Diagnostic Parasitology
THE COLLECTION AND
PRESERVATION OF
PARASITIC MATERIAL FOR
IDENTIFICATION
Dr. Richard A.
French
Connecticut Veterinary Medical
Diagnostic Laboratory
Department of Pathobiology
University of Connecticut,
Storrs, CT 06269-3203
An enhanced
parasitology diagnostic service will be offered by the CVMDL of the
University of Connecticut, Department of Pathobiology. Specimens
from domesticated animals as well as wildlife, aquatic and avian
species are welcome. The service will be headed by Dr. Richard A.
French and is funded by revenues from diagnostic tests and services.
Dr. French has a M.S. in veterinary parasitology, Ph.D. in
veterinary pathology and has extensive training in diagnostic
parasitology. He has published articles on parasitic diseases
including those on parasitic disease of domestic animals, fish and
birds. Dr. French serves in the department as a diagnostic
pathologist, researcher and instructor.
The
support staff and faculty within the department includes
professional laboratory technicians, certified veterinary
technicians, veterinary pathology residents and board certified
diplomates of the American College of Veterinary Pathologists. The
Department of Pathobiology has a long and excellent reputation as a
veterinary diagnostic laboratory serving Connecticut and the New
England states. The Pathobiology Department also is the location of
the Northeastern Research Center for Wildlife Diseases. The
diagnostic laboratory is accredited by USDA/APHIS and is routinely
inspected by the AAVLD.
The
information provided in this publication will aid you in the
collection and preservation of parasitic material for evaluation. In
addition, the information offers techniques you can use in your
practice to aid in the diagnosis of parasitic diseases. We also
provide you with contacts here at the University of Connecticut and
the opportunity to visit our web page, ask questions and/or provide
comments and suggestions.
. Contacts:
Richard
A. French, DVM, MS, PhD
Veterinary Pathologist
Richard.French@uconn.edu
860-486-3738 (CVMDL)
860-486-3936 (fax)
. World Wide Web
site:
Pathobiology
(http://www.lib.uconn.edu/canr/patho/)
Parasitology Diagnostic
Testing Service
(http://www.lib.uconn.edu/canr/patho/parasit.htm)
II.
GENERAL INSTRUCTIONS:
. A complete
signalment and history should be sent with every specimen.
Species and age of the host, clinical signs, treatments
administered (deworming programs), necropsy lesions, specific
location of the parasite, time and manner of onset, number of
animals involved, type of management used, and with wildlife or
zoological specimens, the normal habitat or range of the animal.
. Fecal samples, ideally, should be collected
before the animal has been treated with Kaopectate®,
Pepto-Bismol®, mineral oil, or barium compounds.
These substances in the feces can inhibit detection of
parasites.
. Some parasites (e.g. ectoparasites) leave the host very
soon after its death, and some migrate within the host.
Regardless of how the hosts are handled, the parasites will
recognize and react to any change in their environment.
Some parasites, especially intestinal helminths, will be
eliminated from the host during the early days, or even hours, of
captivity in wildlife. Other
parasites, especially ectoparasites, will multiply and/or transfer
from one host individual or even species, to another. The latter is especially important in zoological specimens
and intermixed wildlife species (e.g. aberrant hosts).
. Freshly collected, living parasites always make
more satisfactory study specimens than those that have undergone
even slight maceration. Parasites
survive longest in cold-blooded hosts and in cyst stages.
. Thorough examination in the proper sequence is important
to prevent losing some parasites while looking for others.
In a freshly killed animal the preferred sequence is (1)
blood, (2) gills and/or skin, (3) internal organs and (4)
musculature.
. Fecal samples
should be submitted in a watertight container.
If the samples can reach the destination (diagnostic
laboratory) in less than 72 hours, no preservatives need to be used.
However some eggs and oocysts may Embryonate/sporulate or
hatch during this time unless air is excluded from the container.
. The recommended method of fecal
fixation is to place the fecal sample in 10-15 volumes of 10%
formalin (the same concentration as for submitting histopathology
samples).
. If coccidia are suspected, place the feces in 2.5% potassium dichromate
solution. This allows
for future sporulation of the coccidia, which is often required for
species identification.
. Fecal larvae
are best recovered with special procedures such as the Baermann
technique (e.g. Strongyloides
spp. or lungworm larvae). Fresh
fecal samples without preservatives or fixatives are required for
the Baermann technique.
. If examination for intestinal
protozoa other than coccidia (particularly Giardia
spp., trichomonads and amoebae) is required, the sample should be
fixed and shipped using sodium acetate-acetic acid-formalin (SAF)
fixative. Mix one
volume of feces with at least three volumes of SAF fixative.
. Blood smears should be made on clean glass slides.
Even pre-cleaned slides should be cleaned thoroughly with a
detergent soap and rinsed or washed in a 1:1 mixture of 95% EtOH and
100% acetone. Both
stained and unstained slides should be submitted.
. Blood samples can and if available should also be
submitted in EDTA. EDTA
blood samples should be refrigerated.
. Heparin should be avoided for mailed samples, as its
anticoagulant activity is short-lived (4-6 hours).
Also, heparin is contraindicated if parasites such as Haemobartonella
spp are suspected.
. Blood specimens for examination for microfilaria should
be fixed in at least 10 volumes of 2% formalin.
Blood samples can also be submitted in EDTA.
V.
ECTOPARASITES/ARTHROPODS:
. Ectoparasites should be fixed and submitted in 70%
alcohol (Exception:
Testing ticks for Borrelia
sp.).
. Fleas and lice
can be collected by placing the freshly dead host in a tightly
closed plastic or paper bag. A
piece of cotton soaked with ether of chloroform added the bag with
mammalian or avian hosts can aid in the recovery of specimens.
Wash out the bag and inspect the washings with a dissecting
scope.
. Furry or feathered
animals may be scrubbed or carefully combed with a small,
fine-bristled brush (toothbrush) which is periodically dipped into
soapy water. (Note: Feather mites do not leave the host
postmortem).
. Mites often
require a skin scraping. Skin
scrapings may be submitted on slides, but must have a sealed
coverslip (seal coverslip with clear nail polish or similar
material). Be sure
there is enough oil under the coverslip so that the material will
not dry out in transit. Send
more than one slide if possible.
Skin scrapings should be taken from the edges of lesions and
try to squeeze the skin while scraping to bring the mites to the
surface. A good, deep
scraping is indicated by the presence of red cells.
. Ticks are
usually embedded in the skin. To
pull from a living or freshly killed host, use fine pointed tweezers
to grasp the tick around its mouthparts, as close to the skin as
possible. Gently pull
the tick straight out with steady pressure, being careful not to
break off the tick's mouthparts.
Avoid twisting the tick.
[Note: Testing ticks for Borrelia
burgdoerferi (the spirochete that causes Lyme
Disease) employing the fluorescent antibody (FA), darkfield
microscopy, or IFA tests requires live or freshly dead specimens.
Dehydrated ticks can be tested employing the PCR method. Place the tick in a sealed container with a water dampened
cotton ball or tissue (do not
place in alcohol). Avoid the use of products such as petroleum jelly or nail
polish remover on the tick prior to removal.
Specimens can be refrigerated.]
. Aquatic hosts
harbor parasites ranging from protozoans to monogeans, leeches,
copopods and isopods and glocchidia.
These parasites can, in general be fixed in 70% EtOH.
Water samples and tank sediment can also aid in the
identification of parasites.
. Animal bedding
and nest materials may be used to collect fleas, lice, and mites
(especially important in burrowing animal and birds). Samples should be submitted in a sealed container.
VI.
ENDOPARASITES/HELMINTHS:
A. PROTOZOANS AND MESOZOANS:
. Mesozoans
are ciliated animals that parasitize the excretory or reproductive
systems of marine invertebrates.
They are uncommon. To
collect them, make tissue smears of the infected organs and treat
like protozoans.
. Intestinal
protozoan forms in fresh fecal samples should be examined within
24 hours (48 hours if samples are tightly covered to maintain
moisture and refrigerated).
. Direct wet smear technique is the first step in the
microscopic examination of a fecal sample.
Examine the preparation at low magnification and very low
light intensity (high contrast) to find amebas,
flagellates, and ciliates.
. Note: Trophozoites are characteristic of loose feces/stools; cysts
of formed stools (ex: Giardia sp.)
. Iodine stained (Lugol's iodine) preparations can be
examined by placing a drop of Lugol's iodine with fecal material
on a slide; add a coverslip. The
cysts will gradually become tinged with yellow, and the nuclei
become dark brown.
. Fecal samples for submission to a diagnostic laboratory
should be fixed in SAF fixative (see "fecal samples," above).
. Protozoans/protozoan
cysts in muscle, internal organs, and skin samples can be
collected by smear techniques, they should be teased from the host;
or the host tissue should be fixed with parasites in situ.
B. NEMATODES:
. Nematodes should be fixed in 70% EtOH but can also be
fixed in 10% formalin. Most
nematodes from domestic animals can be easily identified, but if the
nematodes are from exotic or wild animals, special fixation is
needed. The best method
is to heat 70% EtOH to approximately 60oC,
drop the parasites (live) into the hot alcohol one by one, and
remove them as soon as they are fixed in an extended position.
They should be placed in fresh 70% EtOH for shipping or
collection.
. Nematodes embedded in tissues can be teased out or
submitted fixed in situ using 10% formalin.
Note, nematodes in fresh tissues will sometimes migrate and
leave tissue sections immersed in formalin (e.g. nematodes can live
for several minutes or hours in formalin).
Therefore, formalin should not be changed before submission
or should be examined for parasite specimens.
. Trichinella sp.
requires tissue digestion procedures and samples should be submitted
without fixatives and should be refrigerated.
C. TREMATODES AND
CESTODES:
. Trematodes and cestodes that are recovered at necropsy or
passed in the feces require relaxation before fixation.
Chilling the worms, either in saline or tap water overnight
in the refrigerator, relaxes them with the least handling.
Another method is to place them into 5-10% EtOH at room
temperature. The
relaxed worms can be fixed in 10% formalin or preferably alcohol-formalin-acetic
acid (AFA) fixative.
. Trematode
larvae and cestode larval stages (cysticercus larvae) can be fixed
by first relaxing larvae in hot water or AFA (about 50oC).
Larvae are then fixed in cold AFA or 70% EtOH.
. Note: In the collecting of cestodes the
preservation of the scolex is
critical. In
freshly dead hosts the scolex is embedded in the tissue.
Deep scraping of the mucosa can free the scolex.
Alternatively, excise a small portion of intestinal wall in
which the scolex is embedded, and chill along with the worm in
saline. This will
induce the worm to loosen its hold.
. Note: All domesticated, exotic or wild animals
should be handled with care to prevent the transmission of zoonotic
disease (viral, bacterial, mycotic or parasitic) and to prevent the
transmission of disease to other animals or species. It should also be noted that some parasites might carry or
transmit disease agents [e.g. Ixodes
dammini - Borrelia burgdorferi, fleas - Bubonic plague (Yersinia pestis), ticks
- Rocky Mountain Spotted Fever (Rickettsia
rickettsia), etc.].
. Most zoonotic diseases are reportable when seen in humans
and/or livestock. The state health department will have guidelines specific for
that state.
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