Blood collection techniques and

anesthesia for laboratory animals

By soma sekhar guptha

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.

• GENERAL METHODS FOR BLOODCOLLECTION

Blood samples are collected using the following techniques:

Blood collection not requiring anaesthesia

• Dorsal pedal vein (rat, mice)

• saphenous vein(rat,mice)

Blood collection requiring anaesthesia (local/generalanaesthesia)

• 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

Terminal procedure

• Cardiac puncture (rat,mice,guineapig,rabbit,ferret)

• Posterior vena cava(rat,mice)

Blood collection from saphenous vein

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:

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.

Collection from pedal vein

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 .

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..

Collection from tail vein

Collection from tail snip

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

Orbital sinus

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.

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

Collection from jugular vein

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.

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

Blood vessel cannulation

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

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 bleedingCaution; not more than 6 samples from

both legs of animal

Collection from marginal ear vein

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

Cardiac puncture

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

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

Anesthesia of experimental animals

Definition

• Anesthetics are the drugs which produce reversible loss of sensation and consciousness

• Anaesthesia in four different stages

1. Stage of analgesia2. Stage of delirium3. Surgical anaesthesia4. Medullary paralysis

• Normally local, general anaesthesia used for animals

• Routes for general anaesthesia are

1. Injection2. Inhalation• Normally barbiturates, chloral

hydrate, ketamine ,urethane used for injection

• Chloroform, ether ,cyclopropane, halothane used as inhalation anaesthetics

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

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

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

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

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

Euthanasia for laboratory animals

• it means "painless inducement of a quick death“ of animal by physical or chemical methods

methods

• Physical methods1. Stunning• Electrical stunning• Stunning with capative bolt2.Cervical dislocation3.Decapitation4.Micro wave irradiation5.Concussion

• CHEMICAL METHODS ;1. 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

Cervical dislocation

• 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

Methods not used for animals

• Physical methods like; hyper thermia ,asphyxia, rapid freezing, pithing, strangulation

• CHEMICAL AGENTS;• Co,nitrogen,NO,CHCL3,mgso4,noc

otine, tri chloro ethane

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

NUDE MICE

• 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

• 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

Thank you