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NATURALLY OCCURRING Sarcocystis INFECTION IN DOMESTIC CATS
KAREN D. GILLIS
A THESIS PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE
UNIVERSITY OF FLORIDA
First and definitely foremost, I would like to recognize Dr. Rob MacKay, my boss
and mentor, for his support of both my career and graduate education. I am deeply
grateful for the opportunities Rob has given me in order that I might finally complete my
degree. His high standards and careful planning have allowed this, and other research
projects we've worked on, to be successful. I've worked for Rob since 1996 and have
absolutely enjoyed the experience; I hope to continue on as his technician for some time
I owe thanks to many people involved with this thesis project. Without their help, it
could not have been done. The concept and original design of this project came from Dr.
Rob MacKay, obviously a very important contribution. Although I still strive for
succinctness and clarity, Rob has helped me to become a better communicator; his
language skills are unmatched. Charles Yowell endured many, many interruptions, silly
questions, and requests for help. His patience and knowledge of molecular biology are
admirable. Dr. Julie Levy allowed us to piggy back on her feline heartworm project, and
therefore made the completion of this study that much easier. Dr. Levy was also a great
resource for all things feline. Dr. Ellis Greiner has prodigious expertise in the areas of
parasitology and paternal advice alike. Both of these skills were called upon for this study
and in many instances prior. Dr. John Dame helps me to see "the big picture" and put my
findings into a larger context. This is important, as it's so easy to get bogged down with
the small details. Dr. Andy Cheadle provided assistance with parasitology procedures and
microscopic examinations. Now that he's working in the scientific writing field, I tested
out his editorial skills as well. Dr. Jorge Hernandez was involved at the very beginning of
the project and consulted on its design. Tim Massey, Dwight Schroedter, and Brent
Mayer from the Neogen Corporation developed the methodology for immunoblotting cat
sera and performed the blotting as well. Glenda Eldred provided excellent technical help
and advice for histopathology. Veterinary students Jennifer Hooks, Mike Pegelow, and
Larissa Tavares performed blood collection and FeLV/FIV assays on the cat sera. I thank
them all for their help!
I would like to thank my committee members (Dr. Rob MacKay, Dr. Al Merritt,
Dr. Ellis Greiner, and Dr. Steeve Giguere) for their time and support. They have helped
to make this thesis project a positive experience.
I'd also like to thank Sally O'Connell for her efficient command of all the
administrative details needed to meet the appropriate requirements.
Finally, thanks go to Dr. Charles Courtney, Dean for Research and Graduate
Studies at the College of Veterinary Medicine, for his support of my graduate education.
He didn't give up on me despite the fact that this thesis has been a long time coming.
This study was supported by funding from the Harold R. Morris Trust Fund,
dedicated to cat health care issues, and a University of Florida, College of Veterinary
Medicine Consolidated Faculty Research Development Grant.
TABLE OF CONTENTS
A C K N O W L E D G M E N T S .................................................................................................. ii
LIST OF TABLES ......... .... ..................... ........ .......... ...........vi
L IST O F F IG U R E S .... ...... ................................................ .. .. ..... .............. vii
A B STR A C T ..................... ................................... ........... ................. viii
1 IN TRODU CTION ................................................. ...... .................
2 LITER A TU R E REV IEW ............................................................. ....................... 4
Sarcocy stis ............................................................................................ . 4
T ax o n o m y ........................................................................... 5
L ife C y cle ...................................................................... . 6
M orphology ................................................................................. ........................
P athogenesis ........................................10
Equine Protozoal Myeloencephalitis (EPM) .................................................... 11
S ig n alm e n t ............... ... ......... ................................................................................ 12
H isto ry ...................................................................................................1 2
E p id em biology ...............................................................13
C clinical D diagnosis ....................................................... 13
Immunological Diagnosis.............................. ...............14
T re a tm e n t ....................................................................................................... 1 7
P rev mention ............. ................................................ ................... ............... 18
Sarcocystis neurona ............... ............................... ..........19
L ife C y cle ..................................................... .. .................................... 19
P ath o g en esis ................................................................2 0
D ia g n o sis ............................................ .......................................................... 2 1
Sarcocystis in D om estic C ats................................................................................ 26
Prevalence of Sarcocysts ................. ................................27
M o rp h o lo g y ................................................................................................... 2 7
Sarcocystis felis ..............................................................28
Pathogenesis ....................... ........................ ...............29
Sarcocystis neurona in Domestic Cats ............................................... ............... 29
Bioassays for Sarcocystis neurona in Cats.................................... ............... 30
Serological Surveys for S. neurona Antibody in Domestic Cats ......................32
3 M ATERIALS AND M ETHOD S ........................................ ......................... 34
Collection of Blood and M uscle Samples ...................................... ............... 34
Exam nation for Sarcocysts .............................................. .............................. 34
Molecular Characterization of Sarcocysts.............. ...................... ..............35
Serologic Testing .................... .................... 37
O possum Challenge ....... .......................... ........ ........... .............. .. 39
B radyzoite Culture .. ............... ... ...... ................ ... .................................40
Molecular Analysis of Fixed Tissue from Florida Panther Containing Sarcocysts....41
4 R E S U L T S .......................................................................... 4 3
5 D ISC U SSIO N ........... .............................................. .......................... 55
6 SU M M A R Y ......... .. ............. .................................................................. .....67
L IST O F R EFE R E N C E S ......... .......................... .......... ........................... ............... 68
B IO G R A PH IC A L SK E TCH ..................................................................... ..................76
LIST OF TABLES
4-1 Results of serology and histology for 9 of 50 cats euthanized at the local animal
shelter ............. ... ........ ......................................... .. ......... .. ... .. 52
4-2 Results of serology for 8 of 50 cats taken to trap-neuter-return clinics .................52
LIST OF FIGURES
4-1 Photograph of sarcocyst in cat skeletal muscle, 16x.......................... ............... 43
4-2 Portion of sarcocyst located in quadriceps muscle of cat.................. ...............44
4-3 Photomicrographs from light microscopy of fresh sarcocysts obtained from cat
m u scle ......................................................... .................. 4 4
4-4 Photographs taken from light microscopy of H & E stained sections of cat skeletal
m uscle containing sarcocysts ............................................................................45
4-5 Photomicrographs from light microscopy of the sarcocyst wall of fixed, H & E
stained and fresh sarcocysts obtained from cat muscle, 1000x................................45
4-6 Photomicrograph from TEM of cat sarcocyst .............................46
4-7 Photomicrograph from TEM of cat sarcocyst ........................................................ 47
4-8 Agarose-gel electrophoresis showing results of PCR using ssruRNA gene primer
pair JD26/JD 37 and tem plate DN A .............................................. ............... 48
4-9 Agarose-gel electrophoresis showing results of PCR using ITS-1 primer pair
JNB69/JNB70 and template DNA ...................................................... ............... 48
4-10 Unrooted phylogenetic tree showing the divergence of Sarcocystisfelis ITS-1 gene
sequence to that from Sarcocystis neurona, Sarcocystisfalcatula, and organisms
from related genera: Toxoplasma gondii, Hammondia heydorni, and Neospora
c a n in u m .......................................................................... 5 0
4-11 Im m unoblots of cat serum ............................................................. ..................... 5 1
4-12 Agarose-gel electrophoresis showing results of PCR using ssruRNA gene primer
pair JD26/JD37 and DNA prepared from formalin-fixed, paraffin-embedded
Florida panther tongue containing sarcocysts ................... ......................... 53
Abstract of Thesis Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Master of Science
NATURALLY-OCCURRING Sarcocystis INFECTION IN DOMESTIC CATS
Karen D. Gillis
Chair: Robert J. MacKay
Major Department: Veterinary Medicine
Equine protozoal myeloencephalitis is an important neurological disease of horses
in the United States. Consequently, there is an active research effort to identify hosts
associated with the primary causative agent, Sarcocystis neurona. The purpose of this
study was to determine whether the domestic cat (Felis catus) is a natural host for
S. neurona. Muscle sections from 50 primarily free-roaming domestic cats were
examined for the presence of sarcocysts. Sera from cats in this group and another group
of 50 free-roaming cats were evaluated for the presence of S. neurona antibody.
Sarcocysts were found in 5 of 50 cats (10%), and S. neurona antibody was found in 5 of
100 cats (5%) (one of which was also infected with sarcocysts). Morphological,
molecular (including ribosomal RNA genes), and biological characterization of these
sarcocysts showed that they were not S. neurona or S. neurona-like. Sarcocysts found in
the cats were identified morphologically as Sarcocystisfelis, a common parasite of wild
felids. The life cycle of S. felis is not known; before this study, no molecular marker for
S. felis existed. Although cats were found to be infected with S. felis sarcocysts,
serological data provided evidence of possible infection with S. neurona as well. Further
work is needed to determine the role of the domestic cat in the life cycle of S. neurona.
Sarcocystis spp. are common parasites with a wide range of hosts. Their life cycles
are unusual in that they are perpetuated by using two hosts: a definitive host that eats
sarcocyst-infected muscle, then passes infective sporocysts; and an intermediate host that
ingests these sporocysts and forms muscle sarcocysts as a result (Dubey et al., 1989). A
comprehensive cataloging of Sarcocystis spp. (Odening, 1998) indicates 189 species of
Sarcocystis have been identified to date; however, both definitive and intermediate hosts
are known for only 46% of these species. That report lists domestic cats (Felis catus) as
definitive hosts for 15 species, and as an intermediate host for only one species,
The domestic cat's role as an intermediate host for another species, Sarcocystis
neurona, has implications for the equine industry, resulting in several recent publications
and prompting the writing of this thesis as well. Sarcocystis neurona is the primary
causative agent of equine protozoal myeloencephalitis (EPM), an important neurological
disease of horses (Dubey et al., 1991, reviewed in MacKay et al., 2000). Currently,
effective preventatives for S. neurona infection (and thus EPM) in horses are neither
widely available nor widely used, so there is considerable interest in defining and
controlling the natural hosts of this parasite. Opossums (Didelphis virginiana, Didelphis
albiventris) are definitive hosts for S. neurona (Fenger et al., 1995, Dubey et al., 2001e);
nine-banded armadillos (Dasypus novemcinctus) (Cheadle et al., 2001a, Tanhauser et al.,
2001) and raccoons (Procyon lotor) (Dubey et al., 2001a) have been implicated as natural
intermediate hosts. The striped skunk (Mephitis mephitis) has also been identified as an
intermediate host, but only under laboratory conditions (Cheadle et al., 2001b).
The domestic cat may also be an intermediate host for S. neurona. In 2000, Dubey
and co-workers (2000) reported the first experimental completion of the life cycle of
S. neurona. Sarcocyst formation was induced in four laboratory-raised cats that had been
dosed with large numbers of S. neurona sporocysts obtained from naturally-infected
opossums. Three of these cats were immunosuppressed by corticosteroid administration.
Sarcocysts that formed in cat muscle were infective for two opossums. Sporocysts
produced by the infected opossums in turn caused S. neurona-associated encephalitis in
immune-deficient, interferon-gamma knockout mice. Molecular analysis of merozoites
recovered from the brain of one cat was consistent with S. neurona. Additional studies
have further defined S. neurona infection in cats (Stanek et al., 2001, Turay et al., 2002,
Butcher et al., 2002, Dubey et al., 2002, Rossano et al., 2002).
Before the discovery that the cat is able to function as an intermediate host for
S. neurona, the only other Sarcocystis spp. for which the cat was known to be an
intermediate host was Sarcocystisfelis (Dubey et al., 1992), named for the organism that
formed sarcocysts in the muscle of bobcats (Felis rufus). In that report, it was noted that,
based on ultrastructural morphology, these sarcocysts were identical to those found in the
domestic cat (Felis catus) and in cougars and panthers (Felis concolor). Later, S. felis
was formally identified in cheetahs (Acinonyxjubatus) (Briggs et al., 1993) and lions
(Panthera leo) (Dubey and Bwangamoi, 1994). Other than descriptive morphology of
sarcocysts, little is known about S. felis; no molecular or biological studies have been
reported to date, and the definitive host of this organism is unknown.
In 1990, there were an estimated 60 million feral cats in the United States
(Coleman et al., 1993). An APPMA 2001-2002 National Pet Owners Survey1 indicated
there were 73 million owned cats in the United States. Beyond the laboratory setting,
only one cat has been reported to have S. neurona sarcocysts, and this infection was not
fully characterized (Turay et al., 2002). Because investigations into the relationship
between cats and S. neurona are recent and limited in scope, more information is needed
about the cat's role as a natural intermediate host for Sarcocystis species. Cats are
frequently found around horse barns, so there is concern that cats should be controlled if
they are involved as hosts in the life cycle of S. neurona. If, however, cats are not an
important intermediate host for this Sarcocystis species, the unnecessary removal of large
numbers of cats can be avoided.
This thesis examines Sarcocystis spp. infections in domestic cats. The objective of
the study is to determine if cats are important, natural hosts for S. neurona. This objective
will be achieved by examining cat muscle for the presence of sarcocysts, characterizing
these sarcocysts, and evaluating cat sera for the presence of S. neurona antibodies. The
study hypothesis maintains that cats are not important, natural hosts for S. neurona.
This is the first survey to evaluate relatively large numbers of domestic cats for the
presence of sarcocysts, and the first to test for S. neurona antibodies by criteria used for
commercial-scale testing of horse sera and cerebrospinal fluid (CSF) for EPM. Sarcocysts
found in naturally-infected cats from this study were characterized by morphologic,
molecular, and biological methods. These data were compared to data from other studies
evaluating cats as hosts for Sarcocystis species.
1 ( American Pet Products Manufacturers Association, Inc. Greenwich CT
The name Sarcocystis describes a specific group of obligate intracellular protozoan
parasites. The name is Greek, derived from sarkos, meaning muscle or flesh, and kystis,
meaning bladder, and describes the terminal asexual life stage of the parasite found
encysted in the tissue of its host (Dubey et al., 1989, Dubey and Odening, 2001). The
genus Sarcocystis was named by Lancaster in 1882, however, the parasite was described
40 years earlier by Miescher (Dubey et al., 1989, Dubey and Odening, 2001). Sarcocystis
spp. are found in a large variety of hosts (wild animals, domestic animals, and man) and
are found worldwide. One unique characteristic of Sarcocystis is that it has a
diheteroxenous life cycle; two different types of host, a definitive host and an
intermediate host, succeed one another in the life cycle (Odening, 1998). These different
hosts accommodate different life stages of the parasite; sexual reproduction occurs in the
definitive host, and asexual reproduction in the intermediate host (Dubey et al., 1989,
Dubey and Odening, 2001). Definitive hosts are carnivorous, typically infected by
preying or scavenging on infected intermediate hosts. Sarcocystis may use one or more
similar types of intermediate host and/or definitive host species, although some
Sarcocystis spp. use several unrelated intermediate hosts (or definitive host animals) in
their life cycle (Dubey et al., 1989, Odening, 1998). Currently there are 189 named
species of Sarcocystis; undoubtedly more species will be discovered and named in the
future. Both definitive and intermediate hosts are known for only 89 species (Odening,
Sarcocystis parasites belong to the phylum Apicomplexa, class Sporozoasida, order
Eimeriorina, family Sarcocystidae, and genus Sarcocystis (Dubey and Odening, 2001).
Variations of this categorization, beginning at the family level and above, are recognized
as some parasitologists use subcategories, while others do not (Urquhart et al., 1996). As
Apicomplexans, Sarcocystis spp. possess the characteristic apical complex, located at the
anterior end of certain life stages, that is thought to aid in cell penetration (Dubey and
Odening, 2001). Genera related to Sarcocystis include Eimeria, Isospora,
Cryptosporidium, Toxoplasma, Neospora, Hammondia, Besnoitia, and Frenkelia. These
organisms are biologically distinct from Sarcocystis; location and structure of specific
life stages differ. Most closely related to Sarcocystis are: Toxoplasma, Neospora,
Hammondia, Besnoitia, and Frenkelia, all cyst-forming coccidians capable of using two
vertebrate hosts to complete their life cycle (Urquhart et al., 1996).
Host species names are commonly used to name and classify Sarcocystis species,
for example S. capracanis, and S. suihominis (Dubey et al., 1989, Odening, 1998).
However, it is now known that one species of Sarcocystis may be present in several
species of hosts and one species of host may harbor several Sarcocystis species. Now, the
ultrastucture of the sarcocyst wall is used for the taxonomic classification of Sarcocystis
species in preference to host species names (Dubey et al., 1989, Dubey and Odening,
2001). The structure of the sarcocyst wall may indicate phylogenetic relationships among
hosts; for instance Sarcocystis species found in sheep and goats all possess similar cyst
wall structure (Dubey et al., 1989). Currently there are 37 sarcocyst wall types used to
distinguish species. Ultrastructural cyst wall type is known for 119 species (63%)
Sarcocystis has a diheteroxenous, two-host life cycle in which the definitive host is
infected by eating the sarcocyst-infected muscle or neural tissue of an intermediate host.
The definitive host is carnivorous and the intermediate host is typically a herbivorous
prey animal. However, some definitive host species prey upon other carnivores or
scavenge from dead animals, allowing for both herbivorous and carnivorous intermediate
hosts, including birds and reptiles (Dubey et al., 1989). Some argue that the life cycle of
several Sarcocystis spp. can also be called dihomoxenous; in rare cases, definitive and
intermediate hosts belong to the same species. For instance, mice can function as both
definitive and intermediate hosts for S. muris or S. rodentifelis (Slapeta et al., 2001). Key
however, is that different stages occur in separate animals, even if they belong to the
same species (Odening, 1998). Sarcocystis gallotiae represents a unique case. The life
cycle of this Sarcocystis spp. has been called monoxenous in that different life stages
occur not only in the same species, but possibly the same animal; the host, a Canary
Island lizard named Gallotia galloti, eats its own tail, ingesting sarcocysts in the process
(Matuschka and Bannert, 1987).
When compared to related parasites, several features of the life cycle of Sarcocytis
are unique. Asexual reproduction, schizogony and sarcocyst formation, takes place only
in the intermediate host. Bradyzoites represent a state of "arrested development"; no
further development occurs until they are ingested by a definitive host, where they then
develop into gametocytes rather than schizonts. Sexual reproduction occurs only in the
definitive host and sporogony is accomplished fully within the definitive host; no further
development of sporocysts occurs once they are passed in the definitive host's feces
The following details from the life cycle of Sarcocystis spp. have been summarized
from the text "Sarcocystosis of Animals and Man" by Dubey et al., 1989.
Digestion of sarcocysts releases bradyzoites that invade the small intestinal
epithelium of the definitive host. There, bradyzoites undergo gametogony either
immediately, or after a period of several days (species dependent), differentiating into
both micro (male) and macro (female) gamonts; many more macrogamonts are formed
than are microgamonts. As the microgamont matures, its nucleus divides multiple times;
the resultant nuclei move to the margin of the microgamont, and differentiate into
microgametes that are released and disperse, aided by two flagella, to fertilize
macrogametes. Postfertilization, a thin-walled oocyst is formed. Oocysts undergo
asynchronous sporulation, resulting in two sporocysts, each containing four sporozoites.
Oocysts or sporocysts are passed in the feces, but are also infective even before release
from the small intestinal epithelium. The sexual reproduction process can be completed
in 24 hours and is asynchronous; both gamonts and oocysts are found simultaneously.
Intermediate hosts ingest sporocysts from fecal-contaminated food or water. Aided
by bile salts, excystation of sporocysts occurs in the intestinal tract, releasing sporozoites.
Sporozoites enter the small intestinal epithelium and migrate to the endothelial cells of
the mesenteric lymph node arteries. One or more rounds of asexual multiplication, called
schizogony (or merogony), occur here. Further rounds occur throughout the body in
blood monocytes and vascular epithelium. The location and number of rounds of
schizogony is species-dependent. Schizonts multiply by endopolygeny; numerous
merozoites develop simultaneously within the schizont and bud at its surface. Merozoites
from second (or succeeding) generation schizonts invade striated muscle, or in some
cases nerve cells, and form sarcocysts. The formation and maturation of a sarcocyst is
initiated when the merezoite becomes surrounded by parasitophorous vacuole and
develops a parasitophorous vacuolar membrane. The merozoite then divides by
endodyogeny, producing two metrocytes. Metrocytes rapidly divide, again by
endodyogeny, producing many bradyzoites within the sarcocyst. The parasitophorous
vacuolar membrane transforms into the primary cyst wall. Sarcocysts mature over one to
several months to become infective for the definitive host, and are the terminal asexual
stage, primarily found in skeletal muscle, heart, tongue, esophagus, and diaphragm of the
intermediate host. Factors affecting the number and distribution of sarcocysts include the
number of sporocysts ingested, species of Sarcocystis, species of host, and the
immunological status of the host.
Sarcocysts vary in size and shape depending on species and age. Some species are
always microscopic, for example S. cruzi, whereas others become macroscopic, i.e.
S. gigantea, and S. muris. Common shapes are filamentous, elongated, or globular. Shape
and size of sarcocysts is also dependent upon the type of host cell they are contained in;
long, slender cells contain long, slender sarcocysts. Macroscopic sarcocysts are nearly
always in skeletal muscle or esophageal muscle (Dubey et al., 1989). Because they are
derived from a merozoite having distinct anterior and posterior shapes, sarcocysts often
retain anterior and posterior features as well (Odening, 1998). The sarcocyst consists of a
cyst wall that surrounds metrocyte or bradyzoite stages within. The structure and
thickness of the cyst wall varies among Sarcocystis species; this feature is now the
primary means of differentiating species. A connective tissue capsule, formed by the
host, surrounds the cyst wall of fourteen species (Odening, 1998), as in S. gigantea.
Ultrastructural sarcocyst wall morphology is classified by type based upon a system
imposed by Dubey et al. (1989); 24 specific types have been described in detail.
Recently, an additional 13 types have been described (Dubey and Odening, 2001),
bringing the total to 37. Size, shape and spacing of cyst wall villi, and the presence or
absence of microtubules within the villi, constitute this system of classification. The
sarcocyst wall, ground substance underlying it, and the septae are remnants of the
merozoite. The metrocytes and bradyzoites within the sarcocyst are formed from
endodyogenic divisions of the merozoite nucleus. Other than the sarcocyst wall,
ultrastructural morphology is fairly consistent among species; a few species lack septae,
and the density of bradyzoites within the sarcocysts may vary by species (Dubey and
Odening, 2001). Mature sarcocysts may contain both metrocytes and bradyzoites,
although bradyzoites predominate (Dubey et al., 1989). Bradyzoites are surrounded by
three membranes, forming the pellicle, and have distinct anterior and posterior ends. At
the anterior end are the apical complex and numerous micronemes and several rhoptries.
Micronemes are thought to differentiate into rhoptries; both appear by transmission
electron microscopy (TEM) as dark structures, and are believed to secrete products that
aid in cell penetration (Dubey et al., 1989). Numbers of rhoptries in each bradyzoite may
differ among Sarcocystis species. Although bradyzoites and metrocytes contain the same
typical cell organelles such as a nucleus, endoplasmic reticulum, mitochondria, inclusion
bodies, etc., metrocytes lack the distinctive apical complex, micronemes, and rhoptries
found in bradyzoites (Dubey et al., 1989).
When compared to the sarcocyst, other life stages of Sarcocystis are of little value
in distinguishing among species, although sporocyst size may be useful (Cheadle et al.,
2001c); a typical sporocyst measures 12 |tm X 10 |tm. Dubey and colleagues, again in
their "Sarcocystosis of Animal and Man" text (1989), describe the morphology of the
various Sarcocystis life stages. Sporocysts are surrounded by two membranes, the inner
composed of four fused plates, and contain four sporozoites and a residual body.
Sporozoite morphology is very similar to that for bradyzoites. Once inside the intestinal
epithelial cells of the intermediate host, sporozoites differentiate into schizonts; unlike
sarcocysts and merozoites, a parasitophorous membrane does not surround sporozoites
and schizonts within host cells. Schizonts change morphologically as they mature and
divide by endopolyogeny; the nucleus becomes multi-lobed, each lobe giving rise to two
merozoites. Merozoites vary in size and shape but are typically reported to be
approximately 8 |tm X 2.5 |tm. Merozoites also possess anterior and posterior ends and
an apical complex, but do not contain rhoptries.
Gametogony in the definitive host produces rounded macrogamonts, 10-20 |tm in
diameter, and elongated microgamonts, approximately 7 X 5 |tm. Microgamonts become
multi-nucleated, and eventually contain 3 to 11 slender, bi-flagellated, microgametes
typically measuring 4 X 0.5 |tm. After fertilization, the zygote differentiates into an
oocyst, approximately 24 X 20 |tm and containing two sporocysts within.
Typically there is little or no illness in the definitive host other than gastrointestinal
disease (Dubey et al., 1989, Urquhart et al., 1996, Dubey and Odening, 2001). However,
the pathogenicity of Sarcocystis spp. in intermediate hosts varies by species, from
mildly-pathogenic (S. fayeri in horses, S. gigantea in sheep) to very pathogenic (S. cruzi
in cattle, S. capricanis in goats), and host; disease in small intermediate hosts, i.e.,
S. falcatula in some bird species, and S. idahoensis in deer mice, is more severe than that
seen in large animal hosts (Dubey et al., 1989). In experimental infections, severity of
host disease was dependent upon number of sporocysts given (Dubey et al., 1989). As
Sarcocystis spp. are obligate intracellular parasites, disease is caused by the destruction of
host cells; hemorrhage and inflammatory lesions are commonly seen (Dubey et al., 1989,
Dubey and Odening, 2001). Infection of the central nervous system occurs only rarely
with most Sarcocystis spp. (Dubey et al., 1989) except for S. neurona, which causes
neurological disease in several animal species. In horses, equine protozoal
myeloencephalitis (EPM), discussed in the next section, is the result of such an infection.
In food animals, the presence of sarcocysts, caused for example by S. gigantea, can result
in condemnation of meat, resulting in economic losses for farmers (Dubey et al., 1989).
Sarcocystis infection (ex: S. cruzi, S. ovicanis) has also been implicated in abortions in
farm animal hosts (Dubey et al., 1989, Urquhart et al., 1996).
Equine Protozoal Myeloencephalitis (EPM)
Considerable research effort has been devoted to one species of Sarcocystis,
Sarcocystis neurona, due to the fact that S. neurona infection in horses is the principle
cause of equine protozoal myeloencephalitis (EPM) (Dubey et al., 1991). EPM is seen
only in the Americas, a fact that puzzled researchers until it was found that the
distribution of EPM follows that of the definitive host, the opossum. Both the Virginia
opossum (Didelphis viginiana) and the South American, white-eared opossum (Didelphis
albiventris) are definitive hosts ofS. neurona (Fenger et al., 1995, Dubey et al., 2001e).
Although now likely superceded by West Nile Encephalitis, in 2001, EPM was the most
commonly-diagnosed neurological disease of horses in the United States (Dubey et al.,
2001d). It is estimated that EPM has caused greater than $100 million in losses to the
nation's equine industry (Dubey et al., 2001d). Regarding clinical EPM, costs associated
with diagnosis and treatment alone range from $55.4 to $110.8 million per year (Dubey et
Clinical signs of EPM can be variable due to the location and severity of parasite
damage to the CNS (MacKay et al., 2000). Common signs include head tilt, facial
paralysis, difficulty swallowing, depression (all due to brain or brain stem damage) and
ataxia and gait abnormalities (due to spinal cord damage). Another clinical sign, focal
muscle atrophy, is often characteristic of EPM. Clinical signs may worsen suddenly, or
improve, although relapse is common; EPM is typically a progressive disease (MacKay
et al., 2000). Related parasites from the genus Neospora are implicated in several cases of
EPM. Neospora hughesi is proposed as the causative agent (Marsh et al., 1998).
In the review of EPM by MacKay and co-authors, (2000), the recognition and
emergence of EPM is described. Neurological syndromes designated "segmental
myelitis" and "focal encephalitis-myelitis" were identified in the 1960's (Rooney et al.,
1970). In 1974, protozoal organisms were found to be associated with these lesions, and
were thought to be Toxoplasma gondii (Cusick et al., 1974). Another report attributed the
infections to a Sarcocystis species (Dubey, 1976), later confirmed by electron microscopy
in 1980 (Simpson and Mayhew, 1980). The name "equine protozoal myeloencephalitis"
was proposed in 1976 by Mayhew et al. and widely adapted. In 1991, the causative agent
was named "Sarcocystis neurona"(Dubey et al., 1991).
EPM affects primarily young horses; an analysis of histologically-confirmed cases
of EPM found that most, 62%, were 4 years old or less (Fenger, 1997). Diagnosis by
neurological examination alone found young horses (1-5 years old), and older horses
(>13 years old) were at an increased risk of developing EPM (Saville et al., 2000a). In
addition to age, there are other risk factors for EPM. Immune suppression, linked to
overall health, advancing age, and stress is associated with an increased incidence of
EPM (Saville et al., 2000b). Because of stress, show and racehorses are affected more
than horses kept for breeding or pleasure (Saville et al., 2000a). By neurological
examination, more cases of EPM were diagnosed in summer and fall than were in other
seasons (Saville et al., 2000a), possibly reflective of increased exposure to host animals,
increased survival of S. neurona in the environment (Saville et al., 1997), or stress
induced by timing of competitive events (MacKay et al., 2000). Horses were also more at
risk for developing EPM when opossums were present in the environment or there was
close proximity of opossum habitat, and if rats and mice, possible hosts, were present in
the environment (NAHMS, 1998).
Exposure to S. neurona, as measured by the presence of S. neurona serum
antibodies, indicates that approximately 50% of horses in the United States have been
exposed (MacKay et al., 2000). However, the prevalence of EPM, based upon
postmortem evaluation, is estimated to be less than 1% (Granstrom, 1997, NAHMS,
Diagnosis of EPM can be made by antemortem or postmortem methods. At
necropsy, gross lesions of the CNS are sometimes visible in horses afflicted with EPM;
dark hemorrhagic discolorations of spinal cord sections are frequently depicted. Parasites
are not always found in CNS lesions; only 10 to 36% of hematoxylin and eosin
preparations and 20 to 51% of immunohistological preparations revealed parasites
(MacKay et al., 2000). However, characteristic and consistent lesions are often seen in
preparations when parasites are absent. These lesions include perivascular cuffing by
mononuclear cells, infiltrates of lymphocytes, neutrophils, eosinophils, multinucleate
giant cells, and astrocyte proliferation (MacKay et al., 2000, Dubey et al., 2001d).
Antemortem diagnosis of EPM is best made by a thorough examination of the
horse and suitable clinical laboratory tests. Differentiating EPM from other neurological
diseases can sometimes be difficult. Other diseases to consider include: cervical vertebral
malformation, equine herpesvirus-1 myeloencephalopathy, equine motor neuron disease,
tumors, abscess, migrating metazoan parasites, rabies, West Nile viral encephalitis,
equine degenerative myeloencephalopathy, and vascular and bone malformations
(MacKay et al., 2000).
An immunoblot test to detect S. neurona IgG in serum or CSF is used as an aid to
the diagnosis of EPM. A positive immunblot test of CSF from a neurologic horse
suggests CNS infection, intrathecal production of S. neurona antibody, and EPM.
Collection of CSF must be done by a qualified veterinarian; the procedure poses some
risk to both horse and veterinarian, and sample quality is very important. The procedure
can also be expensive to perform (Dubey et al., 2001d).
The immunoblot test has evolved to some extent since its development in the early
1990's. Initially, eight proteins, of varying molecular weight, were identified as
S. neurona specific (Granstrom et al., 1993). Cross-reactive S. fayeri, S. cruzi, and
S. muris proteins were identified by this test and excluded in the interpretation of positive
results. Versions of this test are currently commercially used for EPM testing at three
laboratories: Neogen Corporation (Lexington, KY), Equine Biodiagnostics Incorporated
(EBI, Lexington, KY), and the Michigan State Animal Health Diagnostic Laboratory,
(East Lansing, MI). Each of these laboratories offers slightly different test formats and
therefore, interpretation of results. The Michigan State test, for example, now preadsorbs
blots with S. cruzi-positive bovine serum to increase test sensitivity and specificity. Due
to differences in test format, reagents, parasite preparations, etc., positive results are
interpreted differently among labs. Serum or CSF reacting against a 17-kDa protein of
S. neurona is considered positive at the Neogen EPM testing lab. At EBI, reactivity
against 14.5, 13, and 7-kDa proteins constitutes a positive result. At Michigan State,
reactivity against both 30-kDa and 16-kDa proteins must be present for a positive result.
Granstrom (1997) describes the use of a non-commercial immunoblot in a
population of 295 horses euthanized for neurological disease. Sensitivity of the test, the
ability of the test to detect antibody in CSF from EPM-positive horses, was 89% (11%
false negative). Specificity, the ability of the test to give negative results for
EPM-negative horses, was also 89% (11% false positive). False positive results can result
from defects in the blood-CNS barrier, or more commonly, from blood contamination of
the sample. The positive predictive value of this test, those horses testing positive that are
truly positive in that population was 85%. The negative predictive value, the percentage
of horses testing negative that are truly free of EPM was 92%. However, because the
prevalence of EPM is estimated to be only approximately 1%, when this test was used in
a normal population of horses (instead of the neurologic horses tested) the positive
predictive value of the test drops to 8%; a test's predictive values are dependent upon the
prevalence of the disease in the population tested. For this reason, the authors suggest that
the immunoblot test not be used for screening CSF from normal horses. From the same
study, serum testing yielded a test sensitivity of 89% (11% false negative) and a
specificity of 71% (29% of neurologic horses did not have EPM, but did have antibody)
confirming the accepted premise, even within this group of neurological horses, that
serum samples testing positive show merely that the horse has been exposed to
S. neurona. Approximately 50% of horses in the U.S. have serum antibodies to
S. neurona, while only 1% have EPM (MacKay et al., 2000).
Immunoblots of CSF and serum from both neurologic and clinically-normal horses
were evaluated by Daft et al. (2002). Immunological results were compared to
immunohistochemistry postmortem. The sensitivity of the immunoblot used in the study
was high, 80%-88%, for both serum and CSF from either neurologic or clinically-normal
horses. However, the specificity was lower: for CSF, 44% for neurologic horses, and
60% for normal horses, and for serum, 38% for neurologic horses, and 56% for normal
horses. False positive results were decreased when weakly reacting or indeterminate
samples were categorized as negative. Daft and co-authors state that the lower specificity
of the test in neurologic horses may reflect damage to the blood-CNS barrier in horses
with neurologic disease other than EPM, or, as indicated previously, these horses could
have been exposed to S. neurona and have a neurologic disease other than EPM. Daft and
coauthors report 40% of horses negative for EPM, had S. neurona antibodies in the CSF,
concluding that weakly-positive CSF may be normal for some seropositive horses. The
value of the test therefore is in ruling out EPM; a negative result is more useful than a
Despite its limitations, the wide acceptance of the immunoblot for detection of
S. neurona antibody in horses has meant that other immunological methods, such as
indirect fluorescent antibody (IFA) testing or enzyme-linked immunosorbent assay
(ELISA) testing, are so far, confined to research applications. The use the polymerase
chain reaction (PCR) for detection of S. neurona in CSF is not yet optimized; the
sensitivity of this method is not as great as hoped (MacKay et al., 2000). This may be due
to the scarcity of parasite in the CSF, or the lack of intact DNA. The use of PCR testing
may be helpful when S. neurona antibody is weak or undetectable in CSF from a horse
with neurologic signs consistent with EPM (Dubey et al., 2001d).
Horses that receive treatment for EPM were 10 times more likely to improve, and
those that improve were 50 times more likely to survive (Saville, 2000a). If started early
after the onset of clinical signs, treatment may be effective in up to 75% of cases (Dubey
et al., 2001d). Traditional EPM treatment involves the use of folate inhibitors such as
sulfadiazine/pyrimethamine combinations. Horses are often treated long-term with this
combination, for months at a time, or even indefinitely. Side effects such as anemia and
leucopenia have been noted (MacKay et al., 2000). Anti-coccidial triazine compounds
such as diclazuril, toltrazuril, and its sulfone metabolite ponazuril, are relatively new
EPM therapies (MacKay et al., 2000). Currently, ponazuril (Marquis, Bayer Animal
Health, Shawnee Mission, KS) is the only product approved specifically for such use.
Triazines must also be adiminstered for weeks at a time (28 days at 5mg/kg for
ponazuril), and are very costly, up to $1000.00 for a course of treatment. Another
medication, nitazoxanide, is a coccidiocidal therapeutic agent currently under review by
the FDA for the treatment of EPM. There have been reports of toxicity associated with
nitazoxanide; however, these effects were lessened when dosages were reduced (MacKay
et al., 2000). As with other EPM treatments, nitazoxanide therapy may be lengthy and
Prevention of EPM has focused on limiting the access of opossums to feed, hay,
water, and bedding supplies. Some horse owners have chosen to feed only pelleted feeds
that have gone through a heat process killing sporocysts which may be present on
feedstuffs or feed ingredients (MacKay et al., 2000). Opossums are sometimes trapped on
horse farms and moved to other locations or killed (personal communication,
T. J. Cutler).
Recently, an EPM vaccine, "designed to aid in the prevention of neurologic disease
due to subsequent infections of S. neurona" (Ft. Dodge Animal Health, Ft. Dodge, IA)
has been introduced. The vaccine is under conditional license from the United States
Department of Agriculture (USDA); results from efficacy studies, currently in progress,
must be satisfactory before full licensure is granted. The vaccine is manufactured from
S. neurona merozoites harvested from cell culture; standardized numbers of merozoites
are chemically-inactivated and mixed with adjuvant. One concern regarding the use of
this vaccine is that serum and CSF samples may test immunoblot positive, particularly
when tested shortly after vaccination, reducing the value of a negative CSF immunoblot
(Fort Dodge Animal Health Bulletin
Named in 1991 for the Sarcocystis spp. isolated from spinal cord lesions of horses
afflicted with EPM (Dubey et al., 1991), S. neurona has now been characterized to some
extent by molecular, biological, and morphological methods. These research efforts aim
to differentiate S. neurona from other Sarcocystis species, identify other possible animal
hosts for this parasite, and improve our understanding of EPM. Sarcocystis neurona was
known to exist before its definitive and intermediate hosts were identified. Morphologic
analysis of merozoites and schizonts isolated from CNS lesions from several horses with
EPM showed that the same parasite was present in each of these lesions, and that it was a
distinct species of Sarcocystis (Dubey et al., 1991, Dubey et al., 2001d). Sarcocystis
neurona was then successfully cultured in vitro (Davis et al., 1991a, 1991b), allowing for
more comprehensive characterization.
Fenger and colleagues (1995) proposed the Virginia opossum (Didelphis
virginiana) as a definitive host because of its status as an omnivore/carnivore endemic in
areas where EPM was found in horses; the study also then provided molecular evidence
that sporocysts shed by opossums were S. neurona sporocysts. The definitive host status
of the opossum was confirmed by Fenger et al., in 1997, when opossum sporocysts
administered to horses induced S. neurona antibody and neurologic disease in these
horses. Another opossum species, Didelphis albiventris, the South American white-eared
opossum was also found to be a definitive host for S. neurona (Dubey et al., 2001e).
Other species of Sarcocystis sporocysts are passed by the opossum as well and can be can
be differentiated by molecular (Tanhauser et al., 1999,Dubey et al., 2001f, Rosenthal et
al., 2001), and possibly morphological, methods (Cheadle et al., 2001c).
The life cycle of S. neurona is completed when the opossum eats sarcocyst-infected
muscle from an intermediate host, most probably from scavenging road-killed host
animals. The intermediate host(s), and thus the complete life cycle were unknown for a
decade or more since the discovery and isolation of S. neurona. In 2000, the life cycle
was completed in the laboratory with domestic cats as experimental intermediate hosts
(Dubey et al., 2000). Shortly thereafter, the armadillo (Dasypus novemcinctus), striped
skunk (Mephitis mephitis), and raccoon (Procyon lotor) were identified as intermediate
hosts as well (Cheadle et al., 2001a, Tanhauser et al., 2001, Cheadle et al., 2001b, Dubey
et al., 2001a). The sea otter (Enhydra lutris) was also found to be naturally-infected
(Lindsay et al., 2000) and capable of infecting opossums. Sea otters likely become
infected by storm water run off contamination of shellfish stocks (Dr. Ellis Greiner,
personal communication); their status as natural intermediate hosts for S. neurona is
unlikely (Dubey et al. 2001b). Once intermediate hosts for S. neurona were identified, the
sarcocyst stage was then described (Cheadle et al., 2001a, Cheadle et al., 2001b, Dubey et
al., 2001a, Dubey et al., 2001b, Dubey et al. 2001c). Sarcocystis neurona therefore
utilizes a single type of definitive host (although others may be identified in the future)
that is host to other Sarcocystis species, and multiple intermediate hosts, each known to
be hosts for other Sarcocystis spp. as well (Odening, 1998).
Sarcocystis neurona was named for its ability to infect neural tissue in a dead-end
host; no muscle sarcocysts have been found in horses (MacKay et al., 2000). Asexual life
stages (merozoites, schizonts, and occasionally sarcocysts) of S. neurona, and EPM-like
signs, are found in neural tissues of other animals: cats (Dubey et al., 1994, Dubey and
Hamir, 2000, Forest et al., 2000), skunks (Dubey and Hamir, 2000), raccoons (Hamir and
Dubey, 2001, Dubey and Hamir, 2000), mink (Mustela vison) (Dubey and Hedstrom,
1993, Dubey and Hamir, 2000), zebra, (Equus burchelli bohmi) (Marsh et al., 2000),
Pacific harbor seal (Phoca vitulina richardsi) (Miller et al., 2001a), sea otter (Miller et al.,
2001b, Lindsay et al., 2000, Lindsay et al., 2001), Straw-necked ibis (Carphibis
spinicollis) (Dubey et al., 2001g), and gannet (Morus bassanus) (Spalding et al., 2002).
Both neural and inflammatory cells in the CNS can become parasitized (Dubey et al.
2001d). Other Sarcocystis spp. are found only rarely in the CNS (Dubey et al., 1989).
Later reports have identified several of the aberrantly-infected animal species listed
above to be intermediate hosts for S. neurona; tissue from these animals was capable of
infecting opossums when fed in an experimental feeding trial (Dubey et al., 2000, Dubey
et al., 2001a, Dubey et al., 2001b, Dubey et al., 2002, Cheadle et al., 2001b).
Morphological. Morphological diagnosis of S. neurona infection has only really
been made feasible since the discovery of its intermediate hosts. Ultrastructural
characterization by TEM has shown the sarcocyst wall of S. neurona as having tapered
villar projections and microtubules within villi (Dubey et al., 2000, Dubey et al. 2001b,
Dubey et al., 2001c, Cheadle et al., 2001b). Cheadle and colleagues (2001b) note that this
morphology correlates to a type 11 per the classification system instituted by Dubey et al.
in 1989. However, the same report notes that TEM of the S. neurona sarcocyst found in
skunks resembles that seen for other Sarcocystis species: S. dasypi, S. kirkpatricki (a
Sarcocystis spp. seen in a raccoon), S. falcatula, and S. fayeri. Ultrastructural
morphology alone may not be sufficient to differentiate species. Other details seen by
TEM include the presence of relatively few rhoptries, and bradyzoites 5 |tm long by 1 |tm
wide (Dubey et al., 2001c). In the same report, S. neurona sarcocysts have measured 700
|tm in length by 50 |tm in width, and in another report, 140 |tm X 20 |tm (Butcher et al.
2002). Although not of significance for species diagnosis, Speer and Dubey (2001) have
described the ultrastructure of S. neurona schizonts and merozoites.
Molecular. Molecular analysis was one of the early tools used to identify and
characterize S. neurona. Random-amplifed polymorphic DNA (RAPD) markers were
developed to differentiate S. neurona from other Apicomplexan parasites and some
Sarcocystis species (Granstrom et al., 1994). Sequencing and PCR of the small subunit
ribosomal RNA (ssurRNA) gene prepared from S. neurona merozoites was used to
identify the Virginia opossum as a definitive host (Fenger et al., 1995). Small subunit
ribosomal DNA sequence was then used to conclude S. neurona and a similar species,
S. falcatula, were synonymous, and therefore birds, particularly the brown-headed
cowbird (Molothrus ater), were likely intermediate hosts (Dame et al., 1995). This
conclusion proved erroneous when additional RAPD-based markers were shown to
differentiate S. neurona from S. falcatula and other species found in the opossum
(Tanhauser et al., 1999). Additional methods such as infectivity of animal hosts (Cutler et
al., 1999) or cell cultures (Lindsay et al., 1999) further differentiated these two
Sarcocystis species. Comparison of sequences from the internal transcribed spacer 1
(ITS-1) region of ribosomal genes, more rapidly evolving than small subunit genes, were
also useful for differentiating S. neurona from S. falcatula (Tanhauser et al., 1999 and
Marsh et al., 1999). The RAPD-derived markers and restriction endonuclease protocols
developed by Tanhauser and co-workers (1999) are now widely used to identify
S. neurona in both new intermediate hosts and in aberrantly-infected animals. Sarcocystis
neurona sequence data from loci amplified by these markers, as well as other regions of
ribosomal RNA gene sequences have been archived in GenBankTM and are proving
useful for phylogenetic analyses of Sarcocystis species.
Immunological. Immuno-based methods are also used to identify S. neurona
exposure or infection and characterize antibody response in the infected host(s). Besides
their use in horses, discussed in the previous section for diagnosis of EPM, serologic
surveys have been used to aid in the identification of natural intermediate hosts for
S. neurona (Tanhauser et al., 2001, Stanek et al., 2001, Rossano et al., 2002, Turray et al.,
2002, Mitchell et al., 2002). Several methods of serologic examination have been used:
direct agglutination testing (Dubey et al., 2001a, Lindsay and Dubey 2001, Dubey et al.,
2000, Dubey et al, 2002, Dubey et al., 2001b), indirect fluorescent antibody testing
(Lindsay et al., 2000, Miller et al., 2001a, Butcher et al., 2002, Ellison et al., 2002,
Rossano et al., 2002), and immunoblotting (Cheadle et al., 2001a, Cheadle et al., 2001b,
Tanhauser et al., 2001, Turay et al., 2002, Butcher et al., 2002, Rossano et al. 2002, Long
et al., 2002). Immunohistochemistry is also widely used, demonstrating the presence of
S. neurona schizonts in the CNS of aberrant and experimental hosts (Dubey et al., 2000,
Lindsay et al., 2000, Dubey et al., 2001a, Dubey et al., 2001b, Cheadle et al., 2001b,
Lindsay and Dubey 2001, Turay et al., 2002, Butcher et al., 2002, Fritz and Dubey,
2002). Sections of brain tissue are usually examined by this method. However,
identification by immunohistochemistry relies on the use of polyclonal S. neurona
antiserum, so infection in these animals is presumptive unless followed by additional
diagnostics (Dubey et al., 2001g, Dubey et al, 2001d). In addition to cross-reactivity
across Sarcocystis species, immunohistochemistry has also demonstrated cross-reactivity
across different strains (i.e., horse spinal cord-derived or intermediate host-derived) and
life stage of the same species (Turay et al., 2002, Butcher et al., 2002). Such variable
immunoreactivity has meant that other techniques, such as biological assay, molecular
characterization, or morphologic identification, are typically used in conjunction with
immunological methods to confirm S. neurona infection.
Biological. Sarcocystis neurona has been successfully cultured in a variety of cell
lines; bovine monocytes (Dubey et al., 1989), equine dermal cells (Murphy and
Mansfield, 1999), and bovine turbinate cells (Speer and Dubey, 2001) are commonly
used for this purpose. Cultures have been established from merozoites (Dubey et al.,
2001d) and sporocysts (Murphy & Mansfield, 1999). Sarcocystis species can be
differentiated biologically from related Apicomplexans by their behavior in vitro (Dubey
and Odening, 2001). Sarcocystis neurona does exhibit certain characteristics in culture
(Lindsay et al., 1999), but these characteristics have not been reported as a means to
differentiate it from other Sarcocystis species. The availability of culture-grown isolates,
however, has facilitated efforts to more fully characterize S. neurona.
In vivo methods, i.e. host specificity and location of infection (aberrant infections
with S. neurona are confined to the CNS) (Dubey et al., 1989, Dubey et al., 2001d) have
been used to diagnose S. neurona infection. Now, other in vivo methods are routinely for
diagnosis and characterization S. neurona infection. An animal model of infection for
studies of the pathogenesis of S. neurona was developed in immune-deficient mice
(Marsh 1997, Dubey and Lindsay, 1998). Both nude and gamma interferon knockout
mice can be inoculated either orally, subcutaneously, or intraperitoneally with sporocysts
or merozoites of S. neurona; these mice develop encephalitic CNS signs within seven to
thirty days postinoculation. Immune-competent mice such as Balb/c, or C57/B1 are not
susceptible to S. neurona infection. Sporocysts or merozoites from other Sarcocystis
species are not infective to nude and gamma interferon knockout mice. This infection
model can therefore be used to test for the presence of viable S. neurona parasite in
preparations of host feces, infected tissues, or cultures established from suspected natural
or aberrant hosts (Dubey et al., 2000, Lindsay et al, 2000, Cheadle et al., 2001b. Dubey et
al., 2001a, Dubey et al., 2001b, Turay et al., 2002, Butcher et al., 2002, Long et al.,
2002). Lack of infection in these mice suggests the sample did not contain S. neurona,
but perhaps some other Sarcocystis species that might be present (Dubey and Lindsay,
1998). Athymic nude mice are thought to be susceptible to infection with protozoal
organisms due to a deficiency of T-cells (Marsh, et al., 1997), while gamma interferon
knockout mice are unable to activate cytotoxic macrophage activity (Deckert-Schluter,
Biological assays utilizing controlled feeding trials have been used to demonstrate
the presence of S. neurona infections in hosts and investigate previously unknown hosts.
Infected tissue from a suspected host is fed to opossums shown to be currently non-
infected. After a period of time, animals fed the suspect tissue are examined for the
presence of S. neurona infection. Tissue from the newly-infected host can be fed back to
the same original host species to demonstrate a complete life cycle (Dubey et al., 2002).
Often times, sporocysts or merozoites produced during the life cycle are inoculated into
immune-deficient mice to verify the infective agent is S. neurona (Dubey et al. 2000,
Dubey et al., 2001a, Dubey et al., 2001b, Cheadle et al., 2001b, Turay et al., 2002,
Butcher et al., 2002).
Experimental models of S. neurona infection utilizing horses have continued to
frustrate EPM researchers. Although horses inoculated with S. neurona develop clinical
signs and S. neurona antibodies, no parasites have been recovered from these inoculated
horses (Fenger et al., 1997, Cutler et al., 2001, Saville et al., 2001); Koch's postulates of
infection have therefore not been fulfilled in horses. Research on this problem continues.
Sarcocystis in Domestic Cats
Historically, intermediate hosts for Sarcocystis species were thought to be
herbivores, definitive hosts carnivores, and life cycles perpetuated through predator/prey
or scavenger/carrion relationships. Cats are listed as definitive hosts for the following 15
species of Sarcocystis (the common name of the intermediate host follows in
parenthesis): S. buffalonis (water buffalo), S. cuniculorum (rabbit), S. cymruensis (rat),
S. fusiformis (water buffalo), S. gigantea (sheep), S. hirsuta (cattle), S. leporum (rabbit),
S. medusiformis (sheep), S. moulei (goat), S. muris (mouse), S. neotomafelis (woodrat),
S. odoi (deer), S. poricifelis (pig), S. rodentifelis (rats, mice), S. wenzeli (chicken)
(Odening, 1998). Cats become infected by preying or scavenging upon these animals.
Compared to other Sarcocystis definitive hosts, infected cats shed low numbers of
sporocysts (Christie et al., 1976, Dubey et al., 1989); cats may not be very good
producers of sporocysts, sarcocysts of feline-transmitted species may be slow maturing,
or some animal species are more resistant to Sarcocystis infection (Dubey et al., 1989).
Cats are able to function as intermediate hosts for Sarcocystis spp. as well. Reports
dating back to as early as 1956 (Eisenstein and Innes) identified muscle sarcocysts in
domestic cats (Kirkpatrick et al., 1986, Everitt et al., 1987, Hill et al., 1988, Edwards et
al., 1988, Fiori and Lowndes, 1988) and wild felids. Among the latter group are bobcats
(Anderson et al., 1992, Dubey et al., 1992), lions (Bhatavdekar and Purohit, 1963, Dubey
and Bwangamoi, 1994, Kinsel et al., 1998), cheetahs (Briggs et al., 1993), and Florida
panthers and cougars (Greiner et al., 1989).
Prevalence of Sarcocysts
It has been suggested that immunosuppression, resulting from lymphoid neoplasia,
corticosteroid therapy, infection with feline immunodeficiency virus (FIV) or feline
leukemia virus (FeLV), or inbreeding, might permit aberrant sarcocystosis by an
opportunistic organism (Kirkpatrick et al., 1986, Hill et al., 1988, Greiner et al., 1989,
Briggs et al., 1993), or extraintestinal infection by a Sarcocystis spp. for which the cat
serves as a definitive host (Edwards et al. 1988). However, sarcocysts have also been
found at in normal, healthy felids (Everitt et al., 1987, Fiori and Lowndes 1988, Greiner
et al., 1989, Anderson et al., 1992, Dubey et al., 1992, Kinsel et al., 1998). Although
typically found at necropsy, surveys for sarcocysts have been conducted using a limited
number of healthy cats. In Indiana, sarcocysts were found in 4 of 12 cats (Everitt et al.
1987), and in Missouri, found in 1 of 9 cats (Turay et al. 2002). In another study,
sarcocysts were collected from muscle biopsies of 5 cats, however no data was given
regarding prevalence (Fiori and Lowndes 1988). Greater rates of sarcocyst infection have
been reported in wild species. Necropsy of four mature lions from Namibia revealed
sarcocysts in each (Kinsel et al., 1998). In Florida, 50% of bobcats (Anderson et al.,
1992), and 83% of free-ranging Florida panthers and cougars (Greiner et al., 1989) were
infected. Sixty-six percent of Arkansas bobcats (Dubey et al., 1992) and 70% of captive
born and raised cheetahs (Briggs et al., 1993) had sarcocysts.
In both domestic cats and wild felid species, sarcocysts have been described
incidental to examination of muscle tissue collected at necropsy from cats afflicted with
neoplasia or other debilitating disease (Bhatavdekar and Purohit 1963, Kirkpatrick et al.
1986, Edwards et al. 1988, Hill et al. 1988, Greiner et al. 1989, Briggs et al. 1993, Dubey
and Bwangamoi 1994), and from apparently healthy cats (Everitt et al. 1987, Fiori and
Lowndes 1988, Greiner et al. 1989, Anderson et al. 1992, Dubey et al. 1992, Kinsel et al.
1998). In these reports, histologic examination revealed sarcocysts of varying sizes, from
24 |tm (Kirkpatrick et al., 1986) to 270 |tm (Everitt et al., 1987) in diameter by 24 |tm
(Everitt et al., 1987) to 2100 |tm in length (Dubey et al., 1992). The ultrastructure of
these sarcocysts appears nearly identical in all cases, having rounded, irregularly-spaced
villi devoid of microtubules, a regularly interrupted electron-dense layer underlying the
parasitophorous membrane, and septae separating bradyzoites that measure
approximately 10 |tm long. It was noted that ultrastructural morphology was inconsistent
with descriptions of sarcocysts for which the cat serves as a definitive host (Kirkpatrick et
In 1992, Dubey and co-workers gave the name Sarcocystisfelis to the organism
which caused sarcocysts in the muscle of bobcats In that report, it was noted that, based
on ultrastructural morphology, these sarcocysts were identical to those found in the
domestic cat and in cougars and panthers. Later, S. felis was formally identified in
cheetahs (Briggs et al., 1993) and lions (Dubey and Bwangamoi, 1994). Sarcocystisfelis
and S. felis-like sarcocysts have been reported in felids from North America, Africa
(Dubey and Bwangamoi, 1994, Kinsel et al. 1998) and India (Bhatavdekar and Purohit
1963). The definitive host of S. felis remains unknown.
The pathogenicity of S. felis infections in cats is also unknown. The majority of
these sarcocysts have been found in skeletal muscle (Kirkpatrick et al., 1986, Everitt et
al., 1987, Hill et al., 1988, Fiori and Lowndes, 1988, Greiner et al., 1989, Anderson et al.,
1992, Dubey et al., 1992, Briggs et al., 1993, Dubey and Bwangamoi, 1994, Kinsel et al.,
1998) and tongue (Greiner et al., 1989, Anderson et al., 1992, Dubey et al., 1992), and
occasionally in diaphragm (Kirkpatrick et al., 1986, Greiner et al., 1989, Anderson et al.,
1992) and cardiac muscle (Bhatavdekar and Purohit 1963, Kirkpatrick et al., 1986,
Everitt et al., 1987, Hill et al., 1988 Greiner et al., 1989, Anderson et al., 1992, Dubey et
al., 1992). Only one report of disseminated sarcocystosis is given; this condition was
associated with immunosuppression due to lymphosarcoma (Edwards et al. 1988). Rarely
described is encephalitis or myelitis in felids caused by infection with Sarcocystis; these
cases have been attributed to infection of the CNS with S. neurona (Dubey et al. 1994,
Dubey and Hamir 2000, Forest et al. 2000).
Sarcocystis neurona in Domestic Cats
The wide distribution and high prevalence of S. neurona antibodies in horses has
prompted the search for equally common and widespread intermediate hosts. In 1994,
Dubey and co-workers published a report of Sarcocystis-associated encephalitis in a
domestic cat (Dubey et al., 1994). That infection was later shown to be S. neurona
(Dubey and Hamir, 2000). As a result, Dubey and colleagues investigated the cat as an
intermediate host for S. neurona (Dubey et al., 2000). That study provided the first
evidence for domestic cats as hosts for S. neurona; the life cycle of S. neurona was
completed through cats and opossums in an experimental bioassay.
Bioassays for Sarcocystis neurona in Cats
Sarcocyst formation was induced in four of five laboratory-raised cats dosed with
100,000-250,000 S. neurona sporocysts obtained from one naturally-infected opossum.
Three of these cats were immunosuppressed by intermittent corticosteroid therapy; one
cat not receiving corticosteroids developed sarcocysts. Sarcocysts that formed in cat
muscle were infective for four of four juvenile opossums; 250 g of cat muscle was fed to
each opossum. The opossums were killed 14 days after eating the infected cat muscle.
Sporocysts recovered from the opossums caused Sarcocystis-associated encephalitis in 12
gamma interferon knockout mice dosed orally with 250,000 sporocysts each. Mouse sera
tested positive for S. neurona antibodies by the S. neurona agglutination test (SAT).
Sarcocysts recovered from the cats were small (700 |tm or less) and none were mature;
they were composed primarily of metrocytes at 144 days postinfection. Sarcocyst
morphology showed slender villar projections containing microtubules. Bradyzoites
within the sarcocyst were 5 to 7 |tm long. Molecular analysis at the 25/396, and 33/54
loci (Tanhauser et al., 1999) of merozoite rRNA genes recovered from the brain of one
cat were consistent with S. neurona.
Turay and colleagues (2002) found S. neurona antibodies by immunoblot in one of
nine feral cats tested from Missouri. Muscle sarcocysts were detected, but not described
morphologically, in this single, seropositive cat. Muscle from the cat was fed to a
juvenile, laboratory-reared opossum, which then shed low numbers of sporocysts (-5000
total) 17 days postinfection that were infective for two, gamma interferon knockout mice
dosed with 300-500 sporocysts each. Merozoites isolated from the mice were used to
establish an in vitro culture. By immunoblot, antigenic variation among isolates of
S. neurona was demonstrated; serum from the infected cat reacted differently with
antigen prepared from each of two horse-derived S. neurona isolates. Molecular data
from the 25/396 and ITS-1 loci of the cat (opossum, mouse)-derived isolate showed it to
be similar to that from S. neurona. Because feral cats were evaluated, this study
suggested domestic cats might be naturally-infected with S. neurona.
In another set of laboratory bioassays, Butcher et al., 2002, characterized infections
induced in cats inoculated with horse-derived (UCD 1) or cat (opossum, mouse)-derived
(Mucat 2) isolates of merozoites harvested from cell cultures. Intravenous inoculations of
1 x 107 merozoites of either UCD 1 or Mucat 2 were given to two young cats. Two
similar cats each received 5 x 107 of the respective isolates by combined intravenous,
subcutaneous, and intramuscular route. Three gamma interferon knockout mice were
each injected intraperitoneally with 4 x 106 of either the UCD 1 isolate or the Mucat 2
isolate to ensure that the inoculums given to cats were viable. Cats were terminated six to
seven weeks postinoculation. Each of five, laboratory-raised opossums received tissue
corresponding to an individual cat. Results from this study demonstrated biological
diversity between the Mucat2 isolate and the UCD 1 isolate. Two of the cats had
developed sarcocysts; each of these cats was inoculated with Mucat 2. One opossum shed
a small number (-200) of sporocysts 23 days postfeeding; this opossum received cat
muscle from a cat inoculated with Mucat 2. Mice inoculated with Mucat2 merozoites or
Mucat2 (opossum) sporocysts developed encephalitis. The UCD 1 isolate did not appear
to be biologically active. In addition to the data provided for biological diversity of
isolates, this study established that life-cycle stages other than sporocysts of (presumed)
S. neurona could be infective to cats.
A series of experiments evaluating domestic cats as hosts for S. neurona was
reported in a single publication by Dubey and colleagues in 2002 (Dubey et al., 2002).
Sera collected from cats in the first life-cycle completion study (2000) were evaluated by
SAT at 0, 7, 20, 35, 49, and 144-167 days postinfection. For the next experiment, sera
collected from three cats dosed subcutaneously with 1 x 106 or more merozoites (isolated
from the brains of gamma interferon knockout mice that had received sporocysts from
naturally-infected opossums) were evaluated at 0, 3, 5, 8, 9, and 14 weeks
postinoculation. These cats were killed and muscle tissue was fed to five opossums, two
laboratory-raised dogs, and two laboratory-raised cats. Antibodies to S. neurona were
found in sporocyst-inoculated cats by 21 days postinfection; antibody titers were 5 to 50
times higher in cats that had received corticosteroid therapy. Sarcocystis neurona
antibodies developed in cats that received merozoites by the subcutaneous route. One of
five opossums fed cat tissue shed sporocysts; for the first time it was possible to produce
sporocysts derived from culture-grown merozoites. Neither dogs nor cats fed cat muscle
shed sporocysts, suggesting cats and dogs are not definitive hosts for S. neurona.
Serological Surveys for S. neurona Antibody in Domestic Cats
Two serological surveys for S. neurona antibody in domestic cats have been
reported. The first utilized the SAT to evaluate the seroprevalence of S. neurona antibody
in 112 sera from cats in Ohio (Stanek et al., 2001). Seventy-six cats were classified as
free roaming. The additional 36 cats came specifically from horse farms. Sarcocystis
neurona antibodies were found in 26 of 112 (23%) of the cats overall, and in 14 of 36
(39%) of the cats from horse farms. Rossano and colleagues (2002) evaluated the
prevalence of S. neurona antibody in 196 cats that had been previously tested for
Toxoplasma gondii antibodies. Both IFA and immunoblot were used in the study. By
immunoblot, 5% of the cats were seropositive. By IFA, 27% were seropositive. The
authors report the IFA test was subject to cross-reactivity when serum dilution was low
(1:20 or less); better agreement between IFA and immunoblot results was seen as
antibody titers increased.
MATERIALS AND METHODS
All animal procedures used were in accordance with protocols approved by the
University of Florida Institutional Animal Care and Use Committee.
Collection of Blood and Muscle Samples
Blood and muscle samples were collected from 50 adult cats euthanized at the local
animal shelter2. Immediately after euthanasia, cats were transported to the College of
Veterinary Medicine for processing. The thoracic cavity was opened, and 10 to 15 mL
blood was aspirated from the caudal vena cava. Clotted blood was centrifuged, and serum
harvested and stored. Tongue, diaphragm, and right quadriceps muscle were excised and
each was divided into three equal parts to be fixed in 10% neutral buffered formalin
solution, stored frozen at -80 C, or kept fresh at 4 C. In addition, blood samples were
collected by jugular venipuncture from 50 adult feral cats admitted to monthly trap-
neuter-return clinics 3 held at the University of Florida, College of Veterinary Medicine.
Examination for Sarcocysts
Three portions of formalin-fixed tissue were taken from each muscle sample,
paraffin-embedded, sectioned at 5-[tm intervals, mounted on slides, and stained with
hematoxylin and eosin (H&E). Stained sections were examined for the presence of
sarcocysts using a dissecting microscope at a magnification of 20x. When possible,
1 Alachua County Animal Services, Gainesville, Fla.
2 Operation CatnipTM
transverse and long dimensions of sarcocysts were obtained. In those cats with sarcocyst
infection, additional fresh muscle tissue was examined with a dissecting microscope at 10
to 40x, and sarcocysts dissected out and placed either in Hank's Balanced Salt Solution
(HBSS, Mediatech, Herndon, VA) for molecular characterization or Trump's fixative for
transmission electron microscopy (TEM). Photomicrographs were taken of both fixed
and fresh specimens. For TEM, samples were transferred to the Electron Microscopy
Core Laboratory (Biotechnology Program, University of Florida). There, samples were
further processed in 1% osmium tetroxide (w/v) and dehydrated in alcohol prior to
embedding in EmBed epoxy resin (Electron Microscopy Sciences Fort Washington, Pa).
Seventy to eighty nm thin sections were double stained in 2% aqueous uranyl acetate and
Reynolds lead citrate and examined with a Hitachi H-7000 transmission electron
microscope (Pleasanton, CA) at 75 keV. Digital photomicrographs were taken with a
Gatan Multiscan camera (Pleasanton, CA).
Molecular Characterization of Sarcocysts
A small portion of quadriceps muscle from a cat with no histological evidence of
sarcocyst infection or S. neurona antibody was used as a negative control for molecular
studies. This tissue, and samples of dissected sarcocysts in HBSS, were pelleted
individually by centrifugation, then mechanically disrupted by repeatedly pressing the
pelleted material against the sides and bottom of the tubes with pipette tips. In each of the
tubes, total genomic DNA was purified through the use of a QIAamp DNA Mini Kit
(QIAGEN Inc., Valencia, CA) and supplied protocol. The polymerase chain reaction
(PCR) was applied to cat sarcocyst DNA, cat muscle DNA (control), and DNA prepared
from culture-derived merozoites of the UFsn-1 S. neurona isolate (Ellison et al., 2002,
Long et al., 2002) and S. falcatula (ATCC 50701, Manassas, VA). Three PCR reactions
were performed. The first utilized the random amplified polymorphic DNA
(RAPD)-derived primer pair JNB25/JD396 used by Tanhauser et al. (1999) to amplify
and differentiate DNA from select species of Sarcocystis. The second reaction was
performed with primer pair JD26/JD37 to amplify a region of the small subunit ribosomal
RNA (ssurRNA) gene (Dame et al., 1995). The third reaction used JNB69/JNB70
primers to amplify the internal transcribed spacer region 1 (ITS-1) of the rRNA gene
(Tanhauser et al., 1999). PCR products were separated by agarose-gel electrophoresis
incorporating ethidium bromide and photographed under ultraviolet light.
Amplified ssurRNA gene DNA, corresponding to two separate samples of cat
sarcocyst, and one sample of cat muscle (control), were purified prior to sequencing by
cutting bands of interest from the agarose gel and utilizing a StrataPrepTM DNA Gel
Extraction Kit (Stratagene Inc., La Jolla, CA) and supplied protocol to remove agarose
and contaminants. The ssurRNA sequencing primers were the same as those used for
Two samples of amplified cat sarcocyst DNA from the ITS-1 region were cloned
directly into the pCR2.1-TOPO vector (Invitrogen, Carlsbad, CA), and five positive
colonies were sequenced with internal, nested primers JNB68/JNB71. ABI Prism Big
DyeTM Terminator Cycle Sequencing Ready Reaction kits (PE Biosystems, Foster City,
CA) and an Applied Biosystems 377 automated sequencing system (Applied Biosystems,
Foster City, CA) were used for sequencing reactions. The BLAST program (Altschul et
al., 1997) was used for to search for sequence homologies. Sequence alignments were
done using CLUSTALW (version 1.8, Thompson et al., 1994) software and nonweighted
parameters and GCGs GAP (Genetics Computer Group, Madison, WI) software
utilizing end-weighted parameters. An unrooted phylogenetic tree was plotted from
aligned ITS-1 sequences with CLUSTALW incorporating the neighbor-joining (N-J)
method to illustrate phylogenetic relationships based upon percent divergence (distance)
of sequence. Sequence data for cat sarcocyst ssurRNA and two representative clones of
ITS-1 DNA were submitted to GenBankTM, accession numbers AY190080, AY190081,
and AY190082 respectively.
A mature SPF cat, born and raised indoors, and fed only processed commercial
food, was used to provide control serum for immunoblot tests. Prior to inoculation with
killed S. neurona merozoites, the cat was anesthetized under isoflurane/02 anesthesia,
and 50 mL blood collected by jugular venipuncture. Serum was then obtained and frozen
at -80 C until further use. Tissue-culture-grown merozoites from the UFsn-1 isolate of
S. neurona, were harvested from culture flasks and purified by differential centrifugation.
Merozoites were then killed by a 30-minute incubation in a 60 C water bath, rinsed in
sterile 0.9% saline solution, and counted on a hemacytometer. For each inoculation, 1 x
105 merozoites were suspended in 0.5 mL sterile 0.9% saline and injected
intramuscularly. Three inoculations were given, each two weeks apart. Seven days after
the 3rd inoculation, 50 mL blood was collected under anesthesia, serum harvested, and
stored frozen at -80 C. Pre and postimmunization serum samples were sent to the EPM
Diagnostic Laboratory at Neogen Corp, Lexington, KY for immunoblot analysis for
antibodies against S. neurona.
The immunoblot procedure, typically performed on equine serum, had previously
been adapted for use with armadillo serum (Tanhauser et al., 2001). The procedure was
further modified for use with cat serum by substituting Protein A/G-peroxidase conjugate
in place of Protein G-peroxidase conjugate (both: Pierce, Rockford, IL). Per Neogen
Corp., immunoblotting techniques are as follows: Culture-grown S. neurona merozoites,
originally derived from a horse with EPM, were solubilized and separated by sodium
dodecyl sulfate polyacrylamide gel electrophoresis (SDS PAGE) on 4 to 20% tris-glycine
gels (Invitrogen, Carlsbad, CA) incorporating Invitrogen Multimark molecular weight
standards. Gels were then placed in an electrophoretic transfer device (BioRad, Hercules,
CA) for protein transfer to 0.2 |tm nitrocellulose membranes (Invitrogen). Membranes
were blocked overnight with blotto (phosphate buffered saline, 5% w/v non-fat dried
milk, 1% antifoam solution, 0.1% sodium azide) then placed in a miniblotter apparatus
(Immunetics, Cambridge, MA). Serum samples, and positive and negative controls, were
diluted 1:10 in blotto and added to the template lanes. Following a 90-minute incubation
at room temperature, the blot was removed from the template and washed 4 times in
phosphate-buffered saline containing 0.1% Tween-20 (PBST). Protein A/G-peroxidase
conjugate, diluted in PBST, is added to the blot for 60 min then washed off, and the blot
developed by the addition of 3, 3', 5, 5' tetramethylbenzidine (TMB) substrate (Vector
Laboratories, Burlingame, CA). Developed blots were dried, interpreted by examination,
and in selected cases, scanned with a BioRad GS-700 imaging densitometer to quantify
Serum samples collected from 50 cats examined for sarcocysts, 50 feral cats from
trap-neuter-return programs, and control cat sera were assayed. Negative controls
included preimmune cat serum and S. neurona-negative horse serum. Immune cat serum
and horse serum, both strongly and weakly reacting, were used as positive controls.
In order to learn if sarcocyst infection might be linked to immunosuppression, cat
sera were tested for immunosuppressive retroviral infections. A commercially available
kit (SNAP FIV Antibody/FeLV Antigen Combo Test kit, IDEXX Laboratories,
Portland, ME) was used to test all serum samples for feline leukemia virus (FeLV)
antigen and feline immunodeficiency virus (FIV) antibody.
A laboratory-raised opossum, fed only a commercial laboratory diet since weaning,
was fed fresh muscle from five cats infected with sarcocysts. Prior to feeding cat muscle,
opossum feces were examined microscopically for the presence of sporocysts using the
Sheather's sugar flotation technique. Muscle from each of the five cats was chopped,
mixed together with commercial, canned cat food and fed daily for four days.
Approximately 30 g of muscle were fed in total. Seven days after the last feeding of cat
muscle, and three times weekly thereafter, opossum feces were checked for sporocysts.
Thirty days after the last feeding of cat muscle, the opossum was euthanized with an
overdose of sodium pentobarbital (Euthasol, Delmarva Laboratories, Midlothian, VA).
The small and large intestines were removed and split along their length with sterile
scissors. A glass microscope slide was used to scrape epithelia and intestinal contents
from the intestinal lumen. Scrapings were collected into a sterile blender cup and
homogenized for 1 minute. The homogenate (approximately 35 mL) was transferred to a
sterile 500 mL flask, 400 mL of deionized water was added, and the mixture stirred for 4
hours. A portion of the mixture was strained through a tea strainer into two centrifuge
tubes. Tubes were centrifuged at 2330 x g for 10 minutes, and the supernatant decanted.
Tubes were filled with Sheathers sugar solution (specific gravity 1.27), and a coverslip
applied to the top of the tubes. Tubes were centrifuged at 2330 x g for 10 minutes.
Coverslips were then applied face down onto glass slides and examined at 100x for the
presence of sporocysts.
Samples of sarcocysts from each of the cats infected with sarcocysts were dissected
from muscle and each placed in 0.5 mL Hank's Balanced Salt Solution (HBSS)
containing 200 U penicillin, 0.2 mg streptomycin, and 0.5 |tg amphotericin B (Antibiotic
Antimycotic Solution, Sigma, St. Louis, MO) per mL. Samples were rinsed twice in this
solution by centifugation at 5000 x g. After the final rinse, a sterile pipette tip was used to
rupture sarcocysts and release bradyzoites. Bradyzoite suspensions were transferred to
individual 25 cm2 flasks containing 75% confluent monolayers of bovine turbinate (BT
1390, ATCC CRL 1390) cells in Dulbecco's Modified Eagle's Medium (DMEM,
Mediatech, Herndon, VA) supplemented with 10% heat-inactivated fetal bovine serum
(Atlas Biologicals, Ft. Collins, CO), and 10 mM HEPES buffer, 2 mM L-glutamine, 100
U penicillin and 0.1 mg streptomycin (each, Mediatech, Herndon, VA) per mL. Twice
weekly, two thirds of the media was aspirated and replaced with fresh media. At 16 and
32 weeks postinfection, cultures were trypsinized with 2mL 0.05% trypsin-EDTA
solution (Mediatech, Herndon, VA) and one third of the cells transferred to a new 25cm2
flask. Cultures were monitored microscopically at 100 and 200x for evidence of
monolayer infection. Cultures were monitored for 60 weeks postinfection.
Molecular Analysis of Fixed Tissue from Florida Panther Containing Sarcocysts
Hematoxylin and eosin stained sections from a 1988 1989 study by Greiner and
colleagues (Greiner et al., 1989) of muscle sarcocysts in Florida panthers were
re-examined microscopically at 40 100x for the presence of sarcocysts. The
formalin-fixed, paraffin-embedded tissue block for panther # 07 tongue was chosen for
molecular analysis as microscopic examination of sections obtained from this specimen
showed this it to contain more sarcocysts (approximately 10) than other specimens
DNA extraction from formalin-fixed, paraffin-embedded tongue was performed.
Eight samples were prepared. For each sample, five slices were shaved from the tissue
block, totaling approximately 50 mg, and placed in a microcentrifuge tube with 800 [l of
xylene. The tube was vortexed for five seconds, and 400 [l ethanol then added. The tube
was again vortexed for five seconds, then centrifuged five minutes at 10,000 x g, and the
supernatant decanted. This de-paraffinization process was repeated twice. After the final
rinse, the tube was inverted and allowed to dry for 20 minutes before 250 [l of lysis
buffer (1% sodium dodecyl sulfate, 100 mM EDTA, 50 mM Tris-HC1, 100 mM NaC1)
were added to the tube, along with 62.5 |tg of proteinase K (Research Products
International, Mt. Prospect, IL) and incubated overnight at 55C. The following day, an
additional 62.5 |tg of proteinase K were added and incubated one hour at 65C. After
cooling to room temperature, and equal volume (375 [Il) of 4M ammonium acetate was
added and the tube vortexed. Twice the volume (750 [Il) of isopropyl alcohol was then
added to the tube to precipitate the DNA. The mixture was cooled to 4C and centrifuged
10 minutes at 14,000 x g. The supernatant was decanted and pellet rinsed twice by
centrifugation at 14,000 x g with 800 [tl of cold (-20C) ethanol. After the final rinse, the
supernatant was decanted and the tube inverted over absorbent paper to dry the pellet.
The pellet was resuspended in 20 [tl of molecular grade water (Invitrogen, Carlsbad, CA)
and allowed to dissolve overnight. Prior to analysis by PCR, the solution was pooled to
form two aliquots and each centrifuged five minutes at 10, 000 x g. Supernatant from one
of the tubes was further purified by standard phenol/chloroform extraction followed by
3M sodium acetate/cold ethanol precipitation of DNA.
PCR with primer pair JD26/JD37 was used for amplification of a region of the
ssurRNA gene (Dame et al., 1995). DNA prepared from culture-derived merozoites of
the UFsn-1 S. neurona isolate (Ellison et al., 2002, Long et al., 2002) and S. falcatula
(ATCC 50701, Manassas, VA) were used as positive controls, and water as negative
control. PCR products were separated by low-melt agarose-gel (SeaPlaque GTC,
Biowhittaker Molecular Applications, Rockland, ME) electrophoresis incorporating
ethidium bromide and photographed under ultraviolet light. The resultant ssurRNA gene
amplicon was cut from the gel, melted, and purified prior to sequencing with a
QiaQuickTM PCR Kit (Valencia, CA) and supplied protocol. ABI Prism Big DyeTM
Terminator Cycle Sequencing Ready Reaction kit (PE Biosystems, Foster City, CA) and
an Applied Biosystems 377 automated sequencing system (Applied Biosystems, Foster
City, CA) were used for sequencing reactions. The ssurRNA sequencing primers were the
same as those used for amplification. The BLAST program (Altschul et al., 1997) was
used for to search for sequence homologies.
Histologic examination of muscle samples revealed sarcocysts in 5 of 50 cats
(10%). Sarcocysts were most abundant in sections of quadriceps muscle. No
inflammatory reaction was seen in tissue surrounding the sarcocysts. In some instances,
sarcocysts were visible grossly in fresh muscle. Sarcocysts appeared filamentous,
sometimes convoluted, and measured approximately 0.1 x 0.5 20 mm (width x length)
(Figs. 4-1 and 4-2). Bradyzoites from fresh sarcocysts measured 3.3 x 8.8 |tm (width x
length) (Fig. 4-3). Additional photomicrographs from light microscopy are shown in
Figures 4-4 and 4-5.
Figure 4-1. Photograph of sarcocyst
(arrow) in cat skeletal muscle, 16x.
Some sarcocysts were visible grossly
and measured 0.1 x 0.5 to 20 mm (width
Portion of sarcocyst located in quadriceps muscle of cat. Hematoxylin and
eosin stain. Scale bar =100 jtm.
- i .
j,. ~ :r: A
~~ ~~ %9 r
Figure 4-3. Photomicrographs from light microscopy of fresh sarcocysts obtained from
cat muscle. A) convoluted, "wavy" shaped sarcocyst filled with bradyzoites
and surrounded by bradyzoites that have spilled out of the ruptured
sarcocyst, 160x. B) close up of same (400x); fresh bradyzoites measure 8.8
x 3.3 pm.
WP- -W -
Figure 4-4. Photographs taken from light microscopy of H & E stained sections of
cat skeletal muscle containing sarcocysts. A) 297 x 113 |tm (160x). B)
500 x 160 itm (100x).
Figure 4-5. Photomicrographs from light microscopy of the sarcocyst wall of A)
fixed, H & E stained and B) fresh sarcocysts obtained from cat muscle,
Images from TEM (Figs. 4-6 and 4-7) show a thin (maximal thickness
approximately 2.5 [tm), relatively simple, sarcocyst wall with short, irregular, villar
protrusions with rounded tips. The electron-dense layer underlying the parasitophorous
vacuolar membrane was interrupted at intervals, especially in areas at the base of, or
between, villi. The ground substance was devoid of microtubules, even as it formed villar
protrusions, and was continuous with the septa that run between bradyzoites and
metrocytes. Bradyzoites measured approximately 6 8 x 1.5 3 |tm. The anterior region
of the bradyzoites contained numerous micronemes. Inclusion bodies, including
amylopectin, lipid, and electron-dense granules, were found in the center region of the
bradyzoites, and cell organelles, including the nucleus, were confined primarily to the
posterior regions. Sarcocyst morphology was compared to published descriptions of
sarcocysts found previously in both domestic cats and wild felid species. Based upon
these comparisons, it was determined that sarcocysts found in this study were Sarcocystis
S. ; Figure 4-6. Photomicrograph
S? from TEM of cat sarcocyst.
Sarcocyst wall shows
S. interruptions of the electron-
V dense layer (EDL, arrow)
V underlying the parasitophorous
EDL -- ED
.GS particularly at the base of, and
t '. Between, villi (V). Villi and
ground substance (GS) lack
microtubules. Bradyzoites (B,
arrow) contain micronemes
(MN), amylopectin granules
(A), electron-dense bodies
(EDB, arrow), and nucleus (N).
Septa (S) separate bradyzoites
within the sarcocyst.
Figure 4-7. Photomicrograph from TEM of cat sarcocyst. Villi (V) have rounded tips
and contain no microtubules. "Hobnailed" appearance of electron dense
layer (EDL) is evident. Ground substance (GS) and a bradyzoite (B, arrow)
are also shown. Bradyzoite contains amylopectin granules (A), electron
dense bodies (EDB) and micronemes (MN). Scale bar = 1 |tm.
Amplification of cat sarcocyst DNA or cat muscle DNA was not seen when PCR
was performed with RAPD-derived primer pair JNB25/JD396. PCR using JD26/JD37
(ssurRNA) primers yielded single PCR products of approximately 410 and 450 bp from
cat sarcocyst, and cat muscle samples, respectively (Fig. 4-8).
Figure 4-8. Agarose-gel
electrophoresis showing bp
results of PCR using 1500
ssruRNA gene primer 1000
pair JD26/JD37 and
template DNA. Lanes
1-5, sarcocyst DNA 400
from cats 1-5 (- 410 300
bp). Lane 6, DNA from 200
uninfected cat muscle too
(- 450 bp). Lane 7, S.
neurona DNA (- 410
bp). Lane 8, S. falcatula
DNA (- 410 bp). Lane
9, water control.
Fragment size ladder (100 bp) included on both sides of gel; ladder legend
(bp) shown at right of photograph. Additional band above primary band in
lane 4 (cat 4) most likely indicates muscle contamination of sarcocyst DNA.
JNB69/JNB70 (ITS-1) amplified an approximately 1080-bp product from cat
sarcocysts and an approximately 1200-bp product from both S. neurona and S. falcatula
DNA prepared from culture-derived merozoites. Uninfected muscle was not amplified
(Fig. 4-9). PCR results were consistent for each of the five samples of cat sarcocyst.
bp Figure 4-9. Agarose-gel
electrophoresis showing results
of PCR using ITS-1 primer pair
JNB69/JNB70 and template
1000 DNA. Lanes 1-5, sarcocyst DNA
from cats 1-5 (- 1080 bp). Lane
500oo 6, DNA from uninfected cat
muscle. Lane 7, S. neurona DNA
300 (- 1200 bp). Lane 8, S. falcatula
DNA (- 1200 bp). Lane 9, water
control. Fragment size ladder
(100 bp) included on both sides
of gel; ladder legend (bp) shown
left of photograph.
Sequence data from the ssurRNA gene of cat sarcocysts showed significant identity
with archived ssurRNA sequence from many Eimeriidea organisms, particularly
S. neurona (GenBank U33149) and S. falcatula (GenBank U35075). By CLUSTALW
and GAP, both of these organisms have a 99% identity with cat sarcocyst ssurRNA
sequence. Archived ssurRNA sequence was available for 6 of the reported 15 species of
Sarcocystis that use a felid definitive host. When compared to ssurRNA sequence
generated from cat sarcocysts, the following identity-based, global alignment scores were
obtained from CLUSTALW for these definitive host species: S. muris 98% (GenBank
M64244), S. hirsuta 92% (GenBank AF017122), S. buffalonis 92% (GenBank
AF017121), S. fusiformis 92% (GenBank U03071), S. gigantea 91% (GenBank L24384),
and S. rodentifelis 82% (GenBank AY015111). After accounting for priming losses and
terminal sequences of 18srRNA (40 bp) and 5.8srRNA (30 bp) DNA, approximately 800
bp of ITS-1 sequence was generated from each of the five cat sarcocyst DNA clones.
Alignment of these cloned ITS-1 sequences showed limited base switching among
clones; by GAP, there was 98% identity of sequence among clones. Cloned cat sarcocyst
DNA from the ITS-1 region showed only 45% identity with S. neurona (UCD1 isolate,
GenBank AF081944) and 46% with S. falcatula (Florida 1 isolate, GenBank AF098244)
sequence when aligned with GAP. Pairwise, global alignment scores from CLUSTALW
for ITS-1 region sequence of cat sarcocyst were only 6% for S. neurona (UCD1 isolate,
GenBank AF081944), 11% for S. falcatula (Florida 1 isolate, GenBank AF098244), 4%
for Hammondia heydorni (CZ-3 isolate, GenBank AF317281), 12% for Toxoplasma
gondii (ME49 isolate, GenBank L49390), and 5% for Neospora caninum (Liverpool
isolate, GenBank L49389). An unrooted, phylogenetic, N-J tree (Fig. 4-10), shows the
evolutionary relationship of ITS-1 sequence from these organisms and that from cat
S. falcatula Toxoplasma
Figure 4-10. Unrooted phylogenetic tree showing the divergence of Sarcocystisfelis
ITS-1 gene sequence to that from Sarcocystis neurona, Sarcocystis
falcatula, and organisms from related genera: Toxoplasma gondii,
Hammondia heydorni, and Neospora caninum. Tree was plotted from
CLUSTALW, incorporating the neighbor-joining (N-J) algorithm.
Inoculation of a cat with the UFsn-1 isolate of S. neurona resulted in
seroconversion against 17-kDa and 30-kDa proteins of S. neurona, shown by immunoblot
(Fig. 4-11). The 17-kDa band had the same apparent molecular mass as an
S. neurona-specific band recognized by equine serum. Pre-immunization cat serum was
negative for reactivity with the 17-kDa antigen. Of the 50 serum samples collected from
cats examined for sarcocysts, 2 samples (4%) were positive by immunoblot for antibody
to S. neurona. One of five cats harboring sarcocysts was seropositive by immunoblot.
Immunoblot testing of the 50 sera collected from feral cats brought to trap-neuter-return
clinics resulted in 3 positives (6%). Combined, 5/100 (5%) cat sera evaluated in this
study were seroreactive with S. neurona.
1 4 5 6 7 8 10 11 12 13141516 1718 1920212223242526
4- 17 kD band (specific)
* 17 kD band (specific)
Immunoblots of cat serum. A) Blot of pre-immune cat serum. Lane 1: no
sample; blotto only. Lane 2: strongly positive horse serum from horse
infected with S. neurona. Lane 3: weakly positive horse serum from horse
infected with S. neurona. Lane 4: horse serum from non-infected animal.
Lane 5: serum from cat prior to immunization with S. neurona. Lane 6: no
sample, blotto only. B) Blot of post-immunisation cat serum. Lane 1: no
sample; blotto only. Lane 2: strongly positive horse serum from horse
infected with S. neurona. Lane 3: weakly positive horse serum from horse
infected with S. neurona. Lane 4: horse serum from non-infected animal.
Lane 5: serum from cat post-immunization with the UFsn-1 isolate of S.
neurona. Lane 6: no sample; blotto only. C) Molecular weight standards
overlay lanes 1-3, which contain blotto solution only. Lane 4: strongly
positive horse serum from horse infected with S. neurona. Lane 5: weakly
positive horse serum from horse infected with S. neurona. Lane 6: horse
serum from non-infected animal. Lanes 7-25: serum from feral cats. Lane
26 contains blotto solution only. Cat serum added to lanes 14 and 15 is
considered positive for antibody to S. neurona.
FeLV antigen and FIV antibody testing of serum collected from the 50 cats
euthanized at the local shelter resulted in 2 cats positive for FeLV antigen (4%), and 2
different cats positive for FIV antibody (4%). One cat positive for FeLV antigen had
sarcocyst infection. No FIV antibody-positive cats had sarcocysts. No FeLV or FIV-
positive cats from this group were positive for antibody to S. neurona. Testing of the
serum from the 50 additional feral cats yielded 2 cats (4%) positive for FeLV antigen and
a different cat (2%) positive for FIV antibody. One cat positive for FeLV antigen was
also positive for antibody to S. neurona. Therefore, of the 100 cats evaluated in this
study, 4 were positive for FeLV antigen (4%), and 3 for FIV antibody (3%). Results are
shown in Tables 4-1 and 4-2 below.
Table 4-1. Results of serology and histology for 9 of 50 cats euthanized at the local
S. neurona F
Cat ID Specific Ag FIV Ab Sarcocysts Sex / Status Source
56710 Pos Neg Neg Pos F, pregnant Stray
57559 Neg Neg Neg Pos M, intact Stray
57605 Neg Neg Neg Pos M, intact Stray
58004 Neg Pos Neg Pos M, intact Stray
57270 Neg Neg Neg Pos M, intact Stray
56709 Pos Neg Neg Neg M, intact Stray
57248 Neg Pos Neg Neg F, intact Stray
57561 Neg Neg Pos Neg M, intact Stray
57601 Neg Neg Pos Neg M, intact Stray
Data for cats not shown was negative.
Table 4-2. Results of serology for 8 of 50 cats taken to trap-neuter-return clinics
S. neurona Fe
Cat ID Specific Ag FIV Ab Sex Source
FO-455 Pos Pos Neg M Trapped
FO-514 Pos Neg Neg M Trapped
FO-505 Pos Neg Neg F Trapped
FO-564 Neg Pos Neg F Trapped
F9-1519 Neg Neg Pos M Trapped
FO-192 Neg Neg Neg F Trapped
FO-205 Neg Neg Neg F Trapped
F9-1416 Neg Neg Neg M Trapped
Data for cats not shown was negative.
Sporocysts were not found in opossum feces that were collected after feeding
sarcocyst-infected cat muscle, and sporocysts were not found in intestinal mucosal
scrapings from the opossum.
No evidence of infection was noted in bovine turbinate cell cultures inoculated with
bradyzoites. Cultures were discarded 60 weeks post-inoculation.
PCR of DNA isolated from formalin-fixed Florida panther tongue yielded a faint
band of approximately 410 bp. Positive control DNA yielded bands of equivalent sizes
1 2 3 4 5 6 7 8
Agarose-gel electrophoresis showing results of PCR using ssruRNA gene
primer pair JD26/JD37 and DNA prepared from formalin-fixed, paraffin-
embedded Florida panther tongue containing sarcocysts. Lane 3, product
amplified from formalin-fixed, paraffin-embedded Florida panther tongue
containing sarcocysts (- 410 bp). Lane 5, S. neurona DNA (- 410 bp).
Lane 6, S. falcatula DNA (- 410 bp). Lane 7, water control. Lanes 2 and 4
are empty, done to minimize DNA contamination when cutting band from
lane 3. Fragment size ladder (100 bp) included on both sides of gel in
lanes 1 and 8; ladder legend (bp) given at right of photograph.
Approximately 360 nucleotides of sequence were generated and submitted for
BLASTn analysis. Sequence homology data from BLAST found the product amplified
from the Florida panther tissue locally homologous with ssurRNA sequence from plant
material, most notably (89%) from the genus Pinus (pine). No homologies to other
Sarcocystis or protozoal organisms were found.
Because of the importance of S. neurona as an equine pathogen, there has been an
active effort to identify relevant hosts for this organism. As a result of the work described
here, it was discovered that sarcocysts were fairly common in cats; however, on the basis
of careful morphological, molecular, and biological characterization, these sarcocysts
could not be classified as S. neurona, or even S. neurona-like.
Our study is the largest reported survey for sarcocysts in domestic cats; five of 50
(10%) cats had sarcocysts. Previously, sarcocysts were found in four of 12 cats in Indiana
(Everitt et al., 1987) and one of nine cats in Missouri (Turay et al., 2002). In another
study, sarcocysts were collected from muscle biopsies of five cats, however no
prevalence data were provided (Fiori et al., 1988). Higher prevalence of sarcocysts has
been reported from surveys of wild felid species. In Florida, 50% of bobcats (Anderson et
al., 1992), and 83% of free-ranging Florida panthers and cougars (Greiner et al., 1989)
were infected. Sixty-six percent of Arkansas bobcats (Dubey et al., 1992) and 70% of
captive born and raised cheetahs (Briggs et al., 1993) had sarcocysts. Sarcocysts were
found in 100% of adult, wild lions from Namibia, but not in juveniles, suggesting that
infection increases with maturity (Kinsel et al., 1998). Sarcocysts are prevalent in Felidae
and appear to have a worldwide distribution.
The population of cats we examined for sarcocysts included (few) surrendered pet
cats, free-roaming cats, socialized strays, and feral cats and thus may be expected to have
a sarcocyst prevalence somewhere between that found in wild felids and that found in
domestic cats kept indoors. In the wild, increased opportunities exist for cats to ingest
sporocysts from definitive host(s); environment and diet no doubt contribute to the
increased prevalence of sarcocyst infection in wild species. Captive wild species are
typically fed raw meat and offal, potentially exposing them to viable sporocysts.
Although the cats in the present study received unknown levels of human care, one may
assume some had access to processed foods, shelter, and preventative health care for at
least part of their lives; factors that may explain the comparatively low prevalence of
sarcocysts found in domestic cats. Data supplied by the local shelter indicated all five
infected cats were listed as "strays", evidence consistent with the notion that stray or feral
cats may be more likely to be infected.
General health and immunity may also be factors for sarcocyst infection. Anderson
and colleagues (1992) could not conclude that the high rate of sarcocyst infection among
bobcats was linked to specific viral immunosuppressive diseases, however, Greiner et al.
(1989) found that 78% of Florida panthers were seropositive for feline panleukopenia
virus, not an immunosuppressive virus on its own, but one that might contribute to poor
health (Dr. Julie Levy, University of Florida, personal communication) and 30% were
positive for FIV. Both Florida panthers and captive populations of cheetahs are reported
to be inbred and to lack genetic diversity, possibly contributing to poor immunity. In a
survey of FeLV and FIV infection in 1143 free-roaming domestic cats sampled from the
same geographical area as those from this study, incidence of FeLV antigen and feline
FIV antibodies was 3.7% and 4.3%, respectively (Lee et al. 2002). In our study, the
apparent prevalence of FeLV was 4% and FIV 3%. Feline leukemia virus antigen was
detected in one of five cats positive for sarcocysts and one different cat positive for
antibody to S. neurona; however, the low prevalence of these potentially
immunosuppressive retroviral infections makes it difficult to draw a conclusion regarding
any association with Sarcocystis infection.
In both domestic cats and wild felid species, sarcocyst morphology has been
described incidental to examination of muscle tissue collected at necropsy from
debilitated and healthy animals. In these reports, histologic examination revealed
sarcocysts of varying sizes: from 24 |tm (Kirkpatrick et al., 1986) to 270 |tm (Everitt et
al., 1987) in diameter by 24 |tm (Everitt et al., 1987) to 2100 |tm in length (Dubey et al.,
1992). The ultrastructure of these sarcocysts was virtually identical in all cases: rounded,
irregularly-spaced villi devoid of microtubules, a regularly interrupted electron-dense
layer underlying the parasitophorous membrane, and septae separating bradyzoites.
Bradyoites measured approximately 10 |tm long. It was noted that ultrastructural
morphology was not consistent with descriptions of sarcocysts for which the cat is known
to serve as a definitive host (Kirkpatrick et al., 1986). Citing the morphological similarity
illustrated for felid sarcocysts, Dubey and colleagues (1992) formally identified them as
Based upon a descriptive classification of Sarcocystis species (Dubey et al., 1989),
the sarcocyst wall of S. felis has been classified as a type 4 (Odening, 1998) or a type 9
without villar microtubules (Dubey et al., 1992). The ultrastructural morphology of
S. neurona sarcocysts has been described for infections in cats (Dubey et al., 2000,
Dubey et al., 2001c), armadillos (Cheadle et al., 2001a), raccoons (Dubey et al., 2001a),
and skunks (Cheadle et al., 2001b). Sarcocysts were less than 200 |tm in length; the
sarcocyst wall was comprised of long, slender villi containing microtubules, consistent
with a classification of type 11 or 12 (Dubey et al., 1989). Bradyzoites were also slender
and 5-7 |tm long. In another report, sarcocysts induced in cats inoculated with S. neurona
or S. neurona-like merozoites were examined by light microscopy only;
photomicrographs show sarcocysts measuring 20 rtm x 140 |jm (Butcher et al. 2002).
Photomicrographs from TEM of the sarcocyst wall of S. neurona obtained from skunk
muscle (Cheadle et al., 2001b) are very different than photomicrographs of TEM from cat
sarcocysts shown in Figs. 4-6 and 4-7.
Sarcocysts seen in the present study are different than S. neurona, but closely
resemble S. felis. Both histologic sections and fresh tissue were examined (Figs. 4-1
through 4-5). Only one previous report (Greiner et al., 1989), describes sarcocysts in
fresh tissue (muscle squash preparations) and reports their length to be comparable (1
cm) to some found in this study. In fresh tissue, some sarcocysts seen in this study were
large enough to be visible grossly, up to 2 cm in length, and many appeared to have a
convoluted or wavy shape.
Compared to other S. neurona intermediate hosts, domestic cats, as companion
animals, are under close observation by their owners and sent in great numbers to
veterinarians for evaluation of health problems, yet few reports in the veterinary literature
describe sarcocystosis, disease associated with widespread sarcocyst infection, or
Sarcocystis-associated neurological disorders in domestic cats. Just one reference
describes encephalitis in domestic cats caused by infection with a Sarcocystis spp.,
(Dubey et al. 1994). A second citation (Dubey and Hamir 2000), further characterizes the
organism found, and identifies it as S. neurona. In experiments designed to demonstrate
and confirm the domestic cat as an intermediate host for S. neurona, cat subjects, even
when dosed with corticosteroids for immune suppression, did not become diseased
(Dubey et al. 2000, Butcher et al. 2002, Turay et al. 2002, Dubey et al. 2002). However,
in similar experiments, both skunks (Cheadle et al. 2001) and raccoons (Dubey et al.
2001a) became diseased when dosed with S. neurona sporocysts. Naturally-occurring
encephalitic infections with S. neurona have been documented in raccoons, skunks, mink,
and sea otter (reviewed in Dubey et al. 2001d). Considering the comparatively huge
numbers of domestic cats, one might expect a greater incidence of morbidity and
mortality associated with S. neurona infection if cats are indeed an important natural
intermediate host for this parasite.
The primer pair JNB25/JD396 does not amplify DNA from all Sarcocystis spp.
(Tanhauser et al., 2001), but readily amplifies both S. neurona and S. falcatula. Lack of
amplification of cat sarcocyst DNA by these primers suggests the sarcocysts found in cat
muscle do not belong to either of these two species. Comparisons among conserved
ssurRNA gene sequence did not readily differentiate cat sarcocyst (S. felis) DNA found in
this study from several other Sarcocystis species, including S. neurona. However, S. felis
ssurRNA gene sequence was not similar to that for Sarcocystis species that use the cat as
definitive host, suggesting sarcocysts seen were not due to an aberrant infection with
these definitive host species.
Based upon a local alignment of 316 nucleotides, ITS-1 sequence analysis was used
to support the conclusion that raccoons are natural intermediate hosts of S. neurona
(Dubey et al., 2001a). Although clearly belonging to Sarcocystis, and 99% similar to
S. neurona ssurDNA, S. felis sarcocyst DNA sequence was quite dissimilar over the
ITS-1 region compared to S. neurona and S. falcatula. Internal transcribed spacer
region 1 primers amplified a smaller-sized product from S. felis sarcocysts than from
cultured S. neurona or S. falcatula merozoites; restriction endonuclease digest of PCR
products, typically done to differentiate species at this locus by PCR alone (Tanhauser et
al. 1999, Turay et al. 2002), was therefore unnecessary. Local alignments of ITS-1
sequence from cat sarcocyst against archived ITS-1 sequences was done by BLAST.
Alignment with many Eimeriidea organisms, most notably S. falcatula (AF389339) and
S. lindsayi (AF387164) occurred only in conserved regions of approximately 40
nucleotides (nt) from the 3' end of the 18srRNA gene and 30 nt from the 5' end of the
5.8S gene. Sequence data for the ITS-1 region of the S. felis was completely non-
homologous with that from S. falcatula or two isolates of S. neurona when conserved
regions were trimmed from the sequence. Although referred to by Marsh et al. 1999,
ITS-1 sequence for S. gigantea and S. muris, definitive host species of the cat, are not
available within the public domain for comparison. The divergence of ITS-1 sequence
from S. felis sarcocysts to those from related coccidian species, even those of the same
genus, is evident in the phylogenetic tree (Fig. 4-10).
Once S. felis was identified as the source of infection found in cats from this study,
an attempt was made to compare ssurRNA genes of sarcocysts in these cats to those from
sarcocysts found previously in Florida panthers. Sarcocysts found in Florida panthers
(Greiner et al., 1989) were assumed to be S. felis as they were morphologically identical
to S. felis found in bobcats (Dubey et al., 1992). Comparable or identical ssurRNA
sequence between panther and domestic cat sarcocysts would have augmented the
findings of the present study. Amplification of DNA from formalin-fixed,
paraffin-embedded tissue, particularly if it is aged, is more difficult than from fresh
tissue. Moreover, degradation of DNA into smaller fragments frequently occurs
(Wickham et al., 2001). Primers JD26/JD37 amplify 410 bp products from both
S. neurona and S. falcatula. It was hoped that this relatively short sequence could be
amplified from the sarcocysts located in the panther tongue; there was less optimism for
amplifying the larger ITS-1 region, 1080 bp from S. felis. Although primers JD26/JD37
did amplify a 410 bp product from the panther tissue (Fig. 4-12), JD26/JD37 are
"universal primers", able to amplify ssurRNA from all eukaryotic organisms. The finding
of sequence homology to plant material (most significantly with Pinus, pine) likely
indicates contamination of the sample at some point prior to sequencing. If comparable
sequence from JD26/JD37 to Sarcocystis had been found, one would still need further
analyses to diagnose S. felis; the ssurRNA gene amplified by JD26/JD37 cannot
differentiate between all Sarcocystis species (Tanhauser et al., 1999). To pursue
corroborating evidence of S. felis in both domestic and previously defined wild felid
hosts, two tactics might be employed: obtain fresh samples from these wild species for
analysis, or re-design primers for the ITS-1 region that amplify shorter, but diagnostic
sections of this gene.
The opossum consumed 30 g of cat muscle containing numerous sarcocysts, yet no
sporocysts were found in feces or gut scrapings. Other researchers have fed greater
quantities of cat muscle to opossums for bioassay purposes (Dubey et al., 2000, Butcher
et al., 2002). The smaller quantity fed in this study may or may not have contributed to
the negative result. By BLAST, ITS-1 sequence obtained from cat sarcocyst was not
similar to any given for Sarcocystis spp. that utilize the opossum as a host. The opossum
is therefore not likely to be a definitive host for S. felis. Although only one opossum was
used, such feeding trials are thought to be a sensitive bioassay for S. neurona, and
comparable studies previously have been carried out with single opossums (Dubey et al.,
2001b, Turay et al., 2002).
Biological diversity exists within Sarcocystis species; completion of the life cycle
of S. neurona through cats, or other intermediate hosts, has not been successful with
some horse CNS-derived isolates of S. neurona. In two studies, cat-derived S. neurona
isolates were able to cause infection in cats or immune-deficient mice, but the
horse-derived UCD-1 isolate of S. neurona was not infective (Butcher et al. 2002, Turay
et al. 2002). This suggests cats may not be intermediate hosts for some isolates of
S. neurona or the pathogenicity of the horse-derived isolates was insufficient. In addition,
merozoites from a different horse-derived isolate of S. neurona, SN2, were not infective
to raccoons (Dubey et al. 2002), whereas opossum-derived isolates were infective.
Two studies have shown cats do not function as definitive hosts for Sarcocystis
spp. considered intermediate host species in cats. Immunosuppressed cats were dosed
with sporocysts from an opossum naturally-infected with S. neurona. Muscle from these
cats was then fed to two laboratory-raised domestic cats, neither of which shed
sporocysts, demonstrating the cat is not a definitive host for S. neurona (Dubey et al.,
2002). Sarcocysts found in Florida panthers and cougars were fed to two
laboratory-reared domestic cats by Greiner and co-workers (1989). No sporocysts were
shed from these cats, indicating the cat is not a definitive host for this species of
Sarcocystis, later found to be S. felis.
The culture of bradyzoites was attempted in order to establish an isolate of S. felis.
Such an isolate could then be used to develop additional molecular markers or foster
further biological study of this little-characterized Sarcocystis species. The cultured
isolate, when fed to candidate host animals, might have been used to identify the
definitive host for S. felis.
Mehlhom and Heydorn (1979) and Fayer (1972) describe the introduction of
Sarcocystis spp. bradyzoites into cell cultures, their subsequent differentiation into macro
and microgamonts, sexual reproduction, and production of oocysts. These processes
occurred quickly; oocysts were formed within one to three days in culture. This result
was not achieved in the present study.
In this study, sera from 100 domestic cats were evaluated by immunoblot for
S. neurona antibody; 5% were found to be positive. Sera from 196 domestic, pet cats in
Michigan were evaluated for S. neurona antibody by both IFA and immunoblot (Rossano
et al., 2002). Twenty-seven per cent of the samples were positive by IFA and 5% by
immunoblot. Because pet cats were tested, seroprevalence may be lower than would be
found in free-roaming cats, although many pet cats also have access to the outdoors. In
the present study, 92% of cats were classified as "free-roaming", the remaining were
surrendered pets. If serologic data is to be considered valid evidence of S. neurona
infection, one might expect a higher prevalence of S. neurona antibodies in wild felid
species or feral domestic cats due to greater exposure to their (unknown) definitive
host(s). No serologic surveys for S. neurona antibodies have been conducted in wild felid
In an abstract presented at the 2001 Conference of Research Workers in Animal
Diseases, Stanek and colleagues describe the use of the S. neurona agglutination test
(SAT) to examine 36 cats living on horse farms where EPM had been diagnosed and 76
cats described as free-roaming. Sarcocystis neurona antibody was found in 39% of farm
cats, and 16% of free-roaming cats (23% total). It is not known if any of the sera
collected for the present study came from cats living on horse farms. Whereas the
immunoblot test used by Rossano and colleagues (2002) utilized a slightly different test
format and interpretation than the one used for the present study, significantly different
test formats, reagents, and interpretation are used for the SAT and IFA tests, both of
which reported higher seroprevalance. It is likely that differences in test formats and
interpretation affect the number of samples that test positive or negative in each case.
To increase the integrity of the serological data, a commercial lab that performs
immunobloting on mass scale was used to test cat sera from this study. The same
laboratory performed immunoblotting of armadillo serum, assisting in the identification
of the armadillo as a natural host for S. neurona (Tanhauser et al., 2001). However, the
immunoblot test is not validated for use in samples from animals other than horses, and
its utility for such samples is unknown. Though in the present study, immunoblotting was
able to clearly demonstrate S. neurona antibody in the cat immunized with S. neurona
merozoites. Histologic examination of postmortem CNS tissue is the "gold standard" for
EPM diagnosis (Daft et al., 2002). According to Lindsay and Dubey (2001), the
immunoblot is considered the "gold standard" of immunological-based testing for S.
neurona IgG, and thus EPM. It is agreed however, that the test is best suited for use with
CSF as an aid to diagnosis of EPM in horses with neurological disease; the test is not
recommended for screening clinically-normal horses, as false positive results occur
(MacKay et al., 2000, Lindsay and Dubey, 2001) in up to 40% of CSF samples from
normal horses (Daft et al., 2002). Antigenic variation within S. neurona has been
reported. In non-commercial, research applications, immunoblotting of cat sera has
demonstrated antigenic variation between several isolates of S. neurona. In contrast to
positive sera from immunized rabbits, and infected mice and horses, serum from a
sarcocyst-infected cat reacted against the 14-kDa band, but not 28-kDa band of antigen
prepared from the UCD-1 isolate of S. neurona. However, when antigen prepared from
another S. neurona isolate, MU1, was used, both 14-kDa and 28-kDa proteins were
recognized by the cat serum (Turay et al. 2002). Further, banding patterns seen on
immunoblots were different for mice inoculated with opossum-cat derived S. neurona
merozoites (Sn-Mucat 2) and mice inoculated with the horse-derived UCD 1 isolate
(Butcher et al. 2002). Specific recommendations for the use of, and limitations associated
with, immunobloting for S. neurona antibody may need to be considered when
interpreting the serological data presented from this study and others evaluating cat sera.
In accordance with protocols for interpretation of equine sera (Neogen, Lexington,
KY), cats from our study were considered positive if they reacted against a specific
17-kDa antigen of S. neurona. Likewise, sera were considered "non-specific" if they
reacted only against a 30-kDa antigen. Inclusion of sera with both nonspecific or specific
antibody reactivity would have yielded 19% cats positive for S. neurona antibody, a
figure comparable with the 23% obtained from SAT as reported by Stanek and colleagues
(2001). Greater seroprevalence reported by IFA may be due to cross-reactivity (Rossano
et al, 2002).
This study found no apparent correlation between positive immunoblot result and
presence of sarcocysts. These results could reflect problems with the sensitivity or
specificity of the immunoblot in cats, or, more likely, visible sarcocysts were not
S. neurona. If the immunoblot was detecting S. neurona infections, S. neurona sarcocysts
may not have been seen either because infections were immature (i.e., presarcocyst,),
eliminated by the cats before encystment, or were too rare to be found by histologic
techniques. It has been reported that S. neurona sarcocyst formation in the cat may take
several months (Dubey et al., 2000). Identification of sarcocyst-infected muscle for
further analysis may have been improved by muscle digestion, and examination of the
digest for bradyzoites (Collins et al., 1980). Alternatively, lysis of muscle tissue and use
of molecular markers specific for S. neurona, i.e. JNB 63/64 PCR primers followed by
restriction enzyme digest (Tanhauser et al., 1999), may have been appropriate means to
search for S. neurona infection in these cats.
Ostensibly, immunoblot results from this study might seem to indicate there is
cross-reactivity of S. felis antibody with S. neurona. However, only one of five S. felis
sarcocyst-positive cats was positive for S. neurona antibody by immunoblot; the other
four cats were antibody-negative, suggesting cross-reactivity is not a factor. The
serological data presented here have not resolved whether or not cats are natural hosts for
S. neurona. However, as pointed out by Rossano and colleagues (2002), one might expect
seroprevalence of S. neurona in intermediate hosts to be similar to that found in horses.
This study found 5% of cats from this area of Florida to have S. neurona antibody. The
seroprevalence in horses from this area is known to be much higher than 5%, perhaps as
high as 50% (MacKay et al., 2000).
SUMMARY AND CONCLUSIONS
Sarcocystis infection was present in the population of domestic cats sampled for
this study. While the results presented here do not confirm a role for cats as intermediate
host for S. neurona, the serologic data suggest that infection with S. neurona may occur;
5% of cats were seropositive. In contrast, sarcocysts commonly infected the cats; 10%
were infected. Morphologic examination showed these sarcocysts to be Sarcocystisfelis.
Molecular analyses showed S. felis to be quite different from other Sarcocystis species for
which comparable sequence data are available. Relatively little is known about the
biology of S. felis. Further research is needed to identify the source of these infections
and their pathogenicity in felids.
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Karen Dunne Gillis was born October 20th, 1961 in Great Barrington,
Massachusetts, the only child of Thomas and Kathleen Gillis. Karen grew up on a small
farm in the village of Monterey, located in the Berkshire Hills of western Massachusetts.
There, she developed a fondness for nature and animals that still continues. She attended
public schools until grade 10, and then enrolled at the Berkshire School in Sheffield,
Massachusetts. Her parents moved to south Florida, and Karen joined them there after
graduation from Berkshire in 1979. Karen attended the University of Florida in
Gainesville, graduating in 1984 with a Bachelor of Science degree from the College of
Agriculture, majoring in both animal science and dairy science. Karen left the University
environment for a short time, working in dairy management at farms in Georgia and
Florida, but returned to Gainesville and the University of Florida in 1985 to work at the
College of Veterinary Medicine. Karen remains happily and successfully employed there;
she is currently a Senior Biological Scientist in the Department of Large Animal Clinical
Sciences. Karen pursued her graduate studies while working full-time. After receipt of
her master's degree in veterinary medical sciences, Karen plans to continue her work at
the College of Veterinary Medicine.