Veterinary parasitology Laboratory Guide Manual.pdf

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JKU, School Of Veterinary Medicine Laboratory Guide Manual On Vet. Parasitology! Page i
JINKA UNIVERSITY
COLLEGE OF AGRICULTURE AND NATURAL RESOURSE
DEPARMENT OF VETERINARY MEDICINE
VETERINARY PARASITOLOGY LABORATORY GUIDE MANUAL
BY
MR. MORODA BOGALA
FEB, 2023
JINKA, ETHIOPIA

JKU, School Of Veterinary Medicine Laboratory Guide Manual On Vet. Parasitology! Page ii
ACKNOWELEDGEMENT
All praises and thanks are due to almighty God for all his glorious and merciful that always
keeps my life confident and peace!
First of all I would like to express my appreciation, Head of veterinary medicine department in
Jinka University Dr. Yebelayhun Mulugeta for giving the opportunity to work this handbook.
Finally I would like to thank all of my staff members of veterinary medicine for they support
advice, guidance and encouragement from the beginning to the end of this lab guide manual.

JKU, School Of Veterinary Medicine Laboratory Guide Manual On Vet. Parasitology! Page iii
PREFACE
Laboratory procedures are an essential diagnostic component of veterinary medicine. The role of
the veterinary technician is to collect and process samples and then analyze and report the
findings. This book provides a useful tool for reference when conducting many common
laboratory tests, and serves as an excellent companion to full textbooks.
The following chapters incorporate discussions on the use of common laboratory equipment and
proper quality control methods of vet. Parasitology and also their laboratory diagnosis.
Useful characteristics of this handbook, whether used in laboratory or a clinic setting, are step-
by-step procedures, discussions of common mistakes and errors, tips and tricks, and plenty of
reference images. Color images are invaluable when trying to identify an unknown structure, and
these are concisely provided in each chapter of this guide manual.

JKU, School Of Veterinary Medicine Laboratory Guide Manual On Vet. Parasitology! Page iv
Contents pages
ACKNOWELEDGEMENT............................................................................................................ii
PREFACE......................................................................................................................................iii
INTRODUCTION .......................................................................................................................... 1
CHAPTER ONE............................................................................................................................. 2
1.1 general guidelines for laboratory safety................................................................................ 2
CHAPTER TWO ............................................................................................................................ 4
2.1 Instruments, equipment and reagents used in veterinary parasitology laboratory................ 4
2.2 chemicals and reagents used in parasitology laboratory....................................................... 8
CHAPTER THREE ........................................................................................................................ 9
3.1 Samples collection and examination..................................................................................... 9
3. 2 Collection and submission of samples:.............................................................................. 10
3.3. Quantitative Fecal Examination Methods.......................................................................... 21
CHAPTER FOUR......................................................................................................................... 26
4.1 Method of detection of protozoan parasites........................................................................ 26
4.2 Protozoa Life Stages ........................................................................................................... 26
4.3. Other techniques for diagnosis of protozoan infection...................................................... 30
CHAPTER FIVE .......................................................................................................................... 32
5.1 Method of detection of external parasites........................................................................... 32
5.2 Skin scraping method.......................................................................................................... 32
5.3 Techniques for preservation and mounting of tick, fleas and lice ...................................... 34
REFERENCES ............................................................................................................................. 39

JKU, School Of Veterinary Medicine Laboratory Guide Manual On Vet. Parasitology! Page 1
INTRODUCTION
 This material discusses the roles that veterinary technicians play in assisting the
veterinarian in diagnosing endoparasites and exoparasities of domestic animals.
 An accurate diagnosis of parasitism is based primary up on the veterinarian’s and the
technician’s awareness of parasites that are prevalent in the immediate geographical area
or ecosystem.
 Heavily parasitized animals often show clinical signs suggestive of the infected organ
system- diarrhea/constipation, Vomiting , anorexia
 Internal Parasites of domestic animals comprise several types of organisms that live
internally in animals feed on their tissue or body fluids or compete directly for their food,
these organism range in size from microscopic to more than 1m in length.
 Parasites also vary in their location within the host and in the means by which they are
transmitted from one host to another.
 Because of these diverse variations, no single diagnostic test can identify all end parasites
 The veterinary technician may be asked to perform a wide variety of diagnostic
procedures to diagnose endoparasitism
 The manual includes essential diagnostic tools and laboratory methods for detection of
parasites in deferent organs, tissues, fluids, secretion, execretion, and how to deal with it
 It also helps the student for studying the morphology, of the parasites, their life cycles,
and their clinical singes and pathogenesis that could produce in intermediate and final
hosts of farm animals.
 This manual also designed to aid the veterinary technicians in diagnosis of ectoparasitism
in domestic animals.
 To diagnose an ectoparasitic infestation, the technician must be able to collect the
ectoparasite and then identify the organism involved.
 This guide manual explains procedures commonly used to collect endoparasite and
ectoparasites from the host and describes those parasites so that a correct diagnosis can be
made.

CHAPTER ONE
1.1 general guidelines for laboratory safety
Parasitology laboratories can be hazardous if the rules are not followed. During a laboratory
Course a student may handle materials which are carcinogenic, poisonous, flammable, and
explosive. Some of these materials and equipment may also cause severe burns, cuts, or
bruises if handled improperly or carelessly.
Most accidents that occur in the laboratory are a result of carelessness, impatience,
improper or unauthorized experimentation, and disregard for safety rules or proper
operating procedures. In order to minimize the chances of an accident in the laboratory
certain rules and regulations must be obeyed at all times when one is working or observing
in vet. Para. Laboratory. Therefore, it is not advisable for anyone to work in a laboratory
without proper knowledge of the dangers involved.
The laboratory cannot accept samples directly from patients; samples must be referred to
the laboratory by a health professional.
Due to the inherent dangers present in a laboratory exercise, it should be understood that
the following rules must be obeyed to minimize the chance of an accident. The student is
expected to exercise proper judgment and extreme caution at all times when working in
the laboratory.
Every person who work in the laboratory should obey to the following rule
 Wear gloves when required
 Never mouth pipette
 No smoking or consuming food or drink anywhere in the laboratory
 Do not works with uncovered opened cuts or broken skin cover with suitable dressing
and latex gloves.
 Do not create aerosols use extreme care when operating centrifuges, stirrers, pipettes etc.
 Wipe of benches in your working area with suitable disinfectant before and after each
day’s work.
 Do not wear lab coats outside the lab
 Do not place personal items such as eyeglasses on workbench
 Beware of reactive and poisonous chemicals and handle them with respect.
 All fixatives and chemicals should be properly labeled

 Know in advance where your nearest fire extinguishers are located.
 Always wash your hands before leaving the laboratory.
 Be aware that all specimens may contain biohazards agents and protect yourself
accordingly.
 Make sure your co-workers are aware of any chemical or biological hazards that exist.
 Don’t forget to use PPE rule
 Disinfect the working area after and before leave in the laboratory
 Do not work with hazardous substances without a second person being present in the lab
 Always label containers with the common known name of the substance and the
appropriate hazard warning sign
 Always secure the tops of reagent bottles immediately after use.
 Always clear up spillages immediately.
 Post laboratory signs, including emergency contacts and chemical inventory, outside each
work area.
 Switch off gas, electric and water points when leaving the laboratory.
 Always Keep the test tubes up right on the rack
 Reagents, stains and laboratory chemicals should be labeled and replaced to their
original position.
Note; PPE rule (Personal Protective Equipment)
 The material must be worn at all times in the laboratory
• Lab coat
• Eye protection: Splash proof chemical goggles. If you do get a chemical in your eye, rinse
your eye immediately using large quantities of water or an eye wash bottle
• Closed shoes with socks must be worn at all times – open-toed shoes, backless shoes, sling
backs, clogs, and sandals are not permitted.
• Always wear gloves when working with unknown substances.
• Always wear the appropriate breathing masks when working with toxic or irritating vapour.

CHAPTER TWO
2.1 Instruments, equipment and reagents used in veterinary parasitology laboratory
Pestle and Mortar; used to crush fecal material to diagnose internal Parasites and to crush
crystalline chemicals.
Mesh Sieve; a metallic net that is used to filter undigested materials of fecal sample.
Graduated Cylinder; is usually of different size and shape (type) and usually used for
measuring of solutions.
Centrifuge Test Tubes; used for the centrifugation of samples.
Rack; a plastic or wooden material used for holding test tubes, pipettes, and glass rods.
Pipettes; are different types and used to take the sample and fill the mc-master or used for the
transport/transfer of supernatant solutions. Ex. Pasteur pipette
MC- Master; Egg Counting Chamber- used for counting the number of parasitic eggs
per gram of faeces.
Glass Rod; sticks like glass rods used for mixing of faecal samples and to take a small
amount of sample to prepare a specimen.
Beakers; they could be made of plastic or glass and used to collect a filtrate part of a faecal
sample and to keep solutions
Glass Slide; used to prepare faecal or blood smears.
Cover Slip; used to cover faecal smear.
Petridish; made up of either plastic or glass material used for culturing fecal sample.
Semi-automatic balance; an instrument used for quantitative measurement of faecal samples.
Baermann apparatus; a material made of Baermann stand, flask, funnel, and rubber tube. It’s
important for extracting of the larval of lungworms. It’s also used to recover larva from cultured
& old faeces.
Centrifuge; and electrical service used for the concentrating of the eggs of parasites from
samples of urine, faeces
Gauze; a cloth used for the Baermann apparatus.
Forceps; used for collecting parasites by picking
Goggles and Apron; used to protect your eyes and clothing from damage.
Reagent Bottle; Used to store, transport, or view reagents such as acids or bases.
Funnel; used to safely transfer substances from one container to another.

Wash Bottle; Handy for rinsing glassware and for dispensing small amounts of dH2O for
chemical reactions.
Test Tube Brushes; used for clean tubes before and after use.
Microscope; is the most important piece of equipment in the veterinary clinic laboratory.
The microscope is used to review fecal, urine, blood, and cytology samples on a daily basis
Microscope can be categorized under different type depend on their function and its parts
Binocular Microscope- Microscope with two eyepieces.
Figure1. Binocular microscope

Monocular Microscope- a kind of microscope to magnify samples using one ocular.
Stereomicroscope; used to magnify the morphology of a directed sample and it has two
magnifying parts used for lower and higher parasitic samples like ticks, lice, flea, and parasitic
larval & eggs.
Figure2. Stereomicroscope

Figure 3. General Laboratory equipment

2.2 chemicals and reagents used in parasitology laboratory
NaCl solution; used for floatation purpose (for the concentrating of parasite eggs)
MgSO4 and ZnSo4; also are used as a floatation fluid
Saturated sugar solution; can also be used as a floatation solution
Iodine solution- important for killing the larvae of parasites to fix it (to restrict the movement
of the larvae) and to identify eggs of protozoa.
Charcoal powder – helps to dry faecal samples
Potassium dichromate – help for preservation and culturing purpose.
Formalin solution- concentration percent can vary according to required purposes. It is used
for the preservation of adult parasites. Ex. it could be 3%, 6%, 10%, or 40%.
Note: Parasite eggs preserved by Formalin will not hatch to larval stage.
Ethanol- used as to preserve parasites at 70% concentratioon.
Xylene- used for cleaning of microscope
Methylene blue- is a stain used to identify eggs of parasites. Ex. Fasciola
Lactophenol blue- is used to stain & make clear structures of parasites especially nematodes
Eg. Makes clear both hooks of spicules of Haemonchus & shapes of spicules of T. axei.
Giemsa stain- a staining material used for fixing blood samples that are prepared for
examination.
Methanol alcohol- used for fixing blood samples that are prepared for examination.

CHAPTER THREE
Laboratory diagnosis of parasitism
Samples collection and examination
3.1 Samples collection and examination
 Diagnostic stages of most parasites can be detected in:
1. Feces: used to diagnose parasite eggs, larvae, oocysts, cysts, Trophozoites,cestode segments
adults.
2. Blood: used to diagnose blood parasites: Babesia, Theileria, Trypanosoma, Dirofilaria
immitis.
3. Sputum: used to diagnose lung parasite eggs, larvae for example Dictyocaulus species (eggs)
is cattle and sheep.
4. Urine: used to diagnose eggs in urinary system for example: Dioctophyma renale (giant
kidney worm), capillaria species in dogs and cats.
5. Skin: used to diagnose external parasites such as: Mange (Sarcoptes, Psorptes).
6. Autopsy: from dead animals
7. Biopsy; from live animals.
 Important factors to be considered in the diagnosis of parasitism & the interpretation of
results are:
 Age of the host
 Previous exposure to parasites (resistance)
 Time of the year (ex. spring rise)
 Physiological relationship (parturient rise)
 Geographical location
 Previous use of anthelmintics
 History of clinical diseases, and other considerations.
 All specimens must be clearly labeled with animal species, location in host, date of
collection, place collected, & collector’s name, & telephone number.
 Always handle faecal samples carefully because some zoonotic parasites, bacteria, &
viruses can threat human health.
 Wear clean laboratory coat & rubber or plastic gloves. But if gloves aren’t available wash
hands thoroughly with disinfectants such as soap.

 Always clean up immediately after the tests have been performed.
 Always keep good records.
 Use the objective lenses with magnification of 4x, 10X & 40X, as they are the most often
used in parasitological diagnostic examination.
3. 2 Collection and submission of samples:
Parasites can infect the oral cavity, oesophagus, stomach, small and large intestine and other
internal organs of animals. Detection for these parasites involves collection and microscopic
examination of feces. Diagnosis is usually by finding life cycle stages of the parasite with in the
feces. These stages include eggs, oocysts, larvae, segments (tapeworms) and adult organisms.
Veterinary technicians may perform the following procedures to detect parasitic infections.
As in many other aspects of laboratory analysis, sample collection in parasitology is Important in
achieving accurate results. In small animal medicine, fecal samples are often brought in by
clients. These samples can certainly be acceptable if proper collection instructions are given to
the owner ahead of time. The sample should be fresh.
If samples cannot be examined within 2 hours, they should be refrigerated. If fecal sample are
submitted to the laboratory after being in the environment for hours or day, fragile protozoan
trophozoites will have died and disappeared.
The egg of some nematodes can hatch within a few days in warm weather, and identification of
nematode larvae is far more difficult than recognizing the familiar egg of common species. Also
free living nematode rapidly invade a fecal sample on the ground, and differentiation of hatched
parasites larvae from these free living species can be time consuming and difficult.
As feces age, a diagnosis can be compromised, as eggs larvae and oocysts sporulate. Samples
obtained later from the yard, pen, or litter box are not acceptable. The volume of sample should
be adequate. A minimum of 10g of fresh feces should be collected.
In large animal medicine, fecal samples are collected directly from an animal’s rectum if
possible.
There are several procedures commonly used to examine feces for internal parasites:

A). Gross examination
Upon receipt of a fecal sample, a gross exam should first be performed to examine the physical
characteristics.
 Consistency
Comment on the consistency of the fecal sample. Is it soft, watery, or hard? Keep in mind
That “normal” consistency varies depending upon the species from which it came.
 Color
Unusual color should be noted; as it could give insight into other underlying conditions, for
example, gray stool can indicate pancreatic insufficiency.
 Blood
Note any blood present. Blood may appear bright red (frank), or black (melena or melanous) and
have a tar-like consistency.
 Mucous
Mucous may be present on the surface of fresh feces. This should be noted.
 Gross parasites
Parasites may be visible upon gross examination of the stool. Most commonly seen indog or cat
feces are roundworms and the proglotids of tapeworms.
Microscopic examination of feces;
 Detect the presence of parasite infection by using microscope. The following
methods are helps us to detect the presence of parasite in the host by using
microscope.
1) Direct fecal smear
One advantage to the direct smear method is that is requires very little feces. This technique can
even be used on the small amount of fecal material found on the end of a thermometer and
collected fecal material into the container. The downside to this technique is that with such a
small sample, the chance of finding evidence of a parasite is greatly reduced.

Materials
 Microscope slides
 Coverslip
 Saline
 Wooden applicator stick
Procedure
 Using a wooden applicator stick, mix a small amount of fecal material into a few drops of
saline in the center of a slide.
 Blend gently until the mixture is homogenous.
 Spread out the mixture to a thin layer.
 Remove any large fecal pieces.
 Place a coverslip over the sample.
 Examine microscopically by 10x,40x( 10x is common)
Figure1. Preparation of fecal smear
2. Concentration methods for fecal examination:
A. Qualitative methods: these methods used for determination the types of infection
1). fecal flotation
There are several types of fecal flotation solutions that are used as a semi-quantitative Method of
evaluating a fecal sample. In these methods, an estimate is made of the number of parasite ova
per gram of feces. The principle behind this method is to use the differences in specific gravity of
parasite eggs and cysts from that of fecal debris and the solution.

A fecal sample is mixed with a flotation solution consisting of various salts or sugars added to
water to increase its specific gravity. Parasite eggs and cysts float to the surface, while most fecal
matter sinks to the bottom.
Specific gravity refers to the weight of an object as compared with the weight of an equal
volume of water.
The SG of most parasitic eggs is between 1.100 & 1.200 g/ml whereas the SG of water is 1.00
To allow for floatation of parasitic eggs, oocytes and other life cycle stages, the floatation
solution must have a higher SG than that of a parasitic material several salt and sugar solution
work well for floatation most have a specific gravity of 1.200 to 1.250.
 Floatation fluids: for general purposes
1. Saturated salt solution
 S.G. 1.18-1.20
 General purpose solution
 Sodium chloride(NaCl): 400 grams
 Water : 1000ml
-Stir thoroughly before use
-May distort eggs.
2. Salt/sugar solution
 S.G.: 1.28
 Sodium chloride : 400 grams
 Water : 1000 ml
 Sugar : 500 grams
 Dissolve the salt in water to make a saturated solution
 Add the sugar to the saturated salt solution
 Stir until the sugar is dissolved.
3. Sodium nitrate
 S.G.: 1.18
 This solution is sometimes used for strongly eggs
 Sodium nitrate : 400 grams
 water : 100 ml
 Add sodium nitrate to water while stirring

 Floatation fluids: used for culture
1. saturated sugar solution
 S.G.: 1.27
 This solution should be used if the eggs are required for culturing as it has little effect
on their viability.
 Sugar ( Sucrose): 454 grams
 Water : 355 ml
 Add sugar to the water until saturated
 Stir solution well before using
 Saturation is indicated by the presence of sugar crystal at the bottom of the container after
stirring for 15 minutes.
 To prevent the growth of mould approximately 2 ml of 37% formaldehyde can be added.
 Floatation Fluids: for specialized requirements
1. Magnisium sulphate
 S.G: 1.2
 This solution gives a better recovery of Trichuris, Capillaria and Ascaris
eggs and is best for Metastrongylus
 MgSo4: 400 grams
 Water : 1000 ml
2. Zinc Sulphate
 S.G. 1.364
 This is the best solution for the recovery of Fasciola eggs
 ZnSo4 : 371 gram
 Water : 1000 ml
Notes;
 Saturated NaCl Solution – Strongyle eggs
 Saturated ZnSO4 Solution- Fasciola eggs
 Saturated MgSO4 Solution – Metastrongylus, Trichris, capillaria,
Ascaris eggs
 Saturated sugar Solution if egg culturing is required later

Materials
 Two specimen containers (or wax paper cups)
 Tongue depressor
 Flotation solution
 Fecal shell vial (or test tube)
 Metal tea strainer
 Coverslip
 Microscope slide.
Procedure
 Emulsify 1 to 2g of fresh feces with saturated salt solution into a container, such as a
specimen container or a wax paper cup.
 Strain this mixture through a metal tea strainer into the second specimen container or
mortar
 This strained mixture is then added to the fecal shell vial (or test tube).
 Add more flotation solution to the shell vial until a meniscus is formed.
 Place a coverslip over the meniscus and let it sit for 10–15 minutes (time will vary
depending upon which type of solution is used).
 After the allotted time, remove the coverslip by lifting it directly upwards.
 When laying the coverslip down, place it at an angle in order to decrease the

number of air bubbles that can become trapped underneath the coverslip.
 Examine under microscopic.
Note: It is important not to delay examination. Depending upon the type of flotation Solution
used, delay can result in parasite egg distortion and the solution can begin to crystallize.
 Centrifugation technique
 In comparison with the simple flotation method, the centrifugation technique is
more efficient at recovering parasite ova from a sample.
 It does, however, require a little more specialized equipment.
Materials
 Two specimen containers (or wax paper cups)
 Metal tea strainer
 Tongue depressor
 Test tubes
 Flotation solution
 Coverslip
 Microscope slide.
Procedure
 Mix 2g of feces in 10mL of flotation solution in a specimen container or wax paper
Cup until a suspension is formed.
 Pour the mixture through the metal tea strainer. Using the tongue depressor, press
the material into the strainer to extract as much liquid as possible.

 Pour the liquid into a centrifuge tube and centrifuge the sample at 1500rpm for 3
Minutes (remember to always counter balance).
 Decant the supernatant, and add flotation solution. Mix well into the sediment. Add
More flotation solution until a meniscus is formed.
 Place a coverslip over the meniscus.
 Return the tube to the centrifuge and centrifuge the sample at 1500rpm for 5
minutes. (remember to counter balance)
Step of fecal floatation by centrifuge. A) Mix 2g of feces with measured ml of
floatation solution in disposable cup. B) Strain the mixture. C) Pour the strained
mixture into 15ml centrifuge tube. D) Fill the tube with floatation solution to form
slight positive meniscus; do not over fill the tube. E) Place the cover slip on the top
of the tube. F) Put the tube in centrifuge; make sure the tube is balanced and adjust
at 1500.r /p for 5 minute. G) Remove the tube and let it stand for 15 minute. H)
Lift the cover slip directly upward and place on microscopic slide.

Note: It is important that you use a centrifuge with swinging buckets, not stationary
buckets. During the centrifugation process, tubes will swing out horizontally and the
coverslip will be held in place.
 After centrifugation, remove the coverslip by lifting straight upwards. Place
the coverslip onto the slide and examine microscopically.
 Sedimentation Techniques
Purpose:
 The fecal sedimentation technique is a qualitative method for detecting trematode eggs
in feces.
Principle:
The majority of trematode eggs are too large and heavy to float reliably in the floatation fluids
normally used for nematode eggs.
They do however sink rapidly to the bottom of a fecal/water suspension and this is the basis of
fecal sedimentation technique.
Procedure
 Take 3 gm of feces in a conical cup and mix with 30ml of water.
 Sieve the mixture through a tea strainer in to a beaker and transfer this in to a centrifuge
tube.
 Centrifuge the filtrate for 3 minutes at 1500 r. p.m
 Discard the supernatant and add a drop of 1% methylene blue drop of the sediment and
examine under the microscope (40X)
 Fasciola eggs appear yellowish and paraphistomum eggs appear grayish with dark
granules in the egg with blue back ground.

 Baermann technique
 In contrast to the previous methods mentioned earlier, the Baermann Technique is used to
Recover parasite larvae, not ova.
Materials;
Baermann apparatus consisting of;
 a funnel
 support structure
 short length of tubing at the end of the funnel
 A clamp at the end of the tubing.
 Gauze or cheesecloth
 Warm water
 Microscope slide
Procedure
 Wrap 5g of feces in gauze or cheesecloth and lay on the support screen inside the
 Funnel of the Baermann apparatus.
 After ensuring the clamp at the end of the tubing is closed, add warm
water until the sample is covered.
 Allow the sample to sit for at least 8 hours (or overnight).
 After the appropriate time has lapsed, loosen the tubing clamp slightly to
withdraw a large drop of liquid onto a slide.
 Add a coverslip to the sample and examine microscopically.
 Fecal culture
 Many nematode eggs are alike athese methods used for determination the types of
infectionnd species such as Haemonchus, Mecistocirrus, Ostertagia, Trichstrongylus,
Cooperia, Bunostomum, and Oesophagostomum cannot be clearly differentiated from
the eggs in faecal samples.
 For these parasites, differentiation can be achieved by the use of fecal cultures.
 They provide a suitable environment for the hatching and development of helminthes
eggs into the infective stage (L3).

Equipment required for fecal culture:
 Fork
 Spoon depressor
 Spatula
 Water
 Jars
 Containers
 Charcoal (It should be added with equal amount with feces and used to make moist feces
to become damp.)
Procedure of fecal culture:
1. Break up collected faeces using a mortar and pestle or stirring device
2. Faeces should be moist and crumbly
3. If faeces are too dry, add water
4. If faeces are too wet, add charcoal until the correct consistence is obtained
5. Transfer the mixture to jar or other containers
6. Leave the culture at room temperature for 14-21 days, at this time all larvae should have
to reached the infected stage
7. If an incubator is available at laboratory, the culture should be placed at 27 oC and left
for 7 -10 days.
8. Add water to culture regularly, every 1-2 days
9. Larvae are identified using baermann technique

3.3. Quantitative Fecal Examination Methods
Quantitative procedure indicate the number of eggs or cyst present in each gram of feces
(severity of infection). Several procedures are used to estimate the numbers of parasite eggs per
gram of feces, including :
1. Mcmaster technique.
2. Stoll΄s technique.
1. McMaster egg counting technique
Purpose;
 The McMaster technique is used for demonstrating and counting helminthes eggs
in fecal samples. It is the most widely employed method for this purpose.
McMaster egg counting technique:
Principle
 The McMaster technique uses a counting chamber which enables a known volume of
faecal suspension (2X0.15ml) to be examined microscopically.
 Thus, if known weight of faeces and a known volume of floatation fluid are used to
prepare the suspension, then the number of eggs per gram of faeces (e.p.g.) can be
calculated.
 The quantities are chosen so that the faecal egg-count can be easily derived by
multiplying the number of eggs under the marked areas a simple conversion factor.
 The McMaster chamber has two compartments, each with a grid etched on to the upper
surface.
 When filled with a suspension of faeces in flotation fluid, much of the debirs will sink
while eggs float to the surface, where they can easily be seen and those under the grid
counted.
Equipment
 Two beakers or plastic containers
 Balance
 Tea strainer, cheesecloth or dental napkin
 Measuring cylinder
 Stirring device (fork, spatula, tongue depressor)
 Pasteur pipettes and rubber teats

 Flotation fluid (choice of solution dependent on species expected to be present and
availability of reagents)
 McMaster counting chamber
 Compound microscope
 If identification is necessary the fecal sample must be cultured to provide L3 larvae for
further examination.
Procedures:
1. Weigh out 3 grams of feces into ajar
2. Add saturated salt solution up to the 45 ml mark (1:15 dilution).
3. Mix contents by glass beads, auto mixer or pestle and mortar.

4. Filter through a mesh sieve screen and collect filtrate.
5. Mix the filtrate well and fill up both the counting chambers of a mac master slide by
using pippet.
6. Count all eggs seen within the ruled areas of both the chambers.
7. The mean of two counts is recommended in calculating egg count

Interpretation:
Volume of each counting chamber is 0.15 ml. 0.15 ml of the solution
of 3 g of faeces in a volume of 45 ml contain, for example x eggs.
 eggs in total volume (45 ml) = x × 45
0.15
 eggs in 1 g of faeces (e.p.g.) = x × 45
0.15 × 3
 Multiply the number of eggs counted by 100 to give egg per gram of feces ( e.p.g)
Advantages
 Egg float up for easy counting
 No interfering fecal material
 Fast and fairly accurate
Dis advantage
 Liable to miss light infestation
 Small quantity of feces examined
 Not good for all fluke egg
 Special slide required
Note: It is not often possible to identify strongly eggs at genus level as the eggs of most
strongylid and trichostrongylid species are similar in appearance and overlapping in size. If
identification is necessary the faecal sample must be cultured to provide L3 larvae for further
examination.

Stoll technique
Procedure
1. Place 5 gm of fresh feces sample in 100 ml graduated measuring cylinder.
2. Add 0.1 N (4%) solution of NaOH (sodium hydroxide) in water up to 75 ml .
3. Shaking the liquid with glass beads.
4. By a graduated pipette, apply 0.15 ml suspension immediately to a microscopic slide and
cover the liquid with a cover slip (22x45) and examine the slide.
It is advisable to check four, preparations the average number of eggs multiplied by 100, equals
the number of eggs per gram feces (EPG,).
Y x 75 x 1 = y x 100
5 0.15

CHAPTER FOUR
4.1 Method of detection of protozoan parasites
4.2 Protozoa Life Stages
Cyst: Infectious form of many protozoan parasites during which they are encapsulated inside a
protective wall; usually found in the feces
Oocyst: Encysted, highly resistant zygotic stage of some sporozoan parasites that may remain
infective for extended periods of time
Trophozoite: Active, motile feeding stage of the flagellate protozoa as well as the postsporozoite
state that is seen in some apicomplexan parasites
Adult: Mature form of protozoan life capable of sexual or asexual reproduction
 To examination of blood parasites, the commonly used techniques are describes below.
A) Wet blood smear
 Collect peripheral blood sample and place immediately a drop of blood on to a
microscopic slide.
 Place a coverslip and examine under 10X objective for localized movement of RBC
which suggested the presence of parasite.
 Trypanosomes and microfilaria may most commonly be suspected.
Figure2. Wet blood smear

Advantage
 Simple and cheap
 If Trypanosomes are found, the diseases are diagnosed on the spot.
Disadvantage
 Unless the animals are brought to the veterinary center, or the blood (an anticoaqulant)
can be taken quickly to the center, a field microscope has to be taken to the herd, as the
parasite lose their mobility after a limited time.
 Limited sensitivity
B) Thin blood smear is a drop of blood that is spread across a large area, thin blood smears
helps doctors discover that species of parasites is causing the infection.
Advantage
 The sensitivity is extremely low, and the main use of thin smears is in fact the specific
identification of trypanosomes found in wet or thick smears.
 But even only a few parasites have been seen in a fresh preparation or a thick film; the
thin smear may be negative.
Geimsa staining
Geimsa stain is commonly used when there is need to examine the blood smears, for the Parasites
but is a good stain for routine examination of blood smear and used to differentiate nuclear and
cyto-plasmic. morphology of the various cells of the blood like platelets, BRCs, WBCs, as well
as the parasites.this stain is the most dependable stain for blood parasites, particularly in thick
blood smears.

Procedure:
1. Take a drop of blood on a grease free clean slide
2. Spread the blood on a slide using cover slip or anothe5r clean slide at an angle of 45o
3. Dry it quickly and fix with methyl alcohol for 2 min
4. Stain with Giemsa diluted 1:10 for 30 min in neutral phosphate buffer.
5. Wash with phosphate buffer at PH 6.8-7.2
6. Allow it to dry by standing up right on the rack
7. Examine under the microscope ( 100X)
Figure 4. Giemsa stained smear
C) Thick Blood Smear
 A thick blood smear examines a slightly greater volume of blood than does a thin blood
smear.
Procedure
 Take a small drop of blood on a clean grease free slide
 Spread it to a size of about 2 cm in such a way that you can read a script.
 Air dry quickly so that it is protected from files
 Dehemoglobinize by gently running distilled water on the smear or by immersing
the smear in distilled water for 5 to 10 minutes.
 Fix with methyl alcohol for 2 minutes
 Stain with Giemsa diluted in buffer distilled water 1:10 for 30 min.
 Wash with buffered distilled water till it assumes bluish purple colors.

 Examine under the microscope ( 40X) and (100X)
Figure 5. Thick blood smear
2. Aspirates
 Aspirates are made from enlarged lymph nodes, skin lesions peritional fluids etc, by
means of a syringe and needle for the detection of parasites.
Procedure
1. Clean and disinfect the part which contains the fluid to be aspirated.
2. Use sterile 5 ml syringe and 18 gauge needle to aspirate the fluid.
3. Draw enough fluid to make a smear and/or wet film
4. Stain the smear with Romanowsky stains and examine the wet film of the aspirate
directly under the microscope
3. Buffy coat method
 The Buffy coat method is a concentration technique used on a small volume of blood.
 When blood is placed in a microhematocrit tube and centrifuged for determining the
packed cell volume (PCV), it separates in to three layers: Plasma, WBC layer (buffy coat)
and RBC layer.
 This technique is quick and may be performed in conjunction with a PCV and total
Protein evaluation.
Material
 Microhematocrit tubes and sealer
 Micrhematocrit centrifuge
 Small file or glass cutter
 Microscpe slide and cover slips

Procedure
1. Fill heparinized or citrated capillary tubes with blood from the animal to be examined.
2. Centrifuge the sample using hematocrit centrifuge
3. Transfer the capillary tube on to a slide
4. Examine the buffy coat in the capillary tube with the microscope (The buffy coat is the
grayish narrow space found between the plasma and the RBC in the capillary seen
flickering at this junction).
5. Cut the capillary tube containing the plasma, the buffy coat and some RBC on a clean
slide.
6. Make a smear if this contain and stain with Giemsa to identify the organisms.
4.3.Other techniques for diagnosis of protozoan infection
 This is used for detection of blood protozoa in the organ and tissues of animals such
as heart, spleen, liver, lung etc.
 Animal inoculation in the diagnosis of protozoa infection such as (Leishmaniasis,
Toxoplasmosis).
 Serological methods for the diagnosis of protozoan infections such as (agglutination,
Immunoflourescence complement fixation, gel diffusion).
i) Impression Organ Smear
Procedures:
 Cut a portion of the organ, e.g. heart, with a pair of scissors.
 The tissue is gently pressed onto a clean slide
 Dry the preparation in the air.
 Stain the impression smear with a preferred staining reagent.
ii) Brain Squash Smear
 A large proportion of specimens submitted from cattle and buffaloes over the years
only gave partial or negative results due to inadequacy of the range of specimens
submitted.
 It is used for detection of blood protozoa in the organs and tissues of animals.

Procedures:
 Remove brain from carcass.
 Cut halfway down into brain-matter using knife, then pull apart forcibly using
knife blade.
 Put on the slide
 Place a pea-sized piece of brain material between two slides, squash and mash
thoroughly.
 Wipe brain material off one slide using the small edge of a second slide.
 Use the first slide to draw out brain matter on the second slide.
 Fix brain smears in methyl alcohol after air drying.
 Stain the smear with a preferred staining reagent.
iii) Intestinal Scrapping
 This technique is conducted for the detection of coccidian oocyst from intestine
samples.
Procedures:
 Cut open the intestine
 Scrape the intestinal lining with a scalpel.
 Put the contents of the scraping onto a slide and add a drop of water.
 Put on the coverslip and examine under 10× objective of a
compound microscope.
Diagnosis of parasitism of urinary system
Collection the urine sample for parasitological examination may be done during normal urination
or catheterization:
• A waxed paper cup 3-5 ml with a lid or other clean container may be used for collection.
• Urine sample should be labeled and refrigerated.
• Methods for diagnosis .
a- Direct method.
b- Urine sedimentation.

CHAPTER FIVE
5.1 Method of detection of external parasites
5.2 Skin scraping method
 It is important to have adequate samples as mites may be absent from a small scraping
from only one area of skin. The hair over the area should therefore be clipped short
and discarded. The area selected should be the moist part on the edge of the lesion.
 Most mites will be on the periphery of active lesions. They will not usually be found
in the thickened dried serous exudate.
 In animals suspected of chorioptic or psoroptic mange, a sharp scalpel should be used
with the blade held at an acute angle, shaving rather than scraping off the outer skin
layer together with hair stumps.
 All specimens should be transferred or scraped directly into a small tube that can be
securely stoppered. The usual samples received for examination is a skin scraping
with some hair or wool.
Procedure:
1. Inspect under a low power of microscope. Mites may be visible.
2. Clip free and remove excess hair or wool.
3. Scrape some material onto a slide, mix a drop of 10% potassium hydroxide, warm and place a
coverslip over the material.
4. Allow the preparation to clear for 5 to 10 min and then examine under medium and high
power.
5. If no mite is seen, place the entire scraping (up to 5 g) in a boiling tube with about 10 ml of
10% potassium hydroxide.
6. Stand the tube in a beaker of water and gradually bring to the boil.
7. When all the crusts and hair have digested after 2 to 5 min in the water bath, allow the liquid in
the tube to cool, and then centrifuge to deposit the mites at 2,000 rpm for 2 min. A longer period
will be required for smaller species. Avoid prolonged boiling in the caustic solution since the
mites will eventually disintegrate.

8. Quickly decant the supernatant and then pipette the deposits off onto microscope slides for
examination.
9. Make permanent mounts (e.g. for notifiable diseases such as sheep scab and parasitic mange of
equines) by mixing the deposit with water, spinning down and decanting the dilute caustic
solution, then adding 0.5 ml of glycerine jelly to the deposit.
10. Melt the jelly in a water bath, pour into a centrifuge and mix with the deposit by rolling the
tube between the hands. With the mixture still warm, pour the fluid onto the slide and cover with
a circular coverslip, 20 ml diameter. The jelly sets rapidly. Now examine the slide almost
immediately under a low power microscope on a mechanical stage. Mark any mites or suspect
mites, ring the mounts off and submit to the CVL for confirmation of the identification.
11. If no mites are seen, treat the wool or hair until stages 5 to 8. Demodex is easily made
transparent by boiling in caustic solution and if present, should be detected in stages 1 to 4.
Where samples from sheep are being examined, a flotation technique can also be used but only if
there is an excessive amount of wool.
12. Immerse the wool in 20% potassium hydroxide in a test tube.
13. Incubate for 3 to 4 hours at 37 °C and centrifuge for 10 min at 2,000 rpm.
14. Tip off the supernatant.
15. Re-suspend in Sheather’s sugar (454 g sucrose, 355 ml tap water, 6 ml 10% formalin).
16. Centrifuge for 10 min at 800 rpm and any mites present will float to the surface. Gently
touch the surface of the sugar solution with the flat-ended glass rod and place the drop of
fluid obtained onto a slide. Mount with a coverslip.
Figure 1. Skin scraping laboratory diagnosis

5.3 Techniques for preservation and mounting of tick, fleas and lice
The correct method of preservation, mounting and labelling of insects is very important for
proper identification and their usefulness for teaching purposes. Specimens of insects can be
mounted in a chloral hydrate medium.
Dehydration is not necessary before mounting in this medium. Small insects like lice can be
mounted while alive. Some large specimens like ticks and fleas require macerating in a
caustic solution before dehydration, clearing and mounting.
All slides made with Berlese’s fluid and other chloral hydrate media must be scaled after drying
by ringing the coverslips with a waterproof substance such as Canada balsam, nail varnish,
Glyceel or any other proprietary substances made for this purpose.
Simple rules in making a mounted slide
1. Use a coverslip of suitable size for the specimen.
2. Place an appropriate amount of mountant on the centre of the slide.
3. Arrange the specimen with head away and tail towards you except for fleas which are
mounted lengthwise.
4. Only one specimen is to be mounted on a slide unless the male and female of the same
species are required or a number of minute forms such as mites.
5. Affix the label with complete data on the specimen at the right side of the slide with the
specimen in the correct viewing position i.e. upside down for a compound microscope
and with head up for a stereoscopic microscope.
Method of macerating with caustic solution
1. Puncture or make nicks in the body of the specimen to allow free penetration of
solution.
2. Boil in 10% potassium or sodium hydroxide solution for about 5 minutes depending on
specimens, or leave the specimen in this solution overnight.
3. Remove from caustic solution into water. Using a blunt instrument and with a gentle
pressure, squeeze out liquefied contents of the abdomen.
4. Wash specimens well in distilled water containing a few drops of glacial acetic acid.

5. Transfer to fresh distilled water.
6. Dehydrate in ascending grades of alcohol e.g. 50%, 70%, 90%. Absolute 1 then
Absolute 2 for 10 minutes in each one.
7. Clear in xylol and mount in DePeX, Canada balsam or any other suitable mountant.
Note: Do not leave specimens too long in xylene as they get hardened quickly and mounting
may be difficult.
Acarine (ticks)
 Over sixty different species are found in Eastern Africa but many of these appear to be of
little or no economic importance. There are two well defined families of ticks, the
Ixodidae or hard ticks and the Argasidae of soft ticks, and the two groups differ from
each other markedly in appearance habits and life histories.
 Members of the family Ixodidae have a hard dorsal shield which covers the entire upper
surface of the male and a relatively small area just behind the head of the female nymph or
larvae.
 This dorsal shield or sctum bears a pattern which is characterstic for each species of tick.
Sometimes the scutum is uniform in colour and the pattern is only made up of the pits
grooves and minute punctuations on it, but in some ticks a colour pattern is also present.
 Even adult ticks usually have to be examining with a stereo microscope or a hand lens
which gives a magnific ation of 10X or more before they can be identified accurately
Larvae and numphae have to be magnified considerably for examination
Preservation: 70% alcohol or 10% formalin
Note: Do not add glycerine in alcohol as this gives the specimens a shine. To preserve the natural
colour of ticks, drop live specimens into a saturated solution of chloroform in 10% formalin.
Transfer specimens to 70% alcohol after a month.
Mounting procedure: it has the same precudure with the above method.

Mites
Examination for mites
Procedure:
 place a drop of mineral oil a slide
 Clean the scalpel blade by wiping it with paper.
 Dip the clean scalpel blade in to the drop of oil on the microscope slide
 Pick up a fold of the patients skin at the edge of the suspected area pinching it firmly
between the thumb and forefinger. Scrape the crest of the fold several times in the same
direction with the oily scalpel. Scrapings will adhere to the blade.
 Stop scraping when a small amount of blood appears.
 Transfer the scraping from the scalpel blade in to the drop of oil on the slide, using a slight
rotary motion.
 Apply a cover grass to the scraping on the slide.
 Examine the preparation under low power (10X) in a methodical manner so that all
portions of the cover glass area seen.
Boil the skin scraping in 10% KOH solution to facilitate identification
Preservation: 70% alcohol with 5% glycerine added to prevent drying out of specimen in
permanent storage.
Mounting: No maceration required. Mount directly in Berlese’s fluid.
Anoplura (Lice)
 Lice may be detected by the presence of either the eggs (nits) which are found cemented to
the host’s hair and adults in the animal’s hair coats. If the hair is pushed in the direction
 Opposite to its growth, lice may be seen moving about and can be caught with a forceps or
the fingers.
 If the light is poor a portion of the hair may be pulled or clipped and sealed in a container
for later examination. This is best done by spreading the hair and debris on a while paper
under a strong light for close observation with a hand lense.
Preservation: 70% alcohol
Mounting procedure: Follow method as described above

Siphonaptera (Fleas)
 These are most easily seen on the less hair parts of the body, but may be found pusing the
hair against its pattern of growth. Fleas may be recovered from an animal by combing or
brushing it over a whikle cloth, especially after dusting it with an insecticide.
 A second method is to enclose the posterior of a small in a plastic bag which contains an
ether soaked cotton pad. The sweepings from the area around the bed of flea infested dog
or cat will usually field flea larvae which are small (3-5mm). These are caterpillar-like
creatures. The browen pupae may also be found.
Procedure: Follow method as described above
Preservation: 70% with 2% glycerine added to prevent drying out.
Note: For rapid clearing of specimens, lactophenol is recommended. After examination,
specimens can be returned into the preservation.
Mounting procedures for Culicoides
1. Place the Culicoides in absolute alcohol for 30 minutes (dehydration).
2. Clear the Culicoides in phenol solution (phenol crystals dissolve in a small amount of absolute
alcohol) overnight.
3.Remove the Culicoides from the phenol solution and place it in a drop of phenol solution on a
microscope slide.
4. Examine the specimen under the dissecting microscope, detach the head and one wing from the
specimen.
5. Place a coverslip over the whole specimen; place a small drop of Canada balsam at the edge of
the coverslip so that it could seep into the specimen.
Mounting procedures for mites
1. Place the mites in lactophenol and leave it overnight.
2. Remove the mites from the lactophenol and rinse in water until the cloudy interface of
lactophenol and water disappears.
3. Place a drop of Hoyer’s medium or other suitable aqueous mountant in the center of a
clean microscope slide.
4. Place the mite in the mountant so that it is at the bottom of the droplet and arrange it on
vertical axis, and place a coverslip over the specimen.

5. Ring the edge of the coverslip with a water-proof substance such as nail varnish (Cutex).
Trichodectes canis (Louse) Ixodes sp. (Tick)

REFERENCES
1. Manual of veterinary parasitological laboratory techniques. (1986). 3rd
edition. Ministry of
Agriculture, Fisheries and Food. HMSO Publications Centre. London
2. Christopher R., Chandrawathani P. and Cheah T.S. (1992). Manual on parasitology. Jabatan
Perkhidmatan Haiwan (in-house publication).
3. Gelma B. (2010). Veterinary parasitology manual for veterinarian technicians. Ambo
University
4. Hendrix, Charles M. and Robinson, Ed. 2012. Diagnostic Parasitology for
Veterinary Technicians, 4th ed. St. Louis: Mosby-Elsevier.
5. Hendrix, Charles M. and Sirois, Margi. 2007. Laboratory Procedures for
Veterinary Technicians, 5th ed. St. Louis: Mosby-Elsevier.
McCurnin, Dennis M. and Bassert, Joanna M. 2006. Clinical Textbook for Veterinary
Technicians, 6th ed. St. Louis: Saunders-Elsevier.
6. Bassert, Joanna M. and McCurnin, Dennis M. 2010. Clinical Textbook for Veterinary
Technicians, 7th ed. St. Louis: Saunders-Elsevier.
7. National Veterinary Drug Formulary (2013). Second Edition Department of Livestock,
Ministry of Agriculture & Forests.Website available at http://www.
ncah.gov.bt/Downloads/File_3.pdf.

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