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Part I
Methods to study primate–parasite
interactions
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978-0-521-87246-1 - Primate Parasite Ecology: The Dynamics and Study of Host-Parasite
Relationships
Edited by Michael A. Huffman and Colin A. Chapman
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1
Collection methods and diagnostic
procedures for primate parasitology
ellis c. greiner and antoinette m c intosh
Photograph by Jessica Rothman
Introduction
A great deal of energy has been expended on studying parasites of free-ranging
primates. Many of these studies have been confined to fecal surveys as it is
both difficult and not practical to gain knowledge of parasites in these hosts by
using the older and normal method of studying the parasites that are recovered at
Primate Parasite Ecology. The Dynamics and Study of Host–Parasite Relationships, ed. Michael
A. Huffman and Colin A. Chapman. Published by Cambridge University Press.
C Cambridge University Press 2009.
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978-0-521-87246-1 - Primate Parasite Ecology: The Dynamics and Study of Host-Parasite
Relationships
Edited by Michael A. Huffman and Colin A. Chapman
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Primate Parasite Ecology
necropsy. Thus, this chapter is designed to aid researchers in being as productive
and efficient as possible in gathering information that is most useful for the
conservation of these interesting creatures. The type of information obtainable
depends upon the restrictions placed on the collection and preservation of
the samples to be examined. If the primates are darted and anesthetized, then
feces, blood, and ectoparasites may be collected. If the primates are not alive,
then parasites and specimens could be recovered from various organs and
tissues at necropsy. We need to be opportunistic with such endeavors as we
cannot make the contributions with only feces that we can from recovery of
parasites at necropsy. The eggs or worms found at necropsy have not been
matched to the adult worm identifications in most cases as they have been with
domestic animals species. Feces, therefore, is the only practical sample that can
be collected and examined with reference to parasites in free-ranging primates
for which capture is not an option. Some general references that might be
useful for aiding in the detection and identification of primate parasites include
those by Melvin & Brooke (1974), Anon. (1979), Zajac & Conboy (2006), and
Garcia (2007). A flow chart of potential diagnostic procedures is depicted in
Figure 1.1.
Collection of fecal and blood samples precautions
Because primate blood and feces are potential sources of organisms infectious
to humans, personal safety of individuals collecting these samples in the field
is imperative and can be achieved by following “universal precautions.” “Universal precautions,” as defined by the Centers for Disease Control (CDC), “are
a set of precautions designed to prevent transmission of human immunodeficiency virus (HIV), hepatitis B virus (HBV), and other bloodborne pathogens
when providing first aid or healthcare. Under universal precautions, blood and
certain body fluids of all patients are considered potentially infectious for HIV,
HBV and other bloodborne pathogens” (CDC website).
Basically, this means wearing gloves and protective clothing, such as disposable aprons, thorough hand washing, proper disposal of needles, scalpels,
syringes and any items contaminated with blood, mucous, feces, and other body
fluids in appropriate leak-proof and puncture-resistant containers for disposal.
Refer to OSHA (Occupational Safety and Health Act) regulations for specifics
on “universal precautions” and recommendations and procedures to follow in
case of animal bites, contaminated needle sticks, or wounds or cuts becoming
exposed to any of these fluids.
Some points to consider in maintaining sample integrity for optimal parasite
detection are:
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- Piroplasms
- Mites
SKIN SCRAPING
- Lice
- Mites
- Ticks
PICK ECTOPARASITES
- Lice
- Mites
BRUSH FUR
Figure 1.1. Flow chart of diagnostic procedures.
GIARDIA ELISA (IDEXX SNAP TM TEST)
COPROCULTURE
- Strongylate nematode larvae
- Organ slide impressions
Prepare blood
smears and
impression smears
of solid organs
Mount lice and mites in
Hoyer’s medium on
microscope slides or
submit to taxonomist
Ticks and lice
examine wet
Fix arthropod in 70%
ethanol
ARTHROPODS
idia
Place in glacial acetic acid, store
in glycerin alcohol, temporarily
mount in lactophenol
Relax flatworms in tap water
for a few hours then fix in
AFA
Allow acanthocephalans to relax
overnight in tap water to evert the
proboscis, fix in AFA
HELMINTHS
- Fecal smear for
Cr
- Sedimentation
yptospor
- Flotation
FIX IN FORMALIN
FIX IN PVA
- Perform Trichrome
stain for amoebae
and gut flagellates
FECAL EXAM
POSTMORTEM EXAM
Fix tissue in formalin
for histological
exams
NECROPSY
PARASITE RECOVERY
- Frozen specimens
- Frozen feces
- Histological sections
BIOPSY
- Drop of blood on filter
paper or FTA elute card
MOLECULAR DIAGNOSTICS
INTEGUMENT EXAM
- Trypanosomes
- Microfilariae
- Malarial parasites
GEIMSA STAIN
BLOOD SMEAR
BAERMANN PROCEDURE
- Nematode larvae
- Adult pinworms
FECAL SMEAR
- Acid fast stain for
Cr
oocysts
- Protozoan cysts
yptosporlarvae
- Nematode
- Coccidian oocysts
- Acanthocephalan eggs
- Cestode eggs
FLOTATION
- Nematode eggs
- All parasites listed
under flotation
- Fluke eggs
SEDIMENTATION
- Motile protozoa
idia
DIRECT SMEAR
FECAL EXAMS
ANTEMORTEM EXAM
PARASITE DIAGNOSIS
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978-0-521-87246-1 - Primate Parasite Ecology: The Dynamics and Study of Host-Parasite
Relationships
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Primate Parasite Ecology
1. Avoid contamination of sample with water or urine “because water
can be contaminated with free-living organisms that can be mistaken
for (human) parasites” (Garcia, 2007).
2. Loss of motility of protozoa may occur if sample is contaminated by
urine.
3. Time frames for examination or fixation (recommended by Garcia,
2007):
a. Liquid feces: Examine or preserve within 30 minutes of passage
(trophozoites).
b. Soft feces: Examine or preserve within 1 hour of passage (trophozoites and cysts).
c. Formed feces: Examine or preserve within 24 hours of passage
(cysts).
4. If quantity allows for attempt at larval culture, do not refrigerate this
portion of sample.
Fecal exams
Examination of feces is useful when attempting to gain information of the
parasite fauna in a given primate. There are a number of procedures that could
be used to detect the different groups of parasites. The number of exams
that can be performed on one sample depends somewhat on the volume of
feces obtained from each host. If at least 2 g of feces can be obtained, then
the following procedures could be done: fecal flotation, fecal sedimentation,
direct smear, a trichrome or iron hematoxylin stain for protozoa from fecal
smears prepared after appropriate fixation, acid-fast stain for Cryptosporidium
sp. on unfixed smears, and Baermann procedures for larval or tiny nematode
recovery. How the samples are collected and fixed will depend on whether
the examinations will be conducted in the field or sent away to a reference
laboratory for processing or taken back into a laboratory setting. Because of
the potential for the presence of infectious organisms, sample containers should
be placed in plastic leak-proof bags before transport and the shipping container
needs to be able to withstand the rigors of postal systems.
Solutions for fecal procedures
Saturated sodium nitrate
Nitrate of soda (commercial grade fertilizer) 5 lb
Hot tap water 1 gallon
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Relationships
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Collection methods and diagnostic procedures
7
OR
Sodium nitrate 400 g
Hot water 1000 ml
Stir ingredients in appropriate-sized container until dissolved. Specific gravity should be 1.20–1.25. The actual specific gravity should be measured with a
hydrometer and recorded on the container when each batch of flotation solution
is prepared.
Sheather’s sugar
Granulated sugar 454 g (1 lb)
Tap water 355 ml (12 fluid oz)
Liquefied phenol crystals or formaldehyde 6.7 ml (a preservative and
mold inhibitor or simply store in the refrigerator)
Dissolve sugar in hot water by stirring over a heat plate. After sugar is
dissolved and the solution has cooled to room temperature, add liquefied phenol
(or formaldehyde) or store in the refrigerator. The specific gravity should be 1.27
and this should be checked with a hydrometer and recorded on the container
when each batch of flotation medium is prepared.
Detergent solution for simple sedimentation
Add 5 ml of liquid dish detergent into 4 liters of tap water. Avoid formation of
bubbles when in use.
Zinc sulfate for detection of Giardia sp. cysts
ZnSO4 · 7H2 O 336 g
Distilled water 1000 ml
Mix and adjust solution to specific gravity of 1.18. While some parasitologists
think they must use zinc sulfate in order to float cysts of this flagellate, sodium
nitrate can be used with good results.
0.85% sodium chloride (normal saline)
NaCl 8.50 g
Distilled water 1000 ml
This is used only for direct smear preparations looking for motility of gut
inhabiting protozoa.
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978-0-521-87246-1 - Primate Parasite Ecology: The Dynamics and Study of Host-Parasite
Relationships
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Primate Parasite Ecology
Saturated sodium chloride
Water 1000 ml
NaCl table grade
Dissolve NaCl in water by stirring until there is a salt residue in the bottom
of the container that will not go into solution.
Procedures
Fecal flotation
Simple sodium nitrate flotation: for recovery of nematode and tapeworm eggs,
coccidian oocysts, mites, and larval nematodes.
Materials:
Saturated sodium nitrate or ∗∗ FecasolTM specific gravity 1.2
15 ml conical centrifuge tubes and tube rack to hold them vertically
(or FecalyzerTM )
Wooden tongue depressors or applicator sticks
50 ml paper cups or small disposable plastic containers
Glass microscope slides and 22 × 22 mm coverslips
Paper towels
Compound microscope.
Procedure using 15 ml centrifuge tubes:
1. Conduct steps 2 through 6 over paper towels.
2. Place 1–3 g of feces into the 50 ml container.
3. Add 15–20 ml sodium nitrate solution and mix well, thoroughly breaking up feces.
4. Place one layer of cheesecloth or gauze over the top of the container. Pour and strain the mixture into a 15 ml conical centrifuge tube
(Figure 1.2) until you have formed a slight positive meniscus at the
top. Set cover slip on top. The liquid should just touch the coverslip,
but not spill over. Many eggs that have already floated to the top will
be lost if you overfill the tube and spillage occurs when the coverslip
is placed on the tube.
5. Let stand vertically for a minimum of 10 minutes (Figure 1.3).
6. Gently lift coverslip straight up without losing any fluid, and place it
on a glass slide.
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Relationships
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Figure 1.2. Setting up a flotation.
Figure 1.3. Flotation running.
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Primate Parasite Ecology
Figure 1.4. How to scan a slide systematically.
Procedure using FecalyzerTM (strictly follow the directions that
come with the FecalyzerTM )
The following is a basic summary of the procedure.
This method assumes you have at least 2–4 g of feces to work with or you
will be picking up any freshly dropped fecal samples. Using the apparatus, pick
up a portion of the fecal sample in the bottom of the white center portion of
the apparatus, and place it in the outer container that is half full of FecasolTM
or sodium nitrate solution. Mix the sample well by turning and twisting the
internal tube in the solution. Then press the white central portion down with
sufficient pressure to seal the edges and fill the tube to the top forming a
slight positive meniscus and place the coverslip on top being careful to avoid
over filling and spilling. Let stand a minimum of 10 minutes (Figure 1.3).
Remove coverslip by picking it straight up and place it on a glass microscope
slide.
Examine the entire area under the coverslip using a low power (10×) objective and adjust light intensity to give good contrast. Begin at one corner of the
coverslip and scan cross to the opposite edge, then move down one field of view
and scan back across to the opposite edge. Continue this method until the entire
area under the coverslip is observed (Figure 1.4). If any suspicious objects
are encountered or you want to observe details of eggs, cysts, or oocysts for
identification and confirmation, move to the high-dry 40× objective (increase
light slightly) and observe details and take measurements. This higher magnification is required to observe the characteristics of smaller protozoan cysts
and oocysts in order to confirm their presence. Do not allow the slide to stand
too long because the salt will begin to crystallize on the edges of the coverslip
making the observation of any eggs or cysts in those areas difficult and may
affect an accurate diagnosis if their presence is overlooked.
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978-0-521-87246-1 - Primate Parasite Ecology: The Dynamics and Study of Host-Parasite
Relationships
Edited by Michael A. Huffman and Colin A. Chapman
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Collection methods and diagnostic procedures
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Fecal sedimentation
This method is used primarily to recover trematode and acanthocephalan eggs,
but other helminth eggs, nematode larvae, mites, protozoan cysts, and oocysts
will sediment as well.
Materials:
Detergent solution, cheesecloth, small plastic beaker, plastic 50 ml conical
centrifuge tubes, appropriate size tube rack to hold tubes vertically, pipettes,
glass slides, and coverslips. (Optional: 5% methyl green or 0.1% methylene blue
stains if available), compound microscope, dissecting microscope if available.
Procedure:
1. In a small beaker or cup, break up and mix 2–4 g of feces in a small
amount of sedimentation solution and stir until uniform mixture is
achieved (no large chunks). Continue adding detergent solution to an
approximate volume of 50 ml avoiding the formation of bubbles.
2. Place one layer of cheesecloth over the beaker and pour mixture
through cheesecloth into the 50 ml tube. If the tube is not full at this
point, add more detergent solution to mixture by pouring it through
the cheesecloth that is still covering the cup (the material trapped in
the cheesecloth will be rinsed back into cup and possibly release more
eggs into the solution), swirl to mix then continue by pouring through
the cheesecloth from the cup into the tube until it is full or contains
approximately 50 ml of filtered mixture.
3. Place the filled tube in rack and allow sedimentation to proceed for 5
minutes in a vertical position.
4. After a sedimentation time of 10 minutes, decant supernatant carefully
and return to vertical position in one movement before any sediment
reaches the lip of the tube or aspirate approximately 75% of the
supernatant with a pipette to avoid mixing or losing the sediment at
the bottom of the tube which contains the eggs.
5. Resuspend the sediment collected at the bottom of the tube by swirling
or carefully tapping the bottom of the tube and refill tube with tap water
by letting the water gently flow down the inside wall of the tube while
avoiding the formation of bubbles.
Note: Avoid formation of bubbles when adding water because eggs
can be trapped in or adhere to detergent bubbles and will not
descend during the next step and will be lost in the decanting
process.
6. Allow to stand and sediment vertically for 5 minutes and decant as in
step 4.
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