Chapter # 4
Blood Examination
4.1. Objective
The main aim of blood examination is to detect the various parasites or
their larvae in the blood stream for accurate diagnosis of the disease or for the
confirmation of a suspected disease.
A large group of parasites may be examined in the blood. Most of the
protozoan and rickettsial parasites that invade the blood are destructive to
erythrocytes. Hence, when patients show clinical symptoms of anemia, blood
should be examined for hemoglobin content, cellular number and the cells and
plasma should be searched for evidence of parasites. A list of important blood
protozoa and rekettsia is as follows:
Blood Disease Host
protozoa
Anaplama spp. Anaplasmosis Cattle, sheep, deer.
Babesia spp Babesiasis Bovines, canines
Eperythrozoon Eperythrozoonosis Cattle, sheep, swine
spp
Theleria spp Theileriasis Cattle
Haemoproteus spp Haemoproteosis Ducks, geese, pigeon, dove
Leucocytozoon Leuccytozoonosis Ducks, geese, turkey
Plasmodium Malaria Wild birds, pigeon, dove
(haemamoeba)
Trypnosomes Trypnosomiasis Equines, bovines, ovines
Toxoplasma spp Toxoplasmosis All domestic & wild animals and
humans
Leishmania spp Leishmaniasis All domestic and wild animals
and humans
In addition we can also observe some nematodes larvae in the blood e.g.,
➢ Dirofilaria immitis (heart worm of dog, cat, fox and wolf)
➢ Dipetalonema reconditum (subcutaneous worm of dog)
➢ Ornithofilaria fallisensis (subcutaneous worm of ducks)
➢ Setaria cervi (peritoneal worm of cattle)
➢ Setaria equina (peritoneal worm of horse)
Mites are usually parasites of skin and adjacent structures but larvae,
nymph and adults of several mites may invade the internal structures e.g., air
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sacs, bony cavities, nasal cavity, paranasal sinuses etc. commonly found mite
in the blood stream is demodex canis in dogs.
4.2. Collection of blood
The blood is collected from the animals through puncture of vein using
syringe and needle. The common sites for blood collection in different
animals are given under:
Animals Sites
Cattle Jugular vein, ear vein
Horses Jugular vein
Camel Jugular vein
Sheep Jugular vein
Goat Jugular vein
Pig Ear vein, anterior vena cava
Dog Cephalic vein, recurrent tarsal vein
Poultry and laboratory animals Directly from heart
Material
➢ Test tubes.
➢ Syringe, needle (16G in large animals, 18 G small animals)
➢ Cotton swabs soaked in antiseptic.
Procedure
➢ Properly shave the area or clean the hairs with scissors.
➢ Apply a cotton swab soaked in the antiseptic for disinfection. This will
help in accurate doiagnosis and prevention of contamination in the sample.
➢ With the help of sterile needle and syringe, draw the blood by gentle
puncturing of vein.
➢ We can also collect the blood directly in the test tube after pricking the
needle in case of large animals.
➢ The blood after collection should be properly labeled with the name of the
owner, patients breed, species and identification, time and date and the
purpose of the examination.
Precautions
➢ Always use sterile apparatus to prevent any contamination.
➢ Collect at least 5 ml of blood.
BLOOD EXAMINATION
Note: For the diagnosis of malaria and filariasis, the dermal capillary method
obtained by squeezing the sacrified skin in the mid scapular region is reported
to give a higher yield of parasites than the blood taken from the veins.
4.3. Preservation of blood
Usually the blood samples are examined immediately after collection in
diseased animals but sometimes, we have to transport the sample to the
laboratory located at a distant place or we have to delay the examination due
to any reason. As the blood has tendency to clot out side the body we must use
some chemicals to store the blood sample for analysis. These chemicals are
called anticoagulants. Different routinely used anticoagulants with their
dosage are as follows:
Sr. Anticoagulant Dose per 10 mL
No.
1 Lithium oxalate 10-15 mg
2 Potassium oxalate 20 mg
3 Sodium oxalate 20 mg
4 Sodium fluoride 40 mg
5 Lithium citrate 30 mg
6 Sodium citrate 60 mg
7 Heparin Few drops of 1% solution
8 EDTA 20 mg
9 Potassium and ammonium [ammonium oxalate 1.2% pot.oxalate
oxalate mixture 0.8%]-1ml
Instead of anticoagulants we can keep the blood samples in the refrigerator at
4C for 24 hours without much alteration.
Precautions
➢ Anticoagulants should be mixed slowly and gently to avoid hemolysis.
➢ The collection vials should be soaked by rotating in between the palms
for proper mixing of the anticoagulant. Vigorous shaking should be avoided as
it may lead to hemolysis and production of foam.
➢ The refrigerated blood samples should be taken out at least 30 minutes
before the start of examination.
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4.4. Methods of Blood examination
There are two methods of blood examination:
1. Direct method
2. Indirect method
1. Direct method
It is also called as “wet film” in which we can observe the movements of
parasites. It is used for immediate obseravtion of microfilaria.
Materials
➢ Glass slides
➢ Cover slip
➢ Dropper
➢ Microscope
Procedure
➢ Place a drop of blood in the centre of the slide.
➢ Put cover slip on it.
➢ Examine under microscope.
Microfilaria are easily detected in peripheral blood due to their large size and
their intentionless lashing movement, which agitates the red cells and
immediately attracts the eye when observe under low power objective. This
enables the detection of a small percentage of infection.
2. Indirect examination
There are two methods of indirect examination.
a. Thin smear.
b. Thick smear.
a. Thin smear
For the demonstration of morphological features of protozoan parasites such
as malaria, trypanosomes, babesia etc.
BLOOD EXAMINATION
Materials
➢ Cover slip
➢ Microscope slides
➢ Glass spreader
➢ Dropper
➢ 95% alcohol
➢ Methyl alcohol fixative
➢ Coplin jar.
Procedure
➢ Clean the glass slides by dipping or rinsing in 95% alcohol.
➢ Place a small amount of blood towards one end of the slide. Only about
two microlitres of blood is required. More than that will result in a thicker
smear with more than the desired monolayer of blood cells.
➢ Place the end of another slide or spreader on the slide containing the
droplet of blood, positioning it about an inch in front of the droplet.
➢ Holding the spreader at an angle of 45, slowly draw it back until it
touches the droplet of blood.
➢ Allow the blood to spread under the spreader due to capillary action.
➢ In a smooth motion, push the slide forward to spread the droplet out in a
layer over the surface of the bottom slide.
➢ Allow it to air dry for one minute.
➢ Most protozoa can be counted in fresh unstained smear but for critical
study they must be stained.
➢ Fix the smear in methyl alcohol (absolute).
➢ Air dry the smear.
➢ Stain the smear with Giemsa stain.
➢ Observe under microscope.
Precautions
➢ The slides should be free from oil or greese.
➢ There should be no bubble formation or holes on the smear.
➢ Don’t grasp either slide, but allow gravity to hold the two in contact.
➢ The smear should taper off to form a tail towards the end of the slide.
➢ While smear preparation, your hands should rest on a table or other steady
surface.
➢ Remember that a thin film should consist of one layer of evenly distributed
blood cells.
➢ Dry the slides in vertical position after removing from fixative or stain.
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➢ Permanent collection should always have the protection of a cover glass, a
mounting medium such as permount should be used.
Note: There are few stains in which the fixative is present i.e., the fixation and
staining of the dried film are accomplished simultaneously. An example is
Wright’s stain in which methyl alcohol acts as a fixative. It gives fair results
and requires a short staining period but is not recommended for parasitological
use. More precise detail is seen in slides prepared with Giemsa stain. Since it
does not contain a fixative, thin films must be fixed in absolute methyl alcohol
and air before staining.
Advantages of thin smear
➢ Morphological features of protozoan parasites can be detected.
➢ It preserves the structures of a parasite with a minimum of distortion.
Disadvantages
The only disadvantage of thin smear may be that
A relatively small amount of blood is examined that may not be sufficient.
b. Thick smear
Objective
It is prepared for the identification of malarial parasites, trypanosomes and
microfilariae in the blood. As a thick layer of blood is used in this method,
many more parasites will be preserved in each field.
Materials
➢ Dropper.
➢ Slides.
➢ Cover slip.
➢ Microscope.
➢ Buffer solution.
➢ Coplin jar.
Procedure
➢ Clean the slides by rinsing or dipping in 95% alcohol.
BLOOD EXAMINATION
➢ Place the drops of blood cells about the size that would be used to make a
thin film. Close together near one side of slide.
➢ Stir the blood with one corner of another absolutely clean slide, mingling
the three drops over an area of 2 cm in diameter.
➢ Allow the film to dry normally.
➢ After the films are thoroughly dry they must be laked to remove the
haemoglobin. This can be done by immersion in buffer solution ,prior to
staining in a coplin jar.
➢ Air dry the film.
➢ Stain with proper staining procedure.
Precautions
➢ Continue stirring for at least 30 seconds to prevent formation of fibrin
strands, which otherwise tend to obscure the parasites.
➢ Do not heat the film for drying because this will fix the blood.
➢ Thick films that cannot be stained immediately should be laked in buffer
solution before storage, because removal of haemoglobin becomes
increasingly difficult with time.
Note: When Giemsa stain is used for thick films, the procedure is exactly
same as that employed with thin films, except that fixation in methyl alcohol
is omitted.
Advantages
➢ Easily recognition of the extracellular parasites.
➢ More sensitive.
➢ Save time.
➢ Comparatively large amount of blood is used that gives more accurate
results.
Disadvantages
➢ Identification of intracellular parasites is difficult due to presence of red
blood cells.
➢ Increased distortion of the parasites.
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BLOOD EXAMINATION
Avian & mammalian smears
The red blood cells of avian are nucleated and oval in shape while those
of mammalian are disk shaped lacking the nucleus. Thus for avian blood we
make a thin smear to minimise the confusion between RBCs and parasites and
for mammalian blood we make a thick smear. Because there are less chances
of confusion of RBCs with the parasites in thick smear due to absence of
nucleus.
4.5. Preparation of thick and thin blood film on the same
slide
For routine malaria microscopy, a thick and thin film are made on the
same slide. The thin film is used as a label but, if well prepared, is also used
for species confirmation. The thick film should be used for examination.
Technique
➢ Apply gentle pressure to the finger and place a single small drop of
blood, on the middle of the slide. This is for the thin film.
➢ Place two or three larger drops, on to the slide about 1 cm from the drop
intended for the thin film as illustrated.
➢ Thin film Using another clean slide as a spreader and with the blood
drops resting on a flat, firm surface, touch the small drop with the spreader
and allow the blood to run along its edges. Firmly push the spreader along the
slide, away from the largest drops, keeping the spreader at an angle of about
45 as shown in the fig 1. Make sure the spreader is in even contact with the
surface of the slide all the time the blood is being spread. The blood film
should not extend to the edges of the slide in order to prevent infection of the
investigator.
➢ Thick film always handle slides by the edges, or by a corner, to make
the thick film as follows:
Using the corner of the spreader, quickly join the larger drops of blood and
spread them to make an even, thick film. The blood should not be excessively
stirred but can be spread in a circular or rectangular form with 3-6
movements.
➢ Allow the thick film to dry in a flat, level position protected from flies,
dust and from extreme heat. Label the dry film with a pen or marker pencil by
writing across the thicker portion of the film the patient’s name or number and
date as shown in the fig 2. Don’t use a ball pen to label the slide.
➢ Wrap the dry slide in clean paper, and dispatch with the patient’s record
form to the laboratory as soon as possible.
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➢ The slide used for spreading the blood films must be disinfected and could
then be used for the next patient, another clean slide from the pack used as a
spreader.
Figure 1. Preparation of Thick and Thin smear on the same slide
Figure 2. Labeling of the slide
BLOOD EXAMINATION
4.6. Blood Concentration Procedures
1. Knot’s concentration technique for microfilariae
Objective
To concentrate the microfilariae for the identification and accurate diagnosis
of the disease when density of microfilariae in blood is very low.
Principle
Centrifugal force is applied on the test tube containing the microfilariae
and lysed erythrocytes. Due to their heavy weight, microfilariae settle down in
the bottom of the tube from where they are isolated and identified.
Materials
➢ Test tubes
➢ Centrifugation machine
➢ Cover slip
➢ Glass slides
➢ Formalin
➢ Methylene blue
➢ Dropper
Procedure
➢ Draw a sample of venous blood into a syringe containing an
anticoagulant such as EDTA or heparin.
➢ Draw 1-2 ml of air into the syringe and mix the blood and anticoagulant
by rocking the syringe in such a way as to run the air bubble back and forth
along the length of the barrel.
➢ Place 1 ml of blood in a 15 ml centrifuge tube. Add 9 ml of 2%
formaline (2 ml of stock 37%formaldehyde solution and 98 ml distilled
water), stopper and mix by inversion and shaking. This lyses the erythrocytes
and fixes the leukocytes and microfilariae.
➢ Wait for two to three minutes.
➢ Centrifuge for about five minutes and pour off the supernatant by
inverting the centrifuge tube only once.
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➢ Add a drop of 1:1000 methylene blue to the sediment, mix, and transfer
some stained sediment to a slide for microscopic examination.
Note: When submitting blood samples to a laboratory for identification of
microfilariae, complete preceeding steps 1,2 and 3 to prepare them for
shipment.
Precautions
➢ Prolonged delay and thermal extremes should be avoided in this
procedure.
➢ Remix blood immediately in step 2 before proceeding with step 3.
Microfilaria in the blood smear
