Blood collection techniques from laboratory animals

1. Blood collection techniques and anesthesia for laboratory animals By soma sekhar guptha

2. Introduction Collection of blood from small laboratory animals is necessary for a wide range of scientific research and there are a number of efficient methods available for that . It is important that blood sample collection from experimental animals should be least stressful because stress will affect the outcome of the study. Various regulatory agencies and guidelines have restricted the use of animals and the techniques used for blood collection in laboratory animals.

3. GENERAL METHODS FOR BLOOD COLLECTION Blood samples are collected using the following techniques : Blood collection not requiring anaesthesia Dorsal pedal vein (rat, mice) saphenous vein( rat,mice )

4. Blood collection requiring anaesthesia (local/general anaesthesia) Tail vein( rat,mice ) Tail snip(mice) Orbital sinus (rat, mice) Jugular vein (rat, mice) Temporary cannula (rat, mice) Blood vessel cannulation (rat, guinea pig, ferret) Tarsal vein (guinea pig) Marginal ear vein/artery (rabbit

5. Terminal procedure Cardiac puncture ( rat,mice,guineapig,rabbit,ferret ) Posterior vena cava( rat,mice )

6. Blood collection from saphenous vein

7. Procedure for saphenous vein blood sample collection Requirements; Animal, rodent handling gloves, towel, cotton, sample collection tubes Lateral saphenous vein is used for sampling while taking aseptic precautions. The back of the hind leg is shaved with electric trimmer until saphenous vein is visible. Hair removal cream can also be used. The animal is restrained manually or using a suitable animal restrainar . gently above the knee joint. The vein is punctured using a 20G needle and enough volume of blood is collected with a capillary tube or a syringe with a needle. The punctured site is compressed to stop the bleeding. While collecting blood:

8. Precautions No more than three attempts are made. continuous sampling should be avoided and collecting more than four samples in a day (24-hour period) is not advisable.

9. Collection from pedal vein

10. Collection from pedal vein The animal is kept in a restrainer . The hind foot around ankle is held and medial dorsal pedal vessel is located on top of the foot. The foot is cleaned with absolute alcohol and dorsal pedal vein is punctured with 23G/27G needle. Drops of blood that would appear on the skin surface are collected in a capillary tube and a little pressure is applied to stop the bleeding .

11. Tail vein blood sample collection Requirements include animal, rodent handling gloves, towel, cotton, sample collection tube and animal warming chamber This method is recommended for collecting a large volume of blood sample (up to 2ml /withdrawal) The animal is made comfortable in a restrainar while maintaining the temperature around at 24 to 27°C. The tail should not be rubbed from the base to the tip as Local aesthetic cream must be applied on the surface of the tail 30 min before the experiment. A 23G needle is inserted into the blood vessel and blood is collected using a capillary tube or a syringe with a needle. In case of difficulties, 0.5 to 1 cm of surface of the skin is cut open and the vein is pricked with bleeding lance or needle and blood is collected with a capillary tube or a syringe with a needle. Having completed blood collection, pressure/silver nitrate ointment/solution is applied to stop the bleeding. If multiple samples are needed, temporary surgical cannula also be used..

12. Collection from tail vein

13. Collection from tail snip

14. Collection from tail snip Requirements include animal, anaesthetic agent, cotton, surgical blade and blood sample collection tubes. This method is recommended for blood collection only in mice. This method should be avoided as far as possible because it can cause potential permanent damage on the animal tail. If needed, it should be done under terminal anaesthesia only. Before collecting the blood local anesthesia is applied on the tail and cut made 1mm from the tip of the tail Blood flow is stopped by dabbing the tail tip

15. Orbital sinus

16. Collection from orbital sinus Requirements are animal,anaesthetic agent,cotton,capillary tube Blood sample collected under general anesthesia Topical ophthalmic anesthetic agent applied to eye before bleeding The animal is scruffed with thumb and fore finger A capillary insert into medial canthus of the eye(30degree angle) Once plexus punctured blood will come through the capillary tube 30 min. after collection check for periorbital lesions.

17. cautions Repeated blood collection avoid. Minor mistake will cause damage to eye 2 weeks allowed between 2 bleedings Adverse effects reported by this method are hemetoma,corneal ulceration,keratitis,damage of optical nerve,intra orbital structures

18. Collection from jugular vein

19. Collection from jugular vein Requirements animal, anesthetic agent,cotton,25G needle, collection tubes It is used to collect micro volumes of blood sample 2 persons are needed to collect blood sample The neck region of the animal is shaved& kept hyper extended position jugular vein appears blue color Needle inserted with draw slowly to avoid collapse this vessel Caution Number of attempts is limited to 3 Apply anaesthetic cream before 30 min.

20. Collection with temporary cannula Requirements animal warming chamber remaining same as above It is made on tail vein & used for many hours Tail cannulated with 25G needle Warming required in order to dilate the vein After this animals to be housed individually in large cages

21. Blood vessel cannulation

22. Blood vessel cannulation Requirements heparin, surgical blade .remaining same as above Usually blood vessel used are femoral vein, carotid artery, jugular vein, vena cava Appropriate analgesia be used to minimize the pain After cannulation animal should housed singly in large spacious cage Blood sample collected over 24 hour at volume of 0.1to 0.2 ml After withdrawing cannula flushed with heparin and with draw volume replaced Caution; this is conducted under aseptic conditions because infections block the cannula

23. Collection from tarsal vein Requirements ; hair remover remaining same as above This vein identified on hind legs of large animals It is visible in blue color Hair removed anesthetic cream applied After 20to30 min blood collected slowly Maximum sample per leg 0.1 to 0.3ml Gentle pressure used to stop bleeding Caution; not more than 6 samples from both legs of animal

24. Collection from marginal ear vein

25. Collection from marginal ear vein Requirements; o-xylene,95%alcohol, 26G needle Animal placed in restrainer Ear cleaned with alcohol and local anesthetic applied before 10 min O- xylene used as topical vasodilator here Surgical blade used to cut the vein After collection clean sterile cotton is kept on the collection site

26. Cardiac puncture

27. Procedure for cardiac puncture Requirements 19&25G needle,1-5 ml syringe It is recommended for terminal stage of study to blood collect large volume of blood from animal Animal is in terminal anesthesia while collection of sample Appropriate needle use Blood sample taken from heart Preferably from ventricle. slowly to avoid collapse Caution ; if animal has dextrocardia sampling may fail

28. Collection from posterior vena cava Animal have to be anesthetize and y or v shape cut in abdomen is made and intestine gently removed Liver pushed so vena cava is identified Needle inserted to collect sample This procedure will repeat 3 or 4 time to collect more blood

29. Anesthesia of experimental animals

30. Definition Anesthetics are the drugs which produce reversible loss of sensation and consciousness Anaesthesia in four different stages Stage of analgesia Stage of delirium Surgical anaesthesia Medullary paralysis

31. Normally local, general anaesthesia used for animals Routes for general anaesthesia are Injection Inhalation Normally barbiturates, chloral hydrate, ketamine ,urethane used for injection Chloroform, ether , cyclopropane , halothane used as inhalation anaesthetics

32. Techniques used for inhalation anaesthetics Technique of insufflation ;(open drop method) Pour liquid anesthesia over a gause in a closed chamber After this place the animal in the chamber for anaesthetize It is a simple procedure without valve,co2 absorber But wastage of compound ,drying of trachea of animals occur

33. through anaesthetic machines Open system; ’ Here in a chamber in that inspired& expired gases are separated by valve Inspired gas having mixture of gases Expired gases reaches directly to atmosphere Normally STEPHAN SLATER is widely used system

34. Half closed &closed systems Here co2 absorber is used So it is removed Inspired gas contain anesthetic compounds. But here change the absorber every 8 hours during anesthesia

35. Closed system In this the animal rebreaths the exhaled gas mixture through sodalime which absorbs co2 Only as much o2 and anaesthetic as have been taken up by the animal Here flow rate is low Used for expensive and explosive agents

36. Pre anaesthetic medication It is recommended prior to anesthesia for easy administration of anesthetics Clonidine used to maintain the general anaesthesia . It relive postoperative shivering Midazolam reducing preoperative anxiety Anti-emetic such as droperidol used Anti cholinergics like atropine used to relive salivary, bronchial secretions Melatonin as anti convulsant , anxiolytic , antinociceptive . Xylazine used as potent sedative and muscle relaxant it is used with ketamine Valium used together with ketamine which calms the patient helps to prevent seizures

37. Euthanasia for laboratory animals it means "painless inducement of a quick death“ of animal by physical or chemical methods

38. methods Physical methods Stunning Electrical stunning Stunning with capative bolt 2.Cervical dislocation 3.Decapitation 4.Micro wave irradiation 5.Concussion

39. CHEMICAL METHODS ; Anesthetics in over dose Anesthetics over dose cause un conscious followed by death Higher concentrations of co2 cause unconsciousness Sodium pento barbiturate widley used for this

40. Cervical dislocation

41. In physical methods electrical stunning common methods for pigs Stunning with capatitive bolt effective for larger animals Cervical dislocation destroys the brain stem Decapitation process is head separated from neck cause interruption to blood supply .this is for worm blooded animals In micro irradiation distraction of brain anatomy

42. Methods not used for animals Physical methods like; hyper thermia ,asphyxia, rapid freezing, pithing , strangulation CHEMICAL AGENTS; Co,nitrogen,NO,CHCL3,mgso4,nocotine, tri chloro ethane

43. Recommended methods Mice; decapitation,cervical dis location,80%co2 in air,sodium pento barbiturate(150mg/kg i.p ) Rat; concansion , cervical dislocation, micro wave irradiation, spb 150 mg/kg i.p Guinea pig;80 % of co2,decapitation,spb 150mg/kg Rabbit; stunning with capataive bolt,covcussion,spb120mg/kg Hamster‘;decapitation,80%co2,spb300mg/kg

44. NUDE MICE

45. It is having genetic mutation leads to detoriated or absent of thumus The main appearance of this without hair It can not generate mature T lymphocytes So unable to mount the most of immune responses like 1.Cell mediated immune response 2. Graft rejection 3. Delayed type of hyper sensitivity reactions 4. Anti body formation require CD4,helper T cells 5. Killing of virus infected

46. Absence of functining T-cells prevents nude mice from allografts Imaging, creating tumors It served in lab to gain in sights into immune system Life span;6months to 1year Disadvantages ; Some are having T cells &leaky So knock out mice with more complete immune defect system constructed USES

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