel down, pierce the skin with the needle directly beside the jugular at a 30° angle to the skin. Provide gentle suction on the needle by retracting the plunger of the syringe and then decisively pierce the jugular vein. (Placement of the needle with the bevel facing down reduces the potential for lacerating the opposite wall of the vein and the risk of causing vessel collapse and vacuum-associated cessation of blood flow).

h. Blood will flow into the syringe if the needle is in the jugular vein. Take care to avoid pushing the needle through the vein.

i. Slowly retract the plunger to allow blood to enter the syringe until the desired volume is collected.

j. Gently remove the needle and apply gentle pressure to the puncture site for 30 seconds to 2 minutes. Look for any excess bleeding through the skin and if note apply additional pressure.

k. Return the bird to the cage or holding bag.

III. Brachial veina. Prepare equipment. Attach appropriate sized syringe to a 22-28 gauge needle

(depending upon size of bird). If an anticoagulant is necessary first rinse the syringe with the anticoagulant and discharge contents such that a small volume of the anticoagulant remains in the hub of the needle.

b. Remove the bird from the crate, pen or holding bag.c. For a two person technique, one person gently places the bird on its side on the table

and with one hand holds the head and with the other hand holds the legs. The other person lifts up the wing.

d. Gently separate the feathers apart to visualize the brachial vein. Apply a small volume of 70% alcohol to the area to wet the feathers, sterilize the area, and expose the brachial vein.

e. With the bevel down, pierce the skin with the needle at a 30° angle to the skin. Provide gentle suction on the needle by retracting the plunger of the syringe and then decisively pierce the brachial vein. (Placement of the needle with the bevel facing down reduces

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the potential for lacerating the opposite wall of the vein and the risk of causing vessel collapse and vacuum-associated cessation of blood flow).

f. Blood will flow into the syringe if the needle is in the brachial vein. Take care to avoid pushing the needle through the vein.

g. Slowly retract the plunger to allow blood to enter the syringe until the desired volume is collected.

h. Gently remove the needle and place the bird wing back down so it is tight to the body to stop the flow of blood. Look for any excess bleeding through the skin and if note apply additional pressure.

i. Return the bird to the crate, pen or holding bag.

5. SafetyPersonal protective equipment (gloves, lab coat) must be worn at all times. Care must be taken to restrain animals properly and prevent injury, particularly needle sticks. Consult the MSDS for isoflurane to ensure the proper safety procedures are followed. Any injuries must be reported to the principle investigator and WSEP.

6. Potential Complications and Troubleshooting

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7. References

Diehl KH et al (2001). A good practice guide to the administration of substances and removal of blood, including routes and volumes. J Appl Toxicol 21:15-23.

Golde WT, Bollobin P, Rodriquez LL (2005). A rapid, simple, and humane method for submandibular bleeding of mice using a lancet. Lab Animal 34(9):39-43.

McGuill MW, Rowan AN (1989). Biological effects of blood loss: Implications for sampling volumes and techniques. ILAR News 4:5(Fall).

First Report of the BVA/FRAME/RSPCA/UFAW Joint Working Group on Refinement (1993). Removal of blood from laboratory mammals and birds. Laboratory Animals 27:1-22.

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Problem ResponseAnimal moves excessively during the procedure

Re-evaluate restraint and/or get help. Operator safety must be the utmost concern when working with large animals. Accidental intra-arterial injection can result in violent reactions and poses a serious danger to both the horse and people around. Do not proceed unless the horse is appropriately restrained.

Failure to obtain blood Re-evaluate needle position and repeat. Get help from trained personnel if you are unsuccessful after 2 attempts.Blood may have clotted in the needle. Use a new needle and repeat the procedure.

Arterial blood is obtained (bright red, squirting from needle)(IV injection)

Re-position the needle. Do not proceed with injection unless the needle is properly seated in the jugular vein. Use a new needle if necessary

Vacuum is lost from the collection tube (jugular collection)

Use a new collection tube and repeat the procedure

Blood flows into the tube at first but stops (jugular collection)

Re-evaluate needle position, especially if the animal has moved. If the needle has been removed from the skin accidentally and vacuum is lost, use a new collection tube and repeat the procedure.Blood may have clotted in the needle. Use a new needle and repeat the procedure.

Blood is seen in the hub of the needle at first but cannot be aspirated once the syringe is attached (IV injection)

Re-evaluated needle position, especially if the animal has moved. This is best done by removing the syringe and repositioning the needle. Do not proceed with injection unless the needle is properly seated in the jugular vein.

Blood may have clotted in the needle. Use a new needle and repeat the procdure.

Hematoma formation (swelling) after needle removal

Apply pressure for 2-3 minutes. Do not allow the horse to lower its head. Inform the principal investigator or UCACS veterinarian.

Needle stick injury Wash your hands with copious amounts of water and soap. Inform the principal investigator and WSEP.


Guidelines for survival bleeding of mice and rats; NIH: http://oacu.od.nih.gov/ARAC/Bleeding.pdf

8. Revision HistoryDate Created: February 26, 2013 Written by: Nelles/ Moroz/

Lohmann/Woodbury/AlcornSOP Review and Revision History

Revision Number Review/Revision Date Reviewer

List the changes made during the most recent revision, as well as the reasons for the changes.

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http://oacu.od.nih.gov/ARAC/Bleeding.pdf

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