Blood Collection

Blood Collection Aulanni’am Animal Care and Use Brawijaya University Aulani_PKH_UB

Agouti Gold Black Chinchilla Brown Black&Tan Silver Himalayan Aulani_PKH_UB

Rabbits --pet animal.

Breeds of the Rabbits identified so far. [ARBA]

The Animal Research Committee (ARC) limits survival blood collection to 1.25% (1.25 ml/100 g) of the animal's current body weight. The frequency of blood collection is dependent upon the volume collected. If the maximum volume is collected, as specified above, blood may be collected once every two weeks. The ARC requires monitoring hematocrit and/or serum protein levels when more frequent collections are necessary. BLOOD COLLECTION LIMITS Aulani_PKH_UB

BLOOD COLLECTION • As a standard recommendation the amount of blood collected every 3 weeks should not exceed 15% of the total blood volume of the animal. Withdrawal of 15% of total blood volume equates to approximately 1% of body weight. • Ear vessels in the rabbit (central ear artery or marginal ear veins) are readily accessible and can be used for collecting blood. • However, in animals with small ears and very small veins, a possible thrombosis of the vessel with subsequent sloughing of the skin may occur. • The most risk of sloughing occurs when the ear artery is used. Aulani_PKH_UB

BLOOD COLLECTION • Rabbits must be tranquilized to reduce handling stress, enhance vasodilation and prevent injury. • The use of Acepromazine at a dose of 1 mg/kg IM or SC is recommended. • The use of topical anesthetic is also recommended. • Should be administered 15 t 30 minutes prior to the procedure to have an effect. • Blood collection from rabbit ears by transecting the vein or the use of xylene or other irritants is not permitted. Aulani_PKH_UB

The Australian Code of Practice for the Care and Use of Animals for Scientific Purposes and the Guideline Animal Research Application Form (Model): • 1 The volume of blood to be collected • 2 The frequency of blood collection • 3 The period over which blood will be collected • 4 The route by which blood will be collected • 5 The technique by which blood will be collected (eg acute • venipuncture vs chronic indwelling intravenous catheter) • 6 The method by which the animal will be restrained for blood collection • 7 The use of anaesthesia and / or analgesia 8 The methods of animal monitoring and frequency with which these methods will be implemented. • 9 The experience of the operator relevant to the species of animal to be used and the blood collection procedures to be undertaken. Aulani_PKH_UB

Blood Collection • The amount of blood needed and other factors will govern the method and sites of collection. • Descriptions of the various techniques for venipuncture in different species is available in the Animal Care Unit (400 ML; 335-7985) in text and videotape format. • Proper insertion of the needle into a vein or other part of the vascular system is normally the most difficult part of the procedure. • Certain guidelines can be given, but only practice provides proficiency. Aulani_PKH_UB

Blood Collection • Veins may be expected to roll, collapse, or shift, making entrance difficult. • A precise, careful introduction of the needle is best and several attempts may be required. • Starting at distal sites will allow repeat attempts more proximally. • The needle is inserted parallel to the vein and the tip directed into the lumen along the longitudinal axis. • When withdrawing blood from a vein, aspiration should be slow so the vessel does not collapse. Aulani_PKH_UB

Site Preparation • The area of injection or incision should be cleaned with alcohol. • Some procedures will require sedation or anesthesia; others may be carried out without anesthesia provided suitable restraint is used. • In order to better visualize veins dilation can be accomplished by immersing the tail in warm water for 5 to 10 seconds or by warming the animal with a low-wattage light bulb for 5 to 15 minutes prior to venipuncture. • This also aids by providing additional light Aulani_PKH_UB

GENERAL PROCEDURES • Amputating digits for the collection of blood is considered unacceptable and incompatible with the humane care and use of laboratory animals. • Requests to use alternative blood collection methods, including transection of vessels or tail clipping may be considered following consultation with a veterinarian. • The alternate method must be described in the ARC protocol. • Survival blood collections may be performed without general anesthesia if peripheral blood vessels are used. • Nonsurvival exsanguinations must be performed under general anesthesia or following carbon dioxide (CO2) or pentobarbital euthanasia, as the heart may continue to function after euthanasia via these methods. Intracardiac bleeding is not permitted without scientific justification and must be conducted under general anesthesia due to the painful nature of blood collection via this route. Aulani_PKH_UB

PROCEDURE • Sedate rabbit with 1 mg/kg of Acepromazine IM 15 minutes prior to blood draw. • Place the rabbit in the restrainer. • Shave or gently pluck the fur over the vessel. • Rub or tap the area with your finger to dilate the vessel. • Clean the skin with alcohol. • Holding the needle with two fingers, penetrate the vein or artery with a small 20 or 22 gauge needle with the bevel up. Sampling of blood from the vein should be performed as close to the base of the ear as possible, whereas sampling from the artery should be performed nearer to the tip of the ear. Additional attempts can be made distally toward the ear tip for the vein and proximally toward the base for the artery. (See diagram.) • Allow the arterial blood to drip from the needle hub and free catch it in an appropriate collection tube. • After collecting the desired amount, apply pressure in the site to create hemostasis for at least 2 minutes. Aulani_PKH_UB

PROCEDURE • Flip the ear back and forth to assure that bleeding will not re-start. • Only release animal from restrainer when bleeding has completely stopped and no hematoma is formed. • Log necessary information in the Rabbit Use Record. • Complete anesthesia-operative report . Complete Anesthesia Monitoring/Surgery/Post Operative Form. Complete post-operative/anesthesia report. • Check rabbit 15 min. to ½ hour later after being returned to cage. • Submit form to area supervisor or veterinary technician office. Aulani_PKH_UB

PROCEDURE • TERMINAL CARDIAC BLOOD COLLECTION • Intracardiac puncture for large volume blood collection is limited to terminal procedures only and is performed under general anesthesia. It is not an acceptable method for blood sampling purposes. • Materials: • Syringes (5-60 ml) with hypodermic needles (20-25G) • Evacuated containers with 16-18G needles • Anesthetic drugs - Ketamine HCl (100 mg/ml) and Xylazine (20 mg/ml) • Isopropyl Alcohol • Gauze Aulani_PKH_UB

Procedure. • Anesthetize rabbit with Ketamine/Xylazine at 35/5 mg/kg IM respectively. • Once the animal is in surgical plane of anesthesia, lay animal on its back using the restraining board. • Prep area with alcohol swab and insert needle at base of sternum (under the Xiphoid) at a 30-45° angle just lateral to the midline (rabbit's left side). See diagram. • Or, the needle can be inserted into the lateral thoracic region toward the area of maximal heart palpitation between ribs of rabbit's left side midway between sternum and back under left elbow. • If using syringe, aspirate slowly the desired amount. If using evacuated container, open clamp on collection set, the blood should flow quickly into the container. The animal succumbs to exsanguination. • Verify that heart has stopped. If it is still beating, euthanize with Fatal Plus IV or intracardiac (100 mg/kg). This will ensure that the heart has stopped prior to disposal. • Submit completed Rabbit Use Record to the veterinary technicians' office, room 204, or area supervisor. • Complete cage card by dating lower right corner. • Complete Animal Removal Card. • Submit completed cage card and animal removal card to area supervisor or leave them on the sink in the animal holding rooms in BEB for floor supervisor to collect. Aulani_PKH_UB

RODENT PROCEDURES Aulani_PKH_UB

RODENT PROCEDURES • Retroorbital blood collection must be performed under general anesthesia, followed by the administration of an antibiotic ophthalmic ointment to both eyes to prevent desiccation and infection; however, when a short-term anesthetic such as isoflurane is used, application of antibiotic ophthalmic ointment is only necessary for the affected eye. • Individuals performing retroorbital blood collection must be adequately trained due to the potential for significant complications. • The ARC does not recommend retroorbital blood collection in rodents for serial sampling. • This procedure has been associated with histopathologic and clinical changes in orbital tissues, including hemorrhage, inflammation, and infection. • As an alternative to retroorbital puncture, the ARC recommends the use of the lateral saphenous vein, located on the hindlimb, for blood collection. This technique may be performed in conscious animals and requires clipping the fur from the area around the vein and swabbing with 70% alcohol prior to puncture.2 Repeated sample collection is permitted, provided the volume does not exceed 1.25% (1.25 ml/100 g) of the animal's current body weight every two weeks. Aulani_PKH_UB

Lateral tail vein venipuncturein rat • Equipment: scalpel blade, 25 to 30 gauge needle • The rat is restrained (picture, picture) using a mechanical device. • The veins may be seen laterally near the base of the tail but good illumination and dilation will normally be required. • Dipping the tail in warm water will help dilate the vessels. • A small blood sample may be collected by capillary action using a microhematocrit tube inserted into the hub of a small needle previously placed into the tail vein (future picture). • Blood pooling can be induced by placing a small rubber band around the base of the tail. Aulani_PKH_UB

Ventral tail artery • Equipment: 22 gauge needle, 3 cc syringe • The rat should be anesthetized and placed in dorsal recumbency. • Bleeding from the tail may be increased by warming it in water at 40 C to 50 C. • Induce arterial dilatation by applying pressure 1 to 2 inches from the tip of the tail with a finger. • Remove the syringe plunger and place the needle bevel up into the tail artery entering at a 20 to 30 degree angle (future picture)6. • If placed properly the syringe will immediately begin to fill with blood. • If blood flow is slow or stops, slowly withdraw the needle to re-establish flow. • Retries should always be done with a new needle in a more proximal location on the tail. • Pressure on the site may be necessary to cease blood flow after needle withdrawal. • Toe clipping or tail clipping to obtain blood samples: Clipping toes is an unacceptable method of blood collection. Tail clipping is not a preferable method for blood collection. Aulani_PKH_UB

Cardiac puncture • Equipment: 0.90 to 0.50 mm needle • Cardiac puncture represents an accepted method of blood collection from rats when more than a few drops are required. However, this method also carries considerable risk to the animal and occasionally deaths occur. • Therefore, it is not recommended as a repetitive blood sampling procedure. • Animals must be anesthetized and restrained in dorsal recumbency. • The needle is inserted under the xyphoid cartilage slightly to the left of midline (picture). • The needle is advanced at a 20 to 30 degree angle from the horizontal axis to the sternum to enter the heart. • Aspirate lightly while advancing. • Blood should be withdrawn slowly, and the amount must be limited (up to 4 ml in an adult rat) unless euthanasia is planned. Aulani_PKH_UB

Orbital sinus venipuncture in rats • Equipment: Capillary tubes • Blood collection from the orbital sinus of rats is frequently used. • Bleeding requires that the tube be directed into the orbital sinus which surrounds the globe. • In the rat, the capillary tube is inserted in the medial canthus with gentle rotation while directing the tube caudally and towards the midline. • Knowledge of the location of the venous structures of the orbit of the rat aids in establishing a successful peri-orbital bleeding technique. • Pressure should be applied after blood collection to prevent hematomas. 0.5 ml of blood can be obtained weekly using this method. • Anesthesia is required for all peri-orbital bleeding procedures Aulani_PKH_UB

REFERENCES • Van Herck, H. et al. Histological Changes in the Orbital Region of Rats After Orbital Puncture. Laboratory Animals (1991b) 26: 53-58. • 2 Hem, A.; Smith, AJ; Solberg P. Saphenous Vein Puncture for Blood Sampling of the Mouse, Rat, Hamster, Gerbil, Guinea Pig, Ferret, and Mink. Laboratory Animals (1998) 32:364-368. • Diehl KH, Hull R, Morton D et al (2001). A good practice guide to the administration of substances and removal of blood, including routes and volumes. Journal of Applied Toxicology 21: 15-23 • Hem A, Smith AJ and Solberg P (1998) Saphenous vein puncture for blood sampling of the mouse, rat, hamster, gerbil, guineapig, ferret and mink Laboratory Animals 32: 364 - 368 Aulani_PKH_UB

REFERENCES • McGill MW and Rowan AN (1989) Biological Effects of Blood Loss: Implications for Sampling Volumes and Techniques ILAR News 31 No 4: 5 – 18 • Morton DB, Abbot D, Barclay R et al (1993/1994) Removal of blood from laboratory mammals and birds - First report of the BVA/FRAME/RSPCA/UFAW Joint Working Group on Refinement Laboratory Animals 27: 1 - 22; 28: 178 - 179 Aulani_PKH_UB

Thank You!! Aulani_PKH_UB

Blood Collection