Collection of Parasitological Samples from Livestock and Birds

Materials to be collected for Parasitological Analysis


  1. Faecal Sample                                     2. Blood Slide
  1. Lymph node smears                            4. Whole Blood ,  5. Serum
  1. Impression Smear                               7. Adult Helminths
  1. Skin scrapings                                     9. Ticks/fleas/lice/any other arthropod


Faecal Sample


a) Fresh faecal material

b) Collect immediately after defecation

c) Collect directly from the rectum of large animals

d) Collect at least several gram of faeces (the size of human’s thumb) for small animals


a)  If collected faecal sample cannot be examined within a few hours, the sample should be refrigerated until it is tested.

b) Faeces should not be frozen

c) Faeces should be examined immediately for suspected case of protozoan infection             (Trophozoites)   

d) Faecal sample containing helminth eggs, can be preserved with an equal volume of 10% formalin

e) Adult Nematode can be preserved in 70% alcohol/ 10% formalin

f) Adult trematodes –first flattened in between two slides and tied up with tags, then preserved in 10% formalin

g) Adult Cestodes- Preserved in 10% formalin or 70% alcohol, precaution should be taken during preservation of scolices.

h) Faecal Sample for Poultry coccidiosis -The dropping may be kept in 2.5% Potassium–dichromate (KMnO4) solution.


a) Faeces should be dispatched/ submitted in a sealed glass or plastic container, clearly marked with the time & date of collection, species of animal, animal’s name, owner’s name, age of the animal and any other information (History) relevant to the case

b) The physical condition of the stool should be noted for color, consistency, and presence of parasites

c) When material is to be sent another laboratory, it should be packaged with ice packs.



Macroscopic Examination

  1. Intact worms or proglottids may be identified on the surface of faecal material
  2. Consistency of faecal material(Liquid/Soft/Formed/Semisolid), soft or liquid faecal material should be examined within one hour of passage to ensure the motility of the parasites
  3. Colour of the faecal material: if muddy colour, it may be due to ascarid worm infestation, greenish color may be due to strongyloidiosis, whitish due to cryptosporidiosis.

Microscopic examination

Direct smear method

  1. Thin smear of  emulsified faeces (to demononstrate the presence of parasite stages-eggs/larvae of helminthes, oocyst of Eimeria sp.)
  2. Small quantity of faeces is placed on a slide
  3. Mixed / dilute with N.S. or ordinary water
  4. A drop of the mixed faecal material spread into a thin layer
  5. Put coverslip on the faecal smear
  6. Observed under the microscope at low (X100) and high (X400) power objective lens to identify the specific parasitic egg
  7. At least 2-3 slides to be examined before giving conclusion.

Parasitic ova found with this method: Coccidian oocyst , helminthic eggs, cestode & trematode eggs (mainly birds)

Concentration Method

  1. Sedimentation by gravitation     
      1. Take approximately 3 g of faeces, mix it with water in a pastle morter
      2. Filter the faecal suspension through a tea strainer or double layer of cheesecloth into urin glass/beaker
      3. Add water to wash strainer until the urine glass is almost full
      4. Keep it standstill for 10-15 min to sediment
      5. Remove (pipette, decant) the supernatant very carefully
      6. Repeat the washing step for 4-5 times till the supernatant becomes clear
      7. Take a drop of sediment to examine under microscope
  2. Sedimentation by centrifugation
      1. Emulsify small amount of faeces in water
      2. Strain it into centreifuge tube
      3. Centrifuge at 2000 rpm for 3-4 min
      4. Discard the supernatant and mix the sediment with water again
      5. Repeat washing by centrifugation followed by discard of supernartant for 2-3 times
      6. Examine the drop of sediment as before


Application: The procedure can be used to assess the presence of trematode infections like liver fluke (Fasciola) and amphistome eggs.


  1. Simple flotation method
  1. Put approximately 3 g of faeces with floatation fluid (saturated sodium chloride solution) in a pestle morter
  2. Mix the contents thoroughly
  3. Pour the faecal suspension through strainer into centrifuge tube erect in stand
  4. Add floatation fluid upto the brim of the tube
  5. put a cover slip on the top of the tube
  6. Allow to stand for 10-20 min.
  7. Gently mount the cover slip on micro slide for microscopic examination of egg /larvae
  1. Flotation by centrifugation

This method provides higher concentration of parasite objects, practically free from detritus by combining the principle of gravitation and floatation.


      1. Follow the step of sedimentation by centrifugation method
      2. After last centrifugation, the sediment resuspended in floatation fluid
      3. Centrifuge again  at 2000 rpm for 1-2 min
  1. Keep the tube in erect stand
  2. Add floatation fluid  with a pipette upto the brim of the tube
  3. put a cover slip on the top of the tube
  4. Allow to stand for 5-10 min.
  5. Gently lift the cover slip and place it on clean slide to examine under microscope


Application: This is simple technique for use in initial surveys & to detect low numbers of helminth eggs especially nematode ova.

  1. Formalin-Ether technique

This method is very useful in the circumstances which require preservation of faecal sample and floatation method is not so effective due to presence of fat and fatty acids.

  1. Emulsify 1-2 gm of faeces in 15 ml water
  2. Strain it and transfer the filtrate into 15 ml centrifuge tube
  3. Centrifuge at 2000 rpm for 2 min and decant the supernatant
  4.  Repeat the washing steps until supernatant is clear
  5. Mix 10 ml of formalin (10%) to the sediment and allow to stand for 10 min or longer for fixation
  6. Add 4 ml of ether and shake vigorously using stopper
  7. Centrifuge at 2000 rpm for 2 min
  8. Carefully loosen the plug at ether-formalin interface with a pipette
  9. Pour off the entire supernatant along with the plug
  10. Take a drop of sediment, mix with a drop of 2% iodine
  11. Put a cover slip and examine under microscope




  1. Stoll’s method
  1. Weigh 1 gm of faecal sample in a balance
  2. Mix with 15 ml of water ( 1 gm in 15 ml)
  3. Strain it, then shake the filtrate to form uniform mixture
  4. Take out 0.15 ml (1/100) on a slide and put coverslip over it
  5. Count the eggs of whole content (0.15 ml)
  6. Multiply the number with 100 to get E.P.G (eggs per gram of faeces) value
  1. McMaster’s method
  1. Weight 1 gm of faecal sample in a balance
  2. Mix with 15 ml of floatation fluid ( 1 gm in 15 ml)
  3. Strain it, then shake the filtrate to form uniform mixture
  4. Charge on special (McMaster slide) where exactly 0.15 ml can be charged into each of the 2 chamber
  5. Count the eggs within ruled areas of both the chamber under low power (X100) microscope
  6. Multiply the number with 100 to get EPG value



Coccidian infections are very much common and important for poultry and ruminant especially in young animals. Many species are responsible for causing coccidiosis in animals and birds and are strictly host specific. When passed in faeces the coccidian oocyst are unsporulated and can not be differentiated. Therefore, culture of faecal sample for sporulation of coccidian oocysts is very much essential for diagnosis as well as for epidemiological study.


  1. Take a small amount (3-5 gm) of faeces in pestle-mortar
  2. Add 2.5% potassium dichromate (KMnO4) to emulsify it
  3. Filter through a tea strainer
  4. Pour the filtrate into petridish to the depth of ½ cm only (if necessary, add KMnO4 solution)
  5. Keep at room temperataure
  6. Examine under microscope at every 6-12 hrs for  complete sporulation
  7. Record shape, size my micrometry and the sporulation time for identification of different species.




  1. Blood Slide
  1. Thin /Thick Smear must be prepared from a single drop of blood
  2. Use Clean grease free slides
  3. Smear can also be prepared from anti-coagulated (EDTA) blood sample submitted in a vacutainer
  4.  Prepare film should be smooth & even, it should be 3-4 cm. long
  5. Labeled the film/ mark the slide
  6. Air dry the film immediately by waving in the air (this avoids cremation of the erythrocytes)
  7. Blood Slide may be sent through slide mailer.



    Preparation of Thin smears

  1. Place a small drop of blood (1.5 –2 cm. from one end of a clean slide).
  2. Hold the spreader at one angle of 45° in front of the drop of blood and bring back to touch the blood allowing it to spread along the edge.
  3. Make the film by pushing the spreader forward a direct and even movement.
  4. The film should be 3-4 cm. long.
  5. Air dry rapidly by waving the film in the air
  6. Labeled the film either by writing with a pencil in the film itself or, if in the laboratory, marks the slide using a diamond tipped pen.
  7. Fix the film in methanol for 2 minutes as soon as the film detoriates rapidly and so it is advisable to carry a small bottle of methanol for this purpose when out doing fieldwork.


Preparation of Thick smears

  1. Place a drop of blood at one end of a slide.
  2. Touch the drop of blood with a spreader & make thin film starting at 3 cm. from the end of the slide.
  3. Spread the remaining blood in the shape of a cube (3 x 20 mm). If the film is too thick, it will peel off the slide.
  4. Air dry and draw a line between two films.
  5. The haemoglobin can be removed from the thick film by inverting the slide and placing it an angle in distilled water (2 min).


Examination of a wet-smear

  1. Take a drop of freshly collected blood on a clean glass slide and put a cover slip on it
  2. And examine under light microscope (first 10X and then 40X objective lens)
  1. By this method, diagnosis of Trypanosoma sp. and microfilaria is very easy, because the viable and motile organisms can easily be seen.
  2. Protozoa in faecal smears and Trichomonas sp. in vaginal smear can also be identified in fresh smear.


Lymph node Smear

  1. Locate the position of lymph node (sub-scapular region and other superficial lymph node) and hold it firmly
  2. By means of syringe and needle pierce the gland & move the needle  up to medulla
  3. Pull the plunger of the syringe to aspirate the sample of the contents of the gland & withdraw the needle
  4. Expel the contents to a clean slide and make the smear
  5. Lymph node smear is required for identification of Koch’s blue body and for diagnosis of bovine tropical theileriosis.


Whole Blood/Serum

  1. Blood may be collected with EDTA and send it through ice-packs
  2. Serum (Blood without EDTA- Serum) may be collected & send in vial packaged with ice packs
  3. Blood should be collected at the time of pyrexia.
  4. Proper identification no. and history of the case must be included during sending of samples





Giemsa’s staining  

This technique varies according to individual preference. The stain usually used in Giemsa diluted 1:9 (timing, dilution and pH may vary slightly with different batches of stain)

  1. At first, the smear is fixed with conc. Methanol and air dried 
  2. Measure 1 ml of concentrated (stock) Giemsa stain  into a  clear empty well ( the stock satin may need to be passed through filter to remove particles)
  3. Add 9 ml of water pH 7.2 ( the ideal pH vary with batches of stain)
  4. Mix well and pour the smear and assure that the whole smear should be covered with stain.
  5. Allow staining for 45 minutes.
  6. Wash the smear in distilled water or tap water
  7. Dry in air and examine under oil immersion objective.



Leishman’s Staining:

  1. Fix air-dried smears in Leishman stain for 2 minutes.
  2. Add an equal volume of buffer distilled water.
  3. Mix by gently rocking the slide to allow even distribution.
  4. Leave staining for 10-15 mints.
  5. Rinse in buffer and immerse until the smear appears pink (1-2 mints) and air dry.
  6. Observe under oil-immersion.





Microfilariae are the larval stage of helminthic parasites, which may occur in the blood, e.g. the microfilariae of canine heart worm, i.e. Dirofilaria immitis. Following methods are applied for diagnosis of microfilaria.


Direct smear

It is very simplest and rapid method sensitivity of the this method is very poor. However, this technique may be used to evaluate the pattern of movement of  microfilariae.


  1. Place one drop of venous blood into a clean microscopic slide.
  2. Place a cover slip over the drop of blood.
  3. Examine the cover slip area under low magnification (x100) of microscope.
  4. Notice for undulating movement of the larvae, which may retain their motility for as long as 24 hrs.

Haematocrit test

This method is slightly more sensitive than direct smear

  1. This technique is not used extensively
  2. Draw fresh whole blood into a micro-haematocrit tube, as for routine packed-cell volume test.
  3.  Spin for three minutes in a haematocrit tube centrifuge.
      1. Examine the plasma portion of the separated blood under low magnification (x100). Swimming microfilariae may be present in the plasma above the buffy coat.

Knotts method

 Concentration method for detection of microfilariae in blood. It is generally considered preferred technique for heartworm screening   because it   is standard, quick, and inexpensive.

      1. Blood is collected into a syringe containing anticoagulant such as heparin or EDTA
      2. Mix 1 ml of the blood with 9 ml of a 2 % formalin solution. If not well mixed, the red cells will not thoroughly lysed, making the test much more difficult to read. Microfilariae, but not red cells, will be fixed by 2% formalin. If 10% formalin is used (the concentration used for fixation of tissues) red cells will also be fixed.
      3. Centrifuge the mixture at 1200 rpm for 5 minutes and discard the supernatant fluid.
      4. Take the sediment on a microscopic slide and add 1 drop of 0.1% of methylene blue using a Pasteur pipette
      5. Examine under low magnification (x10) under microscope. Microfilariae will be fixed an extended position with nuclei stained blue.




Mange is a disease condition of skin caused by wide range of obligatory external parasites, commonly known as mites and classified under the order Acarina, which inhabit different layer of the skin. Mange is a pretty common skin affection encountered in almost all domestic and wild animals, birds and humans. The clinical manifestations included redness, thickening of skin, scaliness and itchiness. Various mange mites encountered in domestic animals and birds are Dedodex spp., Sarcoptes spp., Psoroptes spp., Otodectis spp., Cnemidocoptes spp., Notoedres spp.

Skin Scraping examination

  1. Scraping are taken from the affected area of the body by a scalpel blade or a sharp-edge spoon. The area selected for scraping should be at the edge of a visible lesion and their hair over the area should be clipped away. It is useful to moisten the skin with liquid paraffin so that scrapings adhere to the scalpel. Scraping should be continued until the blood oozes from the surface.
  2. The scraping is transferred on a  clean glass slide and a drop of any mineral oil is added and is mixed properly with a stick.
  3. A cover slip should then be applied and examined under a low objective (10X)
  4.  Demonstration of adult mite or any of the life stages including eggs of mites confirms the diagnosis.
  5. Direct examination of the skin scraping is not very sensitive  for the diagnosis of mange especially, if it is a case of sarcoptic mange in canine.
  6. The following method should be applied to increase the sensitivity.
  7.  The skin scrapping after collection as described above is taken in a centrifuge tube and mixed it with 5-7 ml of 10% KOH and boil the preparation ( while boiling, precautions should be taken to prevent the content of the tube to bump off).
  8. The material is then centrifuged at 1000 rpm for 3-5 min. The supernatant is discarded and the whole sediment is examined microscopically.





  1. Adult ticks and their developmental stages (larva and nymph) remain attached by keeping their mouthparts embedded in skin. Care should be taken during collection, so that no mouthpart may break which is having identification mark.
  2. Mites remain embedded either superficially or deep into the skin of these hosts and can be collected by taking the skin scrapings.
  3. Lice & Fleas pick up with the help of forceps into the container.
  4. Snails are to be collected form water body/pond in a clean air tight container and send it to the laboratory within 24 hr.

Transportation& preservation

  1. Arthropods are usually transported as living specimens in proper containers like plastic /stopper vials
  2. For shorter distances, the arthropods can be transported simply in plastic vials congaing paper labeling and moistened filter paper.
  3. For temporary storage , the arthropods can best be stored at cool temperatures
  4. If stored for a longer period, then it can be stored in wet preservatives like 10% formalin or 70% ethanol.