Absence of Vacuolar Membrane Involving

Absence of Vacuolar Membrane Involving Toxoplasma gondii During Its Intranuclear Localization Author(s): H. S. Barbosa , M. F. Ferreira-Silva , E. V. Guimarães , L. Carvalho , and R. M. Rodrigues Source: Journal of Parasitology, 91(1):182-184. 2005. Published By: American Society of Parasitologists DOI: http://dx.doi.org/10.1645/GE-276R URL: http://www.bioone.org/doi/full/10.1645/GE-276R

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J. Parasitol., 91(1), 2005, pp. 179–182 q American Society of Parasitologists 2005

Zoonotic Genotype of Giardia intestinalis Detected in a Ferret Niichiro Abe, Carolyn Read*, R. C. Andrew Thompson*, and Motohiro Iseki†, Department of Microbiology, Osaka City Institute of Public Health and Environmental Sciences, Tennoji-ku, Osaka 543-0026, Japan; *Division of Veterinary and Biomedical Sciences, Murdoch University, Murdoch, WA 6150, Australia; †Department of Parasitology, Graduate School of Medical Science, Kanazawa University, Takara-machi, Kanazawa 920-8640, Japan. e-mail: [email protected] ABSTRACT:

Giardia intestinalis has been found in a variety of mammals, including humans, and consists of host-specific and zoonotic genotypes. There has been only 1 study of G. intestinalis infection in weasels, but the genotype of its isolate remains unclear. In this study, we report the isolation of Giardia in a ferret exhibited at a pet shop. The isolate was analyzed genetically to validate the possibility of zoonotic transmission. Giardia diagnostic fragments of the small subunit ribosomal RNA, b-giardin, and glutamate dehydrogenase genes were amplified from the ferret isolate and sequenced to reveal the phylogenetic relationships between it and other Giardia species or genotypes

of G. intestinalis reported previously. The results showed that the ferret isolate represented the genetic group A-I in assemblage A, which could be a causative agent of human giardiasis. The flagellate Giardia is a well-known intestinal parasite, which infects a wide range of vertebrate hosts, including humans. At present, 6 species in this genus, i.e., G. intestinalis (syn. G. lamblia, G. duodenalis) in humans, livestock, and other domestic animals, G. microti and G. muris in rodents, G. psittaci and G. ardeae in birds, and G. agilis in amphibians, which can be distinguished in view of the morphology

FIGURE 1. Phylogenetic relationships of the ferret isolate to other Giardia species and G. intestinalis genotypes as inferred by neighbor-joining analysis, based on the nucleotide sequences of the SSUrDNA. Names of the isolates and accession numbers in GenBank are shown in parentheses. 179

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FIGURE 2. Phylogenetic relationships of the ferret isolate to other Giardia species and G. intestinalis genotypes as inferred by neighbor-joining analysis, based on the nucleotide sequences of the b-giardin. Names of the isolates and accession numbers in GenBank are shown in parentheses.

and ultrastructure of their trophozoites, are recognized as valid (Adam, 2001). However, recent molecular studies have shown that G. intestinalis is composed of at least 7 genetically distinct, but morphologically identical, assemblages (assemblages A–G), and, moreover, most of these assemblages appear to have different host preferences, e.g., assemblages C and D in dogs, assemblage E in hoofed livestock, assemblage F in cats, and assemblage G in rats (Monis et al., 1999; Adam, 2001; Monis and Thompson, 2003). On the other hand, assemblage A consists of isolates that can be classified into 2 genetic groups (A-I and A-II) (Thompson et al., 2000). Genetic group A-I consists of a mixture of animal and human isolates. In contrast, group A-II consists entirely of human isolates. Assemblage B consists of a genetically diverse group of mainly human isolates, but some isolates from animals have been included. Therefore, it is supposed that genetic group A-I in assemblages A and B has the potential for zoonotic transmission (Thompson et al., 2000; Monis and Thompson, 2003). At present, there has been only 1 study of Giardia infection in Mustelidae animals, but the genotype of the isolate remains unclear because identification was performed with only conventional microscopy (Williams et al., 1988). In Japan, the ferret is a popular pet sold in many shops, but a detailed survey of zoonotic pathogens in ferrets has not been performed (Abe and Iseki, 2003). Because G. intestinalis is genetically diverse and some isolates from animals appear to have zoonotic potential as mentioned above, it is likely that ferrets harbor ferretspecific or zoonotic genotypes. Therefore, it is important to analyze the isolates from ferrets genetically to elucidate the epizootiology of Giardia infection in animals as well as for the control of human giardiasis. In this study, we obtained an isolate from a ferret in a pet shop and compared it genetically with the multiple genotypes of G. intestinalis reported previously to validate the phylogenetic relationships. A fecal sample was collected from a ferret exhibited at a pet shop in

Kanazawa City, Japan. This animal showed no clinical symptoms, such as diarrhea, when the fecal sample was collected. The purification of Giardia cysts from the fecal sample and the extraction of DNA from cysts were performed following a method reported previously (Abe et al., 2003). Giardia diagnostic fragments were amplified by the polymerase chain reaction (PCR) with the following primer pairs targeting the different gene loci: RH11 and RH4 for the Giardia small subunit ribosomal RNA gene (SSUrDNA) (Hopkins et al., 1997), G7 and G759 for the Giardia b-giardin gene (b-giardin) (Caccio` et al., 2002), and GDH1 and GDH4 for the Giardia glutamate dehydrogenase gene (GDH) (Homan et al., 1998). The area amplified with each primer pair includes a variable region, which can be used to distinguish Giardia species as well as multiple genotypes of G. intestinalis. PCR amplification was performed under conditions reported previously (Hopkins et al., 1997; Homan et al., 1998; Caccio` et al., 2002), except that Ex Taq DNA polymerase, Ex Taq buffer, and deoxynucleoside triphosphate (TAKARA Shuzo Co. Ltd., Otsu, Japan) were used in this study. Amplification products were subjected to electrophoretic separation using 3% agarose gels, stained with ethidium bromide, and observed on a UV transilluminator. The PCR products were gel purified using a QIA quick Gel Extraction kit (QIAGEN GmbH, Hilden, Germany) and sequenced using an ABI PRISM BigDye Terminator Cycle Sequencing FS Ready Reaction kit (PE Applied Biosystems, Foster City, California) on an ABI 310 automated sequencer (PE Applied Biosystems). PCR products were sequenced in both directions using each primer pair mentioned above. Sequences obtained from the ferret Giardia isolate were aligned with available nucleotide sequences reported previously (Baruch et al., 1996; Monis et al., 1996, 1998, 1999; Thompson et al., 2000; Caccio` et al., 2002) from other Giardia species and multiple genotypes of G. intestinalis using Clustal-X (version 1.63b). Evolutionary distance between different isolates was calculated with the Kimura 2-parameter

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FIGURE 3. Phylogenetic relationships of the ferret isolate to other Giardia species and G. intestinalis genotypes as inferred by neighbor-joining analysis, based on the nucleotide sequences of the GDH. Names of the isolates and accession numbers in GenBank are shown in parentheses.

method. Trees were constructed using the neighbor-joining algorithm (Saitou and Nei, 1987) and were drawn using the NJplot program (Perrie`re and Gouy, 1996). The partial sequences of SSUrDNA, b-giardin, and GDH of the ferret Giardia isolate, obtained in this study, have been deposited in the GenBank database as AB159796, AB159797, and AB159795, respectively. The partial SSUrDNA, b-giardin, and GDH were amplified successfully in the ferret isolate (data not shown). The partial SSUrDNA (125 bp) sequence was identical to sequences of the isolates (BAC2 and BAH40C11) found to have the assemblage A. The partial b-giardin sequence (472 bp) of the ferret isolate differed slightly from the sequences of the isolates (WB, KC8) found to have the assemblage A. There were 2 substitutions in the partial b-giardin sequence of the isolate from the ferret as compared with that of the isolate WB or KC8 (data not shown). The partial GDH sequence (592 bp) of the ferret isolate also differed slightly from the sequences of the isolates found to have the assemblage A (Ad-1, Ad-2, Bris-136). There were 1, or 3, substitutions in the partial GDH sequence as compared with that of the isolate Ad-1 or Ad-2 and Bris-136, respectively (data not shown). The close relatedness of the ferret isolate to assemblage A was also reflected in the phylogenetic analysis of b-giardin (Fig. 2) as well as SSUrDNA (Fig. 1): the ferret isolate was clustered with assemblage A. Similarly, the phylogenetic analysis of GDH sequences showed a close relatedness between the ferret isolate and assemblage A, but the ferret isolate was not clustered with the isolates Ad-2 and Bris-136 found to have group A-II but with the isolate Ad-1 found to have group A-I (Fig. 3). At present, the isolates classified into the genetic group A-I have been

found in a variety of mammals, e.g., cattle, pig, horse, cat, dog, beaver, and humans, but the isolates in group A-II have been found only in humans (Adam, 2001; Monis and Thompson, 2003). Therefore, on the basis of the results of the phylogenetic analysis performed in this study and of the molecular epidemiological evidence revealed previously, we place the ferret isolate in genetic group A-I, which appears to have zoonotic potential. Although Giardia infection in Mustelidae had been confirmed already in a black-footed ferret, Mustela nigripes, by light microscopy in 1988 (Williams et al., 1988), since then there have been no reports regarding Giardia infection in weasels. Therefore, our study is the first molecular analysis of an isolate from weasels. Epizootiological surveys of zoonotic pathogens in animals reared in pet shops or by breeders have been overlooked, and thus, periodical examinations of pets are needed to prevent infections with zoonotic pathogens. LITERATURE CITED ABE, N., AND M. ISEKI. 2003. Identification of genotypes of Cryptosporidium parvum isolates from ferrets in Japan. Parasitology Research 89: 422–424. ———, I. KIMATA, AND M. ISEKI. 2003. Identification of genotypes of Giardia intestinalis isolates from dogs in Japan by direct sequencing of the PCR amplified glutamate dehydrogenase gene. Journal of Veterinary Medical Science 65: 29–33. ADAM, R. D. 2001. Biology of Giardia lamblia. Clinical Microbiology Reviews 14: 447–475. BARUCH, A. C., J. ISAAC-RENTON, AND R. D. ADAM. 1996. The molecular

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epidemiology of Giardia lamblia: A sequence-based approach. Journal of Infectious Diseases 174: 233–236. CACCIO`, S. M., M. D. GIACOMO, AND E. POZIO. 2002. Sequence analysis of the b-giardin gene and development of a polymerase chain reaction-restriction fragment length polymorphism assay to genotype Giardia duodenalis cysts from human faecal samples. International Journal for Parasitology 32: 1023–1030. HOMAN, W. L., M. GILSING, H. BENTALA, L. LIMPER, AND F. KNAPEN. 1998. Characterization of Giardia duodenalis by polymerase-chainreaction fingerprinting. Parasitology Research 84: 707–714. HOPKINS, R. M., B. P. MELONI, D. M. GROTH, J. D. WETHERALL, J. A. REYNOLDSON, AND R. C. A. THOMPSON. 1997. Ribosomal RNA sequencing reveals differences between the genotypes of Giardia isolates recovered from humans and dogs living in the same locality. Journal of Parasitology 83: 44–51. MONIS, P. T., R. H. ANDREWS, G. MAYRHOFER, AND P. L. EY. 1999. Molecular systematics of the parasitic protozoan Giardia intestinalis. Molecular Biology and Evolution 16: 1135–1144. ———, ———, ———, J. MACKRILL, J. KULDA, J. L. ISAAC-RENTON, AND P. L. EY. 1998. Novel lineages of Giardia intestinalis identified

by genetic analysis of organisms isolated from dogs in Australia. Parasitology 116: 7–19. ———, G. MAYRHOFER, R. H. ANDREWS, W. L. HOMAN, L. LIMPER, AND P. L. EY. 1996. Molecular genetic analysis of Giardia intestinalis isolates at the glutamate dehydrogenase locus. Parasitology 112: 1–12. ———, AND R. C. A. THOMPSON. 2003. Cryptosporidium and Giardiazoonoses: Fact or fiction? Infection, Genetics and Evolution 3: 233–244. PERRIE`RE, G., AND M. GOUY. 1996. WWW-query: An on-line retrieval system for biological sequence banks. Biochimie 78: 364–369. SAITOU, N., AND M. NEI. 1987. The neighbor-joining method: A new method for reconstructing phylogenetic trees. Molecular Biology and Evolution 4: 406–425. THOMPSON, R. C. A., R. M. HOPKINS, AND W. L. HOMAN. 2000. Nomenclature and genetic groupings of Giardia infecting mammals. Parasitology Today 16: 210–213. WILLIAMS, E. S., E. T. THORNE, M. J. G. APPEL, AND D. W. BELITSKY. 1988. Canine distemper in black-footed ferrets (Mustela nigripes) from Wyoming. Journal of Wildlife Diseases 24: 385–398.


Absence of Vacuolar Membrane Involving Toxoplasma gondii During Its Intranuclear Localization H. S. Barbosa, M. F. Ferreira-Silva, E. V. Guimara˜es, L. Carvalho*, and R. M. Rodrigues, Laborato´rio de Ultra-estrutura Celular, Departamento de Ultra-estrutura e Biologia Celular, Instituto Oswaldo Cruz, Fundaçaõ Oswaldo Cruz, Av. Brasil 4365, 21045-900 Rio de Janeiro, RJ, Brasil; *Laborato´rio de Cultura de Ce´lulas, Departamento de Histologia e Embriologia, Instituto de Biologia, Universidade do Estado do Rio de Janeiro, 20551-170 Rio de Janeiro, RJ, Brasil. e-mail: [email protected] ABSTRACT: Tachyzoites of Toxoplasma gondii were located inside the nucleus of both skeletal muscle cells infected in vitro and peritoneal exudate cells collected from infected mouse in vivo. Ultrastructural analysis demonstrated that T. gondii invades the nucleus of host cells by the parasite apical region and with constriction of its body. We noted that the rhoptry, a secretory organelle of the parasite that is involved in the host cell invasion mechanism, was empty in the intranuclear T. gondii. The parasites were found in the nuclear matrix without evidence of the vacuolar membrane. Frequently, new parasites invaded host cell nucleus, which was already infected. The significance of this nuclear invasion could reflect an alternative route of T. gondii for its transitory survival or an escape mechanism from the host immune response during the in vivo infection (or both).

Toxoplasma gondii is an obligate intracellular parasite, which infects a wide variety of warm-blooded vertebrates, including humans and domestic animals. It is an important pathogen in humans, which can cause a serious and potentially fatal disease in newborn infants and in immunocompromised hosts, such as in acquired immunodeficiency syndrome patients (Dubey, 1997; Liesenfeld et al., 1999). The key for T. gondii survival is its internalization into the host cell. The process of invasion appears to be complex, involving an oriented active process that requires a motile parasite, its attachment to the host cell surface, followed by the discharge of secretory products from the parasite apical organelles (reviewed by Black and Boothroyd, 2000; Carruthers, 2002). In the host cell, the parasite resides within a modified, membrane-bound compartment, the parasitophorous vacuole (PV), which appears to act as a protective interface, avoiding fusion with endosomes and lysosomes, besides supporting the growth and proliferation of the tachyzoites (reviewed by Sinai and Joiner, 1997; Andrade et al., 2001). The available evidences suggest that the PV is nonfusogenic because the surface determinants from the host cell, which are known to be necessary for membrane fusion events, are excluded from the PV during the parasite invasion (Carvalho and De Souza, 1989, 1990; Mordue et al., 1999; Hakansson et al., 2001).

The intranuclear location of T. gondii tachyzoites during the infection of different cell types in vitro has been analyzed by light microscopy (Sourander et al., 1960; Remington et al., 1970; Ogunba, 1972; Azab et al., 1973). The transitory location of microorganisms within the nucleus of host cell, including sporozoites or merozoites from species of Dobellia, Eimeria, Isospora, and Besnoitia has also been described. These parasites have been denominated caryotropics because during the endogenous stages they can reside within the host cell nucleus (Roberts et al., 1971; Peka, 1992, 1993). Despite the frequent intranuclear occurrence among coccidians, little is known about the mechanisms or physiological basis for the reported intranuclear parasitism or the significance of this location in coccidian systematics (Atkinson and Ayala, 1987). The majority of these studies have been performed using light microscopy, which did not allow to observe the details from the invasion mechanisms as well as to characterize whether these parasites were surrounded by membranes. In this article we describe, for the first time at the ultrastructural level, the presence of T. gondii inside the nucleus of both skeletal muscle cells (SKMC) infected in vitro and peritoneal exudate cells collected from infected mouse in vivo. Primary cultures of SKMC were obtained from thigh muscles of 18day-old mouse embryos (for details, see Arau´jo-Jorge et al., 1986; Barbosa et al., 2000). Three- to 6-day-old cultures of SKMC were infected with tachyzoites (RH strain) of T. gondii using a 10:1 parasite–host cell ratio. After 4, 24, 48, and 72 hr of infection, the culture cells were fixed for 30 min at 4 C with 2.5% (v/v) glutaraldehyde in 0.1 M Na-cacodylate buffer (pH 7.2), washed with the same buffer, postfixed for 30 min at 4 C in 1% of osmium tetroxide, routinely processed for transmission electron microscopy, and examined in a Zeiss EM10C. Another ultrastructural approach was analyzing the peritoneal exudate cells obtained from T. gondii–infected Swiss mice after 48–72 hr of infection. Our results showed that intranuclear tachyzoites in SKMC were directly immersed in the nuclear matrix, without any morphological indication of vacuolar membrane surrounding the parasites (Fig. 1). Cells collected from the peritoneal exudates of T. gondii–infected mice also exhibited tachyzoites invading the host cell nucleus (Figs. 2–4), fre-

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FIGURE 1. Ultrastructural aspects of Toxoplasma gondii invasion into SKMC. Nucleus (N) containing parasite (P) immersed in the nuclear matrix, without vacuolar membrane surrounding the tachyzoite. Bar 5 0.5 mm. FIGURES 2–6. Transmission electron micrograph of exudate peritoneal cells obtained from Toxoplasma gondii–infected mice containing intranuclear parasites. 2, 3. Infected nucleus being invaded by new parasite, which presents constriction of its body (arrow). 4. Parasite invasion into the nucleus (N) through the apical region (arrow). Empty rhoptry (R) can be observed. 5. Note the presence of 2 parasites (P) inside the nucleus (N). 6. Shown in detail, the nucleus containing parasite freely in the nuclear matrix (NM) without vacuolar membrane (arrow).

quently with constriction of their body (Fig. 2). Occasionally, the nuclear invasion by the parasites occurred via the parasite apical region, where the conoid is located (Fig. 4). In addition, the rhoptries of the intranuclear parasites were empty (Fig. 4). We noted that infected host cells, which already displayed intranuclear parasites, could have their nuclei reinvaded by new parasites (Figs. 2, 3). Although several host cell nuclei containing 1 or more parasites could be observed (Fig. 5), no dividing parasite was noted. The tachyzoites were also located in direct contact with the nuclear matrix, without vacuolar membrane involving the parasites, as demonstrated in detail in the Figure 6. The observation of intranuclear parasites from the RH strain of T. gondii is not an exclusive event from this parasite strain because it has already been proposed by other authors, who analyzed other T. gondii strains (Remington et al., 1970; Azab et al., 1973). Our group has accumulated evidence for the occurrence of intranuclear bradyzoites in

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another T. gondii strain, the avirulent ME-49, during the cystogenesis process occurring in SKMC in vitro (data not shown). Intranuclear tachyzoites have been observed in host cells undergoing degeneration, in both in vivo and in vitro models. It is well established that after 72 hr of mice infection with T. gondii, their peritoneal exudate cells undergo disruption, liberating the parasites into the peritoneal cavity of the host. Similarly, at this time, T. gondii–infected host cells release parasites into the extracellular medium during their infection in vitro. Our electron microscopy images of intranuclear T. gondii undoubtedly show the absence of plasmatic membrane or host cell organelles surrounding the parasites (or both), as previously reported by Ogunba (1972). Our present results are surely not due to artifactual procedures because both the parasites and host cells, which did not display intranuclear parasites, had their structural organization very well preserved. During the nuclear invasion by T. gondii, we noted the constriction of the parasite body and the presence of empty rhoptries, which suggests an active penetration mechanism. It has been proposed that the nuclear invasion could occur by direct penetration through the nuclear membrane or through the engulfment of PVs containing parasites from the host cell cytoplasm (Remington et al., 1970). However, these authors only analyzed this event by light microscopy, which could generate misinterpretations. The presence of a PV has been demonstrated in some intranuclear coccidians by light microscopy (Shibalova and Morozova, 1979; Mohamed and Molyneux, 1990; Peka, 1992, 1993), except for Eimeria stigmosa, which was found in direct contact with the nucleoplasm of the host cell (Gajadhar et al., 1986). Our ultrastructural analysis did not reveal the presence of a membrane surrounding the intranuclear parasites because they were always found immersed freely in the nucleoplasm. The absence of membrane surrounding intranuclear T. gondii could be explained by the lack of the signaling events, which involve the ability of cell adhesion molecules to initiate the formation of organized structures, as for example, the PV formation. In our study, we regularly noted more than 1 intranuclear parasite per infected nucleus. Some authors described this event as a possible cellular division of intranuclear parasites (Remington et al., 1970; Ogunba, 1972; Azab et al., 1973). However, in our ultrastructural analysis, we did not observe any cellular division of these intranuclear parasites. However, we usually found an already infected nucleus being invaded by new parasites, suggesting that the infected nuclei are not refractory to new infections, instead being, perhaps, more susceptible to new invasions. In conclusion, our ultrastructural analysis of the nuclear invasion by T. gondii showed (1) the participation of the apical region of the parasite, (2) a constriction in the body of the parasite, and (3) empty rhoptries. Taken together, these data suggest active penetration of the T. gondii into the host cell nucleus, without formation of host membranes around the parasites. Moreover, we showed for the first time intranuclear parasites during infection by T. gondii in vivo, suggesting that this event is not restricted to in vitro systems. The significance of this nuclear invasion could reflect an alternative route of T. gondii for its intracellular survival, perhaps transitory, or an alternative escape mechanism from the host immune response during the infection in vivo (or both). The authors thank Maria de Nazare´ Corria Soeiro for critical review ´ vila for processing images, and Gene´sio L. of this manuscript, Bruno A Faria and Jose´ L. Faria for their excellent technical assistance. This work was supported with grants from Conselho Nacional de Desenvolvimento Cientı´fico e Tecnolo´gico (CNPq), Coordenadoria de Aperfeiçoamento de Pessoal de Nı´vel Superior (CAPES), and Fundaçaõ Carlos Chagas Filho de Amparo a` Pesquisa do Estado do Rio de Janeiro (FAPERJ). LITERATURE CITED ANDRADE, E. F., A. C. STUMBO, L. H. MONTEIRO-LEAL, L. CARVALHO, AND H. S. BARBOSA. 2001. Do microtubules around the Toxoplasma gondii-containing parasitophorous vacuole in skeletal muscle cells form a barrier for the phagolysosomal fusion? Journal of Submicroscopic Cytology and Pathology 33: 337–341. ARAU´JO-JORGE, T. C., H. S. BARBOSA, A. L. MOREIRA, W. DE SOUZA, AND M. N. L. MEIRELLES. 1986. The interaction of myotropic and macrophagotropic strains of Trypanosoma cruzi with myoblasts and

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fibers of skeletal muscle. Zeitschrift fu¨r Parasitenkunde 72: 577– 584. ATKINSON, C. T., AND S. C. AYALA. 1987. Isospora manchacensis n. sp., an intranuclear coccidian from the Lousiana ground skink, Scincella lateralis (Say, 1823) (Lacertilia: Scincidae). Journal of Parasitology 73: 817–823. AZAB, M. E., M. A. RIFAAT, S. A. SALEM, I. ZAGHLOUL, AND T. A. MORSY. 1973. The intranuclear development of Toxoplasma. Zeitschrift fu¨r Parasitenkunde 42: 39–42. BARBOSA, H. S., M. C. S. PEREIRA, AND M. N. L. MEIRELLES. 2000. Protocolos de culturas prima´rias. In Doença de Chagas: Manual para experimentaçaõ animal, Tania Arau´jo-Jorge and Solange Lisboa de Castro (eds.). Editora Fiocruz, Rio de Janeiro, Brazil, p. 297–313. BLACK, M. W., AND J. C. BOOTHROYD. 2000. Lytic cycle of Toxoplasma gondii. Microbiology and Molecular Biology Reviews 64: 607– 623. CARRUTHERS, V. B. 2002. Host cell invasion by the opportunistic pathogen Toxoplasma gondii. Acta Tropica 81: 111–122. CARVALHO, L., AND W. DE SOUZA. 1989. Cytochemical localization of plasma membrane enzyme markers during interiorization of tachyzoites of Toxoplasma gondii by macrophages. Journal of Protozoology 36: 164–170. ———, AND ———. 1990. Internalization of surface anionic sites and phagosome-lysosome fusion during interaction of Toxoplasma gondii with macrophages. European Journal of Cell Biology 51: 211– 219. DUBEY, J. P. 1997. Bradyzoite-induced murine toxoplasmosis: Stage conversion, pathogenesis, and tissue cyst formation in mice fed bradyzoites of different strains of Toxoplasma gondii. Journal of Eukaryotic Microbiology 44: 592–602. GAJADHAR, A. A., D. J. RAINNIE, AND R. J. CAWTHORN. 1986. Description of the goose coccidium Eimeria stigmosa (Klimes, 1963), with evidence of intranuclear development. Journal of Parasitology 72: 588–594. HAKANSSON, S., A. J. CHARRON, AND L. D. SIBLEY. 2001. Toxoplasma evacuoles: A two-step process of secretion and fusion forms the parasitophorous vacuole. EMBO Journal 20: 3132–3144.

LIESENFELD, O., H. KANG, D. PARK, T. A. NGUYEN, C. V. PARKHE, H. WATANABE, T. ABO, A. SHER, J. S. REMINGTON, AND Y. SUZUKI. 1999. TNF-alpha, nitric oxide and IFN-gamma are all critical for development of necrosis in the small intestine and early mortality in genetically susceptible mice infected perorally with Toxoplasma gondii. Parasite Immunology 21: 365–376. MOHAMED, H. A., AND D. H. MOLYNEUX. 1990. Developmental stages of Cyclospora talpae in the liver and bile duct of the mole (Talpa europea). Parasitology 101: 345–350. MORDUE, D. G., S. HAKANSSON, I. NIESMAN, AND L. D. SIBLEY. 1999. Toxoplasma gondii resides in a vacuole that avoids fusion with host cell endocytic and exocytic vesicular trafficking pathways. Experimental Parasitology 92: 87–99. OGUNBA, E. O. 1972. Toxoplasma gondii in the nuclei of cells in tissue cultures. Transactions of the Royal Society of Tropical Medicine and Hygiene 66: 811–813. PEKA, Z. 1992. Life cycle and ultrastructure of Eimeria stigmosa, the intranuclear coccidian of goose (Anser anser domesticus). Folia Parasitologica 2: 105–114. ———. 1993. Intranuclear development of asexual and sexual generations of Eimeria hermani Farr, 1953, the coccidian geese. Zentralblatt fu¨r Bakteriologie 278: 570–576. REMINGTON, J. S., E. EARLE, AND T. YAGURA. 1970. Toxoplasma in nucleus. Journal of Parasitology 56: 390–391. ROBERTS, W. L., C. A. SPEER, AND D. M. HAMMOND. 1971. Penetration of Eimeria larimerensis sporozoites into cultured cells as observed with the light and electron microscopes. Journal of Parasitology 57: 615–624. SHIBALOVA, T. A., AND T. I. MOROZOVA. 1979. Intranuclear development of macrogametes in the coccidium Tyzzeria parvula. Tsitologiya (Leningrad) 21: 969–972. SINAI, A. P., AND K. A. JOINER. 1997. Safe haven: The cell biology of nonfusogenic pathogen vacuoles. Annual Review of Microbiology 51: 415–462. SOURANDER, P., E. LYCKE, AND E. LUND. 1960. Observations on living cells infected with Toxoplasma gondii. British Journal of Experimental Pathology 41: 176–180.


Parasitic Helminths of Eurasian Collared-Doves (Streptopelia decaocto) From Florida Diane L. Bean, Edith Rojas-Flores, Garry W. Foster*, John M. Kinsella, and Donald J. Forrester, Department of Pathobiology, College of Veterinary Medicine, University of Florida, P.O. Box 110880, Gainesville, Florida 32610-0880; *To whom correspondence should be addressed. e-mail: [email protected] ABSTRACT: Sixty-three Eurasian collared-doves (ECDs) (Streptopelia decaocto) from Florida were examined for parasitic helminths from June to December 2001. Nine species of helminths were identified (5 nematodes, 2 cestodes, and 2 trematodes). The most prevalent helminths were Ascaridia columbae (73.0%), Fuhrmannetta crassula (28.6%), Ornithostrongylus quadriradiatus (12.7%), and Bruscapillaria obsignata (11.1%). The helminths with the greatest mean intensity were Tanaisia bragai (13.5), A. columbae (9.3), and O. quadriradiatus (7.1). In Florida, the mean intensity of A. columbae in ECDs (9.3) was similar to that found in white-winged doves (Zenaida asiatica) (9.1) (P 5 0.461), and both the intensities were significantly higher than that in the native mourning doves (Zenaida macroura) (3.7) (P 5 0.001 and 0.005, respectively). Fuhrmannetta crassula is reported for the first time in columbids from Florida.

The North American population of Eurasian collared-doves (ECDs), Streptopelia decaocto (Frivaldszky, 1838), is believed to have originated from a group of doves that escaped captivity in the Bahamas in 1974 (Smith, 1987). The exact date of colonization in Florida is unknown because of the misidentification of the ECD as the ringed turtle-dove

FIGURE 1. Community similarity characteristics of the helminth communities of ECDs, MDs, and WWDs in Florida.

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TABLE I. Prevalence and intensity of helminths in Eurasian collared-doves, mourning doves, and white-winged doves in Florida. Eurasian collared-doves, Florida (n 5 63) Prevalence

Nematoda Ascaridia columbae Bruscapillaria obsignata Tetrameres americanus Dispharynx nasuta Ornithostrongylus quadriradiatus Ornithostrongylus spp.§ Aproctella stoddardi

73.0 11.1 3.2 4.8 12.7 — —

9.3 1.1 1.0 2.7 7.1 — —

Cestoda Fuhrmannetta crassula Raillietina sp.\ Killigrewia delafondi Hymenolepididae

28.6 9.5 — —

1.8 (1–6) 1.5 (1–4) — —

— 0.2 0.7 1.3

— 2.0 (2) 1.0 (1) 1.0 (1)

— 3.4 — —

3.2 1.6

13.5 (3–24) 4.0

0.2 0.2

12.0 (12) 1.0 (1)

0.8 0.8

(1–76) (1–2) (1–6) (1–20)

Prevalence

30.5 0.7 4.2 16.0 — 67.3 10.3

Intensity

White-winged doves,† Homestead (n 5 119)

Parasite

Trematoda Tanaisia bragai Brachylaima sp.

Intensity‡

Mourning doves,* Florida (n 5 455)

3.7 5.7 1.6 10.9 — 13.1 6.0

(1–43) (1–15) (1–28) (1–144) (1–160) (1–34)

Prevalence

43.7 0.8 — 4.2 — 79.0 —

Intensity

9.1 (1–47) 2.0 — 1.4 (1–3) — 10.2 (1–104) — — 2.5 (1–7) — — 3.0 2.0

* Data from Forrester et al. (1983). † Data from Conti and Forrester (1981). ‡ Mean values of intensity followed by ranges in parentheses. § As reported by the authors: a complex of 2 species, O. quadriradiatus and O. iheringi, in a ratio of 14:1 based on males only. \ Conti and Forrester (1981) and Forrester et al. (1983) report Raillietina spp. as a complex of at least 2 species; it is possible 1 of these corresponds to F. crassula.

(Streptopelia risoria (Linnaeus, 1758)). The first documented report in southern Florida was in the early 1980s (Romagosa and Labisky, 2000). Within 10 yr of its identification, ECDs had established populations throughout Florida and had been sighted throughout the continental United States by the mid to late 1990s (Romagosa and McEneaney, 1999). The rapid colonization of North America by ECDs is similar to that seen in Europe in the early to mid 1900s. The ECD has a distinct dispersal pattern, known as ‘‘jump’’ dispersal (Romagosa and McEneaney, 1999), which is characterized by long-distance dispersal of individuals and subsequent population coalescence (Pielou, 1979). The dispersal rate of ECDs across North America is skewed by local releases by breeders throughout the United States and an increased awareness and reporting of the species by bird watchers. The ECDs colonization of the United States is enhanced by its varied diet, ability to breed year round, and tolerance of human populations (Romagosa and Labisky, 2000). The effects of ECDs on indigenous dove species, such as the mourning dove (MD) (Zenaida macroura (Linnaeus, 1758)), and recently established species in Florida, such as the white-winged dove (WWD) (Zenaida asiatica (Linnaeus, 1758)) and the ringed turtle-dove, are poorly understood. The only report on helminths in ECDs is based on 5 specimens from Palm Beach, Florida (Forrester and Spalding, 2003). More quantitative data are required to determine the effect that ECDs may have on the spread of parasites to indigenous dove species. This study was initiated to determine the helminth community of ECDs in Florida from 2 areas and to compare our results with the helminths reported from other doves from Florida. A total of 63 ECDs was collected from 4 counties in Florida from June to December 2001, i.e., 32 from Pinellas County (278549N, 828419W), 20 from Okaloosa County (308259N, 868409W), 7 from Santa Rosa County (308219N, 878109W), 3 from Bay County (308119N, 858489W), and 1 from Ramrod Key in Monroe County (248459N, 818209W). ECDs from Pinellas County were collected in June and July; all other ECDs were collected from October through December. All specimens were part of die-offs that occurred in these areas and were the result of a paramyxovirus and to a lesser extent visceral Trichomonas gallinae infections. The dead ECDs selected for this study were in good condition, with most being collected and refrigerated within a

few hours of dying. Identification of the ECDs followed the techniques of Romagosa and McEneaney (1999). The specimens were bagged individually and frozen until examined. Parasite screening techniques used were those described by Kinsella and Forrester (1972). Trematodes and cestodes were preserved in Roudabush’s AFA, stained with either Ehrlich’s hematoxylin or Semichon’s acetocarmine, and mounted in neutral Canada balsam. The nematodes were preserved in 70% ethanol with glycerin, mounted in lactophenol for identification, and then returned to the preservative. ECDs collected in Santa Rosa, Okaloosa, and Bay counties, which are located in the northwestern Panhandle of Florida, were combined as a single ‘‘Florida Panhandle’’ sample (n 5 30) for statistical comparison with the Pinellas County (n 5 32) (west coast of central Florida) collection site. The 2 collecting areas are similar; they are on the Gulf coast of Florida and have large amounts of salt and brackish water surrounding them, a relatively mild climate throughout the year, MDs and rock doves (RD) (Columba livia Gmelin, 1789), and a local human population that provides bird feeders and freshwater, which artificially concentrate a variety of avian species into a small area. Host gender was not identified in the Pinellas sample; thus, no statistical analysis between genders was attempted. Because of our relatively small sample sizes, only parasites with prevalences .10% were compared. For comparisons with parasite data reported for MDs and WWDs (Conti and Forrester, 1981, and Forrester et al., 1983, respectively) in Florida, all 63 ECDs (included is the 1 ECD collected from Ramrod Key, Monroe County) were combined for the statistical analyses. A chi-square test was used to compare prevalences, and a Mann–Whitney rank sum test was used to compare intensities using SigmaStatt for Windows (version 2.03, SPSS, Inc., Chicago, Illinois). Significance was set at P , 0.05. Parasite community similarities were calculated using Jaccard’s similarity index and Sorensen’s percent similarity index, where the latter is calculated using parasite abundance (Magurran, 1988). Terminology used is according to Bush et al. (1997). Helminth voucher specimens have been deposited in the U.S. National Parasite Collection (USNPC), Beltsville, Maryland (accession numbers USNPC 94198, 94400, and 94569–94572). Nine species of helminths were identified in the 63 ECDs that we examined (5 nematodes, 2 cestodes, and 2 trematodes) (Table I). The most prevalent helminths were Ascaridia columbae (73.0%), Fuhrman-

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TABLE II. Prevalence and intensity of helminths in 62* Eurasian collared-doves from Florida. Pinellas County (n 5 32)

Florida Panhandle (n 5 30)

Intensity Parasite

Intensity

Prevalence

Mean

Range

Prevalence

Mean

Range

Nematoda Ascaridia columbae (Gmelin, 1790) (GZ, SI)† Bruscapillaria obsignata (Madsen, 1945) (SI) Tetrameres americanus (Cram, 1927) (ES, PV) Dispharynx nasuta (Rudolphi, 1819) (ES, PV) Ornithostrongylus quadriradiatus (Stevenson, 1904) (SI) Ascaridia (larvae) (GZ, SI) Tetrameres (larvae) (PV) Unidentified larvae (SI)

62.5 21.9 6.3 6.3 3.1 9.4 3.1 43.8

9.0 1.1 1.0 3.5 20.0 1.3 1.0 6.0

1–76 1–2 — 1–6 — 1–2 — 1–17

83.3 — — 3.3 23.3 33.3 — 26.6

9.5 — — 1.0 5.3 2.9 — 3.1

1–48 — — — 1–15 1–8 — 1–8

Cestoda Fuhrmannetta crassula (Rudolphi, 1819) (SI) Raillietina sp. (SI)

50.0 15.6

1.9 1.6

1–6 1–4

6.6 3.3

1.0 1.0

— —

Trematoda Tanaisia bragai dos Santos, 1934 (KD) Brachylaima fuscatum (Rudolphi, 1819) (SI)

6.3 —

13.5 —

3–24 —

— 3.3

— 4.0

— —

* An additional Eurasian collared-dove was collected on Ramrod Key, Monroe County, in the Florida Keys and was found to have only A. columbae with an intensity of 63. This information is not included in this table but is in Table I. † Location in host: ES 5 esophagus, GZ 5 gizzard, KD 5 Kidney, PV 5 proventriculus, SI 5 small intestine.

netta crassula (28.6%), Ornithostrongylus quadriradiatus (12.7%), and Bruscapillaria obsignata (synonym of Capillaria obsignata) (11.1%) (Table I). The helminths with the greatest mean intensity were Tanaisia bragai (13.5), A. columbae (10.4), and O. quadriradiatus (7.1). Prevalences and intensities of parasites for the Pinellas County and the Florida Panhandle collecting sites are given in Table II. Only 5 parasite species had prevalences .10%. There was no significant difference in prevalences or intensities for A. columbae between the Panhandle site and the Pinellas County site (P 5 0.120 and 0.689, respectively). The Panhandle site had a significantly higher prevalence of O. quadriradiatus (23.3%) than the Pinellas site (3.1%) (P 5 0.024) but a lower prevalence of B. obsignata (0.0%) than the Pinellas site (21.9%) (P 5 0.010). The prevalence of F. crassula was significantly higher at the Pinellas site (50%) than the Panhandle site (6.6%) (P 5 0.001); however, the prevalence of Raillietina sp. was statistically similar at both sites (P 5 0.197). Jaccard’s index (0.55) and Sorensen’s percent similarity index (44.7%) indicate some similarity between the Pinellas and Panhandle ECD helminth community compositions, but these were not identical. All the species of parasites found in ECDs have been identified previously in MD and WWD populations in Florida (Table I), except for F. crassula, which is reported for the first time in columbids from Florida. A comparison of prevalences and intensities for parasites of the 63 ECDs and those reported in WWDs and MDs are given in Table I, and helminth community similarities are given in Figure 1. Prevalence of A. columbae was significantly higher in ECDs (73.0%) than those reported for WWDs (43.7%) and MDs (30.5%) in Florida (P , 0.001 for both). The prevalence of A. columbae reported for WWDs was significantly higher than that reported for MDs in Florida (P 5 0.009). The mean intensity of A. columbae in ECDs (9.3) was similar to that found in WWDs (9.0) (P 5 0.461) in Florida; however, both were significantly higher than the intensity reported for MDs (3.6) in Florida (P 5 0.001 and 0.005, respectively). The parasite community similarities among ECDs, MDs, and WWDs in Florida are presented in Figure 1. Jaccard’s index indicates some similarity in the helminth species present in all 3 dove species. However, Sorensen’s percent similarity index, which uses parasite abundance, indicates a low similarity between the parasite communities of ECDs and MDs but a high similarity between MD and WWD parasite communities. The significant differences in the prevalences between the 2 collection sites for several of the ECD parasites may be due, in part, to the

jump dispersal of the host as described by Romagosa and McEneaney (1999). The ECDs for these 2 areas of Florida may have different origins and, therefore, had slightly different parasite faunas to start with, especially if many of them were released by breeders, pet owners, or both. The 2 collection areas are ecologically similar. The differences may also be attributed to seasonal variations in some of the helminths, as reviewed by Bush (1990). Ascaridia columbae seems to be 1 of the core parasite species in the ECDs throughout Florida. It was found in ECDs from the Florida Keys (Ramrod Key, Monroe County) in the south to Santa Rosa County in the northern Panhandle. Although Ramrod Key was the only location on the east coast of Florida where ECDs were collected, it is likely that A. columbae can be found in any ECD population in Florida. It is interesting to note that the prevalence and intensity of A. columbae were highest in the introduced columbids, ECDs, and WWDs and lowest in the native MDs. We did not compare prevalences and intensities for Ornithostrongylus sp. or Raillietina sp. in the ECDs with those reported in WWDs or MDs. Both Conti and Forrester (1981) and Forrester et al. (1983) reported Ornithostrongylus spp. as ‘‘a complex of 2 species, O. quadriradiatus and O. iheringi in a ratio of 14:1 based on males only.’’ Confounding this further is the fact that there has been some ambiguity among the data due to taxonomic changes over the years. Ornithostrongylus crami was once thought to be synonymous with O. quadriradiatus until Durette-Desset et al. (2000) gave evidence for it being a separate species. Therefore, some of the Ornithostrongylus spp. reported previously in the MDs and WWDs in Florida may have been O. crami. Similarly we did not compare the cestodes in the ECDs with those reported in MDs and WWDs in Florida by Conti and Forrester (1981) and Forrester et al. (1983). The Raillietina spp. reported in MDs and WWDs by these authors were ‘‘a complex of at least 2 species.’’ Because at the time of these reports Fuhrmannetta was considered a subgenus in the genus Raillietina, it is possible that F. crassula may have been present in the MDs, WWDs, or both and grouped and reported as Raillietina spp. with the others. However, 1 of us (J.M.K.) examined the voucher specimens deposited in the USNPC by both Conti and Forrester, and none was F. crassula (5R. (F.) crassula). The Raillietina sp. we report in the ECDs appears to be a single species. Mollhagen (1976) described Tetrameres columbicola as a separate species from the previously recognized Tetrameres americanus (Cram, 1927). According to his description, several articles written in the early 1980s identified the Tetrameres sp. found in MDs and RDs as T. col-

RESEARCH NOTES

umbicola (Conti and Forrester, 1981; Forrester et al., 1983; Simpson et. al., 1984). The Tetrameres sp. from the ECDs also fit Mollhagen’s description of T. columbicola more closely than the previously described T. americanus. However, Mollhagen’s description has no taxonomic validity under the International Code because his data were reported only in a Ph.D. dissertation and never published in a refereed journal. Therefore, the Tetrameres species found in the ECDs, WWDs, and MDs is T. americanus. In Florida, the ECDs have a helminth fauna similar to those found in the WWD and native MD populations. The higher prevalences and intensities for some of the helminths in the ECDs may be because of their being the most recent of the introduced exotic dove species in Florida. The effect of the paramyxovirus and visceral T. gallinae infections on the helminth fauna of the ECDs we sampled is unknown. Because of the small sample sizes for our ECDs, the statistical analyses presented in this study may be rather weak. A larger sampling of the ECD population in Florida, including samples from the interior regions of the peninsula, which are lacking in this study, and the inclusion of immature as well as adult birds as found in the MD and WWD surveys, is needed to substantiate our conclusions. We thank the personnel of the Florida Fish and Wildlife Conservation Commission for collecting the doves in Pinellas County and the Health Departments of Santa Rosa, Okaloosa, and Bay counties for sending us the doves from the Florida Panhandle. We also thank Ellis Greiner and Patrick Meeus for reading earlier drafts of the manuscript and making useful suggestions for its improvement. This research was supported by the Florida Agricultural Experiment Station and approved for publication as Journal Series R-10041. LITERATURE CITED BUSH, A. O. 1990. Helminth communities in avian hosts: Determinations of pattern. In Parasite communities: Patterns and processes, G. W. Esch, A. O. Bush, and J. M. Aho (eds.). Chapman and Hall, London, U.K., p. 197–232. ———, K. D. LAFFERTY, J. K. LOTZ, AND A. W. SHOSTAK. 1997. Par-

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asitology meets ecology on its own terms: Margolis et al. revisited. Journal of Parasitology 83: 575–583. CONTI, J. A., AND D. J. FORRESTER. 1981. Interrelationships of parasites of the white-winged doves and mourning doves in Florida. Journal of Wildlife Diseases 17: 529–536. DURETTE-DESSET, M. C., R. A. GUERRERO, AND J. BOYER. 2000. Two Trichostrongylina (Nematoda) from Venezuela: A new species of Ornithostrongylus (Heligmosomoidea), parasitic in birds (Columbiformes) and a new species of Molineus (Molineoidea), parasitic in snakes (Squamata). Zoosystemia 22: 5–10. FORRESTER, D. J., J. A. CONTI, J. D. SHAMIS, W. J. BIGLER, AND G. L. HOFF. 1983. Ecology of helminth parasitism of mourning doves in Florida. Proceedings of the Helminthological Society of Washington 50: 143–152. ———, AND M. G. SPALDING. 2003. Parasites and diseases of wild birds in Florida. University Press of Florida, Gainesville, Florida, 1132 p. KINSELLA, J. M., AND D. J. FORRESTER. 1972. Helminths of the Florida duck, Anas platyrhynchos fulvigula. Proceedings of the Helminthological Society of Washington 39: 173–176. MAGURRAN, A. E. 1988. Ecological diversity and its measurement. Princeton University Press, Princeton, New Jersey, 179 p. MOLLHAGEN, T. 1976. The study of the systematics and hosts of the parasite nematode genus Tetrameres (Habronematoidea: Tetrameridae). Ph.D. Dissertation. Texas Tech University, Lubbock, Texas, 546 p. PIELOU, E. C. 1979. Biogeography. John Wiley and Sons, New York, 351 p. ROMAGOSA, C. M., AND R. F. LABISKY. 2000. Establishment and dispersal of the Eurasian collared-dove in Florida. Journal of Field Ornithology 71: 159–166. ———, AND T. MCENEANEY. 1999. Eurasian collared-dove in North America and the Caribbean. North American Birds 53: 348–353. SIMPSON, C. F., J. W. CARLISLE, AND J. A. CONTI. 1984. Tetrameres columbicola (Nematoda: Spiruridae) infection of pigeons: Ultrastructure of the gravid female in glands of the proventriculus. American Journal of Veterinary Research 45: 1184–1192. SMITH, P. W. 1987. The Eurasian collared-dove arrives in the Americas. American Birds 42: 1370–1379.


Clinical Muscular Sarcocystosis in a Dog J. Chapman, M. Mense, and J. P. Dubey*†, Department of Veterinary Pathology, Armed Forces Institute of Pathology, Washington, D.C. 20306-6000; * Animal Parasitic Diseases Laboratory, Animal and Natural Resources Institute, Beltsville Agricultural Research Center, United States Department of Agriculture, Beltsville, Maryland 20705-2350; † To whom correspondence should be addressed. e-mail: [email protected] ABSTRACT: Muscular sarcocystosis is a rare infection in dogs. Clinical myositis associated with an unidentified species of Sarcocystis was diagnosed in an adult dog from Canada. There was granulomatous myositis associated with numerous immature sarcocysts in a muscle biopsy obtained from the dog. The sarcocysts were up to 550 mm long and up to 45 mm wide. The sarcocyst wall was approximately 1 mm thick and contained short, stubby, villar protrusions that lacked microtubules. This is the first report on clinical muscular sarcocystosis in a dog.

Species of Sarcocystis have a 2-host, prey–predator, life cycle, with herbivores as intermediate hosts and carnivores as definitive hosts (Dubey et al., 1989). The intermediate host becomes infected with Sarcocystis spp. by ingesting sporocysts or oocysts, or both, excreted in the feces of the definitive host. After a brief period of schizogony, the parasite encysts in muscles and forms sarcocysts. The definitive host becomes infected by ingesting sarcocysts in infected muscles of intermediate hosts. Some animals act both as intermediate and definitive hosts but usually not for the same species of Sarcocystis (Dubey et al., 1989). The domestic dog (Canis familiaris) is a definitive host for nu-

merous species of Sarcocystis (Dubey et al., 1989), but only sexual stages are known to occur, and they are restricted to the intestine. In addition, there are reports on sarcocysts of unknown species in the muscles of 4 dogs (Sahasrabudhe and Shah, 1966; Hill et al., 1988; Blagburn et al., 1989; Bwangamoi et al., 1993). Findings of sarcocysts in a dog from India (Sahasrabudhe and Shah, 1966), from Georgia, U.S.A. (Hill et al., 1988), from Alabama, U.S.A. (Blagburn et al., 1989), and from Kenya (Bwangamoi et al., 1993) were incidental, apparently without any clinical signs. The dogs from Georgia and Kenya also had carcinoma. We report clinical myositis associated with numerous sarcocysts in a dog from Canada. A 5-yr-old, neutered, male Labrador cross dog with a history of lethargy, anorexia, and vomiting of 1 day’s duration was admitted in May 2003 to a veterinary hospital in British Columbia, Canada. The owners reported that the dog was hiking with them in high-range country 5 days earlier and had disappeared for a short time during the hike. Clinical evaluation indicated a fever (rectal temperature 41.5 C), dehydration, abdominal pain, and ataxia, with occasional petit mal seizure activity. Initial therapy included intravenous fluids and a broad-spectrum

188


FIGURE 1. Sarcocystis sp. sarcocysts in muscle biopsy of the dog. HE stain. (A) Necrosis and inflammation (arrow) and sarcocysts (arrowheads). (B) Giant cell (arrow), mononuclear cells, and sarcocysts (arrowheads). (C) Longitudinal section of a sarcocyst with metrocytes (small arrows). Note septa (arrowheads) and smooth sarcocyst wall (large arrow).

antibiotic. Blood was collected for hematologic evaluation (CellDyn 3500, Abbott Diagnostics, Santa Clara, California, and Dade Dimension RXL, Dade Behring Inc., Newark, Delaware). Results of the blood evaluation revealed a left shift characterized by 2% bands, moderate toxic change, and marginal total white blood cell (WBC) count (4.1 3 109/ L [reference 4.0–15.0 3 109/L]) as well as lymphopenia (0.492 [reference 0.960–4.800 3 109/L]). Although the hematocrit was within reference range (0.517 [0.390–0.560 L/L]), the serum total protein was low (51 g/L [reference 54–71 g/L]). Other abnormalities included a high alanine aminotransaminase (ALT) (3,436 IU/L [reference 0–113 IU/L]), alkaline phosphatase (312 IU/L [reference 04–113 IU/L]), creatinine kinase (CK) (841 IU/L [reference 0–314 IU/L]), and aspartate aminotransferase (AST) (2,829 IU/L [reference 8–56 IU/L]). The urine specific gravity was 1.032, with marked bilirubinuria and a 41 hematuria. Therapy was continued, and a repeat panel revealed a decrease in platelets but an improved WBC count and differential. The ALT/AST had declined, but the total bilirubin was markedly increased at 257 mmol/L (reference 0–7 mmol/L), with persistent bilirubinuria and hematuria. A serologic test for canine infectious hepatitis was consistent with previous vaccination. The dog was treated with dietary support, broad-spectrum antibiotics, nonsteroidal anti-inflammatory drugs, and glucocorticoids. The dog was reevaluated 5 wk after the initial episode for progressive inability to walk and dysphagia. The dog had muscle atrophy, weight loss, and generalized pain on palpation. Hematologic evaluation revealed a low albumin (24 g/L [reference range 31–42 g/L]), ALT (764 IU/L), AST (1,544 IU/L), and CK (13,206 IU/L). Joint fluid analysis was within normal limits and serologic tests for antinuclear antibody (antigen substrate slides [Hep-2] Antibodies Inc., Davis, California) and Toxoplasma gondii IgG titer (T. gondii substrate slides, Biomerieux, St. Laurent, Quebec, Canada) were negative. An indirect fluorescent antibody test (IFAT) titer to Neospora caninum using whole tachyzoites (N.

caninum antigen slides, VMRD, Pullman, Washington) revealed a titer of 1:800 or more. Muscles from shoulder and biceps regions were removed surgically and fixed in neutral buffered 10% formalin. Tissue was processed routinely, sectioned at 5 mm thickness, stained with hematoxylin and eosin (HE) or periodic acid–Schiff (PAS) reaction, and examined microscopically, revealing granulomatous myositis with protozoan tissue cysts. Deparaffinized sections of muscle were stained initially at the Central Laboratory for Veterinarians, Langley, British Columbia, Canada, using commercially available antibodies to T. gondii, and N. caninum (VMRD). The protozoan tissue cysts reacted with N. caninum antibodies. The dog was treated with clindamycin (Antirobe, Pfizer Canada Inc., Kirkland, Quebec, Canada, 12.5 mg/kg twice daily) the day after biopsy, which continued for 9 wk. Improvement in the dog’s clinical condition was noted within 1 wk. A repeat hematological evaluation 3.5 mo after the initial episode revealed a decline in ALT to 128 IU/L, CK to 752 IU/L, and antibody titer to N. caninum to 1:200. The dog has remained clinically normal during the past year. Slides from this case were submitted to the Armed Forces Institute of Pathology, Washington, D.C., for teaching purposes. Further immunohistochemical examination was performed at the Animal Parasitic Diseases Laboratory, Beltsville, Maryland. Deparaffinized sections were reacted with antibodies to T. gondii, N. caninum, Sarcocystis neurona, S. cruzi, and BAG-1 rabbit antibodies using the procedures described by Dubey et al. (2001). The specificity and preparation of antibodies to S. neurona (Dubey et al., 1999), and N. caninum and T. gondii (Lindsay and Dubey, 1989) were as described. The BAG1 antibodies were a gift from McAllister et al. (1996); these are specific to bradyzoites of cyst-forming coccidia (McAllister et al., 1996; Dubey and Sreekumar, 2003). The S. cruzi serum was prepared against bradyzoites of S. cruzi, which is genus specific (Granstrom et al., 1991).

RESEARCH NOTES

189

FIGURE 2. Transmission electron micrographs of sarcocysts from an unidentified species of Sarcocystis from a dog. (A) The host cell (Hc) is degenerate (arrow). The sarcocyst wall consists of short villar protrusions(Vp) and a homogenous ground substance layer (Gs), which is continued in to the sarcocyst interior as septa (Se). One bradyzoite (Br) and several metrocytes fill the sarcocyst. (B) Higher magnification of the sarcocyst wall shows villar protrusions (Vp) that abut the host cell (Hc). The Vp are lined by an electron-dense layer (Edl), which is interrupted at irregular distances (arrowheads). The ground substance (Gs) is homogenous.

A portion of the biopsy tissue was deparaffinized and processed for transmission electron microscopy. Examination of numerous sections confirmed necrotizing myositis with pyogranulomatous inflammation (Fig. 1). Skeletal muscle bundles were surrounded and separated by neutrophils, macrophages, and multinucleated giant cells, which occasionally formed larger accumulations

that replaced muscle bundles (Fig. 1A, B). There was myocyte degeneration and necrosis, characterized by individualized myocytes with hypereosinophilic sarcoplasm, vacuolation, loss of cross-striations, and pyknosis. Regenerating myocytes had lightly basophilic sarcoplasm, with multiple central nuclei. Numerous sarcocysts of an unidentified species of Sarcocystis were

190


associated with lesions (Fig. 1A, B). These sarcocysts were immature, septate, and contained a few to numerous metrocytes (Fig. 1C). The sarcocyst wall was thin (,1 mm thick) and without striations. They were up to 550 mm long and up to 45 mm wide. Sarcocysts contained metrocytes that were PAS positive. Four sarcocysts were examined ultrastructurally. The sarcocyst wall had short, stubby villar protrusions (Vp) on the entire wall (Fig. 2). The Vp were up to 0.8 mm long and up to 0.7 mm wide, narrow at the base, and expanded laterally (Fig. 2B). The wall of the sarcocyst (parasitophorous vacuolar membrane, PVM) had minute undulations. The PVM was lined by a 70-nm-thick electron-dense layer that was interrupted at irregular distances. The interior of the Vp was homogenous, without any microtubules. Underneath the Vp was the homogenous granular substance (Gs) layer that was approximately 0.25 mm thick. The total sarcocyst wall, including the Vp and the Gs, was approximately 1.0 mm thick in longitudinal section. The Gs was continued into the interior of the sarcocyst as septa. The sarcocysts contained numerous metrocytes that lacked mylopectin granules. Only 1 bradyzoite was found among organisms in 4 sarcocysts examined ultrastructurally (Fig. 2A). Protozoans in muscles did not react with antibodies to N. caninum, T. gondii, S. neurona, or BAG-1 antibodies but reacted positively with S. cruzi antibodies. These results support the morphological diagnosis of sarcocystosis due to an unidentified species. The lack of staining by BAG-1 antibodies indicates the absence of bradyzoites in the sarcocyst. The species of the Sarcocystis in the dog in this study was not determined because sarcocysts were immature although the Vp on the sarcocyst wall appeared to be unique. The structure of the sarcocyst wall is considered as 1 of the most useful criteria to distinguish Sarcocystis species within a given host. Dubey et al. (1989) and Dubey and Odening (2001) divided the structure of the cyst wall into 37 types based on their ultrastructure. In general, the structure of the sarcocyst walls in 2 dogs from Georgia and Alabama was similar to that in the present case, although few details were provided by Hill et al. (1988) and Blagburn et al. (1989) to permit a species determination. Finding of numerous immature sarcocysts at about the same stage of development along with inflammatory lesions suggests that the dog ingested many Sarcocystis sp. sporocysts at one time (Dubey et al., 1989). Clinical sarcocystosis is rarely diagnosed even though subclinical infections are common in livestock. One reason for this is that the clinical disease is usually associated with the rupture of second-generation intravascular schizonts, and by the time tissues are submitted for diagnosis, the second-generation schizonts have disappeared. The merozoites released from schizonts invade myocytes and may incite host response. By the time sarcocyst mature in host tissues, inflammation usually has subsided; therefore, it is difficult to diagnose clinical sarcocystosis. In most Sarcocystis species, bradyzoites are formed by 75 days postinoculation (Dubey et al., 1989). If this were to apply in the present case, the dog probably became infected with Sarcocystis sp. within 3 mo of being biopsied. Whether the dog was immunosuppressed at the time of acute sarcocystosis is not known. Little is known of antemortem diagnosis and chemotherapy for sarcocystosis in dogs. Elevated CK values indicated muscle injury, and the results of biopsy examinations indicated that the dog had severe myositis related to parasitism with Sarcocystis sp. Bilirubinuria and lowered protein values might have been due to hepatic injury, probably associated with schizogony preceding the formation of sarcocysts (Dubey et al., 1989). The dog in this study responded favorably to

treatment with clindamycin. The clindamycin therapy was initiated because of the high serum antibody titer to N. caninum and reactivity of the parasite in sections to VMRD N. caninum antibodies prepared in a goat. The cross-reactivity of the sarcocysts with VMRD N. caninum serum might have been natural Sarcocystis sp. infection in the goat used to prepare antiserum to N. caninum. The parasite did not react with N. caninum antibodies prepared in a rabbit. The presence of an IFAT N. caninum titer of 1:800 only indicates exposure, and the dog might not have clinical neosporosis; N. caninum was not detected in muscle biopsy. The authors thank Sally Lester, Central Laboratory for Veterinarians, Langley, British Columbia, Canada, for her help in preparation of this paper and Sean Hahn for technical assistance. LITERATURE CITED BLAGBURN, B. L., K. G. BRAUND, K. A. AMLING, AND M. TOIVIO-KINNUCAN. 1989. Muscular Sarcocystis in a dog. Proceedings of the Helminthological Society of Washington 56: 207–210. BWANGAMOI, O., T. A. NGATIA, AND J. D. RICHARDSON. 1993. Sarcocystis-like organisms in musculature of a domestic dog (Canis familiaris) and wild dogs (Lycaon pictus) in Kenya. Veterinary Parasitology 49: 201–205. DUBEY, J. P., M. M. GARNER, M. D. STETTER, A. E. MARSH, AND B. C. BARR. 2001. Acute Sarcocystis falcatula-like infection in a carmine bee-eater (Merops nubicus) and immunohistochemical cross reactivity between Sarcocystis falcatula and Sarcocystis neurona. Journal of Parasitology 87: 824–831. ———, D. E. MATTSON, C. A. SPEER, R. J. BAKER, D. M. MULROONEY, S. J. TORNQUIST, A. N. HAMIR, AND T. C. GERROS. 1999. Characterization of Sarcocystis neurona isolate (SN6) from a naturally infected horse from Oregon. Journal of Eukaryotic Microbiology 46: 500–506. ———, C. A. SPEER, AND R. FAYER. 1989. Sarcocystosis of animals and man. CRC Press, Boca Raton, Florida, p. 1–215. ———, AND C. SREEKUMAR. 2003. Redescription of Hammondia hammondi and its differentiation from Toxoplasma gondii. International Journal for Parasitology 33: 1437–1453. GRANSTROM, D. E., R. C. GILES, P. A. TUTTLE, N. M. WILLIAMS, K. B. POONACHA, M. B. PETRITES-MURPHY, R. R. TRAMONTIN, T. W. SWERCZEK, C. B. HONG, G. B. REZABEK, E. T. LYONS, AND J. H. DRUDGE. 1991. Immunohistochemical diagnosis of protozoan parasites in lesions of equine protozoal myeloencephalitis. Journal of Veterinary Diagnostic Investigation 3: 75–77. HILL, J. E., W. L. CHAPMAN, AND A. K. PRESTWOOD. 1988. Intramuscular Sarcocystis sp. in two cats and a dog. Journal of Parasitology 74: 724–727. LINDSAY, D. S., AND J. P. DUBEY. 1989. Immunohistochemical diagnosis of Neospora caninum in tissue sections. American Journal of Veterinary Research 50: 1981–1983. MCALLISTER, M. M., S. F. PARMLEY, L. M. WEISS, V. J. WELCH, AND A. M. MCGUIRE. 1996. An immunohistochemical method for detecting bradyzoite antigen (BAG5) in Toxoplasma gondii-infected tissues cross-reacts with a Neospora caninum bradyzoite antigen. Journal of Parasitology 82: 354–355. SAHASRABUDHE, V. K., AND H. L. SHAH. 1966. The occurrence of Sarcocystis sp. in the dog. Journal of Protozoology 13: 531.

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Detection of Antibodies to Neospora caninum in Cattle by Enzyme-Linked Immunosorbent Assay with Truncated NcSRS2 Expressed in Escherichia coli Irungu Gaturaga*, Bayin Chahan*, Xuenan Xuan†, Xiaohong Huang, Min Liao, Shinya Fukumoto, Haruyuki Hirata, Yoshihumi Nishikawa, Yasuhiro Takashima‡, Hiroshi Suzuki, Kozo Fujisaki, and Chihiro Sugimoto, National Research Center for Protozoan Diseases, Obihiro University of Agriculture and Veterinary Medicine, Inada-cho, Obihiro, Hokkaido 080-8555, Japan; *Contributed equally to this work; †To whom correspondence should be addressed; ‡Department of Global Agricultural Science, Graduate School of Agriculture and Life Science, University of Tokyo, Bunkyo-ku, Tokyo, Japan. e-mail: [email protected] ABSTRACT: The surface antigen 1–related sequence 2 of Neospora caninum (NcSRS2) is considered as an immunodominant antigen. In this study, the gene encoding truncated NcSRS2 (NcSRS2t) lacking an Nterminal signal peptide and C-terminal hydrophobic regions was expressed in Escherichia coli, and its diagnostic potential in an enzymelinked immunosorbent assay (ELISA) was evaluated. ELISA could discriminate clearly between known N. caninum–positive and –negative sera from cattle. Field serum samples collected from cattle in Brazil were examined for the diagnosis of N. caninum infection using ELISA. Of the 197 samples analyzed, 64 (32.5%) samples were positive for antibodies to N. caninum. Of the 64 ELISA-positive samples, 58 (90.6%) were confirmed as positive by Western blot analysis with whole-parasite antigens. These results suggest that ELISA with recombinant NcSRS2t is an effective method for diagnosis of N. caninum infection in cattle.

Neospora caninum is an apicomplexan parasite, which causes abortion, neonatal neurological diseases, or both, in a variety of domestic animals, such as cattle, dogs, goats, sheep, and horses (Dubey, 1999). Dogs are the only definitive host known to excrete the oocysts of N. caninum (McAllister et al., 1998). To date, many serological methods, i.e., immunofluorescent antibody test (IFAT), enzyme-linked immunosorbent assay (ELISA), Western blotting, and direct agglutination test using intact tachyzoites or tachyzoite-derived antigens, have been developed for the detection of antibodies to N. caninum in animals (Jenkins et al., 2002). However, the use of whole tachyzoites or tachyzoite-derived antigens sometimes may result in false positives due to cross-reaction with other closely related parasites, such as Toxoplasma gondii. It is therefore necessary to develop a specific diagnostic test using N. caninum–specific antigens. The surface antigen 1–related sequence 2 of N. caninum (NcSRS2) is an important candidate for developing a diagnostic reagent for the detection of specific antibodies to N. caninum (Howe et al., 1998; Nishikawa, Kousaka et al., 2001). In this study, the gene encoding truncated NcSRS2, which lacks an N-terminal signal peptide and C-terminal hydrophobic regions, was expressed in Escherichia coli, and its diagnostic potential in ELISA was evaluated. The genomic DNA extracted from Nc-1 strain of N. caninum (Dubey et al., 1988) was used as template for polymerase chain reaction (PCR). The truncated NcSRS2 (NcSRS2t) gene (Howe et al., 1998) without sequences encoding a hydrophobic signal peptide (amino acids 1–53) and a C-terminus (amino acids 383–400) was amplified by PCR using oligonucleotide primers, 59-ACGAATTCTGCGCCGTTCAAGTCG-39 and 59-ACGAATTCAAGGCAACTCGTCGTC-39, that contain introduced EcoRI sites to facilitate cloning. The PCR product was digested with EcoRI, cloned into the EcoRI site of the bacterial expression vector pGEX-4T-3 (Amersham Biosciences, Piscataway, New Jersey), and then expressed as a glutathione S-transferase (GST) fusion protein (GST– NcSRS2t) in E. coli (DH5a strain). The GST–NcSRS2t was purified with Glutathione Sepharose 4B, according to the manufacturer’s instructions (Amersham Biosciences). ELISA with GST–NcSRS2t was performed as described previously (Chahan et al., 2003). GST–NcSRS2t and control GST antigen were diluted 5 mg/ml, and all serum samples were diluted 1:100. The ELISA result was determined for each sample by taking the mean optical density value of 2 readings with GST– NcSRS2t and subtracting the mean value of 2 readings with GST control antigen. A sample was considered positive if the calculated absorbance value was greater than or equal to 0.1.

A portion of the recombinant GST–NcSRS2t was expressed as a soluble form, in contrast to GST–NcSRS2 (full length), which was completely expressed as an insoluble form (data not shown). This result indicates that removal of the highly hydrophobic signal peptide and Cterminus had improved its hydrophilicity. The molecular mass of reduced GST–NcSRS2t was estimated as 61 kDa (GST, 26 kDa, 1 NcSRS2t, 35 kDa) as expected (Fig. 1A). Western blot analysis shows that NcSRS2t reacted strongly with sera from N. caninum–infected cattle (Fig. 1B), dog (Fig. 1C), and mouse (Fig. 1D) but not with sera from uninfected animals or from T. gondii–infected mice (data not shown). In addition, similar results were obtained in Western blot analysis when nonreduced GST–NcSRS2t was used. This result indicates

FIGURE 1. Sodium dodecyl sulfate–polyacrylamide gel electrophoresis and Western blot analyses of reduced recombinant NcSRS2t. A. Antigens stained with amido black 10B. B. Antigens reacted with sera from Neospora caninum–infected cattle. C. Antigens reacted with serum from experimentally N. caninum–infected dog. D. Antigens reacted with serum from experimentally N. caninum–infected mouse. Lane 1, GST–NcSRS2t; lane 2, GST; lane 3, N. caninum tachyzoite lysate; and lane 4, Vero cell lysate.

TABLE I. Prevalence of bovine Neospora caninum infection on 6 farms in Mato Grosso do Sul, Brazil.

Farm

No. examined

A B C D E F Total

26 32 21 15 49 54 197

No. seropositive* 14 14 10 4 11 11 64

(53.8) (43.8) (47.6) (26.0) (22.4) (20.4) (32.5)

* Antibodies to N. caninum were detected by ELISA with GST–NcSRS2t. The ELISA was considered positive when an optical density at 415 nm greater than or equal to 0.1 was observed at dilutions of 1:100 and more. Values in parentheses are percentages.

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TABLE II. Comparison of ELISAs with GST–NcSRS2t and GST–NcSAG1t for detection of antibodies to Neospora caninum in cattle.

GST–NcSAG1t1† GST–NcSAG1t2 Total

GST– NcSRS2t1*

GST– NcSRS2t2

61 (31.0) 3 (1.5) 64 (32.5)

5 (2.5) 128 (65.0) 133 (67.5)

Total 66 (33.5) 131 (66.5) 197 (100.0)

* The ELISA using GST–NcSRS2t as antigen. Values in parentheses are percentages. † The ELISA using GST–NcSAG1t as antigen (Chahan et al., 2003).

that the recombinant NcSRS2t retained its immunoreactivity after deleting both N- and C-terminuses. To evaluate whether recombinant NcSRS2t expressed in E. coli can be a suitable antigen for the diagnosis of N. caninum infection, the purified GST–NcSRS2t was tested by ELISA using sera from mice experimentally infected with either N. caninum or T. gondii and sera from uninfected mice. All sera from BALB/c mice infected with N. caninum Nc-1 strain (n 5 10, 2 mo postinfection) were positive, with optical density ranging from 0.5 to 2.0, whereas all sera from uninfected BALB/c mice (n 5 10) or from BALB/c mice infected with T. gondii Beverley strain (n 5 10, 2 mo postinfection) were negative, with optical density ranging from 0 to 0.05. In addition, the ELISA with GST– NcSRS2t was evaluated using sera from dogs experimentally pre- and postinfected with N. caninum Nc-1 strain (Beagle, n 5 4, 1 mo postinfection). All sera from infected dogs were positive, with optical density ranging from 0.8 to 2.0, whereas all sera from dogs before infection were negative, with optical density ranging from 0 to 0.06. ELISA with GST–NcSRS2t was also evaluated using known seropositive and seronegative bovine sera previously diagnosed by IFAT (Nishikawa, Inoue et al., 2001). All IFAT-positive sera (n 5 20) were positive, with optical density ranging from 0.15 to 2.0, whereas all IFAT-negative sera (n 5 20) were negative, with optical density ranging from 0 to 0.08. Serum samples collected from cattle in Brazil were then tested for detection of antibodies to N. caninum by ELISA with GST–NcSRS2t. As shown in Table I, of the 197 sera tested, 64 (32.5%) were positive. The positive rates among 6 farms ranged from 20.4 to 53.8%. Of the 64 ELISA-positive samples, 58 (90.6%) were confirmed as positive by Western blot analysis with whole parasite as antigen. The antibody to NcSRS2t was detected in cattle aged from a few months to 11 yr (data not shown). Recently, we reported that ELISA with GST–NcSAG1t was useful for the detection of antibodies to N. caninum in cattle (Chahan et al., 2003). In this study, the ELISA with GST–NcSRS2t was compared with the ELISA with GST–NcSAG1t (Table II). Although there was a good correlation between the 2 ELISAs, 3 negative ELISA samples with GST–NcSAG1t were positive by ELISA with GST–NcSRS2t, whereas 5 negative ELISA samples with GST–NcSRS2t were positive by ELISA with GST–NcSAG1t. This result indicates that a combination of the 2 ELISAs or ELISA with a mixture of the 2 recombinant antigens may provide a more effective tool for the detection of antibodies to N. caninum in cattle. Baszler et al. (2001) has recently validated that a commercially available monoclonal antibody–based competitive inhibition ELISA (cELISA) is useful for the detection of antibodies to N. caninum in cattle. Our next step will be to compare ELISA using both GST– NcSAG1t and GST–NcSRS2t with commercial cELISA to validate the former to be a useful method for the detection of serum antibodies to N. caninum in cattle. Previous studies have shown that mice vaccinated with recombinant vaccinia virus expressing NcSRS2 were protected from N. caninum infection (Nishikawa, Inoue et al., 2001; Nishikawa, Xuan et al., 2001). Recently, Cannas et al. (2003) reported that the mice immunized with NcSRS2-based recombinant antigen and DNA vaccine were protected from N. caninum infection. The next step will be to implement immunization trials with animals to determine the potency of recombinant NcSRS2t expressed in E. coli as a potential subunit vaccine to control N. caninum infection.

This study was supported by a grant from The 21st Century COE Program (A-2), Ministry of Education, Culture, Sports, Science, and Technology, Japan, and Grants-in-Aid for Scientific Research from the Japan Society for the Promotion of Science. LITERATURE CITED BASZLER, T. V., S. ADAMS, J. VANDER-SCHALIE, B. A. MATHISON, AND M. KOSTOVIC. 2001. Validation of a commercially available monoclonal antibody-based competitive-inhibition enzyme-linked immunosorbent assay for detection of serum antibodies to Neospora caninum in cattle. Journal of Clinical Microbiology 39: 3851–3857. CANNAS, A., A. NAGULESWARAN, N. MULLER, S. EPERON, B. GOTTSTEIN, AND A. HEMPHILL. 2003. Vaccination of mice against experimental Neospora caninum infection using NcSAG1- and NcSRS2-based recombinant antigens and DNA vaccines. Parasitology 126: 303– 312. CHAHAN, B., I. GATURAGA, X. HUANG, M. LIAO, S. FUKUMOTO, H. HIRATA, Y. NISHIKAWA, H. SUZUKI, C. SUGIMOTO, H. NAGASAWA, K. FUJISAKI, I. IGARASHI, T. MIKAMI, AND X. XUAN. 2003. Serodiagnosis of Neospora caninum infection in cattle by enzyme-linked immunosorbent assay with recombinant truncated NcSAG1. Veterinary Parasitology 118: 177–185. DUBEY, J. P. 1999. Neosporosis—The first decade of research. International Journal for Parasitology 29: 1485–1488. ———, A. L. HATTEL, D. S. LINDSAY, AND M. J. TOPPER. 1988. Neonatal Neospora caninum infection in dogs: Isolation of the causative agent and experimental transmission. Journal of the American Veterinary Medical Association 193: 1259–1263. HOWE, D. K., A. C. CRAWFORD, D. LINDSAY, AND L. D. SIBLEY. 1998. The p29 and p35 immunodominant antigens of Neospora caninum tachyzoites are homologous to the family of surface antigens of Toxoplasma gondii. Infection and Immunity 66: 5322–5328. JENKINS, M., T. BASZLER, C. BJORKMAN, G. SCHARES, AND D. WILLIAMS. 2002. Diagnosis and seroepidemiology of Neospora caninum-associated bovine abortion. International Journal for Parasitology 32: 631–636. MCALLISTER, M. M., J. P. DUBEY, D. S. LINDSAY, W. R. JOLLEY, R. A. WILLS, AND A. M. MCGUIRE. 1998. Dogs are definitive hosts of Neospora caninum. International Journal for Parasitology 28: 1473–1478. NISHIKAWA, Y., N. INOUE, X. XUAN, H. NAGASAWA, I. IGARASHI, K. FUJISAKI, H. OTSUKA, AND T. MIKAMI. 2001. Protective efficacy of vaccination by recombinant vaccinia virus against Neospora caninum infection. Vaccine 19: 1381–1390. ———, Y. KOUSAKA, K. TRAGOOLPUA, X. XUAN, L. MAKALA, K. FUJISAKI, T. MIKAMI, AND H. NAGASAWA. 2001. Characterization of Neospora caninum surface protein NcSRS2 based on baculovirus system and its application for serodiagnosis of Neospora infection. Journal of Clinical Microbiology 39: 3987–3991. ———, X. XUAN, H. NAGASAWA, I. IGARASHI, K. FUJISAKI, H. OTSUKA, AND T. MIKAMI. 2001. Prevention of vertical transmission of Neospora caninum in BALB/c mice by recombinant vaccinia virus carrying NcSRS2 gene. Vaccine 19: 1710–1716.

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Effects of a- and b-Adrenergic Agonists on Toxoplasma gondii Infection in Murine Macrophages Julie Getz and Fernando P. Monroy*, Department of Biological Sciences, Northern Arizona University, P.O. Box 5640, Flagstaff, Arizona 86011; *To whom correspondence should be addressed. e-mail: [email protected] ABSTRACT: We investigated the effects of a- and b-adrenergic receptor agonists on the ability of Toxoplasma gondii to infect and proliferate in cultured murine macrophages. Macrophages pretreated in vitro with varying concentrations of a- and b-adrenergic agonists and incubated with the RH strain of T. gondii did not result in a significant increase in the percentage of infected macrophages compared with negative controls. When parasites were pretreated with L-phenylephrine, an a-agonist, and L-isoproterenol, a b-agonist, before infection, there was no significant change in the percentage of infected macrophages. Clonidine, an a2-adrenergic agonist, led to a significant decrease in the number of infected macrophages at all concentrations tested. The effects of clonidine were blocked by yohimbine, a specific a2-adrenergic antagonist, but not by phentolamine, an a1-adrenergic antagonist. These results suggest that the antiparasitic effects exhibited by clonidine (a2-adrenergic agonist) are mediated through an a2-adrenoreceptor found on the surface of T. gondii.

Toxoplasma gondii is an opportunistic protozoan parasite capable of developing acute and chronic infections in mice and humans. During the acute stage, T. gondii tachyzoites infect and destroy host cells; in the chronic stage, tissue cysts are formed within the host (Dubey, 1998). Chronic infections can persist for months, years, or even the lifetime of the host, causing reactivation when cysts rupture and bradyzoites are released, infecting the surrounding host organs (Dubey, 1998). During infection with T. gondii, macrophage secretion of interleukin (IL)-12, and tumor necrosis factor (TNF)–a are crucial to the control of T. gondii infection in the host. Macrophages infected with T. gondii secrete IL12 and TNF-a, which in turn stimulates a T helper cell–mediated response, promoting the secretion of interferon (IFN)-g by T cells (Gazzinelli et al., 1996). Macrophages activated by IFN-g use phagocytosis, phagolysosomal degradation, nitric oxide (NO) intermediate production, and respiratory burst activity to mediate the inhibition of parasite proliferation within the host (Hunter et al., 1996). Host susceptibility to infection has been shown to be modulated by stress (Elenkov and Chrousos, 1999). Cold-water stress (CWS), a physical stressor, has been shown to modulate macrophage immune responses during T. gondii infection (Banerjee et al., 1999; Aviles and Monroy, 2001). Macrophages from CWS mice secreted lower levels of TNF-a and showed low levels of nitric oxide synthase and were unable to prevent parasite proliferation (Monroy et al., 1999). Catecholamines, norepinephrine, and epinephrine are naturally released in the body by the sympathetic nervous system in response to stress (Broug-Holub et al., 1998). In addition, the effects of catecholamines on immune cells are dependent on the type of adrenergic receptor that is engaged. Activation through a-adrenergic receptors leads to the activation of macrophages, whereas activation through b-adrenergic receptors leads to downmodulation in macrophage activation (Spengler et al., 1990). Therefore, it is important to determine what type of receptor is involved during T. gondii infection because both types of receptors are present in macrophages. We investigated the effects of a- and b-adrenergic receptor agonists on T. gondii’s ability to infect and proliferate in murine macrophages. Macrophage pretreatment with a- and b-adrenergic agonists did not result in any significant effect when compared with controls. Parasite pretreatment with clonidine, an a2-adrenergic agonist, resulted in a significant decrease in the number of infected macrophages at all concentrations tested. The specific effects of clonidine were blocked by yohimbine, an a2-adrenergic antagonist, but not by phentolamine, an a1adrenergic antagonist. These results suggest that the antiparasitic effects exhibited by clonidine are mediated through an a2-adrenoreceptor found on the surface of T. gondii. Monolayers of RAW 264.7 macrophages (American Type Culture

Collection, Manassas, Virginia) were maintained in Dulbecco minimal essential medium (DMEM) supplemented with 10% fetal calf serum, 1% glutamine, and 100 mg/ml streptomycin at 37 C, 5% carbon dioxide (CO2). The RH strain of T. gondii was maintained by routine passages of tachyzoites in human foreskin fibroblast (HFF) cells in 25-cm2 tissue culture flasks. Parasites were purified by serial passage through 20, 23, and 27 gauge needles, followed by passage through a 3.0-mm Nucleopore filter (Costar Corporation, Pleasanton, California). Parasite concentrations were determined using a hemocytometer and resuspended at 5 3 105 parasites/ml to obtain a ratio of 5:1 parasites to macrophages. The adrenergic agonists, L-isoproterenol (b), L-phenylephrine (a1), and clonidine (a2), and the a-adrenergic antagonists, phentolamine (a1) and yohimbine (a2), were purchased from Sigma (Sigma Chemical Co., St. Louis, Missouri). Concentrations to be tested were 0.1, 1.0, and 10 mM in DMEM (Shen et al., 1994). Macrophages (1 3 105/ml) were cultured in 8-well chamber slides (Nalge NUNC Int., Naperville, Illinois). The chamber slides were incubated for 1 hr at 37 C, 5% CO2, to allow macrophages to adhere and washed 3 times with Ca21-free phosphate buffered saline (PBS). When agonists and antagonists were tested, they were incubated with macrophages or parasites for 1 hr at 37 C, 5% CO2, and washed with Ca21-free PBS by centrifugation at 2,200 g, 4 C for 8 min. When antagonists (yohimbine [a2 specific] and phentolamine [a1 specific]) were used to treat parasites, they were present in the same molar concentrations as the agonist (clonidine). After incubation and washing, cells were infected with 5 3 105/ml parasites and incubated for 18–24 hr at 37 C, 5% CO2. The cultures were terminated by washing the chamber wells with Ca21-free PBS and then fixed and stained with Diff-Quick (Dade Behring Ag, Dundingen, Switzerland). Slides were examined microscopically at 31,000, recording the total number of macrophages and the number of cells infected with 1 or more parasites. These results are representative of the means 6 SD of 3 different experiments for each protocol. Differences were determined to be significant if P , 0.05, determined by the Student’s t-test (P , 0.05). When macrophages were pretreated with the adrenergic agonists clonidine, L-phenylephrine, or L-isoproterenol at concentrations 0.1, 1.0, or 10 mM, there was no significant difference in the percentage of infected macrophages by the treatments compared with negative controls (Fig. 1A). Furthermore, no difference was also found among the treatments in the number of intracellular parasites, which ranged from 7.8 6 0.7 to 9.2 6 0.8. To determine the effect adrenergic agonists have on T. gondii, parasites were pretreated with clonidine, L-isoproterenol, or Lphenylephrine at 0.1, 1.0, or 10 mM concentrations in the same 5:1 macrophage to parasite ratio used above. As shown in Figure 1B, parasite incubation with 1 or 10 mM L-phenylephrine (a1) resulted in a slight increase in the percentage of infected macrophages by 5.55 and 10.13%, respectively, but overall there was no significant change in the percentage of infected macrophages or in the number of intracellular parasites (8.1 6 0.8 to 9.6 6 1.0) for any concentration. Pretreatment of T. gondii with L-isoproterenol, a b-agonist, resulted in no significant effect, although there was a slight but not significant decrease in intracellular parasites ranging from 7.1 6 0.6 to 8.2 6 0.8. Clonidine, an a2-adrenergic agonist, led to a significant decrease in the number of infected macrophages at 10 and 1.0 mM concentrations (P , 0.05) but not at 0.1 mM. Similarly, the number of intracellular parasites decreased from 6.8 6 0.4 at 0.1 mM to 4.6 6 0.3 (P , 0.05) and 3.0 6 0.2 (P , 0.05) for 1 and 10 mM, respectively. To determine whether a1 or a2 receptors were being used during the antiparasitic effect of clonidine, specific a-blockers, yohimbine (a2) and phentolamine (a1), were used in comparable molar concentrations with clonidine. Coculture of T. gondii with clonidine and phentolamine, a specific a1-blocker, showed no effect. However, when yohimbine, a specific a2-blocker, was used in

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FIGURE 2. Effects of a-adrenergic antagonists on the ability of Toxoplasma gondii to infect and replicate within macrophages. Parasites (5: 1 macrophage to parasite ratio) were incubated with clonidine and equimolar concentrations of yohimbine (a2) or phentolamine (a1) for 1 hr at 37 C, as described above. Values represent mean 6 SD of 3 repeat experiments.

FIGURE 1. Effects of pretreatment of macrophages (A) and parasites (B) with adrenergic receptor agonists on intracellular replication of Toxoplasma gondii. Macrophages or parasites were pretreated with varying molar concentrations of adrenergic agonists at 37 C for 1 hr, washed, incubated with parasites at a 5:1 macrophage to parasite ratio for 24 hr, and prepared for microscopy. Values represent means 6 SD of 3 repeat experiments.

the coculture, the changes mimicked those of control trials (Fig. 2). Once again, the number of intracellular parasites significantly decreased from 9.5 6 0.8 to 3.6 6 0.4 when clonidine at 0.1 and 10 mM was incubated with phentolamine. No significant difference was found when concentrations of clonidine were incubated with the a2-blocker yohimbine (range from 8.3 6 0.8 to 9.2 6 0.9). To control T. gondii infection, macrophages require activation by IFN-g to induce killing of the parasite (Suzuki et al., 1989). Macrophages produce IL-12 and TNF-a, which are crucial in the control of T. gondii infection by the host (Hunter et al., 1996). Our laboratory has previously shown that CWS modulated macrophage immune responses during T. gondii infection, resulting in an increased percentage of infected cells and increased number of intracellular parasites. This decreased ability of macrophages to control T. gondii replication was associated with low levels of nitric oxide (NO) production (Banerjee et al., 1999) and decreased cytokine production (Monroy et al., 1999; Aviles and Monroy, 2001). Adrenergic agonists mimic catecholamines that are naturally released in the body by the sympathetic nervous system in response to stress; these agents have been reported to have an effect on parasite infection both in vitro and in vivo (Connelly et al., 1988; Benedetto et al., 1993; Broug-Holub et al., 1998; Aviles and Monroy, 2001). In this report, we investigated the effects of a- and b-adrenergic receptor agonists on T. gondii’s ability to infect and proliferate in murine macrophages. Previous reports have shown that macrophages (Shen et

al., 1994; Szelenyi et al., 2000), human umbilical vein endothelial cells (HUVEC) (Benedetto et al., 1997), and dendritic cells (Fermin et al., 1999) possess a- and b-like adrenergic receptors that can influence immune response to infection. Pretreatment of HUVEC with a2-agonists resulted in increased killing of T. gondii. These anti–T. gondii effects in HUVEC were mediated through a2-adrenoceptors, and parasite killing was associated with increased NO production (Benedetto et al., 1997). In the hemoprotozoan parasite, Trypanosoma cruzi, Connelly et al. (1988) reported the presence of both a- and b-adrenergic receptors in trypomastigotes. Nevertheless, preincubation of T. cruzi trypomastigotes with L-isoproterenol (b-agonist) decreased the percentage of infected macrophages and the number of intracellular parasites, whereas pretreatment with the a-adrenergic agonist, L-phenylephrine, resulted in increased percentage of infected macrophages as well as the number of intracellular parasites. In this report, pretreatment of parasites with clonidine, an a2-adrenergic agonist, reduced macrophage infection with T. gondii. The abolishment of clonidine’s ability to decrease macrophage infection when cocultured with yohimbine (specific a2-adrenergic blocker) further established the presence of a2-like adrenergic receptors on the surface of T. gondii. In conclusion, our results show the following (1) a-adrenergic agonists can have significant effects on the ability of T. gondii to multiply within macrophages, (2) the antiparasitic effects of clonidine seem to be mediated through a2-adrenergic receptors on the surface of T. gondii, and (3) although the function of a2-adrenoceptors on T. gondii is unknown, we can speculate that activation of a2-adrenceptors leads to decreased cyclic adenosine monophosphate (cAMP) levels (Exton, 1981). It is possible that this effect on T. gondii infectivity resulted from decreased cAMP levels required for parasite growth and differentiation in mammalian cells (Choi et al., 1990; Kirkman et al., 2001). Further studies should include determining the role of stress on an adrenergic receptor expression and the effects of a2-adrenergic agonist on cytokine production during infection with T. gondii. This research was supported by the NIGMS-IMSD Program and Extramural Research Support at Northern Arizona University. LITERATURE CITED AVILES, H., AND F. P. MONROY. 2001. Immunomodulatory effects of cold stress on mice infected intraperitoneally with a lethal dose 50 (LD50) of Toxoplasma gondii. Neuroimmunomodulation 9: 6–13. BANERJEE, S., H. AVILES, M. FOX, AND F. P. MONROY. 1999. Cold stressinduced modulation of cell immunity during acute Toxoplasma gondii infection in mice. Journal of Parasitology 85: 442–448.

RESEARCH NOTES

BENEDETTO, N., A. FOLGORE, C. FERRARA, M. MOLITIERNO, AND F. GALDIERO. 1997. Effects of a-adrenergic agonists on Toxoplasma gondii replication in human umbilical vein endothelial cells. Pathologie Biologie 45: 9–18. ———, ———, AND M. GALDIERO. 1993. Impairment of natural resistance to Toxoplasma gondii infection in rats treated with b adrenergics, b blockers, corticosteroids or total body irradiation. Pathologie Biologie 41: 404–409. BROUG-HOLUB, E., J. H. A. PERSOONS, K. SCHORNAGEL, S. C. MASTBERGEN, AND G. KRAAL. 1998. Effects of stress on alveolar macrophages: A role for the sympathetic nervous system. American Journal of Respiratory Cellular and Molecular Biology 19: 842–848. CHOI, W. Y., H. W. NAM, J. H. YOUN, D. J. KIM, W. K. KIM, AND W. S. KIM. 1990. The effect of cyclic AMP on the growth of Toxoplasma gondii in vitro. Kisaengch’ung Hak Chapchi 28: 71–78. CONNELLY, M. C., A. AYALA, AND F. KIERSZENBAUM. 1988. Effects of a- and b-adrenergic agonists on Trypanosoma cruzi interaction with host cells. Journal of Parasitology 74: 379–386. DUBEY, J. P. 1998. Advances in the life cycle of Toxoplasma gondii. International Journal for Parasitology 28: 1019–1024. ELENKOV, I. J., AND G. P. CHROUSOS. 1999. Stress hormones, Th1/Th2 patterns, pro/anti-inflammatory cytokines and susceptibility to disease. Trends in Endocrinology and Metabolism 10: 359–368. EXTON, J. H. 1981. Molecular mechanisms involved in a-adrenergic responses. Molecular and Cellular Endocrinology 23: 233–264. FERMIN, Z., D. BOUT, P. RICCIARDI-CASTAGNOLI, AND J. HOEBEKE. 1999. Salbutamol as an adjuvant for Nasal vaccination. Vaccine 17: 1936–1941.

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GAZZINELLI, R. T., D. AMICHAY, T. SHARTON-KERSTEN, E. GRUNWALD, J. M. FARBER, AND A. SHER. 1996. Role of macrophage-derived cytokines in the induction and regulation of cell-mediated immunity to Toxoplasma gondii. Current Topics in Microbiology and Immunology 219: 127–139. HUNTER, C. A., Y. SUZUKI, C. S. SUBAUTE, AND J. S. REMINGTON. 1996. Cells and cytokines in resistance to Toxoplasma gondii. Current Topics in Microbiology and Immunology 219: 113–125. KIRKMAN, L. A., L. M. WEISS, AND K. KIM. 2001. Cyclic nucleotide signaling in Toxoplasma gondii bradyzoite differentiation. Infection and Immunity 69: 148–153. MONROY, F. P., S. BANERJEE, T. DUONG, AND H. AVILES. 1999. Cold stress-induced modulation of inflammatory responses and intra cerebral cytokine mRNA expression in acute murine toxoplasmosis. Journal of Parasitology 85: 878–886. SHEN, H. M., L. X. SHA, J. L. KENNEDY, AND D. W. OU. 1994. Adrenergic receptors regulate macrophage secretion. Journal of Immunopharmacology 16: 905–910. SPENGLER, R. N., R. M. ALLEN, D. G. REMICK, R. M. STRIETER, AND S. L. KUNKEL. 1990. Stimulation of a-adrenergic receptor augments the production of macrophage-derived tumor necrosis factor. Journal of Immunology 145: 1430–1434. SUZUKI, Y., F. K. CONLEY, AND J. S. REMINGTON. 1989. Importance of endogenous IFN-g for prevention of toxoplasmic encephalitis in mice. Journal of Immunology 143: 2045–2050. SZELENYI, J., J. P. KISS, E. PUSKAS, M. SZELENYI, AND E. S. VIZI. 2000. Contribution of differently localized a- and b-adrenoreceptors in the modulation of TNF-a and IL-10 production in endotoxemic mice. Annals of the New York Academy of Sciences 17: 145–153.


Detection of Paragonimus heterotremus Eggs in Experimentally Infected Cats by a Polymerase Chain Reaction–Based Method Pewpan M. Intapan, Chaisiri Wongkham, Kanokwan J. Imtawil, Wilawan Pumidonming*, Thidarat K. Prasongdee, Masanao Miwa†, and Wanchai Maleewong, Departments of Parasitology and Biochemistry, Faculty of Medicine, Khon Kaen University, Khon Kaen 40002, Thailand; *Department of Microbiology and Parasitology, Faculty of Medical Science, Naresuan University, Pitsanuloke 65000, Thailand; †Institute of Basic Medical Sciences, University of Tsukuba, Tsukuba City, Ibaraki 305-8575, Japan. e-mail: [email protected] ABSTRACT: A polymerase chain reaction (PCR) procedure for the detection of Paragonimus heterotremus eggs in stool samples was developed and compared with Stoll’s egg count method. The primers were designed on the basis of a previously constructed pPH-13–specific DNA probe, which produced an approximate 0.5-kb amplified product. This PCR method could detect as few as 5 eggs in 0.6 g of artificially inoculated feces of a healthy control cat or as little as 1 3 1024 ng of P. heterotremus genomic DNA. The assay had 100% sensitivity in all infected cats. The method did not yield an approximate 0.5-kb product with DNA from other parasites such as Gnathostoma spinigerum, Trichinella spiralis, Fasciola gigantica, Echinostoma malayanum, Opisthorchis viverrini, Dirofilaria immitis, and Taenia saginata; exceptions were Paragonimus siamensis and Paragonimus westermani. In addition, no genomic DNA from Escherichia coli, Burkholderia pseudomallei, Acinetobacter anitratus, Mycobacterium tuberculosis, Staphylococcus aureus, b-Streptococcus grA, and Proteus mirabilis or from the vertebrate and invertebrate hosts of P. heterotremus was amplified in the PCR assay. This assay has great potential for application in clinical epidemiological studies.

The lung fluke Paragonimus sp., a most harmful parasite, causes paragonimiasis in humans and other animals. It is estimated that more than 20 million people are infected worldwide (World Health Organization, 1995). Although Paragonimus westermani is the most common species, P. heterotremus is the main etiological agent of human paragonimiasis in China, Laos, and Thailand (Maleewong, 2003). Currently, the diagnosis of paragonimiasis is based on the demon-

stration of Paragonimus sp. eggs in feces, sputa, or both (Yokogawa, 1965). However, the microscopic examination for the parasite’s eggs is laborious and time consuming and reliable only in the hands of experienced personnel. Furthermore, it is sometimes subject to bias and confusion with other digenean flukes. Immunological techniques have been used with varying success to diagnose P. heterotremus infections by detecting specific antibodies in the sera of infected individuals (Indrawati et al., 1991; Maleewong et al., 1991, 1992; Dekumyoy et al., 1998). However, false-positive results were found because of the crossreactions caused by other parasites, e.g., Fasciola gigantica (Maleewong et al., 1992). Another problem of antibody detection is the persistence of antibodies after treatment, thus making the distinction between present and past infection difficult. Moreover, an antigen capture– enzyme-linked immunosorbent assay developed to detect P. heterotremus excretory–secretory antigen in feces (Maleewong et al., 1997) had a low sensitivity. Another diagnostic effort was the construction of a specific DNA probe (pPH-13) to detect P. heterotremus eggs in feces (Maleewong et al., 1997). This specific probe was shown not to cross-hybridize with genomes of related parasites, bacteria, or vertebrate and invertebrate hosts of P. heterotremus (humans, cats, crabs, and snails). However, the technique had a low sensitivity as well. The probe’s DNA sequence (1,473 bp) data were submitted to GenBank (AZ254640) and showed no significant homology to any other stored parasite DNA. In this study, we report the development of a polymerase chain reaction (PCR) technique based on the above DNA sequence for the detection of P. heterotremus in experimentally infected cats.

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FIGURE 1. Ethidium bromide staining patterns of the PCR products on a 2% agarose gel. The arrow indicates location of an approximate 0.5kb amplified product. A. Sensitivity of the PCR for the detection of Paragonimus heterotremus genomic DNA. Lane M, DNA size markers (100bp ladder from Promega Corporation, Madison, Wisconsin); lane N, negative control containing no DNA. Lanes corresponding to PCR products from the 10-fold serial dilutions of P. heterotremus DNA: 1 ng (lane 1), 1 3 1021 (lane 2), 1 3 1022 (lane 3), 1 3 1023 (lane 4), 1 3 1024 (lane 5), and 1 3 1025 (lane 6). B. Capability of the PCR for the detection of P. heterotremus eggs artificially inoculated into feces from healthy control cat. Lane M, DNA size markers (100-bp ladder from Promega Corporation); lanes N and P, negative and positive controls containing no DNA and 1 ng P. heterotremus DNA, respectively. PCR products from 40 (lane 1), 20 (lane 2), 10 (lane 3), 5 (lane 4), and 1 (lane 5) P. heterotremus eggs. C. Representative specificity of the PCR for the detection of P. heterotremus. Lane M, DNA size marker (100-bp ladder from Promega Corporation); lanes N and P, negative and positive controls containing no DNA and 1 ng of P. heterotremus DNA, respectively. PCR product with DNA from Gnathostoma spinigerum (lane 1), Trichinella spiralis (lane 2), Fasciola gigantica (lane 3), Echinostoma malayanum (lane 4), Opisthorchis viverrini (lane 5), Dirofilaria immitis (lane 6), and Taenia saginata (lane 7).

Adult P. heterotremus worms were obtained from the lungs of experimentally infected cats and used for the preparation of DNA. The worms were identified as P. heterotremus as described previously (Miyazaki and Vajrasthira, 1967). Fresh or frozen adult worms of P. heterotremus were extracted and purified for DNA using the Nucleospin Tissue kit (Macherey-Nagel GmbH&Co., Duren, Germany). The DNA was resuspended in 5 mM Tris–HCl, pH 8.5, and used as positive control as well as for sensitivity evaluation of the PCR amplification. For sensitivity of the detection assay, 10-fold serial dilutions of P. heterotremus genomic DNA were prepared in duplicate with distilled water, starting from 1 ng to 1 3 1025 ng/10 ml. Each dilution was used for sensitivity evaluation of the PCR. Different numbers (1, 5, 10, 20, 40) of P. heterotremus eggs, artificially inoculated into 0.6 g of feces from a healthy control cat, were also used to test the sensitivity. For specificity, 1 ng of genomic DNA from other organisms was tested. The genomic DNA used for specificity testing was prepared from other lung flukes, i.e., P. siamensis and P. westermani, as well as a number of other parasites, i.e., Gnathostoma spinigerum, Trichinella spiralis, F. gigantica, Echinostoma malayanum, Opisthorchis viverrini, Dirofilaria immitis, and Taenia saginata. Additional DNA samples were also extracted from several species of bacteria, i.e., Escherichia coli, Burkholderia pseudomallei, Acinetobacter anitratus, Mycobacterium tuberculosis, Staphylococcus aureus, b-Streptococcus grA, and Proteus mirabilis, and from vertebrate and invertebrate hosts of P. heterotremus (humans, cats, crabs [Tiwaripotamon beusekomae], and snails [Onchomelania nosophora]). Domestic cats of both sexes, in groups of 3–5, were examined and confirmed to be worm free before P. heterotremus infection. Stool examinations of all cats were done using formalin–ether sedimentation technique (Beaver et al., 1984). Animals harboring any parasites, i.e., hookworms or Toxocara cati, were treated with albendazole (10 mg/kg body weight), whereas Spirometra spp. and Dicrocoelium spp. infections were treated with niclosamide (100 mg/kg body weight) and praziquantel (40 mg/kg body weight), respectively. The cats were kept in the animal unit of the Faculty of Medicine, Khon Kaen University. They were housed separately in clean cages and reared on cooked fishes, boiled rice, canned feline food, and water. Veterinary care was provided throughout the study. Finally, through a stomach tube, the cats were each fed P. heterotremus metacercariae, which were obtained from naturally infected crabs (T. beusekomae). In the experimental study, 3 groups of cats were infected with 5, 10, and 40 metacercariae, and 1 group was kept as a healthy control group.

The feces of each cat was collected every 2 wk, for 12 wk. Stoll’s egg count (Beaver et al., 1984) was used together with PCR amplification to evaluate the detection assay. The egg counts of each group were expressed as mean values 6 SD, respectively. To ensure infectivity after the experiment, the infected cats were terminally anesthetized, and the lungs were dissected for worm recovery. The fecal slurry made with 5 ml of 0.1 M phosphate-buffered saline (PBS), pH 7.4, and 3 ml of ether was centrifuged at 1,500 g for 5 min. The pellet was resuspended in 10 ml of 0.1 M PBS, pH 7.4, and centrifuged at 1,500 g for 5 min. This washing–centrifugation step was repeated thrice or until the supernatant was clear. The pellet was homogenized with disposable polypropylene pestles (Belco Glass Inc., Vineland, New Jersey) followed by an extraction step using the QIAampt DNA Stool Mini Kit (QIAGEN, Hilden, Germany). The eluted DNA was resuspended in 100 ml Tris-EDTA buffer, pH 9.0 (QIAGEN), and 10 ml of the resulting suspension was used in the PCR reaction. For PCR amplification, the forward primer (59-GGG TAG CAC CAT CTC GT-39) and the reverse primer (59-GAT CAC GAG GAC CCC AT-39) used in the assay were designed upon the sequences of the pPH13 probe, using the Gene Fisher software program (http://bibiserv. techfak.uni-bielefeld.de/genefisher/). The PCR was performed using a DNA thermal cycler (2400; Perkin–Elmer, Norwalk, Connecticut). The reaction was carried out in a 25-ml volume containing PCR buffer (20 mM Tris–HCl, pH 8.4, 50 mM KCl, 2 mM MgCl2), 200 mM of each deoxynucleoside triphosphates, 1 mM of each of the primers, 10 ml of the sample, and 1.5 U of Taq DNA polymerase. The DNA template was initially denatured at 94 C for 5 min. The PCR procedure comprised 35 cycles at 94 C for 30 sec (denaturation), 56 C for 30 sec (annealing), 72 C for 45 sec (extension), and a final extension at 72 C for 7 min. Each 10 ml of the amplified products was analyzed by electrophoresis in a 2% agarose gel using 0.53 Tris-Borate-EDTA buffer, pH 8.0. To confirm the amplicons to be the P. heterotremus DNA, the PCR products were purified by extraction from agarose gel using Nucleospin (Macherey-Nagel) and sequenced in both directions using an Autoread Sequencing Kit (Amersham Biosciences, Piscataway, New Jersey) and an automated sequencer (AlFexpress, Pharmacia Biotech, Uppsala, Sweden). Sequence alignments were facilitated using the multilin program (http://prodes.toulouse.inra.fr/multalin/multalin.html). The results of a typical experiment in which 10-fold serial dilutions of adult P. heterotremus DNA were amplified are shown in Figure 1A. The primer set yielded approximate 0.5-kb product and did not show

RESEARCH NOTES

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TABLE I. Stoll’s egg count of fecal samples in a study of Paragonimus heterotremus–infected cats. Number of metacercariae given to each group

4

5 10 40 0‡

2(0/3) 2(0/3) 2(0/5) 2(0/5)

Intensity of the eggs as detected by Stoll’s egg count at week* (number of positive/total cats tested) 6 2(0/3) 2(0/3) 2,833 6 801 (2/5) 2(0/5)

8 2(0/3) 2(0/3) 40,900 6 25,312 (5/5) 2(0/5)

10

12

4,222 6 2,823 (3/3) 3,075 6 800 (3/3) 122,075 6 64,668 (4/4)† 2(0/5)

3,339 6 2,024 (3/3) 3,955 6 1,520 (3/3) 130,375 6 29,148 (4/4)† 2(0/5)

* The intensity of the egg count was expressed as mean values 6 SD. † One cat died after the eighth week of infection. ‡ This group was left as control. No infection was performed.

any amplification in the absence of the target P. heterotremus DNA (Fig. 1A, lane N). Cat fecal specimens, artificially inoculated with 1, 5, 10, 20, and 40 P. heterotremus eggs, were amplified for 35 cycles. As few as 5 eggs could be clearly detected as an amplicon band in the ethidium bromide–stained gel (Fig. 1B, lane 4). The primers could also detect as little as 1 3 1024 ng of P. heterotremus genomic DNA (Fig. 1A, lane 5), as well as P. siamensis and P. westermani DNA (data not shown). However, the PCR with the primer set did not show an approximate 0.5-kb amplified product with DNA from other parasites, i.e., G. spinigerum, T. spiralis, F. gigantica, E. malayanum, O. viverrini, D. immitis, and T. saginata (Fig. 1C, lanes 1–7). The PCR did not amplify the genomic DNA of other bacteria or of the vertebrate and invertebrate hosts of P. heterotremus (data not shown). The results of the Stoll’s egg count in experimentally infected cats are shown in Table I. During the fourth week of infection, both Stoll’s egg count and PCR methods were negative for all groups. After the fourth week of infection, both methods showed the same sensitivity. All fecal specimens from the infected cats that had P. heterotremus eggs during the periods of study were positive by the PCR method, whereas all noninfected controls were negative. Thus, the results gave high sensitivity and specificity. In addition, the obtained PCR products were

sequenced, and the sequenced data demonstrated approximately 90% identity with the pPH-13–specific DNA sequence of P. heterotremus (Fig. 2). The significant base changes were found, which could be due to the natural strain variation in the population of P. heterotremus because the P. heterotremus metacercariae used to infect cats in the former study (Maleewong et al., 1997) were recovered from crabs collected from different areas than our recent samples. This reasoning is supported by a study on the intraindividual variations in the first internal transcribed spacer (ITS) of the ribosomal gene sequence of P. westermani and related species (van Herwerden et al., 1999). Paragonimiasis, caused by P. heterotremus, is an important endemic disease in southeastern Asia. The lack of an accurate, rapid, sensitive, and specific diagnostic method suitable for epidemiological studies and for monitoring the results of antihelminthic treatment hampers the design of an effective eradication program. Because the PCR method is an extremely sensitive and specific technique, with potentially widespread use in the diagnosis of infectious diseases, it has also been successfully used for the detection of helminths in feces, e.g., for the detection of Echinococcus multilocularis (Mathis et al., 1996; Dinkel et al., 1998), O. viverrini (Wongratanacheewin et al., 2001, 2002; Maleewong et al., 2003), and Schistosoma mansoni (Pontes et al., 2002,

FIGURE 2. Representative alignment of the nucleotide sequences of the PCR products with the sequence of pPH-13. Alignment gaps are indicated by hyphens. Arrows indicate positions and directions of gene-specific primers used for the PCR. AZ254640 represents the GenBank accession number of pPH-13. Representative PCR products are indicated as amplicon1, amplicon2, and amplicon3, respectively.

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2003). This study describes the first use of a PCR-based technique for the detection of P. heterotremus eggs in feline feces. Our method gave reliable results as early as in the sixth week of infection with 40 metacercariae. A P. heterotremus–specific DNA probe constructed previously (Maleewong et al., 1997) could detect as little as 50 pg P. heterotremus DNA or approximately 1,500–2,000 eggs/g in feces. The current method is thus approximately 150–500 times more sensitive than the older probe. It was estimated that under these conditions, specimens containing approximately 8–10 eggs/g could be readily detected by PCR. However, additional experiments for detection in lightly infected cats with 1 or 2 P. heterotremus metacercariae are needed in the future. No amplification of an approximate 0.5-kb band occurred with DNA from other parasites, bacteria, or hosts, nor did the PCR react with any negative control samples, thus showing a high specificity. Because of its high degree of sensitivity and specificity, the PCR protocol described in this study could be used for distinguishing Paragonimus sp. from the other parasites examined in the present study that are common causes of human infection, i.e., G. spinigerum, T. spiralis, F. gigantica, E. malayanum, O. viverrini, D. immitis, and T. saginata (Beaver et al., 1984), and could provide an alternative to the available classic microscopic and more modern molecular or immunological methods for the detection of Paragonimus spp. in fecal samples. However, this technique cannot discriminate between species of P. heterotremus and other lung flukes, i.e., P. siamensis and P. westermani, as has been done in the previous reports (Ryu et al., 2000; Sugiyama et al., 2002). Ryu et al. (2000) developed two specific primers from nucleotide sequences of the ITS2 gene of Paragonimus spp., which could identify P. westermani and P. ohirai. Sugiyama et al. (2002) established primers from ITS2 that could discriminate between P. westermani and P. miyazakii metacercariae. However, these primers have not been adapted for use in fecal samples. Nevertheless, our PCR assay can be used as a screening test for the reservoir host of the parasites in epidemiological studies looking at eradication programs because the treatment of the different lung fluke infections is quite similar. For such field studies, the discrimination of the species in the genus Paragonimus would not be important. We are currently working to refine the specific primers that can differentiate P. heterotremus from other Paragonimus spp., i.e., specific primers from nucleotide sequences of the ITS2 gene of ribosomal DNA of Paragonimus spp. Once species-specific primers have been created, the present PCR should have great value not only for the detection of P. heterotremus in cats but also in humans and intermediate hosts. Another advantage of this assay is its speed because a large number of samples can be processed at the same time and results can be obtained within 1 day. This study received financial support from Khon Kaen University. The authors wish to thank Mark Roselieb for his assistance in the manuscript preparation. LITERATURE CITED BEAVER, P. C., R. C. JUNG, AND E. W. CUPP. 1984. Clinical parasitology, 9th ed. Lea and Febiger, Philadelphia, Pennsylvania, 825 p. DEKUMYOY, P., J. WAIKAGUL, AND K. S. EOM. 1998. Human lung fluke Paragonimus heterotremus: Differential diagnosis between Paragonimus heterotremus and Paragonimus westermani infections by EITB. Tropical Medicine and International Health 3: 52–56. DINKEL, A., M. VON NICKISCH-ROSENEGK, B. BILGER, M. MERLI, R. LUCIUS, AND T. ROMIG. 1998. Detection of Echinococcus multilocularis in the definitive host: Coprodiagnosis by PCR as an alternative to necropsy. Journal of Clinical Microbiology 36: 1871–1876.

INDRAWATI, I., W. CHAICUMPA, P. SETASUBAN, AND Y. RUANGKUNAPORN. 1991. Studies on immunodiagnosis of human paragonimiasis and specific antigen of Paragonimus heterotremus. International Journal for Parasitology 21: 395–401. MALEEWONG, W. 2003. Paragonimus species. In International handbook of foodborne pathogens, M. D. Miliotis and J. W. Bier (eds.). Marcel Dekker, Inc., New York, p. 601–611. ———, P. M. INTAPAN, C. WONGKHAM, S. WONGRATANACHEEWIN, P. TAPCHAISRI, N. MORAKOTE, AND W. CHAICUMPA. 1997. Detection of Paragonimus heterotremus in experimentally infected cat feces by antigen capture-ELISA and by DNA hybridization. Journal of Parasitology 83: 1075–1078. ———, ———, ———, T. WONGSAROJ, T. KOWSUWAN, W. PUMIDONMING, P. PONGSASKULCHOTI, AND V. KITIKOON. 2003. Detection of Opisthorchis viverrini in experimentally infected bithynid snails and cyprinoid fishes by a PCR-based method. Parasitology 126: 63–67. ———, C. WONGKHAM, P. INTAPAN, S. PARIYANONDA, AND N. MORAKOTE. 1992. Excretory-secretory antigenic components of Paragonimus heterotremus recognized by infected human sera. Journal of Clinical Microbiology 30: 2077–2079. ———, ———, S. PARIYANONDA, P. INTAPAN, V. PIPITGOOL, W. DAENSEEGAEW, AND N. MORAKOTE. 1991. Antigenic components of Paragonimus heterotremus recognized by infected human serum. Parasite Immunology 13: 89–93. MATHIS, A., P. DEPLAZES, AND J. ECKERT. 1996. An improved test system for PCR-based specific detection of Echinococcus multilocularis eggs. Journal of Helminthology 70: 219–222. MIYAZAKI, I., AND S. VAJRASTHIRA. 1967. Occurrence of the lung fluke Paragonimus heterotremus Chen & Hsia 1964, in Thailand. Journal of Parasitology 53: 207. PONTES, L. A., E. DIAS-NETO, AND A. RABELLO. 2002. Detection by polymerase chain reaction of Schistosoma mansoni DNA in human serum and feces. American Journal of Tropical Medicine and Hygiene 66: 157–162. ———, M. C. OLIVEIRA, N. KATZ, E. DIAS-NETO, AND A. RABELLO. 2003. Comparison of a polymerase chain reaction and the KatoKatz technique for diagnosing infection with Schistosoma mansoni. American Journal of Tropical Medicine and Hygiene 68: 652–656. RYU, J. S., U. W. HWANG, D. Y. MIN, K. S. SHIN, S. J. NAM, AND O. R. LEE. 2000. Molecular identification of Paragonimus ohirai and P. westermani from Anhui Province, China. Parasite 7: 305–309. SUGIYAMA, H., Y. MORISHIMA, Y. KAMEOKA, AND M. KAWANAKA. 2002. Polymerase chain reaction (PCR)-based molecular discrimination between Paragonimus westermani and P. miyazakii at the metacercarial stage. Molecular and Cellular Probes 16: 231–236. VAN HERWERDEN, L., D. BLAIR, AND T. AGATSUMA. 1999. Intra- and interindividual variation in ITS1 of Paragonimus westermani (Trematoda: Digenea) and related species: Implications for phylogenetic studies. Molecular Phylogenetics and Evolution 12: 67–73. WONGRATANACHEEWIN, S., W. PUMIDONMING, R. W. SERMSWAN, AND W. MALEEWONG. 2001. Development of a PCR-based method for the detection of Opisthorchis viverrini in experimentally infected hamsters. Parasitology 122: 175–180. ———, ———, ———, V. PIPITGOOL, AND W. MALEEWONG. 2002. Detection of Opisthorchis viverrini in human stool specimens by PCR. Journal of Clinical Microbiology 40: 3879–3880. WORLD HEALTH ORGANIZATION. 1995. Control of foodborne trematode infections. World Health Organization Technical Report Series 849: 1–157. YOKOGAWA, M. 1965. Paragonimus and paragonimiasis. Advances in Parasitology 3: 99–158.

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Disseminated Visceral Coccidiosis and Cloacal Cryptosporidiosis in a Japanese White-Naped Crane (Grus vipio) Yongbaek Kim*, Elizabeth W. Howerth†, Nam-Sik Shin‡, Soo-Whan Kwon§, Scott P. Terrell\, and Dae-Yong Kim#¶, *Laboratory of Experimental Pathology, National Institute of Environmental Health Sciences, Research Triangle Park, North Carolina 27709; †Department of Pathology, College of Veterinary Medicine, University of Georgia, Athens, Georgia 30605; ‡Department of Wild Animal Medicine, College of Veterinary Medicine, Seoul National University, Seoul 151-742, Korea; §Everland Zoological Gardens, Yongin 449-715, Korea; \Walt Disney World Animal Programs, Bay Lake, Florida 32830; #Department of Veterinary Pathology, College of Veterinary Medicine, Seoul National University, Seoul, 151-742 Korea; ¶To whom correspondence should be addressed. e-mail: [email protected] ABSTRACT: A 4-mo-old male Japanese white-naped crane (Grus vipio) kept in an outdoor exhibit at the Everland Zoological Gardens in Korea became depressed and developed anorexia, weight loss, and diarrhea. Death of this bird was associated with an overwhelming systemic infection by an intracellular coccidian parasite, which resulted in necrosis and granulomatous inflammation in a number of major organs, including the intestine, liver, spleen, and kidney. Coccidian parasite–laden macrophages were commonly found in the blood vessels of these organs. Using electron microscopy and polymerase chain reaction assays, the parasite was identified as Eimeria sp. The bird was also infected with Cryptosporidium sp., which suggests an immunosuppressed state, although the cause of such suppression was not identified. Our findings suggest that an initial Eimeria sp. intestinal infection spread to other organs through the blood vessels, with the immunosuppressed state possibly contributing to a rapid hematogenous transmission. To our knowledge, this is the first report of disseminated visceral coccidiosis caused by Eimeria sp. in a captive Japanese white-naped crane.

Disseminated visceral coccidiosis associated with Eimeria spp. was first reported in captive sandhill cranes (Grus canadensis) and whooping cranes (G. americana) at the Patuxent Wildlife Research Center in 1977 (Novilla et al., 1981). Fecal examination identified E. reichenowi and E. gruis in most of those cases. Visceral coccidiosis caused by E. reichenowi and E. gruis was also reported in demoiselle cranes (Anthropoides virgo) and sarus cranes (G. antigone) (Yakimoff and Matschoulsky, 1935; Pande et al., 1970). Because of its potential to cause mortality, management of visceral coccidiosis is regarded as an important issue for captive and wild cranes. This article describes a case of fatal disseminated visceral coccidiosis in a captive Japanese white-naped crane (G. vipio) with concurrent cloacal cryptosporidiosis. To our knowledge, this is the first such case reported in this crane species. A 4-mo-old male Japanese white-naped crane kept in an outdoor exhibit at the Everland Zoological Gardens, Yongin-city, Korea, became depressed and exhibited anorexia, weight loss, and diarrhea. The crane was unresponsive to fluid therapy and died within 3 days of the commencement of treatment. The crane was submitted to the College of Veterinary Medicine, Seoul National University for postmortem examination. At necropsy, the crane was very thin and dehydrated. All air sacs were cloudy and contained numerous 1- to 2-mm white foci. The liver and spleen were enlarged 2–3 times their normal sizes. White foci, similar in size and shape to those noted in the air sacs, were also present on the surface and throughout the parenchyma of the liver, spleen, and kidneys. Representative tissue samples from the major organs were collected and fixed in 10% phosphate-buffered formalin, routinely processed, and stained with hematoxylin and eosin (HE) for light microscope examination. For electron microscopy, formalin-fixed liver and spleen samples were minced into 1-mm cubes and postfixed in 2% glutaraldehyde with 2% sucrose in a 0.1 M sodium cacodylate buffer, and subsequently in osmium tetroxide. Fixed specimens were epon–araldite embedded, and ultrathin sections were stained with lead citrate and uranyl acetate and examined under a transmission electron microscope. Tissues from the liver, spleen, kidney, and small intestine were collected aseptically; routine aerobic and anaerobic bacterial cultures were performed. Polymerase chain reaction (PCR) testing was conducted on formalinfixed, paraffin-embedded tissues, which included liver, kidney, lung,

proventriculus, and intestines. Distilled water was used as a negative control. Sandhill cranes with visceral coccidiosis and goat feces heavily infected with Eimeria spp. were used as positive controls. DNA from the paraffin-embedded blocks was extracted using a Puregene DNA isolation kit (Gentra Systems, Minneapolis, Minnesota) according to manufacturer’s instructions. Consensus regions within the 18S ribosomal DNA (rDNA) of Eimeria spp. were identified by aligning available sequence data from 8 Eimeria spp. Consensus sequences unique to Eimeria spp., which flanked a region containing a previously recognized interspecific variation, were used to design primers EIMF (59-39: ACC ATGGTAATTCTATG) and EIMR (59-39: CTCAAAGTAAAAGTTC C). These primers were predicted to amplify a 621-bp fragment of Eimeria spp. 18S rDNA. PCR was performed in 100-ml reaction volumes containing approximately 150–350 ng DNA, 7.5 units Taq polymerase (United States Biochemical Corp., Cleveland, Ohio), 1.2 mM each of deoxynucleotide triphosphate, 1.0 mM of each oligonucleotide primer, 8.5 ml 103 PCR buffer (United States Biochemical Corp.), and 2.5 mM MgCl2. The conditions of amplification were 34 cycles of denaturation at 94 C for 30 sec, annealing at 45 C for 30 sec, and extension at 72 C for 45 sec in a DNA thermal cycler (Hybaid Instruments, Holbrook, New York). Amplification products were observed after separation on 1.0% agarose gels, ethidium bromide staining, and UV transillumination. Histological changes in the major parenchymal organs were similar, although with differing lesion severity. The villi of the small intestine were mildly atrophied and fused to each other, and the lamina propria expanded by mild, to moderate, infiltration of lymphocytes and plasma cells admixed with fewer macrophages. A coccidian parasite at various stages of development including merozoites, schizonts, and gamonts, was present in the cytoplasm of intestinal epithelial cells (Fig. 1). Coccidia-laden macrophages also were present in the blood vessels of the lamina propria, tunica muscularis, and serosa and were often associated with perivascular inflammation. The coccidian parasites were round to oval or elongated, 10–25 mm in length, and frequently formed lobulated clusters composed of 3–6 organisms. Multifocal randomly distributed areas of necrosis accompanied by a heavy infiltration of macrophages, lymphocytes, and a few heterophils, and perivascular lymphoplasmacytic infiltration were noted in the liver (Fig. 2). Coccidian parasites were present in the cytoplasm of Kupffer cells and infiltrating macrophages associated with the necrotic foci. Changes in the pancreas, kidney, spleen, and lung were similar to those seen in the liver. The blood vessels in most organs examined, including the brain and spinal cord, were often filled with macrophages containing intracytoplasmic organisms similar to those seen in the liver. In addition, the apical surface of the epithelium lining the submucosal glands of the cloaca was covered by numerous basophilic round organisms, ranging from 3 to 5 mm in diameter. These organisms were consistent with Cryptosopidium sp. (Fig. 3). No significant bacterial pathogen was isolated. Electron microscopy revealed coccidian parasites at various developmental stages present within parasitophorous vacuoles in macrophages. Merozoites were oval to pear shaped and surrounded by a bilayered capsule. The anterior ends of the coccidian parasites had a conoid with peripherally arranged micronemes, which were continuous with dense tubular micronemes. A few electron-dense globules and glycogen granules were scattered throughout the cytoplasm. A pericentric nucleus was also observed.

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FIGURE 3. Cloaca; Japanese white-naped crane. Myriads of round basophilic organisms consistent with Cryptosporidium sp. (arrowheads) on the luminal surface of the epithelial cells of mucosa and tubular glands. Note many Eimeria sp.–laden macrophages (thick arrows) in the blood vessels of the lamina propria. HE. Bar 5 50 mm.

FIGURE 1. Small intestine; Japanese white-naped crane. Merozoites, schizonts, and gamonts of Eimeria sp. within the cryptal epithelial cells (arrowheads) associated with necrosis (thick arrows). Note numerous Eimeria-laden macrophages (thin arrows) within the blood vessel of lamina propria. HE. Bar 5 40 mm.

FIGURE 2. Liver; Japanese white-naped crane. Multifocal randomly distributed necrotic foci accompanied by infiltration of macrophages, lymphocytes, and fewer heterophils. Note Eimeria-laden macrophages in the necrotic area (arrowheads) and blood vessels (thin arrow). HE. Bar 5 150 mm.

PCR analysis showed that all paraffin-embedded tissues contained Eimeria sp.–specific rDNA according to consensus sequences. A 621bp PCR product was obtained from all paraffin-embedded tissues from this bird as well as from a sample from a sandhill crane infected with visceral coccidiosis and from a goat fecal sample containing Eimeria sp. (Fig. 4). On the basis of light microscope findings, electron microscopy, and PCR, the death of this crane was due to an overwhelming disseminated Eimeria spp. infection. The organism appeared to have disseminated throughout the body through the circulatory system on the basis of the findings of perivascular inflammatory responses and the presence of organism-laden macrophages in the blood vessels of multiple organs.

FIGURE 4. Agarose gel electrophoresis of PCR products separated on 1% agarose gel. The expected Eimeria sp.–specific product is 621 bp. From right to left: 100-bp DNA ladder; neg 5 water; pos 5 goat feces infected with Eimeria sp.; SHC 5 sand hill cranes with visceral coccidiosis; paraffin-embedded blocks from Japanese white-naped crane, 117 5 liver and kidney, 217 5 intestine, proventriculus, and cloaca, 417 5 lung and liver.

RESEARCH NOTES

Although eimeriid coccidia are generally site specific for the intestinal tract, some species that infect mammals and birds are also known to be capable of developing in organs other than the intestine. Eimeria stiedai normally occurs in the liver of the rabbit and E. truncata in the kidneys of geese and ducks (Overstreet, 1981). The coccidians, E. gruis and E. reichenowi, are common parasites of whooping cranes and sandhill cranes, which are associated with granulomatous necrotizing hepatitis, pneumonia, and splenitis (Carpenter et al., 1980; Novilla et al., 1981). One of these species is likely the cause of visceral coccidiosis in this case, but fecal examination was not performed, so the organism could not be further identified. To our knowledge, this is the first report of Cryptosporidium sp. infection in cranes. Cryptosporidiosis is often associated with an immunosuppressed state (Tzipori, 1988), so immunosuppression may have contributed to the systemic spread of the eimerian parasite in this case. However, there was no conclusive evidence of immunosuppression in this bird. Alternatively, the eimerian infection may have caused immunosuppression allowing Cryptosporidium sp. infection in this crane. Two other cranes that had been kept in the same exhibits died about 1 mo before submission of this case. According to the attending veterinarian, although coccidia were identified as Eimeria spp. on fecal floatation, the species could not be determined. The remaining live cranes in the exhibit were treated with a coccidiostatic drug, and no clinical coccidiosis has been noted since.

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Disseminated visceral coccidiosis is an occasional cause of mortality in wild and captive cranes. Management of disseminated visceral coccidiosis in cranes is crucial to prevent fatality. This study indicates that anticoccidian programs directed against visceral coccidiosis should be undertaken in zoos. This study was supported by the Brain Korea 21 Project. LITERATURE CITED CARPENTER, J. W., T. R. SPRAKER, AND M. N. NOVILLA. 1980. Disseminated visceral coccidiosis in whooping cranes. Journal of American Veterinary Medical Association 177: 845–848. NOVILLA, M. N., J. W. CARPENTER, T. R. SPRAKER, AND T. K. JEFFERS. 1981. Parental development of Eimerian coccidia in sandhill and whooping cranes. Journal of Protozoology 28: 248–255. OVERSTREET, R. M. 1981. Species of Eimeria in nonepithelial sites. Journal of Protozoology 28: 258–260. PANDE, B. P., B. B. BHATIA, P. P. S. CHAUHAN, AND R. K. GARG. 1970. Species composition of coccidia of some of the mammals and birds at the zoological gardens, Lucknow (Uttar Pradesh). Indian Journal of Animal Science 40: 154–166. TZIPORI, S. 1988. Cryptosporidiosis in perspective. Advances in Parasitolology 27: 63–129. YAKIMOFF, W. L., AND S. N. MATSCHOULSKY. 1935. Die Kokzidiose der Kraniche. Zeitschrift fur Parasitenkde 8: 239–240.


Significant Morphological but Little Molecular Differences Between Trypanosoma of Rodents From Alaska Juha Laakkonen, Andrew Smith*, Kyndall Hildebrandt†, Jukka Niemimaa, and Heikki Henttonen, Finnish Forest Research Institute, P.O. Box 18, FIN-01301, Vantaa, Finland; *Population Biology Research Group, School of Biological Sciences, University of Liverpool, Biosciences Building, Liverpool, L69 7ZB, U.K.; †University of Alaska Museum, 907 Yukon Drive, Fairbanks, Alaska, 99775. e-mail: [email protected] ABSTRACT: We examined blood smears of 173 rodents and 33 shrews captured at 4 sites in the Gates of the Arctic National Park, northern Alaska, in summer 2002. Trypanosoma spp. were detected in the plasma of 5 Microtus oeconomus, 4 Microtus miurus, and 1 Lemmus trimucronatus. The trypomastigote morphology from different individuals of M. oeconomus caught at the same site and of M. miurus from different sites varied significantly. The 4 DNA sequences obtained from the blood smear positive samples contained 2 different haplotypes very similar to each other and to that of Trypanosoma microti. Of possible vectors of blood parasites, the flea Amalaraeus dissimilis was collected from M. miurus.

Despite the increasing interest for biodiversity assessments, baseline information on the parasite diversity in the Arctic and subarctic regions is minimal or absent for many mammal species. The Beringian Coevolution Project (BCP) was designed as a comprehensive inventory of mammals and parasites across Beringea to document the effects of historical and contemporary global changes on faunal assemblages in Arctic Region (Hoberg et al., 2003). As a part of this project, a wide array of parasite samples were collected from small mammals in the Gates of the Artic National Park during summer of 2002. In this article, we describe the Trypanosoma spp. infections found in rodents during that survey and compare the trypomastigote morphology with those recorded from arctic rodents in previous studies. We also provide molecular data from 3 Trypanosoma-infected rodent species. The material consisted of blood smears of 173 rodents and 33 shrews captured at 4 sites in the Gates of the Arctic National Park, Alaska, U.S.A, in July and August of 2002. The number and species caught are shown in Table I. Three of the sites (Agiak Lake, 68804948.40N, 152856944.10W, altitude 960 m above sea level; Fortress Mountain, 68834948.40N, 152857935.60W, altitude 750 m; and Nanushuk River, 68816925.40N, 150839917.60W, altitude 990 m), are arctic tundra habitat.

The unnamed lake site (678279170N, 150851949.50W, altitude 320 m) North of Koyukuk River is taiga forest habitat close to the tree line. Animals were caught with snap-traps (Museum Special and the Finnish Type) and plastic pitfalls. The locations of all trapping lines were recorded by global positioning system. Each animal was assigned a permanent archival record number (AF in appendices), and voucher specimens (skeletons, skulls, skins, and frozen tissues) were deposited in the University of Alaska Museum at Fairbanks. Animals were dissected immediately after capture in the field. On necropsy, animals were combed for ectoparasites, which were stored in 70% ethanol for later identification. A drop of blood was obtained from the heart for preparation of thin blood smears, which were air-dried, fixed in methanol, and stained later in the laboratory with Giemsa’s stain. Each smear was examined with a microscope for 5 min at 3400 and for 10 min at 31,000. DNA was extracted from 10 blood smear positive samples collected on Isocodet STIX (Schleicher and Schuell, Keene, New Hampshire) using an alkaline digest method (Bown et al., 2003). In brief, the bloodstained Isocodet blotting papers were boiled at 100 C in a 500 ml solution of 1.25% NH3 on a dry hot plate in 1.5-ml Sure-Lock microcentrifuge tubes (Fisher Scientific, Loughborough, U.K.). After 20 min, tubes were removed, centrifuged briefly, and replaced with the lid open for a further 25 min, or until half of the original volume remained. A nested polymerase chain reaction (PCR) was used which specifically targets a variable region of the Trypanosoma 18S ribosomal RNA (rRNA) gene (Noyes et al., 1999, 2000). Each 50-ml reaction contained 5 ml of a 1:10 dilution of the DNA solution, 25 ml 23 PCR Master Mix (Abgene, Surrey, U.K.), 18 ml ddH20, and 1 ml each of the following external primers: TRY927F 59 GAAACAAGAAACACGGGAG and TRY927R 59 CTACTGGGCAGCTTGGA, for 30 cycles of 94 C 30 sec; 55 C 60 sec; 72 C 90 sec. The products from the first-stage reaction were diluted 1:10 and 2 ml used as a template for the second

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TABLE I. Hemoparasites of small mammals caught at 4 sites at the Gates of the Arctic National Park, Alaska, in 2002. N 5 number of blood smears examined.

N

Trypanosoma spp.

Bartonella spp.

Agiak Lake 68804950.90N, 152856931.40W Microtus oeconomus Pallas Microtus miurus Osgood Lemmus trimucronatus Davis Clethrionomys rutilus Pallas Sorex monticolus Merriam

18 18 6 16 22

4 0 1 0 0

3 0 3 3 1

Unnamed lake, Koyukuk 678279170N, 150851949.50W Microtus oeconomus Microtus miurus Lemmus trimucronatus Clethrionomys rutilus Synaptomys borealis Richardson Sorex cinereus Kerr Sorex hoyi Baird

2 1 12 16 31 1 5

0 1 0 0 0 0 0

0 1 6 2 3 0 0

Fortress Mountain 68834948.40N, 152857935.60W Microtus oeconomus Microtus miurus Dicrostonyx groenlandicus Traill Sorex monticolus Sorex cinereus Sorex yukonicus Dokuchaev

3 28 3 1 3 1

1 2 0 0 0 0

2 10 0 0 0 0

Nanushuk River 68816938.50N, 150839919.60W Microtus oeconomus Microtus miurus

4 15

0 1

0 4

round using 1 ml of the following internal primers: SSU561F 59 TGGGATAACAAAGGAGCA and SSU561R 59 CTGAGACTGTAACCTCAAAGC, 1 ml ddH20, 45 ml ReddyMixt PCR Master Mix (Abgene) and the same cycling conditions as outlined above. DNA extraction, primer preparation, and the first- and second-stage PCR preparation were all carried out in separate dedicated rooms to reduce the risk of contamination. Twelve microlitres of the second-round PCR product were loaded onto a 1.5% agarose gel stained with ethidium bromide and observed under UV light. Samples producing a band of ;540 kb were considered positive (AF 59919; AF 61877; AF 61380; AF 59915). Amplified DNA was purified with a QIAquick PCR Purification Kit (QIAGEN, Sussex, U.K.) according to manufacturer’s instructions and sequenced commercially (ABC, Imperial College, London, U.K.) using an ABI 377 automated sequencer. Trypomastigotes were detected in the blood smears of 5 Microtus oeconomus, 4 Microtus miurus, and 1 Lemmus trimucronatus (Table I). All infected M. oeconomus from Agiak Lake were mature females (2 were pregnant). The 2 infected M. miurus from Fortress Mountain were immature males. The rest were all mature males. The number of hosts was too low for statistical comparisons between sex and age groups. The number of trypomastigotes found in the plasma of the infected rodents ranged from 3 to 22 per 1,000 erythrocytes. In the infected L. trimucronatus, only 5 trypomastigotes were found, even after examination of the entire smear. In a few samples, the morphology of the trypomastigotes was poorly preserved most likely because the blood smears had not dried properly in the wet field conditions. Trypomastigotes from 7 infected animals were measured (as in Laakkonen et al., 2002) for morphological comparison (Voucher specimens in U.S. Na-

tional Parasite Collection, USNPC nos. 94139, 94140, 94141, 94142, 94143, 94144, 94145). The mean total length of trypomastigotes from M. oeconomus ranged from 21.85 to 26.5 mm and that of the flagellum from 4.35 to 6.50 mm. In M. miurus the corresponding values varied from 22.8 to 29.8 mm and from 3.45 to 6.80 mm. The detailed measurement data of the 9 trypomastigote parameters measured are available from the corresponding author on request. The trypomastigote morphology from different individuals of M. oeconomus caught at the same site (Agiak Lake) varied significantly (P , 0.001, not shown). In M. miurus, similar, significant variation in the trypomastigote morphology was detected between vole individuals from different sites (P , 0.001, not shown). The trypomastigotes of L. trimucronatus were large (maximum total length 36 mm), and had a relatively long flagellum (mean 11.35) compared with that of trypomastigotes from other host species. The trypomastigotes found in M. oeconomus were larger than those seen in this host species from Toolik Lake, Alaska (Laakkonen et al., 2002), but similar in size and other morphological characteristics to trypomastigotes of M. oeconomus from Lower Ugashik Lake, Alaska Peninsula (Fay and Rausch, 1969). The trypomastigotes of lemmings appear to be slightly larger, and to have longer flagella than those of Microtus spp. voles (Quay, 1955; this study). Our Trypanosoma sp. findings in M. miurus and L. trimucronatus appear to be new host records (Podlipaev, 1990). The considerable differences in the morphology of trypomastigotes from the same host species within, and between, study sites indicates that morphological comparison alone is not a reliable method for the identification of Trypanosoma species of arctic rodents. This result supports the conclusions of the only previous comprehensive biometrical comparison of T. lewisi–like trypomastigotes, revealing significant variations in size between parasite populations from the same host and no significant differences between parasite populations from different host species (Davis, 1952). Other blood parasites (Babesia spp.) are known to be morphologically pleomorphic depending on the host species or stage of infection (Kreier and Baker, 1987). The 18S rRNA haplotypes from the M. miurus (AF 61877 and AF 61380) differed from that of Trypanosoma microti at 2/15 polymorphic positions (not shown). The haplotypes from the L. trimucronatus (AF 59915) and M. oeconomus (AF 59919) differed from that of T. microti at 1/15 polymorphic positions and from that of the M. miurus at 3/15 positions (GenBank AY 586621, AY 586622, AY 586623, and AY 586624). The sequence data obtained from the 3 host species caught at 3 different locations showed that the Trypanosoma spp. found from different rodent species were very similar to each other and to that of T. microti. However, the European Trypanosoma species T. microti and T evotomys differ from each other at only 2/15 polymorphic positions (Noyes et al., 2002), consequently the small differences between haplotypes may be markers for significant biological differences in these parasites. Nevertheless, the data indicated that the haplotypes did not correlate with morphological types. Additional molecular analyses are needed to determine whether the identical haplotypes found in L. trimucronatus and M. oeconomus (caught at the same site) can be taken as evidence for possible geographic populations. The record of Trypanosoma sp. in Alaskan rodents consists of only a few reports on wild rodents (Quay, 1955; Fay and Rausch, 1969; Laakkonen et al., 2002) and of 1 on the introduced Rattus norvegicus (Schiller, 1956). Of wild rodents, trypanosomes have been detected in a few Dicrostonyx torquatus (Quay, 1955) but not in any of the few Microtus pennsylvanicus and Synaptomys borealis examined for blood parasites (Quay 1955; Laakkonen et al., 2002; Table I). Trypanosomes have not been reported from arctic shrews, but sample sizes in this and in previous, similar studies have been low (Laakkonen, 2000; Laakkonen et al., 2002). Rod-shaped organisms resembling Bartonella sp. bacteria were detected in all examined rodent species, except Dicrostonyx groenlandicus (Table I), which had a low sample size (N 5 3). Because several morphologically similar microorganisms are found inside, or on the surface of red blood cells of wild rodents, definite identification was not attempted based on morphology. Of the Trypanosoma sp.–infected rodents, fleas were found only in 1 M. miurus from the unnamed lake site at Koyukuk. It had 2 females

RESEARCH NOTES

of Amalaraeus dissimilis in it. This old male vole was also infected with the Bartonella sp.–like rods in erythrocytes. The number of ectoparasites found in the infected hosts was low at least partly because the collections were conducted on dead hosts. Ectoparasites tend to leave the host shortly postmortem, once the core temperature drops to ambient levels. The present information on ectoparasite fauna of arctic rodents (Holland, 1985; Murrell et al., 2003) indicates that although some ectoparasite species may be primarily associated with certain host species, many arctic rodent species share several ectoparasite species. This provides interesting avenues for the transmission of Trypanosoma spp. and other blood parasites of these, and other hosts (Noyes et al., 2002). Further molecular analyses of Trypanosoma spp. from both fleas, and their host species are needed to gain insight into the host–parasite coevolution of hemoparasites of (arctic) small mammals. We are grateful for Joe Cook, Eric Hoberg, and Sam Telford for the research collaboration on BCP, and Amy Runck, Steve MacDonald, and other members of the BCP for organizing the field expedition. We thank Ralph Eckerlin for the identification of the fleas, and Harry Noyes for comments on the sequence data. This study was partly funded by the NSF 0196095 (BCP), the National Park Service Inventory Funding, and the Finnish Academy (Grant 50474 to H.H.). The work was carried out under the Scientific Collecting Permits from the Alaska Department of Fish and Game. LITERATURE CITED BOWN, K. J., M. BEGON, M. BENNETT, Z. WOLDEHIWET, AND N. H. OGDEN. 2003. Seasonal dynamics of Anaplasma phagocytophila in a rodent-tick (Ixodes trianguliceps) system, United Kingdom. Emerging Infectious Diseases 9: 63–70. DAVIS, B. S. 1952. Studies on the trypanosomes of some California mammals. University of California Publications in Zoology 57: 145–248. FAY, F. H., AND R. L. RAUSCH. 1969. Parasitic organisms in the blood of arvicoline rodents in Alaska. Journal of Parasitology 55: 1258– 1265. HOBERG, E. P., S. J. KUTZ, K. E. GALBREATH, AND J. COOK. 2003. Arctic

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biodiversity: From discovery to faunal baselines—Revealing the history of a dynamic ecosystem. Journal of Parasitology 89: S84– S95. HOLLAND, G. P. 1985. The fleas of Canada, Alaska and Greenland (Siphonaptera). Memoirs of the Entomological Society of Canada, No. 130, Entomological Society of Canada, Ottawa, Canada, 631. p. KREIER, J. P., AND J. R. BAKER. 1987. Parasitic protozoa. Allen and Unwin, Boston, Massachusetts, 241 p. LAAKKONEN, J. 2000. Microparasites of three species of shrews from Finnish Lapland. Annales Zoologici Fennici 37: 37–41. ———, H. HENTTONEN, M. W. HASTRITER, J. NIEMIMAA, AND G. H. JARRELL. 2002. Hemoparasites and fleas of shrews and rodents from Alaska. Acta Parasitologica 47: 173–176. MURRELL, B. P., L. A. DURDEN, AND J. A. COOK. 2003. Host associations of the tick, Ixodes angustus (Acari: Ixodida) on Alaskan mammals. Journal of Medical Entomology 40: 682–685. NOYES, H. A., P. AMBROSE, F. BARKER, M. BEGON, M. BENNETT, K. J. BOWN, AND S. J. KEMP. 2002. Host specificity of Trypanosoma (Herpetosoma) species: Evidence that bank voles (Clethrionomys glareolus) carry only one T (H.) evotomys 18S rRNA genotype but wood mice (Apodemus sylvaticus) carry at least two polyphyletic parasites. Parasitology 124: 185–190. ———, J. R. STEVENS, M. TEIXEIRA, J. PHELAN, AND P. HOLZ. 1999. A nested PCR for the ssrRNA gene detects Trypanosoma binneyi in the platypus and Trypanosoma sp. in wombats and kangaroos in Australia. International Journal for Parasitology 29: 331–339. ———, ———, ———, ———, AND ———. 2000. Corrigendum to: A nested PCR for the ssrRNA gene detects Trypanosoma binneyi in the platypus and Trypanosoma sp. in wombats and kangaroos in Australia. International Journal for Parasitology 30: 228. PODLIPAEV, S. A. 1990. Catalogue of world fauna of Trypanosomatidae (Protozoa). Proceedings of the Zoological Institute of the USSR Academy of Sciences 217: 1–177. SCHILLER, E. L. 1956. Ecology and health of Rattus at Nome, Alaska. Journal of Mammalogy 37: 181–188. QUAY, W. B. 1955. Trypanosomiasis in the collared lemming, Dicrostonyx torquatus (Rodentia). Journal of Parasitology 41: 562–565.


Genotyping of Giardia duodenalis From Humans and Dogs From Mexico Using a b-Giardin Nested Polymerase Chain Reaction Assay Marco Lalle, Enedina Jimenez-Cardosa*, Simone M. Caccio`, and Edoardo Pozio†, Department of Infectious, Parasitic and Immunomediated Diseases, Istituto Superiore di Sanita`, viale Regina Elena 299, 00161 Rome, Italy; *Laboratorio de Investigacioń en Parasitologıá, Hospital Infantil de Me´xico ‘‘Federico Go´mez,’’ Me´xico, 06720 D.F. Me´xico; †To whom correspondence should be addressed. email: [email protected] ABSTRACT:

Cysts of Giardia duodenalis were collected in Mexico from symptomatic children (n 5 9) and from pet dogs (n 5 5), and they were directly characterized by nested polymerase chain reaction (PCR) amplification of the b-giardin gene. Eight isolates of human origin established as in vitro cultures and 2 reference strains, representing assemblages A and B of G. duodenalis, were also analyzed. PCR–restriction fragment length polymorphism showed that all isolates belonged to assemblage A. Sequence analyses indicated that the large majority of isolates were of the A1 genotype; interestingly, 2 human isolates displayed the A3 genotype, which has been previously identified in human isolates from Italy. The presence of cysts of the A1 and A3 genotypes in isolates from pet dogs is consistent with their role as reservoirs for human infection, although further studies are needed to confirm the occurrence of zoonotic transmission. Remarkably, cysts of assemblage B have not been found in any of the Mexican isolates studied to date. Giardia duodenalis (syn. Giardia intestinalis, Giardia lamblia) is an

enteric protozoan parasite, which infects a wide variety of mammals, including humans. Individuals with acute infection generally present with diarrhea, abdominal pain, nausea, and weight loss, although the infection is asymptomatic in a significant proportion of cases (Faubert, 2000). Giardiasis is 1 of the leading causes of epidemic gastroenteritis worldwide. In Africa, Asia, and Latin America, approximately 200 million people have symptomatic giardiasis, with some 500,000 new cases per year (WHO, 1996). With specific regard to Mexico, epidemiological data suggest that giardiasis is the most common enteric parasitic infection, with the highest prevalence observed among children (SoteloCruz, 1998), a finding related to the immature immunological status of children and poor hygiene practices (Thompson, 1994). Despite their morphological homogeneity, G. duodenalis isolates from different mammalian species display considerable genetic variability, and at least 7 groups, or assemblages (designated as ‘‘assemblages A–G’’), have been described (Monis et al., 2003). Assemblages

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A and B are associated with human infection yet have also been detected in other mammalian hosts. Within assemblage A, the A1 genotype comprises genetically related isolates from humans and animals, whereas genotypes A2 and A3 have mainly been identified in humans (Monis et al., 1998; Caccio` et al., 2002). Assemblage B isolates form another genetic group that is quite variable and which is associated with human and animal infections (Thompson, 2000; Monis et al., 2003). The remaining assemblages are considered to be host specific; for example, assemblages C and D have only been recovered in dogs and cats (Hopkins et al., 1997; Monis et al., 1998; van Keulen et al., 2002; McGlade et al., 2003). In light of this information, the zoonotic potential seems to be restricted to assemblages A and B. Whereas a considerable number of human and animal isolates from Europe, Asia, North America, and Australia have been genetically typed, much less information is available from other regions of the world. In this article, the results of a molecular analysis of 22 G. duodenalis isolates from Mexico using the b-giardin gene as marker are presented. Fecal samples were collected from May to July 2002 at the Federico Go´mez Hospital of Me´xico City from 9 children (5 males and 4 females, 5- to 9-yr-old) with symptomatic giardiasis. Positive fecal samples were also collected from 5 puppies of private owners in the region of Sinaloa, northwestern Mexico. The presence of cysts was assessed by microscopic analysis. Furthermore, 8 isolates of G. duodenalis from symptomatic children of the IMSS Hospital and of the Pediatric Institute of Mexico City were established as axenic cultures. The Portland-1 (assemblage A1) and Nij5 (assemblage B) strains were used as controls. DNA was extracted by standard phenol–chloroform procedure from 5 mg of feces or from 1 3 105 trophozoites obtained from the axenic cultures. A 511-bp fragment of the b-giardin gene was amplified using a nested polymerase chain reaction (PCR) assay. The primary amplification was performed using 2 ml of DNA and the forward primer G7 and the reverse primer G759, as previously described (Caccio` et al., 2002). Five microliters of the primary PCR was used in the nested amplification with primers bGiarF (59-GAACGAGATCGAGGTCCG-39) and bGiarR (59-CTCGACGAGCTTCGTTGTT-39). Amplification of a 292-bp fragment of the small-subunit ribosomal RNA (SSU-rRNA) was performed as described by Hopkins et al. (1997). Reactions were carried out on a Perkin–Elmer GeneAmp 2400 (Applied Biosystems, Foster City, California) thermocycler under the following conditions: after an initial denaturation cycle of 5 min at 94 C, 35 cycles were run, each consisting of 30 sec at 94 C, 30 sec at 53 C (59 C for SSU-rRNA), and 1 min at 72 C, followed by a final extension cycle of 7 min at 72 C. After purification with Qiaquick kit (Qiagen, Hilden Germany), PCR products were fully sequenced on both strands using the ABI Prism BigDye Terminator Cycle Sequencing Kit (Applied Biosystems). The sequencing reactions were analyzed on an ABI 310 automatic DNA sequencer (Applied Biosystems) and assembled with the program SeqMan II (DNASTAR, Madison, Wisconsin). For the analysis of restriction fragment length polymorphism (RFLP), aliquots of the PCR products (7– 15 ml) were digested with 10 U of HaeIII (New England Biolabs, Beverly, Massachusetts) for 4 hr at 37 C in a final volume of 20 ml. The digested products were separated by electrophoresis on 2% agarose gel. Amplification of G. duodenalis DNA using the nested b-giardin PCR yielded the expected fragment (about 500 bp) from all isolates (data not shown). For the initial determination of the assemblage, a PCR–RFLP assay was used. In this assay, HaeIII digestion of the b-giardin gene fragment yielded specific patterns for assemblage A (fragments of 186, 150, 110, and 50 bp) and for assemblage B (fragments of 150, 116, 110, 85, 27, and 23 bp), and these patterns were distinguishable by gel electrophoresis (Fig. 1). The results indicate that all isolates belonged to assemblage A, with the exception of a single dog isolate, which showed a mixed pattern (Fig. 1, lane 3). To determine the genotype(s) within assemblage A, the b-giardin gene fragments were sequenced and aligned with the homologous sequences available in the GenBank database. The comparison showed that all axenic isolates were of the A1 genotype, as were most (7 of 9 isolates) of the human fecal samples. However, the remaining 2 human isolates displayed the A3 genotype, which has only been described for 2 human isolates collected in Italy (Caccio` et al., 2002). The genotypes A1 and A3 were found to differ by only 3 nucleotide substitutions. Among dog isolates, 3 were identified as the A1 genotype, 1 as the A3 genotype, and 1 as a mixed A1/ B3 infection, as judged from the heterogeneous sequence profile; of

FIGURE 1. Electrophoretic separation of b-giardin nested PCR products (511 bp) after restriction with the endonuclease HaeIII. Lanes M, 50-bp ladder; lane 1, isolate HIM3; lane 2, isolate HIM7; lane 3, isolate dog 3; lane 4, isolate dog 7; lane 5, isolate HIM15; lane 6, isolate HIM17; lane 7, reference strain Portland-1 (assemblage A); lane 8, reference strain Nij5 (assemblage B).

note, the dog-specific assemblages C and D were not identified. Because this was rather unexpected, the dog samples were also typed at the SSUrRNA locus. Sequencing of the PCR products confirmed the results obtained at the b-giardin locus. Moreover, the homologous b-giardin gene fragments of assemblages C and D (GenBank AY545646 and AY545647, respectively) are very different from those of assemblages A and B, both in term of nucleotide sequence and HaeIII restriction patterns. The predominance of the A1 genotype in the human isolates studied is consistent with the results of Cedillo-Rivera et al. (2003). These authors typed a panel of 26 axenic isolates of human origin from various Mexican regions using PCR–RFLP at the variant-specific surface protein locus and found that all isolates were of the A1 genotype. In contrast, Ponce-Macotela et al. (2002) only found the A2 genotype in a panel of 22 axenic isolates established from human samples, also collected in various Mexican regions. When considering the typing data from these 2 studies, together with the data from the present study, the A1 genotype is the most prevalent (40 of 62, 65%), followed by A2 (19 of 62, 30%) and A3 (3 of 62, 5%). Remarkably, cysts of assemblage B were not identified in any of the human samples. This may in part be explained by the selection pressure of the axenization process, which is known to favor the growth of organisms of assemblage A over those of assemblage B (Andrews et al., 1992; Karanis and Ey, 1998). However, cysts of assemblage B were not found in the 9 isolates for which typing data were obtained directly from the fecal material. Studies on a larger number of isolates are needed to clarify this issue. The dog-specific assemblages C and D were not identified in the 5 dog isolates studied here, nor they were identified in the 2 isolates studied by Ponce-Macotela et al. (2002). Although this may well depend on the fact that small samples were tested, the presence of cysts of assemblages A and B in the dog isolates indicates the occurrence of cross-transmission between dogs and humans (or other infected animals). In fact, there is increasing evidence that infection with the dogspecific assemblages prevails in areas where the main pattern of trans-

RESEARCH NOTES

mission is dog-to-dog, as occurs in rural regions (Hopkins et al., 1997; Monis et al., 1998; Thompson, 2000). However, a predominance of assemblages A and B was reported in dogs living in urban environments in North America and Europe (Monis et al., 1998; van Keulen et al., 2002). The identification of the A1 and A3 genotypes in dogs is suggestive of a zoonotic nature of these genotypes. Because the samples were collected in different regions of Mexico, the evidence is only circumstantial, and the occurrence of zoonotic transmission must be studied in more defined epidemiological conditions. We thank D. Tonanzi for his technical assistance. This study received financial support from Research Project 2156/RI, entitled ‘‘Infezioni da Cryptosporidium e Giardia attraverso alimenti e acque: metodi di identificazione ed epidemiologia molecolare’’ of the Istituto Superiore di Sanita`, Rome, Italy. LITERATURE CITED ANDREWS, R. H., N. B. CHILTON, AND G. MAYRHOFER. 1992. Selection of specific genotypes of Giardia intestinalis by growth in vitro and in vivo. Parasitology 105: 375–386. CACCIO`, S. M., M. DE GIACOMO, AND E. POZIO. 2002. Sequence analysis of the beta-giardin gene and development of a polymerase chain reaction-restriction fragment length polymorphism assay to genotype Giardia duodenalis cysts from human faecal samples. International Journal for Parasitology 32: 1023–1030. CEDILLO-RIVERA, R., J. M. DARBY, J. A. ENCISO-MORENO, G. ORTEGAPIERRES, AND P. L. EY. 2003. Genetic homogeneity of axenic isolates of Giardia intestinalis derived from acute and chronically infected individuals in Mexico. Parasitology Research 90: 119–123. FAUBERT, G. 2000. Immune response to Giardia duodenalis. Clinical Microbiology Reviews 13: 35–54. HOPKINS, R. M., B. P. MELONI, D. M. GROTH, J. D. WETHERALL, J. A. REYNOLDSON, AND R. C. THOMPSON. 1997. Ribosomal RNA sequencing reveals differences between the genotypes of Giardia isolates recovered from humans and dogs living in the same locality. Journal of Parasitology 83: 44–51.

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KARANIS, P., AND P. L. EY. 1998. Characterization of axenic isolates of Giardia intestinalis established from humans and animals in Germany. Parasitology Research 84: 442–449. MCGLADE, T. R., I. D. ROBERTSON, A. D. ELLIOT, AND R. C. THOMPSON. 2003. High prevalence of Giardia detected in cats by PCR. Veterinary Parasitology 110: 197–205. MONIS, P. T., R. H. ANDREWS, G. MAYRHOFER, AND P. L. EY. 2003. Genetic diversity within the morphological species Giardia intestinalis and its relationship to host origin. Infection Genetics and Evolution 3: 29–38. ———, ———, ———, J. MACKRILL, J. KULDA, J. L. ISAAC-RENTON, AND P. L. EY. 1998. Novel lineages of Giardia intestinalis identified by genetic analysis of organisms isolated from dogs in Australia. Parasitology 116: 7–19. PONCE-MACOTELA, M., M. N. MARTINEZ-GORDILLO, R. M. BERMUDEZCRUZ, P. M. SALAZAR-SCHETTINO, G. ORTEGA-PIERRES, AND P. L. EY. 2002. Unusual prevalence of the Giardia intestinalis A-II subtype amongst isolates from humans and domestic animals in Mexico. International Journal for Parasitology 32: 1201–1202. SOTELO-CRUZ, N. 1998. Giardiasis en ninõs. Aspectos clinicos y terapeúticos. Boletino Medico Hospital Infantil de Mexico 55: 47. THOMPSON, R. C. 2000. Giardiasis as a re-emerging infectious disease and its zoonotic potential. International Journal for Parasitology 30: 1259–1267. THOMPSON, S. C. 1994. Giardia lamblia in children and the child care setting: A review of the literature. Journal Paediatric Children Health 30: 202–209. VAN KEULEN, H., P. T. MACECHKO, S. WADE, S. SCHAAF, P. M. WALLIS, AND S. L. ERLANDSEN. 2002. Presence of human Giardia in domestic, farm and wild animals, and environmental samples suggests a zoonotic potential for giardiasis. Veterinary Parasitology 108: 97– 107. [WHO] WORLD HEALTH ORGANISATION. 1996. The World Health Report 1996. World Health Organization, Geneva, Switzerland, 68 p.


Functional Expression of a Recombinant Copper/Zinc Superoxide Dismutase of Filarial Nematode, Brugia malayi W.-G. Lee, J.-H. Hwang*, B.-K. Na†, J.-H. Cho, H.-W. Lee, S.-H. Cho, Y. Kong†, C.-Y. Song‡, and T.-S. Kim§, Department of Tropical and Endemic Parasitic Diseases, National Institute of Health, Seoul 122-701, Korea; *Laboratory of Animal Resources, National Institute of Toxicological Research, Korea Food and Drug Administration, Seoul 122-704, Korea; †Department of Molecular Parasitology and Center for Molecular Medicine, Sungkyunkwan University School of Medicine and Samsung Biomedical Research Institute, Suwon 440-746, Korea; ‡Department of Biology, College of Natural Science, Chung-Ang University, Seoul 156-756, Korea; §To whom correspondence should be addressed. e-mail: [email protected] ABSTRACT: A gene encoding a copper/zinc superoxide dismutase (Cu/ Zn-SOD) of a filarial nematode, Brugia malayi, has been isolated and the biochemical properties of a functionally expressed recombinant enzyme were investigated. The cloned complementary DNA contained a single open reading frame of 477 bp encoding 158 amino acids (aa), which conserved metal-binding residues as well as residues specific for Cu/Zn-SODs. Comparison of the deduced aa sequence of the enzyme with that of other helminthes species, including filarial worms, exhibited high degree of similarities (49–98%). Recombinant enzyme of 32 kDa had an isoelectric point of 6.6 and was shown to consist of 2 subunits linked by interchain disulfide bonds. Enzyme activity of the recombinant protein was inhibited by potassium cyanide and hydrogen peroxide but not by sodium azide. It showed a wide range of pH optima, i.e., 7.0–11.0 and was highly resistant to heat inactivation.

Brugia malayi is a filarial parasite that causes lymphatic filariasis, a disease that affects millions of people in the Southeast Asia. Most of

the lymphatic filariasis includes asymptomatic microfilaremia, but some patients develop lymphatic dysfunction causing lymphedema and elephantiasis, which are frequently involved in the lower extremities (Michael and Bundy, 1997). The host–parasite relationships are dynamic and require the parasite to adapt to, or circumvent, a series of host responses, many of which are quite hostile. Because of an oxygen stress in many organisms, antioxidant systems are used to cope with reactive oxygen species (ROS) generated during cellular metabolism. In addition, parasites are exposed to additional ROS produced by host effector cells. To defend against oxidative killing by the hosts, parasites have evolved a number of defenses including DNA repairing systems, scavenging substrates, and antioxidant enzymes (Callahan et al., 1988). Prominent among antioxidant enzymes are superoxide dismutases (SODs), which catalyze the decomposition of superoxide anion into hydrogen peroxide (H2O2) and molecular oxygen (Fridovich, 1986; Bannister et al., 1987). SODs are postulated to play a major role in the protection of filarial parasites

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against the cellular, oxygen-mediated killing mechanisms of the hosts (Callahan et al., 1991; Henkle et al., 1991; James et al., 1994; HenkleDu¨hrsen et al., 1997; Selkirk et al., 1998). SODs have been cloned or partially characterized from several filarial nematodes, i.e., Dirofilaria immitis (Callahan et al., 1991), Onchocerca volvulus (Henkle et al., 1991; James et al., 1994; Henkle-Du¨hrsen et al., 1995), and Brugia species (Tang et al., 1994; Qu et al., 1995). However, little information is available on enzymatic characteristics of the SODs, especially those of recombinant forms. We herein report on the cloning of a cytosolic copper/zinc superoxide dismutase (Cu/ZnSOD) of B. malayi and expression of the enzyme as an active form in Escherichia coli, and the characterization in part of its biochemical properties. Adult B. malayi was recovered from the peritoneal cavity, heart, lung, and testis of Mongolian jirds (Meriones unguiculatus) experimentally infected with the microfilariae 5 mo before. Worms were harvested, washed with phosphate-buffered saline (0.1 M, pH 7.4) 5 times, and stored at 270 C until use or immediately used for RNA preparation. Total RNA and messenger RNA (mRNA) were isolated using Trizol reagent (GIBCO-BRL, Grand Island, New York) and Oligotex mRNA purification kit (Qiagen, Valencia, California), respectively. Reverse transcription–polymerase chain reaction was performed to synthesize complementary DNA (cDNA) using the sense primer SL1 (59GGGCGGCCGCGGGTTTAATTACCAAGTTTGTA-39) derived from the spliced leader 1 (SL1) sequence of nematodes (Krauser and Hirsh, 1987) and the oligo-dT primer. Polymerase chain reaction (PCR) was performed using 2 specific primers designed on the basis of the Brugia pahangi cytosolic Cu/Zn-SOD sequence (Tang et al., 1994). The forward primer was 59-TTATCATGAGTGCGAAT-39 and the reverse primer was 59-AAAATCAATTCAAGAAGCAG-39. Thermal cycler profile included 94 C (10 min) and 35 cycles of 94 C (1 min), 55 C (1 min), and 72 C (1 min), followed by a 72 C extension (10 min). After being analyzed, the PCR product was ligated into pCR2.1 vector (Invitrogen, Carlsbad, California). The plasmid was transformed into competent E. coli TOP10 cells (Invitrogen), and the nucleotide sequence was determined using the BigDye Terminator Cycle Sequencing Ready Reaction kit (PE Biosystem, Beaconsfield, U.K.) and ABI automated DNA sequencer. The nucleotide sequence is available in GenBank databases under the accession number AY428604. The entire coding region of B. malayi Cu/Zn-SOD gene was amplified by PCR using forward primer 59-GGATCCATGAGTGCGAAT39, which contains a BamHI site upstream of the start codon, and reverse primer 59-CTCGAGTCAAGAAGCAGCAC-39, which harbors XhoI site downstream of the stop codon. The PCR product was finally transformed into competent E. coli TOP10 cells. The plasmid DNA was digested with BamHI and XhoI and placed on pGEX-4T-2 vector (Amersham Biosciences, Uppsala, Sweden) using the same enzyme sites. The recombinant plasmid was transformed into competent E. coli BL-21 cells (Pharmacia) and spread on Luria–Bertani (LB) agar plate containing ampicillin (100 mg/ml). Expression of fusion protein was induced by adding isopropyl-b-D-thiogalactoside (IPTG; a final concentration of 1 mM) for 3 hr at 37 C. The fusion protein was purified using Glutathione Sepharose 4B column (Amersham Biosciences). The GST carrier domain was further cleaved by treating the fusion protein with thrombin (Sigma, St. Louis, Missouri) in cleavage buffer (50 mM Tris– HCl, pH 8.0, 100 mM NaCl, 2.5 mM CaCl2, 0.1% 2-mercaptoethanol) for 4 hr at room temperature. SOD activity was determined by the neotetrazolium chloride (NTC) reduction assay (Kim et al., 2000). One unit of enzyme activity was defined as the amount of the enzyme required to cause 50% inhibition in the rate of reduction of NTC under the assay condition. Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) was performed on 12% gel. Electrophoresis under the nonreducing condition was performed at 4 C in the absence of 2-mercaptoethanol and SDS. SOD activity staining in nonreducing PAGE was done by riboflavin or nitro blue tetrazolium method (Choi et al., 2000). For inhibitor assay, the purified recombinant enzyme (1 mg) was mixed with potassium cyanide (KCN) (6 mM), sodium azide (NaN3) (10 mM), or H2O2 (10 mM) and incubated at 37 C for 30 min, after which the remaining activity in each sample was assayed spectrophotometrically or the samples were electrophoresed on nonreducing gel and stained for SOD activity as described above. To determine the pH optima for the SOD activity, the activity was measured by standard assay in 50 mM sodium

phosphate buffer (pH 7.0), 50 mM Tris–HCl buffers (pH 8.0–9.0), and 50 mM glycine–NaOH buffers (pH 10.0–11.0), instead of the standard buffer. After 30 min of incubation at 37 C, enzyme activity was detected. The enzyme was also incubated at various temperatures, 37, 50, 70, and 100 C, for indicated times, and the residual activities were measured. The isoelectric point of the enzyme was determined by isoelectric focusing on an ampholine PAGplate gel (Amersham Biosciences), pH range 3–9, following the manufacturer’s instruction. After the focusing, the pH gradient was measured with a surface electrode and then the PAGplate divided into 2 parts: the first half was stained with Coomassie Brilliant Blue and the second half was stained for SOD activity as described above. To analyze the presence of disulfide bonds in the active enzyme, the purified enzyme (10 mg) was incubated in various concentrations (0–14.4 mM) of 2-mercaptoethanol for 30 min at room temperature before being subjected to SDS-PAGE without additional boiling (Choi et al., 2000). After electrophoresis, the gel was stained and destained as described above. The Cu/Zn-SOD of B. malayi contained a single open reading frame of 477 bp encoded by 158 amino acids (aa) and possessed metal-binding residues as well as residues specific for Cu/Zn-SODs. A significant degree of sequence similarities was observed with those of B. pahangi (GenBank X76284), O. volvulus (X57150), Acanthocheilonema viteae (AJ240082), D. immitis (AF004949), Fasciola hepatica (AF071229), and Schistosoma mansoni (M97298) by 96.8, 89.2, 89.2, 79.1, 52.7, and 49.0%, respectively (data not shown). The aa residues responsible for binding to copper and zinc (His-46, His-48, His-63, His-71, His-80, His120, and Asp-83) were present in the middle region (Tainer et al., 1983; Fridovich, 1986; Bannister et al., 1987). The Arg residue at position 146 necessary to guide the superoxide anion to the active site (Fridovich, 1986; Bannister et al., 1987) and the 2 Cys residues at positions 57 and 149 that might form disulfide bond were also conserved. The aa residues possibly involved in Cu/Zn-SOD dimmer formation (Gly37, Leu-38, Gly-41, His-43, Arg-79, Gly-85, and Ile-113) were also found with some modification (Deng et al., 1993); Leu-147 and Val151 were replaced by Val and Ile. Neither a putative signal peptide sequence nor a transmembrane-spanning domain was recognized in the sequence. The expression of recombinant B. malayi Cu/Zn-SOD was induced 3 hr after adding IPTG and monitored by SDS-PAGE. A total of 1.5 mg of active enzyme was obtained from 1 L culture. As shown in Figure 1A, the molecular weight of the expressed protein was approximately 42 kDa, which corresponded well with that predicted by cDNA sequence (lane 1): 26 kDa from GST carrier and 16 kDa from B. malayi Cu/Zn-SOD (lane 2). When the fusion protein was further cleaved by thrombin treatment, the recombinant protein migrated to 16 kDa on SDS-PAGE analysis (lane 3). The native molecular mass of the enzyme was about 32 kDa by Ultrogel AcA34 molecular sieve chromatography (data not shown). The isoelectric point of the recombinant SOD was determined to be approximately 6.6 (Fig. 1B). The enzyme activity of the recombinant protein was observed by gel activity staining or spectrophotometrically by NTC reduction assay. As shown in Figure 2A, B, the activity was significantly reduced in the presence of KCN (6 mM) and H2O2 (10 mM) by 82 and 80%, but not in the presence of NaN3 (10 mM). The enzyme retained high activity over a broad pH range between 7.0 and 11.0; however, maximum level of activity was at pH 7.0 (Fig. 3A). The enzyme was highly stable at 37 and 55 C, maintaining activity up to 95% for at least 72 hr, whereas it was highly unstable at 100 C, at which the activity was rapidly inactivated (Fig. 3B). Figure 4 showed the effect of reducing agent. A 32 kDa protein was observed in the absence of 2-mercaptoethanol; however, as the concentration of the reducing agent increased, the 32 kDa band became fainter and a band of approximately 16 kDa became darker and prominent. The protein was completely reduced at the final concentration of 14.4 mM of 2-mercaptoethanol and showed a single band of 16 kDa. Molecular sieve chromatography and reducing property analyses suggested that the active enzyme was dimeric in structure. Considering the fact that each subunit of the enzyme has only 2 Cys residues and the dimeric form is disrupted by reducing agent, but not by SDS, the 2 subunits appeared to be linked by 2 interchain disulfide bonds. There are 2 forms of Cu/Zn-SODs, i.e., extracellular and cytosolic forms. A partial analysis on localization and differential expression of extracellular and cytosolic Cu/Zn-SODs in B. malayi has been reported previously (Qu et al., 1995). It is assumed that B. malayi should express

RESEARCH NOTES

FIGURE 1. Expression, purification, and determination of pI of Brugia malayi recombinant Cu/Zn-SOD. (A) Proteins analyzed by SDSPAGE. Lane 1, GST/SOD fusion protein (3 mg); Lane 2, GST/SOD fusion protein after treatment of thrombin (5 mg); Lane 3, purified recombinant Cu/Zn-SOD of B. malayi (2 mg). Mr, molecular weights in kDa. (B) Isoelectrofocusing gel. Lane 1, stained with Coomassie blue; Lane 2, stained for SOD activity.

FIGURE 2. Inhibitor profile of recombinant Cu/Zn-SOD of Brugia malayi. (A). In gel activity, SOD staining revealed that the enzyme activity was significantly inhibited in the presence of KCN (6 mM) and H2O2 (10 mM) but not in the presence of NaN3 (10 mM). Control lane contained no inhibitor. (B) Quantification of SOD activity of the recombinant enzyme by spectrophotometric assay in the absence or presence of each inhibitor (mean 6 SD, n 5 3).

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FIGURE 3. Effects of pH and heat inactivation on the enzyme activity of recombinant Cu/Zn-SOD of Brugia malayi. (A) Effect of pH. SOD activity was assayed in 50 mM sodium phosphate buffer (pH 7.0), 50 mM Tris–HCl buffers (pH 8.0–9.0), and 50 mM glycine–NaOH buffers (pH 10.0–12.0). Maximal activity was presented as 100%. (B) Thermal stability. The enzyme was incubated at various temperatures for indicated times and then the residual enzyme activities were measured. Maximal activity was shown as 100%. +, 37 C; ●, 50 C; ▫, 70 C; and n, 100 C. considerable amount of SOD to survive in lymphatics and blood vessels where oxidants from host protective cells, including macrophages, prevail. Because superoxide anions are not able to penetrate biological membranes (Lynch and Fridovich, 1978), extracellular form might be more effective than cytosolic form as a defense mechanism used by the parasites against oxidative killing of the hosts. Extracellular SOD was found predominantly in the hypodermis of adult parasite and thought to be associated with the surface of the parasite. Moreover, the enzyme has been detected in peritoneal lavage fluid of B. malayi–infected gerbils, indicating that this enzyme is also secreted in vivo (Tang et al., 1994). In the database of Brugia EST initiatives (http://nema.cap.ed.ac.uk/

FIGURE 4. Effect of a reducing agent on separation of recombinant Cu- and Zn-SOD. Brugia malayi recombinant Cu- and Zn-SOD (10 mg) was treated with various concentrations of 2-mercaptoethanol (0–14.4 mM) and separated by 12% SDS-PAGE. Mr, molecular weights in kDa.

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nematodeESTs/), we found cDNA clusters putatively encoding cytosolic Cu/Zn-SODs (cluster BMC00677). When aligned with these sequences, our sequence exhibited high sequence identity that ranged from 97 to 98%. The gene cloned in this study might represent the Cu/Zn-SOD of B. malayi. Because we could not find any evidence that cytosolic Cu/ Zn-SOD of B. malayi is membrane bound, it might not play a major role in protecting parasites from oxygen-mediated killing mechanisms of the hosts; instead, it might be involved in the survival of parasites by detoxifying intracellular superoxide radicals generated by cellular metabolism. Overexpression of enzymatically active recombinant B. malayi Cu/ Zn-SOD would help us to investigate the enzymatic characteristics and molecular structure of the enzyme in detail. The pathophysiological and biological role of the enzyme in host–parasite interaction should also be further elucidated. This work was supported by a grant from the National Institute of Health, Ministry of Health and Welfare, Republic of Korea (NIH-3486112-156). LITERATURE CITED BANNISTER, J. V., W. H. BANNISTER, AND G. ROTTILIO. 1987. Aspects of the structure, function and application of superoxide dismutases. Critical Review of Biochemistry 22: 111–180. CALLAHAN, H. L., R. K. CROUCH, AND E. R. JAMES. 1988. Helminth antioxidant enzymes: A protective mechanism against host oxidants? Parasitology Today 4: 218–225. ———, ———, AND ———. 1991. Dirofilaria immitis superoxide dismutase: Purification and characterization. Molecular and Biochemical Parasitology 49: 245–252. CHOI, D. H., B. K. NA, M. S. SEO, H. R. SONG, AND C. Y. SONG. 2000. Purification and characterization of iron superoxide dismutase and copper-zinc superoxide dismutase from Acanthamoeba castellanii. Journal of Parasitology 86: 899–907. DENG, H.-X., A. HENTATI, J. A. TAINER, Z. IQGAL, A. CAYABYAB, W.-Y. HUNG, E. GETZOFF, P. HU, B. HERFELDT, R. P. ROSS, C. WARNER, G. DENG, E. SORINO, C. SMYTH, H. E. PARGE, A. AHMED, A. D. ROSES, R. A. HALLWELL, M. A. PERICAK-VANCE, AND T. SIDDIQUE. 1993. Amyotrophic lateral sclerosis and structural defects in Cu, Zn superoxide dismutase. Science 261: 1047–1051.

FRIDOVICH, I. 1986. Superoxide dismutases. Advances in Enzymology 58: 62–97. HENKLE, K. J., E. LIEBAU, S. MULLER, B. BERGMANN, AND R. D. WALTER. 1991. Characterization and molecular cloning of a Cu/Zn superoxide dismutase from the human parasite Onchocerca volvulus. Infection and Immunity 59: 2063–2069. HENKLE-DU¨HRSEN, K., R. S. TUAN, G. WILDENBERG, M. L. ESCHBACH, W. TAWE, P. ZIPFEL, AND R. D. WALTER. 1997. Localization and functional analysis of the cytosolic and extracellular CuZn superoxide dismutases in the human parasitic nematode Onchocerca volvulus. Molecular and Biochemical Parasitology 88: 187–202. JAMES, E. R., D. C. MCLEAN JR., AND F. PERLER. 1994. Molecular cloning of an Onchocerca volvulus extracellular Cu-Zn superoxide dismutase. Infection and Immunity 62: 713–716. KIM, T. S., Y. JUNG, B. K. NA, K. S. KIM, AND P. R. CHUNG. 2000. Molecular cloning and expression of Cu/Zn-containing superoxide dismutase from Fasciola hepatica. Infection and Immunity 68: 3941–3948. KRAUSER, M., AND D. A. HIRSH. 1987. A trans-spliced leader sequence on actin mRNA in C. elegans. Cell 49: 753–761. LYNCH, R. E., AND I. FRIDOVICH. 1978. Permeation of the erythrocyte stroma by superoxide radical. Journal of Biological Chemistry 253: 4696–4699. MICHAEL, M., AND D. A. P. BUNDY. 1997. Global mapping of lymphatic filariasis. Parasitology Today 13: 472–476. QU, X., L. TANG, M. MCCROSSAN, K. HENKLE-DU¨HRSEN, AND M. E. SELKRIK. 1995. Brugia malayi: Localization and differential expression of extracellular and cytoplasmic CuZn superoxide dismutases in adults and microfilariae. Experimental Parasitology 80: 515–529. SELKIRK, M. E., V. P. SMITH, G. R. THOMAS, AND K. GOUNARIS. 1998. Resistance of filarial nematod parasites to oxidative stress. International Journal of Parasitology 28: 1315–1332. TAINER, J. A., E. D. GETZOFF, J. S. RICHARDSON, AND D. C. RICHARDSON. 1983. Structure and mechanism of copper, zinc superoxide dismutase. Nature (London) 306: 284–287. TANG, L., X. OU, K. HENKLE-DU¨HRSEN, AND M. E. SELKIRK. 1994. Extracellular and cytoplasmic CuZn superoxide dismutases from Brugia lymphatic filarial nematode parasites. Infection and Immunity 62: 961–967.


Modified Sugar Centrifugal Flotation Technique for Recovering Echinococcus multilocularis Eggs From Soil Kayoko Matsuo and Haruo Kamiya*, Department of Parasitology, Hirosaki University School of Medicine, Zaihu-cho, Hirosaki 036-8562, Japan; *To whom correspondence should be addressed. e-mail: [email protected] ABSTRACT: Among soil-transmitted parasitic diseases, alveolar hydatidosis due to the ingestion of Echinococcus multilocularis eggs is becoming a serious problem in Hokkaido, the northern most island of Japan. Dissemination of the infection far from the endemic areas can occur if motor vehicles transmit soil contaminated with eggs. No appropriate and validated method for recovering the taeniid eggs from soil is available. A modified sugar centrifugal flotation technique, using a sucrose solution of specific gravity 1.27 and 0.05% Tween-80, was evaluated as a method to successfully recover eggs from soil. Contamination levels as low as 10 eggs per gram could be detected. This method may be useful to determine the prevalence of E. multilocularis, its transmission, and the potential for by monitoring soil contamination with eggs.

Soil is contaminated with various kinds of helminth eggs, which can be transmitted to humans and other animals (Mizgajska, 1997; Chongsuvivatwong et al., 1999). Echinococcus multilocularis is the causative agent of alveolar hydatidosis and is distributed widely in the Northern

Hemisphere. Echinococcus multilocularis eggs are deposited in the environment by defecation from the infected definitive hosts, mainly foxes. Human infection with E. multilocularis is more likely to occur by accidental ingestion of eggs in soil, contaminated vegetables, water etc., rather than through direct contact with definitive hosts, except dogs. Various techniques have been developed for ascarid egg detection from the soil (World Health Organization Chronicle, 1968; Uga et al., 1989; Ruiz de Ybanez et al., 2000). However, few reports for recovering taeniid eggs from soil are available. A method for recovering ascarid eggs from sand (Uga et al., 1989) was adapted to a sugar centrifugal flotation technique (Ito, 1980) using a sucrose solution of specific gravity 1.27 (Nonaka et al., 1998). Soil was collected from the ground of Hirosaki University School of Medicine located in a nonendemic area for E. multilocularis. Twenty grams of soil mixed with 20 (equivalent to 1 egg per gram [EPG]), 200 (10 EPG), or 2,000 (100 EPG) E. multilocularis eggs was placed in 50-ml conical tube to which was added 40 ml of 0.05% Tween-80. The eggs were not infective, having been preserved in 70% ethanol since 1969.

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TABLE I. Modified sugar centrifugal flotation technique for recovering Echinococcus multilocularis eggs from 20 g of soil. Number of eggs detected (mean 6 SD)† Time of incubation 2 hr Overnight Total * † ‡ §

20‡ (1 EPG)

200‡ (10 EPG)

2,000‡ (100 EPG)

0.3 6 0.5 (3/10)§ 0.2 6 0.4 (2/10) 0.5 6 0.5 (5/10)

1.5 6 0.7 (10/10)** 8.1 6 3.7 (10/10)*** 9.6 6 4.0 (10/10)***

64 6 18.7 (10/10)*** 137 6 40.8 (10/10)*** 201 6 54.5 (10/10)***

P , 0.05; ** P , 0.01; *** P , 0.001. Number of detected eggs were evaluated by Student’s t-test and found significant, where P , 0.05, compared with that of sample containing 1 EPG. Number of eggs added. Number of tubes detected eggs/examined (n 5 10).

The mixture was stirred vigorously and sieved through 100-mm mesh. The suspension was centrifuged at 1,000 g for 5 min, and the supernatant was discarded. The sediment was resuspended in 7–8 ml sucrose solution (1.27 specific gravity). The suspension was transferred to 15ml tubes and centrifuged again at 1,000 g for 15 min. Tubes were filled to the top, and a coverslip (24 3 24 mm) was placed on the tube. Coverslips were examined microscopically 2 hr later. A new coverslip was placed on the top of tube and left overnight to detect the remaining eggs. Statistical significance of the results was determined using Student’s t-test. Data were expressed as mean 6 SD, and a P value of less than 0.05 was taken as the minimum level of significance. Eggs were detected in all soil samples added with 100 EPG after 2 hr of incubation. More eggs were detected after overnight incubation compared with 2 hr of incubation. As shown in Table I, the egg recovery rate was 2.5, 4.8, and 10% in soil samples containing 1, 10, and 100 EPG, respectively. The egg detection rate significantly increased with increase in EPG. The present method could detect contamination of soil with taeniid eggs at a concentration of 10 EPG. An average of 300 eggs per gravid segment was counted in E. multilocularis collected from the fox (Zeyhle and Bosch, 1982). Therefore, if only 1 gravid segment was mixed with 20 g of soil, the EPG in this locality will be more than 10. A total of 34,000 E. multilocularis adults were recorded from a heavily infected fox in endemic area of Japan (Morishima et al., 1999). Echinococcus multilocularis eggs deposited in feces and dispersed into soil could clearly contaminate a wide area. Furthermore, if fox heavily infected with E. multilocularis was involved in a motor accident, a huge number of eggs could be disseminated into the environment from injured viscera. According to Tsukada et al. (2000), infected foxes frequently encountered traffic accidents in Hokkaido, Japan. Therefore, motorcars contaminated with eggs could disperse the parasite to nonendemic areas. It is somewhat difficult to evaluate the sensitivity of present techniques for application to field survey. However, the contamination of soil with E. multilocularis is not the same in different areas. The contamination of eggs in soil should be more intense around fox dens, showing that the wild rodents, its intermediate host, captured near the dens were heavily infected (Kamiya et al., 1977). Therefore, the sensitivity level of 1 EPG is still valuable for detection of eggs in certain instances such as in the soil of highly endemic areas or soil adhered to motorcars run over the injured fox viscera infected with E. multilocularis. Recently, the present technique adapted for 2 kg of soil was used in a survey for detecting E. multilocularis eggs in soil left in the Hokkaido to Aomori ferryboat to monitor the transmission of eggs from endemic to nonendemic area. Although no helminth egg was detected, Isospora oocysts, mites, and eggs of mites were found (Matsuo et al., 2003). Of course, the soil collected from the endemic area must be incubated at 70 C for 12 hr (Nonaka et al., 1998) or frozen at 280 C for 3 days or more (Veit et al., 1995) to inactivate egg infectivity.

LITERATURE CITED CHONGSUVIVATWONG, V., S. UGA, AND W. NAGNAEN. 1999. Soil contamination and infections by soil-transmitted helminths in an endemic village in southern Thailand. Southeast Asian Journal of Tropical Medicine and Public Health 30: 64–67. ITO, S. 1980. Modified Wisconsin sugar centrifugal-flotation technique for nematode eggs in bovine feces. Journal of Japan Veterinary Medical Association 33: 424–429. KAMIYA, H., M. OHBAYASHI, K. SUGAWARA, AND K. HATTORI. 1977. An epidemiological survey of multiloculara echinococcoisis in small mammals of eastern Hokkaido, Japan. Japanese Journal of Parasitology 26: 148–156. MATSUO, K., T. INABA, AND H. KAMIYA. 2003. Detection of Echinococcus multilocularis eggs by centrifugal flotation technique: Preliminary survey of soil left in the ferryboats commuting between Hokkaido Island, where E. multilocularis is endemic, and mainland Japan. Japanese Journal of Infectious Diseases 56: 118–119. MIZGAJSKA, H. 1997. The role of some environmental factors in the contamination of soil with Toxocara spp. and other geohelminth eggs. Parasitology International 46: 67–72. MORISHIMA, Y., H. TSUKADA, N. NONAKA, Y. OKU, AND M. KAMIYA. 1999. Evaluation of coproantigen diagnosis for natural Echinococcus multilocularis infection in red foxes. Japanese Journal of Veterinary Research 46: 185–189. NONAKA, N., H. TSUKADA, N. ABE, Y. OKU, AND M. KAMIYA. 1998. Monitoring of Echinococcus multilocularis infection in red foxes in Shiretoko, Japan, by coproantigen detection. Parasitology 117: 193–200. RUIZ DE YBANEZ, M. R., M. GARIJO, M. GOYENA, AND F. D. ALONSO. 2000. Improved methods for recovering eggs of Toxocara canis from soil. Journal of Helminthology 74: 349–353. TSUKADA, H., Y. MORISHIMA, N. NONAKA, Y. OKU, AND M. KAMIYA. 2000. Preliminary study of the role of red foxes in Echinococcus multilocularis transmission in the urban area of Sapporo, Japan. Parasitology 120: 423–428. UGA, S., T. MATSUMURA, N. AOKI, AND N. KATAOKA. 1989. Prevalence of Toxocara species eggs in the sandpits of public parks in Hyogo Prefecture, Japan. Japanese Journal of Parasitology 38: 280–284. VEIT, P., B. BILGER, V. SCHAD, J. SCHAFER, W. FRANK, AND R. LUCIUS. 1995. Influence of environmental factors on the infectivity of Echinococcus multilocularis eggs. Parasitology 110: 79–86. WORLD HEALTH ORGANIZATION CHRONICLE. 1967. Control of ascariasis. Technical Report Series No. 379. World Health Organization, Geneva, Switzerland, p. 155–159. ZEYHLE, E., AND D. BOSCH. 1982. Comparative experimental infection of cats and foxes with Echinococcus multilocularis. Zentralblatt fu¨r Bakteriologie 277: 117–118.

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Spatial Variation of Trichinella Prevalence in Rats in Finnish Waste Disposal Sites T. Mikkonen, J. Valkama*, H. Wihlman†, and A. Sukura, Department of Basic Veterinary Sciences, Faculty of Veterinary Medicine, P.O. Box 66, University of Helsinki, FIN-00014 Helsinki, Finland; *Ringing Centre, Finnish Museum of Natural History, University of Helsinki, FIN-00014 Helsinki, Finland; †The National Food Agency, Vanha talvitie 5, P.O. Box 28, 00581 Helsinki, Finland. e-mail: [email protected] ABSTRACT: Trichinellosis is 1 of the most widespread parasitic zoonoses in the world and can be lethal to humans. Trichinella spp. are also parasites of considerable economic importance. Because rats may play a role in the transmission of trichinellosis to swine and farmed wild boar, 767 brown rats (Rattus norvegicus Berkenhout) from 13 Finnish waste disposal sites were examined for Trichinella spp. by a HCl–pepsin digestion method. Trichinella spp. were found to be a common parasite in trapped rats (overall prevalence, 19%) detected in 12 of 13 dumps. Significant differences were observed between sites in the prevalence (0–49%) of Trichinella spp. Female rats were more often and more heavily infected than males, but age was not shown to be a risk factor for trichinellosis. In addition, positive correlation was demonstrated between rat population density and prevalence. Trichinella spiralis was identified by multiplex polymerase chain reaction in 28 rats. The median density of infection was 42 (range, 0.5–6,925) larvae/ g of host tissue, but neither the occurrence nor the density of the parasite was related to the physical condition of the animal.

Trichinella spp. are cosmopolitan nematodes, with the largest range of host species of any parasitic nematode (Dupouy-Camet, 2000; Pozio, 2000). Trichinellosis is 1 of the most widespread parasitic zoonoses worldwide and can be lethal to humans (Capo and Despommier, 1996). Considerable financial losses are caused in controlling the parasite (Pozio, 1998). The medical and social costs of human outbreaks of trichinellosis can also be notable (Ancelle et al., 1990). Trichinella spp. are transmitted passively when a host ingests raw or undercooked meat infected with the organism. The life cycle of Trichinella spp. is divided into a sylvatic cycle among wildlife hosts and a synanthropic cycle when it is associated with domestic animals and humans (Campbell, 1988). These 2 cycles can coexist. Studies on trichinellosis among wildlife in Finland have demonstrated the parasite to be common in raccoon dogs (Nyctereutes procyonoides), foxes (Vulpes vulpes), and other carnivores (Oksanen et al., 1998; Oivanen et al., 2002). The prevalence varies geographically, being higher in southern than in northern Finland (Oksanen et al., 1998; Oivanen et al., 2002). Interestingly, Trichinella spiralis, typical in the synanthropic cycle, has also been found in sylvatic animals in Finland (Oivanen et al., 2002). Finland used to be, in addition to Spain, the only country in the European Union where porcine trichinellosis was found frequently (Pozio, 1998), and during the 1980s, the prevalence of trichinellosis in domestic pigs and wildlife increased dramatically (Oivanen et al., 2002). However, the situation has changed since 2004, when new, high endemic nations, such as Poland and Baltic countries, have joined the European Union. Rats are thought to play an important role in the transmission of trichinellosis to swine (Schad et al., 1987; Smith and Kay, 1987) and farmed wild boar (Oivanen et al., 2000). However, rat trichinellosis has seldom been studied under natural conditions and often only with small sample sizes. Our aim in this article was to study trichinellosis in natural rat populations at waste disposal sites. Rat populations can rapidly increase on small communal waste disposal sites because of the lack of appropriate preventive measures. Because these sites are relatively homogeneous, study of these allows accurate comparisons to be made on the distribution of Trichinella spp., and risk factors for infection and their effects on the host to be evaluated. Rats were collected from 13 waste disposal sites; 12 of these sites belonged to a rat control experiment conducted in 1994. The 13th site was included in the study after a rat control hazard that resulted in the exodus of the rats after the site’s closure in the autumn of 2000. The original 12 sites were situated in southwestern Finland. The sites were of small or medium size, each handling the refuse of 1,400–10,000

people. In addition to the number of people using the sites, the size of each waste disposal site was also determined according to the amount of waste (m3/yr) and area (ha) combined (m3/yr/ha) (Anonymous, 1992). No rat poison was used during or 3 mo before the study (sites 1–12). The rats were caught with live traps (Raina-Tuote, Naantali, Finland). Three separate trappings, for 5 days each, were carried out at each site between 8 April and 16 September 1994. The captured rats were anesthetized with ether and marked with toe clipping such that all individuals caught during the same day received a similar marking. Numbers of marked and unmarked individuals were used when determining the estimate for the population size during each trapping (for methods, see Krebs, 1989). After the fourth trapnight, during the first and second trapping series, 3–6 adult female rats were randomly killed. During the third effort, all rats (a maximum of 50) caught on the last night were killed. The population density (rats/ha) was calculated by dividing the estimated number of rats by the area (ha) of the site. Because of the nature of the original study, the rats killed during spring and summer were mainly adult females, whereas on the final trapnight all the captured rats were killed, irrespective of their sex or age. Eight rats caught from the 13th waste disposal site in southern Finland in the autumn of 2000 were captured alive in traps, as described above, and killed. Local hunters collected the rest of the sampled rats (190). Site 13, which disposed waste for a community of 13,000 people, was closed at the end of September 2000. Because of insufficient preventative actions before the closure, such as the use of adequate rat poisoning, covering of the site with soil, and leveling of the site, the rat population had not decreased. Under weakened conditions, i.e., lack of nutritional resources, rats migrate in mass to surrounding areas. The invasion started on the night of 7–8 October 2000. On 9 October, there were still too many rats that hunters shot the 190 used in this study in 2 hr. Rats killed in 1994 were stored in a freezer (218 C), but rats killed from 2000 were not frozen before examination. Sex, body weight (to the nearest 5 g), and length (without tail) were recorded. After skinning and evisceration, the animals were weighed again. Individuals ,170 mm were classified as juveniles, and individuals $170 mm were considered adults (Bishop and Hartley, 1976). Samples (1 g) were taken from the diaphragm or abdominal muscles (or both) to examine for the presence of Trichinella spp. (sites 1–12). In addition, the tongue and flexor muscles (5-g samples) were used from rats of site 13. A total of 767 rats, including 294 males and 454 females (19 animals not sexed), were examined for Trichinella spp. The samples were artificially digested by the HCL–pepsin Stomacher method, according to modified Commission Directive 84/319/EEC (Anonymous, 1984). The larvae were counted in a petri dish under a stereomicroscope (330). The density of larvae was expressed as larvae per gram (LPG). Muscle larvae were stored at 220 C in 70% ethyl alcohol in distilled water and rehydrated in a descending alcohol series before DNA extraction. Trichinella species were identified on pooled samples of 2–40 larvae by multiplex polymerase chain reaction (multiplex-PCR), slightly modified from Zarlenga et al. (1999). The reference strains used were T. spiralis (ISS004), Trichinella nativa (ISS042), Trichinella britovi (ISS100), and Trichinella pseudospiralis (ISS013). Species determination was made from 825 muscle larvae from 45 individual rat hosts from 8 sites. Oivanen et al. (2002) analyzed a separate stock of larvae from 29 rat hosts, 28 of which overlapped with this study. The prevalences between the sites were compared with Pearson chisquare test. Logistic regression was used to determine the risk factors for trichinellosis and to analyze the associations between the occurrence of Trichinella spp. and the rat population density, as well as the amount of waste handled in the waste disposal site.

RESEARCH NOTES

211

TABLE I. Density of the rat population and distribution of rats examined for Trichinella spp. by sex and waste disposal site.

Site 1 2 3 4 5 6 7 8 9 10 11 12 13 Total

Density (no./ha) 157 533 52 618 242 423 73 950 187 426 516 331 ND

Infected/examined Male

Female

2/34 8/19 0/17 4/9 7/18 0/13 0/4 1/17 1/3 2/15 7/19 4/26 0/100 36/294

3/53 13/35 4/28 21/43 22/41 7/19 1/16 2/44 5/14 9/27 11/26 8/29 0/79 106/454

ND*

Total

Prevalence (%)

19 19

5/87 21/54 4/45 25/52 29/59 7/32 1/20 3/61 6/17 11/42 18/45 12/55 0/198 142/767

5.7 39 8.8 48 49 21.9 5.0 4.9 35 26 40 22 0 19

95% CI† 1.0–11 26–52 0.6–17 34–62 36–62 7.7–36 0–15 0–10 12–57 13–39 26–54 11–33 0

LPG (median) 3 90 1 101 68 69 1 17 36 17 63 28 0 42

* ND, not determined. † CI, confidence interval.

Body condition was measured as the residual of the regression of body length (tip of nose to anus) against body mass. Because the relationship between body weight and length was nonlinear, logarithmic transformations were made. Analysis of variance (ANOVA) was used to determine whether the occurrence of Trichinella spp. and the density of the parasite were related to the body condition, sex, or age of rat hosts from different sites. Probabilities (P) of less than 0.05 were considered to be statistically significant. The ratio between the variance and the mean density (LPG) indicates whether the parasite population is aggregated. The statistical analyses were performed using software packages SPSS 10.0 for Windows (SPSS Inc., 1999) and Statistics for Windows (Analytical Software, 1996). The overall prevalence of Trichinella spp. was 19% (142/767), and parasites were present at 12 of the 13 sites. The prevalence (Pearson x2 5 168, df 5 12, P , 0.001) of Trichinella spp. differed significantly between sites. Interestingly, the 198 rats caught from site 13 were all negative for Trichinella spp. The data are presented in detail in Table I. The data were analyzed to determine the probability of an individual rat being infected with Trichinella spp. and to identify factors affecting

FIGURE 1. Regression between the prevalence of Trichinella spp. and population densities of rats (no. of rats/ha) in 12 Finnish waste disposal sites. The filled circle represents outlier area 8, which was not used in the calculation of the regression line.

infection at the site (population) level. The ages of 757 rats were recorded. Fifteen percent of young rats (16/110) and 19% of adult rats (126/647) were infected; 23% (106/454) of female rats were found to carry Trichinella spp., whereas only 12% (36/294) of male rats were infected. Both sex and age were determined for 741 rats; 14% (34/249) of adult males and 4% (2/45) of juvenile males were Trichinella spp. positive, whereas 27% (14/52) of juvenile female rats and 23% (92/395) of adults had an infection. The density of parasites varied considerably; the minimum was 0.5 LPG of muscle tissue, but 1 adult female with 6,925 LPG was also found (median, 42 LPG). However, 40% of the infected rats had a density .100 LPG. The median density of infected adult males (n 5 34) was 59 (range, 1–2,733) LPG and females (n 5 92) was 35 (range, 0.5–6,925) LPG. Juvenile males (n 5 2) had a median density of 560 (30 and 1,089) LPG and females (n 5 14) had 42 (range, 3–596) LPG. The density of the rat population and their sex and age were selected as variables to explain the differences in the occurrence of Trichinella spp. in a logistic regression model. Because the amount of communal waste correlated positively with the size of the human population in the area, human population was excluded from the analysis. Area 8 was excluded from the comparison because the rat population density was exceptionally high because of enormous amounts of vegetable waste contributed by a nearby food factory. Density was not measured for site 13. Overall, in the main effect model, the probability of a rat being infected was significantly dependent on population density (odds ratio [OR], 1.003; 95% confidence interval [CI], 1.002–1.004; P , 0.01) and host sex (OR, 1.846; 95% CI, 1.177–2.895; P , 0.01). The higher the rat population density, the more prevalent was the parasite (Fig. 1). Females were more often infected than males, whereas age had no significant influence on infection probability (OR, 1.077; 95% CI, 0.576– 2.015; P . 0.5). The amount of waste did not affect the prevalence of Trichinella spp. (data not shown). The residual of the regression of body length and body mass was used as an index in evaluating the physical condition of the rats. Body condition was unaffected by infection density of Trichinella spp. or the occurrence of the parasite (ANOVA, P . 0.05; data not shown) in different areas. Female and adult rats were in better conditions than males and juveniles, and the condition differed significantly between areas (P , 0.05). The variance of the data was higher than the mean (s2 5 258,766, x¯ 5 106.7), indicating that the Trichinella spp. population was highly aggregated among the hosts. The observation that the LPG number of 1 heavily infected individual (6,925 LPG) accounts for 11% of the cumulative LPG value (60,720 LPG) in all rats supports this finding. In multiplex-PCR, only T. spiralis was identified in 28 rats that originated from 8 different sites.

212


Compared with other studies on wild rats, the overall Trichinella spp. prevalence of 19% found here was quite high. A study of 944 rats from dumps and farms in Iowa, for instance, yielded 5% of infected animals (Zimmermann and Hubbard, 1969). However, higher prevalences have also been reported elsewhere. In an American city dump, the prevalence of Trichinella spp. in rats was quite similar to our results (Robinson and Olsen, 1960), reaching 42% on 1 farm (Leiby et al., 1990) and challenging our highest prevalence of 49% from site 5. The probability of a rat becoming infected with Trichinella spp. in this study was dependent on the area. Trichinella spp. was identified at 12 of the 13 sites studied. At 8 of the infected sites, the prevalence was high, and at 4 sites it was low. Possibly, the infection pressure from the wild and synanthropic cycles combined could be the source of the infection in waste disposal sites. Despite the ban of meat waste at all sites, 1 meat-processing factory ignored this edict and disposed of meat at a waste disposal site. Some farmers also disposed of dead pigs. Because meat is officially inspected and pork trichinellosis is rare (prevalence, 0.004%), it seems unlikely that the rats at the 12 infected sites were exposed to Trichinella spp. through pork. However, dead foxes and raccoon dogs were occasionally seen at the sites. Raccoon dogs and foxes are abundant in southern Finland, and hunting them is common. Hunters may discard the carcasses of prey in waste disposal sites together with household garbage, and traffic-killed animals may also end up in these sites. The improper disposal of the carcasses of hunted wolves has been established to increase the prevalence of Trichinella spp. (Pozio et al., 2001). In southern parts of Finland, Trichinella spp. is often found in wild carnivores, i.e., 40% of lynx (Oksanen et al., 1998), 49% of raccoon dogs (Mikkonen et al., 1995), and 74% of foxes (Oivanen et al., 2000) have been reported to carry Trichinella spp. In Finland, T. spiralis has been identified commonly in raccoon dogs as well as in red fox, although T. nativa is the most prevalent species in the sylvatic cycle (Oivanen et al., 2002). Abundance of the species in the study area leaves open the possibility of transmission of Trichinella spp. from sylvatic animals to waste disposal site rats. In experimental infections, T. nativa has shown only low infectivity and persistency in rats (Pozio et al., 1992; Malakauskas et al., 2001), which may explain why T. spiralis, but no T. nativa, was detected in these rats. Host density is assumed to be connected with the opportunity of a parasite to invade a host population, and colonization by a parasite is dependent on the number of hosts available in an area (Morand and Poulin, 1998). Rats are known to exhibit cannibalistic behavior (Calhoun, 1962; Boice, 1972), especially in high-density populations where social stress is strong. We recorded high population densities (950 individuals/ha), and the prevalence of Trichinella spp. increased with rat population density. The disposal sites with low, 0, or high prevalence were quite similar in size (area and amount of waste/ha), and the surrounding environments were also similar. All positive sites were located in the countryside enclosed by fields and forests. No animal husbandry was practiced in the close vicinity of the 12 positive sites. The Trichinella-negative site was situated in a village, where the closest houses were 200 m from the site. However, fields and forests bordered it on 1 side. Surrounding areas, such as large forests, could act as natural barriers to rats and their parasites. Because of the lack of an enclosing fence, wildlife had free access to the waste disposal sites. Similar to many other helminth parasites (Shaw and Dobson, 1995), T. spiralis in rats is distributed in an aggregated manner, with the parasites being concentrated only in a small proportion of hosts and most hosts having few or no parasite. The overdispersion of parasites in the host population influences the pathogenicity (Anderson and May, 1979; May and Anderson, 1979) and affects the parasite-induced morbidity and mortality of the hosts (Anderson, 1987; Smith and Guerrero, 1993). In our study, neither the occurrence nor the intensity of T. spiralis affected the animal’s body condition. Our finding that female rats are more often and more heavily infected with Trichinella spp. than males is consistent with the study of Leiby et al. (1990). Nevertheless, there are conflicting results concerning the correlation of sex with the prevalence and abundance of parasites. Male raccoon dogs were demonstrated to be more often infected with Trichinella spp. than females (Mikkonen et al., 1995); however, no such difference was observed in foxes (Roneus and Christensson, 1979; Prestrud et al., 1993). The intensity of infection depends on exposure as

well as susceptibility, i.e., innate and acquired immunity. The nutritional habits of male and female rats are similar. However, reproduction is costly to the female. The cost of the high reproductive effort of female rats, which are often pregnant and weaning, could decrease the female’s ability to defend against parasites. We expected adult rats to be infected more often than juveniles, but this was not the case in this study. This may be because of the short life span of rats; they seldom live beyond 1 year of age (Glass et al., 1988). Summing up, this study recorded high but variable prevalences of T. spiralis at waste disposal sites, which can increase the risk of Trichinella spp. invading the domestic cycle. The higher the rat population density, the higher the prevalence. Female rats were more often and more heavily infected than males. The authors wish to thank Tapani Va¨ha¨vahe, Jorma Hirn, and Mikko Vesanen for providing study material and Sakari Mykra¨ and all others who assisted in the fieldwork. Voitto Haukisalmi is acknowledged for his helpful advice, and Carol Ann Pelli is thanked for editing the language of the manuscript. This project was financially supported by the Marjatta and Eino Kolli Foundation, the Emil Aaltonen Foundation, and the Farmos Research and Science Foundation. LITERATURE CITED ANALYTICAL SOFTWARE. 1996. Statistics for Windows. Analytical Software, Tallahassee, Florida, 333 p. ANCELLE, T., G. RENAUD, J. DUPOUY-CAMET, AND G. FOULON. 1990. Evaluation du couˆt me´dical et social de deux e´pide´mies de trichinose survenues en France en 1985. (Evaluation of the medical and social cost of 2 trichinosis outbreaks in France in 1985.) Revue d’Epide´miologie et de Sante´ Publique 38: 179–186. ANDERSON, R. M. 1987. The role of mathematical models in helminth population biology. International Journal for Parasitology 17: 519– 529. ———, AND R. M. MAY. 1979. Population biology of infectious diseases: Part 1. Nature 280: 361–367. ANONYMOUS. 1984. Commission Directive 84/319/EEC. Journal of European Communities 167: 34–43. ANONYMOUS. 1992. Suunnittelukeskus Oy. Turun ja Porin laä¨nin yhdyskuntien ja¨tehuollon kehitta¨missuunnitelma. (Turku and Pori Countys waste development plan.) Varsinais-Suomen Seutukaavaliiton Offset, Turku, Finland, 168 p. BISHOP, J. A., AND D. J. HARTLEY. 1976. The size and age structure of rural populations of Rattus norvegicus containing individuals resistant to the anticoagulant poison warfarin. Journal of Animal Ecology 45: 623–646. BOICE, R. 1972. Some behavioral tests of domestication in Norway rats. Behaviour 42: 198–231. CALHOUN, J. B. 1962. The ecology and sociology of the Norway rat. Publications of U.S. Department of Health and Education Welfare, Publications of Health Services No. 1008. Bethesda, Maryland, 288 p. CAMPBELL, W. C. 1988. Trichinosis revisited, another look at modes of transmission. Parasitology Today 4: 83–86. CAPO, V., AND D. D. DESPOMMIER. 1996. Clinical aspects of infection with Trichinella spp. Clinical Microbiology Reviews 9: 47–54. DUPOUY-CAMET, J. 2000. Trichinellosis: A worldwide zoonosis. Veterinary Parasitology 93: 191–200. GLASS, G. E., G. W. KORCH, AND J. E. CHILDS. 1988. Seasonal and habitat differences in growth rates of wild Rattus norvegicus. Journal of Mammalogy 69: 587–592. KREBS, C. J. 1989. Ecological methodology. Harper & Row, New York, 654 p. LEIBY, D. A., C. H. DUFFY, K. D. MURRELL, AND G. A. SCHAD. 1990. Trichinella spiralis in an agricultural ecosystem: Transmission in the rat population. Journal of Parasitology 76: 360–364. MALAKAUSKAS, A., C. M. O. KAPEL, AND P. WEBSTER. 2001. Infectivity, persistence and serological response of nine Trichinella genotypes in rats. Parasite 8: 216–222. MAY, R. M., AND R. M. ANDERSON. 1979. Population biology of infectious diseases: Part 2. Nature 280: 455–461. MIKKONEN, T., V. HAUKISALMI, K. KAUHALA, AND H. WIHLMAN. 1995. Trichinella spiralis in the raccoon dog (Nyctereutes procyonoides)

RESEARCH NOTES

in Finland. Bulletin of the Scandinavian Society for Parasitology 2: 100. MORAND, S., AND R. POULIN. 1998. Density, body mass and parasite species richness of terrestrial mammals. Evolutionary Ecology 12: 717–727. OIVANEN, L., C. M. O. KAPEL, E. POZIO, G. LA ROSA, T. MIKKONEN, AND A. SUKURA. 2002. Associations between Trichinella species and host species in Finland. Journal of Parasitology 88: 84–88. ———, T. MIKKONEN, AND A. SUKURA. 2000. Survey of trichinellosis in wild boar farm. Acta Pathologica, Microbiologica et Immunologica Scandinavica 108: 814–818. OKSANEN, A., E. LINDGREN, AND P. TUNKKARI. 1998. Epidemiology of trichinellosis in lynx in Finland. Journal of Helminthology 72: 47– 53. POZIO, E. 1998. Trichinellosis in the European Union: Epidemiology, ecology and economic impact. Parasitology Today 14: 35–38. ———. 2000. Factors affecting the flow among domestic, synanthropic and sylvatic cycles of Trichinella. Veterinary Parasitology 93: 241– 262. ———, A. CASULLI, V. V. BOLOGOV, G. MARUZZI, AND G. LA ROSA. 2001. Hunting practices increase the prevalence of Trichinella infection in wolves from European Russia. Journal of Parasitology 87: 498–501. ———, G. LA ROSA, P. ROSSI, AND K. D. MURRELL. 1992. Biological characterization of Trichinella isolates from various host species and geographical regions. Journal of Parasitology 78: 647–653. PRESTRUD, P., G. STUVE, AND G. HOLT. 1993. The prevalence of Trich-

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inella sp. in arctic foxes (Alopex lagopus) in Svalbard. Journal of Wildlife Diseases 29: 337–340. ROBINSON, H. A., AND O. W. OLSEN. 1960. The role of rats and mice in the transmission of the porkworm, Trichinella spiralis (Owens, 1835) Raillett, 1835. Journal of Parasitology 48: 589. RONEUS, O., AND D. CHRISTENSSON. 1979. Presence of Trichinella spiralis in free living red foxes (Vulpes vulpes) in Sweden related to Trichinella infection in swine and man. Acta Veterinaria Scandinavica 20: 583–594. SCHAD, G. A., C. H. DUFFY, D. A. LEIBY, K. D. MURRELL, AND E. W. ZIRKLE. 1987. Trichinella spiralis in an agricultural ecosystem: Transmission under natural and experimentally modified on farm conditions. Journal of Parasitology 73: 95–102. SHAW, D. J., AND A. P. DOBSON. 1995. Patterns of macroparasitic abundance and aggregation in wildlife populations: A quantitative review. Parasitology 111: S111–S133. SMITH, G., AND J. GUERRERO. 1993. Mathematical models for the population biology of Ostertagia ostertagi and the significance of aggregated parasite distribution. Veterinary Parasitology 46: 243–257. SMITH, H. J., AND E. K. KAY. 1987. Role of rats in the transmission of Trichinella spiralis to swine. Canadian Veterinary Journal 28: 604. SPSS INC. 1999. SPSS 10.0 for Windows. SPSS, Chicago, Illinois, 233 p. ZARLENGA, D. S., M. B. CHUTE, A. MARTIN, AND C. M. O. KAPEL. 1999. A multiplex PCR for unequivocal differentiation of six encapsulated and three non-encapsulated genotypes of Trichinella. International Journal for Parasitology 29: 1859–1867. ZIMMERMANN, W. J., AND E. D. HUBBARD. 1969. Trichiniasis in wildlife of Iowa. American Journal of Epidemiology 90: 84–92.


Helminths of the Virginia Opossum Didelphis virginiana (Mammalia: Didelphidae) in Mexico Anne Monet-Mendoza, David Osorio-Sarabia, and Luis Garcıá-Prieto*, Laboratorio de Helmintologıá, Instituto de Biologıá, Universidad Nacional Autońoma de Me´xico, AP 70-153, 04510, Coyoacań, D.F., Me´xico; *To whom correspondence should be addressed. e-mail: [email protected] ABSTRACT: The goal of this study was to provide further information about helminth parasites of Virginia opossum Didelphis virginiana Kerr, 1792 from Mexico. During routine faunal investigations between 1958 and 2001, 101 opossum were necropsied. Nineteen taxa of helminths were collected, representing 13 genera from hosts in 27 localities from Mexico. There are 58 new locality records, with 6 species recorded in Mexico for the first time: Brachylaima virginiana Dickerson, 1930; Cruzia americana Mapleston, 1930; Didelphonema longispiculata (Hill, 1939); Didelphostrongylus hayesi Prestwood, 1976; Viannaia didelphis Travassos, 1914; and Viannaia viannai Travassos, 1914. This increases the number of helminth taxa previously known for this host in Mexico to 28.

The opossum, Didelphis virginiana Kerr, 1792, occurs from southern Canada through the United States and Mexico, down to Costa Rica (Gardner, 1982). Its helminth fauna has been the subject of extensive investigation mostly in North America, where 75 taxa have been described (Potkay, 1977; Harris, 1983; Alden, 1995). However, helminthological studies of this host in Mexico are limited, making available information scarce and fragmentary (Lamothe, 1981; Prado, 1993; Can˜eda, 1997; Salgado and Cruz, 2002). Thus far, 18 taxa have been reported from Mexico. The aim of this study was to compile and provide further information concerning helminth parasites of D. virginiana in several localities of Mexico. The specimens studied were collected during routine faunal investigations developed in the Laboratorio de Helmintologıá, Instituto de Biologıá, Universidad Nacional Autońoma de Me´xico (UNAM) over several years (1958–2001) and localities from Mexico. In total, 101 opossum were examined. Most of the hosts were captured using Tom-

ahawk traps under permission number FAUT.0056, Secretarıá de Medio Ambiente y Recursos Naturales, Me´xico. A few road-kills were necropsied, as well as several animals shot by local hunters. Trapped opossum were killed with an overdose of sodium pentobarbital and examined by standard procedures. Platyhelminths were relaxed with hot tap water, fixed in Bouin’s fluid for 8 hr under cover glass pressure, and then placed in vials containing 70% alcohol; they were stained with Mayer’s paracarmine, Delafield’s hematoxylin, or Gomori’s trichrome and mounted in permanent slides with Canada balsam. Acanthocephalans were placed in cold physiological saline solution to evert the proboscis. These helminths and the nematodes were fixed with hot 4% formalin and studied without the aid of a permanent mount. Voucher specimens were deposited in the Coleccioń Nacional de Helmintos (CNHE), Biology Institute, Mexico City, and in the Harold W. Manter Laboratory of Parasitology (HWML), Lincoln, Nebraska. Prevalence, mean intensity, and range of intensity follow the definitions outlined by Margolis et al. (1982). Nineteen taxa of helminths, including 5 digeneans, 1 cestode, 2 acanthocephalans, and 11 nematodes were collected from opossum from 10 states of Mexico. The most widely distributed species were: Turgida turgida (Rudolphi, 1819) (in 9 states) and Cruzia americana Mapleston, 1930, and Cruzia tentaculata (Rudolphi, 1819) (in 4 states); these 3 species reached the highest levels of prevalence and mean abundance (Table I). There were 58 new locality records. Six species are recorded in Mexico for the first time: Brachylaima virginiana Dickerson, 1930; C. americana; Didelphonema longispiculata (Hill, 1939); Didelphostrongylus hayesi Prestwood, 1976; Viannaia didelphis Travassos, 1914; and Vian-

(1–3)

12.5/6

94/20

(1–11)

(7–43)

56/22.4 (4–70) 62/28.3 (5–31)

19/1.6

Colima (n 5 16) %/MI (Range)

17/8 (8)

Nayarit (n 5 6) %/MI (Range)

14/1.3 (1–2)

Oaxaca (n 5 22) %/MI (Range)

14/2.5 (1–4)

14/1.5 (1–2) 7/5 (5) 7/1 (1)

7/1 (1) 93/19.8 (4–32)

33/6 (6) 100/7 (3–11) 28/8.5 (3–14)

82/16.3 (2–54)

(6–94)

68.1/22 33/12 (12)

Michoacań (n 5 7) %/MI (Range)

7/14 (14) 21/13.3 (1–23)

50/10 (8–12)

Me´xico (n 5 4) %/MI (Range)

14/8.5 (8–9)

(1)

Jalisco (n 5 3) %/MI (Range)

32/1.6 (1–3) — 4.5/0.4 (1)

7.1/1

Guerrero (n 5 14) %/MI (Range)

* n, sample size; %, prevalence; MI, mean intensity; (Range), range of infection.

Brachylaima virginiana Brachylaima sp. Paragonimus mexicanus Rhopalias coronatus Rhopalias macracanthus Mathevotaenia sp. 14/3 (3) Oligacanthorhynchus tortuosa Oncicola luehei Didelphostrongylus hayesi Cruzia sp. Cruzia americana Cruzia tentaculata 57/36.5 (7–60) Gnathostoma turgidum Turgida turgida Didelphonema longispiculata Trichuris sp. Viannaia sp. Viannaia didelphis Viannaia viannai

Species

Chiapas (n 5 7) %/MI (Range)

Geographic distribution

TABLE I. Helminths collected from 101 opossum Didelphis virginiana in 10 states of Mexico.*

5/7

65/6

(7)

(5–13)

79/168 (3–315)

35/7.6 (6–27) 20/7 (4–10)

Veracruz (n 5 20) %/MI (Range)

50/8 (8)

50/22 (22)

Yucatań (n 5 2) %/MI (Range)

214 THE JOURNAL OF PARASITOLOGY, VOL. 91, NO. 1, FEBRUARY 2005

RESEARCH NOTES

215

TABLE II. Helminthological record of the Virginia opossum Didelphis virginiana in Mexico.

Helminth Digenea Brachylaemidae Brachylaima virginiana* Brachylaima sp.*

Paragonimidae Paragonimus mexicanus†

Rhopaliasidae Rhopalias coronatus*

Rhopalias macracanthus*

Strigeidae Duboisiella proloba* Cestoda Anoplocephalidae Anoplocephalidae gen. sp.* Mathevotaenia sp.* Proteocephalidae Proteocephalinae gen. sp.*

Acanthocephala Oligacanthorhynchidae Oligacanthorhynchus tortuosa*

Locality

Estado de Me´xico Tequesquinahuac** Guerrero Laguna Tres Palos** Zacatula** Colima Coahuayana** Comala La Barragana La Esperanza Madrid Michoacań Agua Blanca Nayarit El Guayabito El Mamey Veracruz Laguna Escondida Los Tuxtlas

Georeference

CNHE Coll. No.

HWML Coll. No.

Reference

198329300N, 998129020W

3687

This study

168499220N, 998439500W 188009380N, 1028109420W

4658 4657

This study This study

188449320N, 198199370N, 198239040N, 198109200N, 198059110N,

4650

This study Lamothe et Lamothe et Lamothe et Lamothe et

1038429170W 1038459310W 1038399000W 1038529000W 1038529190W

al. al. al. al.

(1981) (1981) (1981) (1981)

198289520N, 1008299430W

Lamothe (1981)

218329000N, 1058069300W 218269150N, 1058109100W

Lamothe et al. (1986) Lamothe et al. (1986)

188359380N, 958059160W 188379170N, 958059350W

Can˜eda (1997) Lamothe et al. (1997)

Veracruz Alvarado** Ejido La´zaro Ca´rdenas Laguna Escondida Los Tuxtlas Los Tuxtlas Veracruz Alvarado** Ejido La´zaro Ca´rdenas Laguna Escondida Los Tuxtlas Los Tuxtlas Playa Escondida Playa Escondida Oaxaca Temascal**

188169010N, 968249390W

Veracruz Playa Escondida

188359120N, 958039470W

188479050N, 188369100N, 188359380N, 188379170N,

958450359W 958069100W 958059160W 958059350W

4620

4621 188479050N, 188369100N, 188359380N, 188379170N,

958450359W 958069100W 958059160W 958059350W

4622

This study Can˜eda (1997) Can˜eda (1997) Can˜eda (1997) This study

4624

This study Can˜eda (1997) Can˜eda (1997) Can˜eda (1997) This study Can˜eda (1997) This study

4623

This study

4625 188359120N, 958039470W

Lamothe et al. (1997)

Colima \ Chiapas Lagos de Coloń**

158509000N, 918529470W

Veracruz Laguna Escondida Los Tuxtlas

188359380N, 958059160W 188379170N, 958059350W

Can˜eda (1997) Can˜eda (1997)

178159170N, 928079300W

Prado (1993)

198289520N, 1008299430W

Prado (1993)

188539000N, 998099260W


Chiapas Cascadas Agua Azul Michoacań Agua Blanca Morelos Progreso Oaxaca Temascal**


188169010N, 968249390W

4600

4497

This study

This study

216


TABLE II. Continued

Helminth

Oncicola luehei*

Pachysentis gethi* Plagiorhynchidae Porrorchis nickoli*‡‡

Nematoda Angiostrongylidae Didelphostrongylus hayesi†

Cruziidae Cruzia sp.* Cruzia americana‡

Cruzia tentaculata‡,§

Locality Veracruz Balzapote Los Tuxtlas Playa Escondida Oaxaca Mixtequilla** Veracruz Ejido La´zaro Ca´rdenas Los Tuxtlas Playa Escondida Colima La Esperanza Chiapas Cascadas Agua Azul Cascadas Agua Azul Veracruz Balzapote Catemaco Ejido La´zaro Ca´rdenas Ejido La´zaro Ca´rdenas Las Cabanãs San Andre´s Tuxtla Sontecomapan

Guerrero Laguna Tres Palos** Taxco El Viejo** Oaxaca Temascal** Guerrero Coyuquilla** Colima Colima** Comala** Madrid** La Esperanza** Guerrero Laguna Tres Palos** La Estancia** Taxco El Viejo** Oaxaca Mixtequilla** Nizanda** Temascal** Veracruz Las Cabanãs** Chiapas Cascadas Agua Azul** Colima ‡‡ Comala** Dos Amates** Jalisco Autlań-Melaque** Juntas-Palmas** Veracruz Balzapote Ejido La´zaro Ca´rdenas Laguna Escondida Los Tuxtlas Playa Escondida

Georeference

CNHE Coll. No.

HWML Coll. No.

188369390N, 958058020W 188379170N, 958059350W 188359120N, 958039470W

Reference Can˜eda (1997) Prado (1993) Can˜eda (1997)

168329000N, 948349000W

45821

This study

188369100N, 958069100W 188379170N, 958059350W 188359120N, 958039470W

Can˜eda (1997) Prado (1993) Can˜eda (1997)

198109200N, 1038529000W


178159170N, 928079300W

Salgado & Cruz (2002) Prado (1993) Salgado & Cruz (2002) Salgado & Cruz (2002) Salgado & Cruz (2002); Can˜eda (1997) Prado (1993) Salgado & Cruz (2002) Salgado & Cruz (2002)

188369390N, 958059020W 188249050N, 958069340W 188369100N, 958069100W 188369000N, 958039150W 188279000N, 958129430W 188319180N, 958019380W

168499220N, 998439500W 198289380N, 998359120W

4206 4270


188169010N, 968249390W

4271

This study

188009330N, 1028019450W

4596

This study

198139450N, 198199370N, 198059110N, 198109200N,

4593

1038439100W 1038459310W 1038529190W 1038529000W

168499220N, 998439500W — 198289380N, 998359120W 168320000N, 948349000W 168399300N, 958009250W 188169010N, 968249390W

45802 45804 45807 4204 45803 4537

This This This This

study study study study

This study This study This study

45810


188369000N, 958030150W

45811

This study

178159170N, 928079300W

45812

This study

198199370N, 1038459310W 198199500N, 1038459300W

45813 45814

Lamothe et al. (1981) This study This study

208180009N, 1048300009W 208450009N, 1058060009W 188369390N, 188369100N, 188359380N, 188379170N, 188359120N,

958058020W 958069100W 958059160W 958059350W 958039470W

4594 4590

ND ND

45815

This study This study Can˜eda (1997) Can˜eda (1997) Can˜eda (1997) Can˜eda (1997) This study

RESEARCH NOTES

217

TABLE II. Continued

Helminth

Locality Rancho Tebanca Yucatań Me´rida**

Gnathostomatidae Gnathostoma sp.\ Gnathostoma procyonis\

Gnathostoma turgidum\,#

Gongylonematidae Gongylonema mexicanum¶

Physalopteridae Turgida turgida‡,\

Spirocercidae Didelphonema longispiculata\ Trichuridae Trichuris sp.‡

Tabasco Villahermosa Morelos Cuernavaca Tepoztlań Guerrero Laguna Tres Palos** Oaxaca Temascal Temascal Veracruz Balzapote Ejido La´zaro Ca´rdenas Laguna Escondida Colima Colima** Comala** Dos Amates** La Barragana** La Esperanza** ‡ Madrid** Estado de Me´xico Tequesquinahuac** Guerrero Coyuquilla** Laguna Tres Palos** Taxco El Viejo** Jalisco Autlań-Melaque** Michoacań El Ortigal** Nayarit Pen˜itas** Oaxaca Mixtequilla** Nizanda** Temascal** Veracruz Ejido La´zaro Ca´rdenas Laguna Escondida Las Cabanãs** Los Tuxtlas** Playa Escondida Playa Escondida Yucatań Me´rida**

Georeference

CNHE Coll. No.

HWML Coll. No.

Reference Can˜eda (1997) Can˜eda (1997)

188249570N, 958009170W 208589520N, 898369360W

45816

This study

188009280N, 928559580W

Lamothe (1997)

188559150N, 998149330W 188599030N, 998079000W

Lamothe (1997) Lamothe (1997)

168499220N, 998439500W

4261

This study

188169010N, 968249390W

Lamothe et al. (1998) Almeyda et al. (2000)

188369390N, 958058020W 188369100N, 958069100W 188359380N, 958059160W

Can˜eda (1997) Can˜eda (1997) Can˜eda (1997)

198139450N, 198199370N, 198199500N, 198239040N, 198109200N,

1038439100W 1038459310W 1038459300W 1038399000W 1038529000W

4589 3421 3419 3420 4573–4574

198059110N, 1038529190W

4575

This study This study This study This study This study Lamothe et al. (1981) This study

198329300N, 998129020W

4585

This study

188009330N, 1028019450W 168499220N, 998439500W 198289380N, 998359120W

4586 4208 4541


208450009N, 1058060009W

45817

This study

198269300N, 1028039550W

4582

This study

218029000N, 1058159000W

4592

This study

168320009N, 948340009W 168399300N, 958009250W 188169010N, 968249390W

4591 5037

188369100N, 188359380N, 188369000N, 188379170N, 188359120N,

958069100W 958059160W 958030150W 958059350W 958039470W

45818

4584 45819 4580

208589520N, 898369360W

45820

This study This study This study Can˜eda (1997) Can˜eda (1997) This study This study Can˜eda (1997) This study This study

Guerrero Taxco El Viejo**

198289380N, 998359120W

4538

This study

Guerrero Taxco El Viejo**

198289380N, 998359120W

4540

This study

218


TABLE II. Continued

Helminth Trichuris didelphis†,§

Viannaiidae Viannaia sp.*

Viannaia didelphis* Viannaia viannai*

Locality Veracruz Laguna Escondida Los Tuxtlas Rancho Tebanca Guerrero Laguna Tres Palos** Veracruz Balzapote Los Tuxtlas** Playa Escondida Colima La Esperanza** Madrid** Guerrero Taxco El Viejo**

Georeference

CNHE Coll. No.

188359380N, 958059160W 188379170N, 958059350W 188249570N, 958009170W

HWML Coll. No.

Reference Can˜eda (1997) Can˜eda (1997) Can˜eda (1997)

168499220N, 998439500W

4205

This study

188369390N, 958058020W 188379170N, 958059350W 188359120N, 958039470W

4652

Can˜eda (1997) This study Can˜eda (1997)

198109200N, 1038529000W 198059110N, 1038529190W

4651 4649


198289380N, 998359120W

4543

This study

* Intestine. ** New locality record. † Lungs. ‡‡ Locality not specified. ‡‡ Recorded by Prado (1993) and Can˜eda (1997) as Longisoma marsupialis Prado, 1993, specific name only published in a Bachelor thesis. CNHE, Accession number to Coleccioń Nacional de Helmintos; HWML, Accession number to Harold W. Manter Laboratory of Parasitology; ND, voucher specimens not deposited. ‡ Caecum. \ Stomach. # Liver. ¶ Esophagus. § Rectum.

naia viannai Travassos, 1914, increasing the number of helminth taxa known for this host in Mexico to 28 (Table II). It was not possible to identify 5 of the 19 helminth taxa found in this study because of several reasons, i.e., the poor conditions of the material (Cruzia sp., Brachylaima sp.), the small number of specimens collected (Viannaia sp.), or immaturity (Mathevotaenia sp.). Trichuris sp. is similar in some ways to the 5 species described in American marsupials, i.e., Trichuris minuta Rudolphi, 1819; Trichuris marsupialis Foster, 1939; Trichuris urichi Wolfgang, 1951; Trichuris reesali Wolfgang, 1951; and especially Trichuris didelphis Babero, 1960. However, the analysis of paratypes of this species (USNPC 039038) showed that our specimens could not be assigned to T. didelphis because of the shape of the spicular sheath (tubular in our material vs. pear shaped in Babero’s [1960] specimens) and the egg size (0.062–0.069 3 0.030–0.031 vs. 0.070–0.108 3 0.035–0.045, respectively). Records of Paragonimus rudis (Diesing, 1850) from Mexico, attributed by Alden (1995) to Lamothe (1981) and Lamothe et al. (1986), are erroneous. In both records, the specimens were assigned originally to Paragonimus mexicanus Miyazaki & Ishii, 1968. Moreover, P. rudis is now considered as species inquirenda (Tongu et al., 1995). The Mexican helminthological record for D. virginiana, constituted by 28 taxa (10 recorded in this study and 18 reported previously), reflects the same trend as the one reported by Alden (1995); thus, nematodes are the most prevalent group of helminths in both studies (15 and 38 taxa, respectively). Digeneans are the second most prevalent group represented in both regions (22 taxa in Alden’s study vs. 6 from Mexico); in North America, the other 2 groups (cestodes and acanthocephalans) comprise 9 and 7 taxa, respectively, whereas in Mexico these possess 3 and 4 taxa. The helminth parasites recorded in this host from Mexico add 7 taxa to the helminth fauna of D. virginiana in North America: Duboisiella proloba Baer, 1938 and Rhopalias coronatus (Rudolphi, 1819) (Digenea); Mathevotaenia sp., (Cestoda); Porrorchis nickoli Salgado and Cruz, 2002, Oncicola luehei (Travassos, 1917), and Pachysentis gethi (Machado, 1950) (Acanthocephala); and Gongylonema mexicanum (Caballero and Zerecero, 1944) (Nematoda) (Table II). In addition, 6 helminth species have been registered in several localities from Mexico

from hosts identified as Didelphis mesamericana tabascensis (Allen, 1901): Aspiododera raillieti Travassos, 1913, C. tentaculata, Globocephalus marsupialis (Teixeira de Freitas and Lent, 1936), R. coronatus, Rhopalias macracanthus Chandler, 1932, and T. turgida (Caballero, 1937, 1951; Caballero and Zerecero, 1944; Caballero et al., 1944). Because the name D. mesamericana tabascensis has been used indistinctly for Didelphis marsupialis Linnaeus and D. virginiana (see Gardner, 1973) and because there are no host voucher specimens, these records cannot be considered as part of Mexican helminth fauna of D. virginiana. Of the 28 taxa reported in Mexico, 20 were identified to specific level. The distribution of this 20 species includes 7 Pan-American species (R. macracanthus; D. hayesi; Gnathostoma turgidum Stossich, 1902; T. turgida; Trichuris didelphis; V. didelphis; and V. viannai), 6 from South America (P. mexicanus; R. coronatus; D. proloba; O. luehei; P. gethi; and C. tentaculata), 5 from North America (B. virginiana; Oligacanthorhynchus tortuosa (Leidy, 1850); C. americana; Gnathostoma procyonis Chandler, 1942; and D. longispiculata), and 2 endemic species (P. nickoli and G. mexicanum). This mixed composition can be explained on one hand by the confluence of the Nearctic and Neotropical biogeographical zones in Mexico. On the other hand, the generalist character of all 20 helminth species could promote their exchange among different hosts (other marsupials, felids, mustelids, and procyonids, indiscriminately), thereby enriching the helminth fauna of the Virginia opossum in Mexico. However, scarce knowledge regarding the helminth fauna of D. virginiana in Mexico makes both hypotheses premature. Further studies are necessary to make more accurate generalizations. The authors thank Florencia Bertoni, Elisa Cabrera, Cristina Castilla, Luis F. Gopar, and Rogelio Rosas for their assistance in field work, Georgina Ortega for bibliographic assistance, and Antonieta Arizmendi, Agustin Jimeńez, Rafael Lamothe, and Berenit Mendoza for their helpful comments and corrections that improved the manuscript. This study was partially funded by grant IN204600-3 under the Programa de Apoyo a Proyectos de Investigacioń e Innovacioń Tecnolo´gica (PAPIITUNAM) to Virginia Leoń from Instituto de Biologıá, UNAM, and BSR 8612329, BSR 9024816, DEB 9496263, and DBI 10097019 to Scott

RESEARCH NOTES

Gardner from H. W. Manter Laboratory of Parasitology, University of Nebraska, Lincoln. LITERATURE CITED ALDEN, K. J. 1995. Helminths of the opossum, Didelphis virginiana, in Southern Illinois, with a compilation of all helminths reported from this host in North America. Journal of the Helminthological Society of Washington 62: 197–208. ALMEYDA, A. J., M. D. BARGUES, AND S. MAS-COMA. 2000. ITS-2 rDNA sequencing of Gnathostoma species (Nematoda) and elucidation of the species causing human Gnathostomiasis in the Americas. Journal of Parasitology 86: 537–544. BABERO, B. B. 1960. Further studies on helminths of the opossum, Didelphis virginiana, with a description of a new species from this host. Journal of Parasitology 46: 455–463. CABALLERO, E. C. 1937. Nema´todos de algunos vertebrados del Valle del Mezquital, Hidalgo. Anales del Instituto de Biologıá, Universidad Nacional Autońoma de Me´xico 8: 189–200. ———. 1951. Estudios helmintolo´gicos de la Regioń Oncocercosa de Me´xico y de la Repu´blica Mexicana. Nematoda 6a Parte. Y algunas consideraciones en torno a los geńeros Onchocerca Diesing, 1841 y Acanthospiculum Skjrabin y Schikhobalowa, 1948. Anales del Instituto de Biologıá, Universidad Nacional Autońoma de Me´xico 22: 141–158. ———, M. BRAVO, AND C. ZERECERO. 1944. Estudios helmintolo´gicos de la Regioń Oncocercosa de Me´xico y de la Repu´blica de Guatemala. Trematoda I. Anales del Instituto de Biologıá, Universidad Nacional Autońoma de Me´xico 15: 59–72. ———, AND M. C. ZERECERO. 1944. Estudios de la Regioń Oncocercosa de Me´xico y de la Repu´blica de Guatemala. Nema´toda. 2a Parte. Anales del Instituto de Biologıá, Universidad Nacional Autońoma de Me´xico 15: 389–407. CAN˜EDA, G. C. 1997. Para´sitos de tres especies de marsupiales de la Estacioń ‘‘Los Tuxtlas’’ y algunas zonas cercanas, Veracruz, Me´xico. B.S. Thesis. Universidad Nacional Autońoma de Me´xico, Mexico City, D.F., 193 p. GARDNER, A. L. 1973. The systematics of the genus Didelphis (Marsupialia: Didelphidae) in North and Middle America. Special Publications the Museum Texas Tech University 4: 1–81. ———. 1982. Virginia opossum (Didelphis virginiana). In Wild mammals of North America, J. A. Chapman and G. A. Feldhamer (eds.). John Hopkins University Press, Baltimore, Maryland, p. 3–36. HARRIS, E. A. 1983. A checklist of the helminths parasites of marsupials

219

and monotrems. British Museum (Natural History), London, U.K., 170 p. LAMOTHE, A. R. 1981. Hospederos definitivos e intermediarios de Paragonimus mexicanus, Miyazaki e Ishii, 1968, en Me´xico. Anales del Instituto de Biologıá, Universidad Nacional Autońoma de Me´xico 52: 39–44. ———. 1997. Hospederos definitivos, intermediarios y parateńicos de Gnathostoma en Veracruz y Oaxaca, Me´xico. Cuadernos Mexicanos de Zoologıá 3: 22–28. ———, H. AKAHANE, D. OSORIO, AND L. GARC´ıA. 1998. Hallazgo de Gnathostoma turgidum en Didelphis virginiana de Temascal, Oaxaca, Me´xico. Anales del Instituto de Biologıá, Universidad Nacional Autońoma de Me´xico 69: 225–229. ———, J. L. ALONSO, AND R. LO´PEZ. 1986. Una nueva zona ende´mica de Paragonimiasis en Me´xico. Anales del Instituto de Biologıá, Universidad Nacional Autońoma de Me´xico 57: 415–418. ———, L. GARC´ıA, D. OSORIO, AND G. PE´REZ-PONCE DE LEOŃ. 1997. Cata´logo de la Coleccioń Nacional de Helmintos, 1st ed. Instituto de Biologıá, Universidad Nacional Autońoma de Me´xico and Comisioń Nacional para el Conocimiento y Uso de la Biodiversidad, Mexico City, D.F., 211 p. ———, R. PINEDA, AND O. MEAVE. 1981. Infeccioń natural de Paragonimus mexicanus en Didelphis virginiana californica en Colima, Me´xico. Anales del Instituto de Biologıá, Universidad Nacional Autońoma de Me´xico 52: 45–50. MARGOLIS, L., G. W. ESCH, J. C. HOLMES, A. M. KURIS, AND G. A. SCHAD. 1982. The use of ecological terms in parasitology (report of an ad hoc committee of the American Society of Parasitologists). Journal of Parasitology 68: 131–133. POTKAY, S. 1977. Diseases of marsupials. In The biology of marsupials, D. Hunsaker II (ed.). Academic Press, New York, p. 415–506. PRADO, A. D. 1993. Estudio taxono´mico de 10 especies de acantoce´falos (Acanthocephala Rudolphi, 1801) de vertebrados de Me´xico. B.S. Thesis. Universidad Nacional Autońoma de Me´xico, Mexico City, D.F., 86 p. SALGADO, G., AND A. CRUZ. 2002. Porrorchis nickoli n. sp. (Acanthocephala: Plagiorhynchidae) from mammals in Southeastern Mexico, first known occurrence of Porrorchis in the Western Hemisphere. Journal of Parasitology 88: 146–152. TONGU, Y., H. HATA, Y. ORIDO, M. PINTO, R. LAMOTHE, M. YOKOGAWA, AND M. TSU. 1995. Morphological observations of Paragonimus mexicanus from Guatemala. Japanese Journal of Parasitology 44: 365–370.


Effects of Heparin Administration on Trypanosoma brucei gambiense Infection in Rats Kazuhiko Nishimura, Kensuke Shima, Masahiro Asakura, Yoshihiro Ohnishi, and Shinji Yamasaki, Division of Veterinary Science, Graduate School of Agriculture and Biological Sciences, Osaka Prefecture University, 1-1, Gakuencho, Sakai, Osaka 599-8531, Japan. e-mail: [email protected] ABSTRACT: We examined whether heparin administration influences in vivo trypanosome proliferation in infected rats. Administration of heparin every 8 hr via cardiac catheter inhibited growth of Trypanosoma brucei gambiense and prolonged survival of treated rats. Heparin administration increased lipoprotein lipase activity, high-density lipoprotein (HDL) concentration in the blood, and haptoglobin messenger RNA content of the liver. The presence of heparin in culture media did not directly affect proliferation of trypanosomes in vitro. However, the addition of plasma from infected rats treated with heparin to culture media decreased the number of trypanosomes. This effect was decreased by incubating the trypanosomes with benzyl alcohol, a known inhibitor of receptor-mediated endocytosis of lipoprotein. These data suggested that heparin administration reduced the number of trypanosomes in infected rats. Trypanosome lytic factor, a HDL and haptoglobin-related protein,

protects humans and some animals from infection by Trypanosoma brucei brucei. In rats, increases in HDL and haptoglobin may affect the proliferation of T. b. gambiense. Trypanosome lytic factor, including components of high-density lipoprotein (HDL), haptoglobin-related protein, apolipoprotein L-1, etc., protects humans and certain animals from Trypanosoma brucei brucei infection (Ortiz-Ordonez et al., 1994; Smith and Hajduk, 1995; Hager and Hajduk, 1997; Drain et al., 2001; Vanhamme et al., 2003). Other animals produce xanthine oxidase, which also exhibits trypanocidal effects (Muranjan et al., 1997; Wang et al., 2002). However, it is unclear why animals, such as rats, which have similar components, are not resistant to trypanosome infection. In addition to its anticoagulant properties, heparin exhibits various

220


physiological activities and influences a wide range of components in the blood (Swaney and Orishimo, 1989; Goldberg and Chajek-Shaul, 1990; O’Meara et al., 1994; Tornvall et al., 1995). Heparin activates lipoprotein lipase (LPL) in vascular endothelial cells and influences HDL composition (Swaney and Orishimo, 1989; Goldberg and ChajekShaul, 1990; O’Meara et al., 1994; Tornvall et al., 1995). In this study, we examined whether heparin treatment of trypanosome-infected rats influences trypanosome proliferation in vivo. Experiments were performed on 10-wk-old male Wistar rats (Nippon SLC Co., Shizuoka, Japan). Cardiac catheterization was completed 3 days before experimental infection with Trypanosoma brucei gambiense Wellcome strain (WS). This strain is noncloned, old monomorphic laboratory trypanosome. Trypanosomes were subcutaneously inoculated into rats at 1 3 106 cells per rat; viability, as determined by light microscopy, was greater than 95%. After infection, blood was collected from the catheter at 0600 hr daily. Heparin was dissolved in 0.4 ml saline and administered via the catheter after blood collection and at 1400 and 2200 hr. Control rats were administered pure saline. The number of trypanosomes in the blood was determined by light microscopy using a hemocytometer. Seven days postinfection (PI), blood and liver were collected. Blood was collected from the abdominal aorta using a syringe containing heparin. Blood samples were immediately cooled to 4 C, and serum was prepared by centrifugation at 900 g for 10 min. Liver samples were immediately frozen in liquid nitrogen. HDL cholesterol in the serum was measured as described by Burstein et al. (1989). LPL activity in the serum was determined with the Lipoprotein Lipase Activity Kit (Roar Biomedical, Inc., New York, New York). Primers for reverse transcription–polymerase chain reaction (RTPCR) were obtained from SIGMA Genosys Japan Co. (Ishikari, Japan). We used the primer pair 59-AAGATGGTGAAGGTCGGT-39 and 59GAAGGGGCGGAGATGATG-39 for rat glyceraldehyde phosphate dehydrogenase (GAPDH) messenger RNA (mRNA), positions 28–381 based on the report of Tajima et al. (1999), and 59-CTCACCACTGGGGCCAC-39 and 59-CATTCCGCGCCCCCAAC-39 for rat haptoglobin mRNA, positions 376–669 based on the report of Lee et al. (2002). RNA was extracted from the liver using the Gen Elute Mammalian Total RNA Kit (Sigma Chemical Co., St. Louis, Missouri). Levels of haptoglobin mRNA were determined by semiquantitative RT-PCR using the SuperSCRIPT One-Step RT-PCR with PLATINUM Taq (Invitrogen Corp., Carlsbad, California). The housekeeping gene, GAPDH, was used as a control. Each sample was serially diluted before amplification so that amplification of GAPDH was equivalent among samples under conditions where samples were in the exponential phase of am-

FIGURE 1. Abundance of WS trypanosomes in the blood. Trypanosomes were injected on day 0. Heparin was administrated every 8 hr from day 0. Each value and bar represents the mean 6 SD of results from 4 rats. Crosses indicate death.

plification. The relative amounts of haptoglobin mRNA were then determined by electrophoresis of samples through a 1.5% agarose gel containing ethidium bromide; fluorescence was analyzed using the gel documentation analysis system GelDoc 2000 (Bio-Rad Laboratory, Tokyo, Japan). Seven days PI, blood was collected from infected rats that were not treated with heparin for harvesting of trypanosomes. The blood was centrifuged at 900 g for 10 min. Trypanosomes were washed twice with phosphate–saline–glucose buffer, pH 8.0, consisting of 57 mM Na2HPO4, 3 mM NaH2PO4, 44 mM NaCl, and 56 mM glucose and separated by chromatography on diethylaminoethyl cellulose (DE-52; Whatman, Maidstone, U.K.) as described by Lanham (1968). The trypanosomes were resuspended in Iscove’s modified Dulbecco’s medium (IMDM; Invitrogen Corp.) for in vitro assay. Seven days after the initiation of heparin treatment, blood was collected from uninfected and infected rats 1 hr after heparin treatment. The blood was collected into nonheparinized tubes for plasma separation. Postheparin plasma was used to determine survival rate of trypanosomes in vitro. Trypanosomes were suspended to 3 3 104 cells/ml and incubated at 37 C for 24 hr with postheparin plasma, and numbers of trypanosomes were determined. Trypanosomes used in this experiment were cultured in modified IMDM containing 5% fetal bovine serum in 95% O2 and 5% CO2 at 37 C (Yabu et al., 1998; Nishimura et al., 2004). Standard laboratory chemicals and reagents were purchased from the Wako Pure Chemical Co. (Osaka, Japan). Student’s t-test was used to determine significant differences. Heparin treatment every 8 hr inhibited trypanosome proliferation, and treated rats survived longer than untreated rats (Fig. 1). Regression curve of saline treatment was log(y) 5 8.7 1 1.9x, and those of heparin treatment were for 20 U/100 g body weight (BW), log(y) 5 9.6 1 1.5x; 200 U/100 g BW, log(y) 5 8.6 1 1.1x; and 2,000 U/100 g BW, log(y) 5 9.3 1 1.1x. Maximal effects of heparin were observed at 200 U/100 g BW every 8 hr. The addition of heparin to culture media did not affect trypanosome numbers after 24 hr (Fig. 2), which suggested that heparin did not directly affect the trypanosomes. The number of trypanosomes in the case of addition of heparin to the plasma of infected rats treated with saline also did not differ. However, culture media that contained plasma from infected rats treated with heparin inhibited trypanosome proliferation. Plasma from uninfected rats treated with heparin also inhibited trypanosome proliferation (data not shown). The inhibitory effect of plasma from rats treated with heparin on proliferation was negated when the trypanosomes were incubated with benzyl alcohol, a known inhibitor of receptor-mediated endocytosis of lipoproteins

FIGURE 2. Effect of heparin and plasma from infected rats on trypanosomal abundance in vitro. Trypanosomes were suspended to 3 3 104 cells/ml and were cultured for 24 hr at 37 C with heparin and plasma isolated from infected rats 7 days PI. Each column and bar represents the mean 6 SD of results from 4 rats. Asterisks indicate significant differences from untreated samples (P , 0.05).

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FIGURE 3. Effect of heparin administration on LPL activity, HDL concentration, and haptoglobin mRNA levels. Heparin (200 U/100 g BW) was administrated every 8 hr. Blood and liver samples were harvested from rats that had been infected for 7 days PI. (A) LPL activity after no treatment. (B) Concentration of HDL cholesterol in the blood. (C) Representative RT-PCR results for haptoglobin mRNA in liver. M: fX174 HaeIII digest (20 ng/ml, 5 ml). (D) Ratio of haptoglobin mRNA expression to GAPDH-mRNA expression per sample. Each column and bar represents the mean 6 SD of results from 5 rats. Asterisks indicate significant differences from untreated rats (P , 0.05).

(Sainte-Marie et al., 1990; O’Meara et al., 1994; Tornvall et al., 1995). These data suggested that a change in plasma components in response to heparin administration was related to the decrease in the number of trypanosomes. Heparin administration increased total lipolytic activity in the serum, and this augmentation was predominantly due to LPL (Goldberg and Chajek-Shaul, 1990). We also confirmed that heparin treatment of uninfected rats increased LPL activity and serum HDL concentration (Fig. 3A, B). LPL activity of infected rats decreased upon trypanosome infection, although heparin treatment counteracted this effect (Fig. 3A). Plasma HDL cholesterol concentration increased in response to heparin treatment of infected rats (Fig. 3B). Haptoglobin mRNA levels increased in response to heparin treatment in both infected and uninfected rats (Fig. 3C, D). These results suggest that heparin treatment of infected rats prevented the decrease in LPL activity. A strong positive correlation was observed by Tornvall et al. (1995) between LPL levels in preheparin plasma and HDL cholesterol levels. In this study, we also confirmed that HDL concentration increased in response to heparin. Heparin-induced lipolysis produces atypical HDL in rats (Swaney and Orishimo, 1989) and humans (O’Meara et al., 1994). We demonstrated that plasma from rats treated with heparin inhibits trypanosome proliferation. Heparin modulates cytokine production and affects immunity (Call and Remick, 1998; Anastase-Ravion et al., 2003), and cytokines affect proliferation of trypanosomes (Shi et al., 2003; Nishimura et al., 2004). Therefore, changes in cytokine production in response to heparin may be related to the inhibition of proliferation in trypanosomes.

LITERATURE CITED ANASTASE-RAVION, S., C. BLONDIN, B. CHOLLEY, N. HAEFFNER-CAVAILLON, J. J. CASTELLOT, AND D. LETOURNEUR. 2003. Heparin inhibits lipopolysaccharide (LPS) binding to leukocytes and LPS-induced cytokine production. Journal of Biomedical Materials Research 66A: 376–384. BURSTEIN, M., A. FINE, V. ATGER, E. WIRBEL, AND A. GIRARD-GLOBA. 1989. Rapid method for the isolation of two purified subfractions of high density lipoproteins by differential dextran sulfate-magnesium chloride precipitation. Biochimie 71: 741–746. CALL, D. R., AND D. G. REMICK. 1998. Low molecular weight heparin is associated with greater cytokine production in a stimulated whole blood model. Shock 10: 192–197. DRAIN, J., J. R. BISHOP, AND S. L. HAJDUK. 2001. Haptoglobin-related protein mediates trypanosome lytic factor binding to trypanosomes. Journal of Biological Chemistry 276: 30254–30260. GOLDBERG, D. M., AND T. CHAJEK-SHAUL. 1990. Effect of chronic heparin administration on serum lipolytic activity and some aspects of lipid metabolism. Biochimica et Biophysica Acta 1047: 103–111. HAGER, K. M., AND S. L. HAJDUK. 1997. Mechanism of resistance of African trypanosomes to cytotoxic human HDL. Nature 385: 823– 826. LANHAM, S. M. 1968. Separation of trypanosomes from the blood of infected rats and mice by anion-exchangers. Nature 218: 1273– 1274. LEE, M. Y., S. Y. KIM, J. S. CHOI, I. H. LEE, Y. S. CHOI, J. Y. JIN, S. J.

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PARK, K. W. SUNG, M. H. CHUN, AND I. S. KIM. 2002. Upregulation of haptoglobin in reactive astrocytes after transient forebrain ischemia in rats. Journal of Cerebral Blood Flow and Metabolism 22: 1176–1180. MURANJAN, M., Q. WANG, Y. L. LI, E. HAMILTON, F. P. OTIENO-OMONDI, J. WANG, A. VAN PRAAGH, J. G. GROOTENHUIS, AND S. J. BLACK. 1997. The trypanocidal Cape buffalo serum protein is xanthine oxidase. Infection and Immunity 65: 3806–3814. NISHIMURA, K., K. HAMASHITA, F. KAWAHARA, H. IHARA, S. KOZAKI, Y. OHNISHI, AND S. YAMASAKI. 2004. Differential effects of interferon g on production of trypanosome-derived lymphocyte triggering factor by Trypanosoma brucei gambiense and Trypanosoma brucei brucei. Journal of Parasitology 90: 740–745. O’MEARA, N. M., V. G. CABANA, J. R. LUKENS, B. LOHARIKAR, T. M. FORTE, K. S. POLONSKY, AND G. S. GETZ. 1994. Heparin-induced lipolysis in hypertriglyceridemic subjects results in the formation of atypical HDL particles. Journal of Lipid Research 35: 2178– 2190. ORTIZ-ORDONEZ, J. C., J. B. SECHELSKI, AND J. R. SE ED. 1994. Mechanism of lysis of Trypanosoma brucei gambiense by human serum. Journal of Parasitology 80: 924–930. SAINTE-MARIE, J., M. VIGNES, M. VIDAL, J. R. PHILIPPOT, AND A. BIENVENUE. 1990. Effects of benzyl alcohol on transferrin and low density lipoprotein receptor mediated endocytosis in leukemic guinea pig B lymphocytes. FEBS Letters 262: 13–16. SHI, M., W. PAN, AND H. TABEL. 2003. Experimental African trypanosomiasis: IFN-gamma mediates early mortality. European Journal of Immunology 33: 108–118. SMITH, A. B., AND S. L. HAJDUK. 1995. Identification of haptoglobin as

a natural inhibitor of trypanocidal activity in human serum. Proceedings of the National Academy of Sciences of the United States of America 92: 10262–10266. SWANEY, J. B., AND M. W. ORISHIMO. 1989. Effects of heparin-induced lipolytic activity on the structure of rat high-density lipoprotein. Biochimica et Biophysica Acta 1002: 338–347. TAJIMA, H., K. TSUCHIYA, M. YAMADA, K. KONDO, N. KATSUBE, AND R. ISHITANI. 1999. Over-expression of GAPDH induces apoptosis in COS-7 cells transfected with cloned GAPDH cDNAs. Neuroreport 10: 2029–2033. TORNVALL, P., G. OLIVECRONA, F. KARPE, A. HAMSTEN, AND T. OLIVECRONA. 1995. Lipoprotein lipase mass and activity in plasma and their increase after heparin are separate parameters with different relations to plasma lipoproteins. Arteriosclerosis Thrombosis and Vascular Biology 15: 1086–1093. VANHAMME, L., F. PATURIAUX-HANOCQ, P. POELVOORDE, D. P. NOLAN, L. LINS, J. VAN DEN ABBEELE, A. PAYS, P. TEBABI, H. VAN XONG, A. JACQUET, N. MOGUILEVSKY, M. DIEU, J. P. KANE, P. DE BAETSELIER, R. BRASSEUR, AND E. PAYS. 2003. Apolipoprotein L-I is the trypanosome lytic factor of human serum. Nature 422: 83–87. WANG, J., A. VAN PRAAGH, E. HAMILTON, Q. WANG, B. ZOU, M. MURANJAN, N. B. MURPHY, AND S. J. BLACK. 2002. Serum xanthine oxidase: Origin, regulation, and contribution to control of trypanosome parasitemia. Antioxidants and Redox Signaling 4: 161–178. YABU, Y., T. KOIDE, N. OHTA, M. NOSE, AND Y. OGIHARA. 1998. Continuous growth of bloodstream forms of Trypanosoma brucei in an axenic culture system containing a low concentration of serum. Southeast Asian Journal of Tropical Medicine and Public Health 223: 591–595.


Development of Neospora caninum Cultured with Human Serum In Vitro and In Vivo Y. Omata, R. Kano, Y. Masukata, Y. Kobayashi*, M. Igarashi, R. Maeda, and A. Saito, Laboratory of Veterinary Physiology, Obihiro University of Agriculture and Veterinary Medicine, Obihiro 080-8555, Japan; *Department of Veterinary Pathology, Obihiro University of Agriculture and Veterinary Medicine, Obihiro 080-8555, Japan. e-mail: [email protected] ABSTRACT: Because there has been no report of symptomatic Neospora caninum infection in humans, we examined the effect of human serum on the parasite’s growth in either a bovine angioendothelial cell or Caco-2 cell culture in vitro and in immunocompromised mice in vivo. There was no difference in intracellular parasite numbers between cells incubated with human serum at 24 hr after challenge and those incubated with fetal bovine serum (FBS), which has no titer for the anti–N. caninum agglutination antibody test. Serum of sheep infected with N. caninum, which has the anti–N. caninum antibody, reduced the numbers of the intracellular parasite significantly. We also showed that there was no inhibitory effect on the intracellular multiplication of the parasite in cells incubated with human serum through incorporation of 3H-uracil. CB-17 scid mice administered human serum daily and challenged with N. caninum died on day 20 or 22 after challenge, when large numbers of parasite clusters were found in the brain, oviduct, adrenal gland, lung, stomach, spleen, skeletal muscle, pancreas, and mesenteric lymph nodes. Scid mice administered FBS survived until the end of the experiment. These results suggest that adult human serum may have no inhibitory effect on the development of N. caninum in vitro and in vivo.

Neospora caninum, a Toxoplasma-like apicomplexan protozoan parasite, causes severe neuromuscular disease and repeated abortion through transplacental transmission in livestock and companion animals. This parasite can penetrate many kinds of mammalian host cells, and it has been shown to be capable of infecting a wide range of species. After being orally infected, dogs (the definitive host) shed oocysts in the feces, and this is recognized as 1 source of infection (McAllister et al., 1998; Lindsay et al., 1999). The other major means of infection is transplacental transmission. Oral infection may occur through the in-

gestion of raw meat contaminated with the parasite or its sporulated oocysts. Although serological evidence suggests that humans are potentially susceptible to N. caninum infection (Tranas et al., 1999), the numbers of serologically positive persons in the population seems to be very low, and there has been no report of symptomatic human infection (Graham et al., 1999). Therefore, it is still not known whether N. caninum has zoonotic potential (Dubey and Lindsay, 1996). Sundermann and Estridge (1999) showed that N. caninum tachyzoites can penetrate human-foreskin cells, suggesting that the parasite has the potential to infect humans. We supposed that if humans have natural resistance against N. caninum, the mechanism for this would involve inhibition of the process by which the parasite penetrates the host cells. It has been reported that antimicrobial peptides, such as lactoferrin, which has an inhibitory effect on toxoplasmosis, are present in body fluids (Tanaka et al., 1997). However, it is still not known whether the various types of antimicrobial peptide in human body fluids have any effect on N. caninum, and to our knowledge, no study has been done on the effect of human serum on the development of N. caninum in vitro or in vivo. To determine whether human serum has an inhibitory effect on either the invasion of host cells or the growth of N. caninum, human serum was added to a cell culture medium infected with N. caninum. In 1 experiment, we used Caco-2 cells from a human colonic adenocarcinoma, on the basis of the assumptions that oral ingestion is 1 of the infection routes and intestinal epithelial cells would be exposed to the parasite. We also examined the effect of inoculating scid mice with human serum. The results of this study indicated that adult human serum may have no inhibitory effect on the growth of N caninum in host cells and suggested that the zoonotic potential of N. caninum cannot be ignored.

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FIGURE 2. Incorporation of 3H-uracil into BAE cells at 42 hr after inoculation, with or without Neospora caninum, expressed as mean growth index (count per minute [cpm] test/cpm control) of 3 samples.

FIGURE 1. Effect of human serum on infectivity of Neospora caninum with respect to BAE or Caco-2 cells. Infection rates are expressed as numbers of N. caninum per 100 cells, calculated from their numbers in a total of 400 cells. Mean 6 SD of triplicate samples. Medium, culture medium; Nor. shp, noninfected sheep serum; Inf. shp, infected sheep serum; BS, bovine serum; Human, human serum; FBS, fetal bovine serum. (a) all sera 10%; (b) FBS 10%, human 10 and 40%.

Five-week-old female CB-17 scid mice were purchased from Japan CLEA (Tokyo, Japan) and housed in a microisolator. Bovine angioendothelial cells (BAE cells) and Caco-2 cells were grown in Dulbecco modified Eagle medium (D-MEM) containing 10% fetal bovine serum (FBS) (D-MEM10FBS) and nonessential amino acids (ICN Biomedicals. Inc., Aurora, Ohio). Tachyzoites of N. caninum were isolated from sheep (Kobayashi et al., 2001; Koyama et al., 2001) and maintained by continuous passage in BAE cell cultures. They were then collected from the cell cultures and suspended in D-MEM10FBS. To remove the host cells and debris, the suspension was passed through a 3-mm polycarbonate filter (Nucle-

pore; Corning Costa Corporation, Tokyo, Japan), and then the parasite suspension was adjusted to a concentration of 1 3 106 parasites/ml with D-MEM10FBS. Heat-inactivated adult bovine serum (BS) and FBS were purchased from Biofluids International Inc. (Rockville, Maryland). Human serum samples were collected from 10 healthy adult volunteers, inactivated by heat treatment at 56 C for 30 min, and then pooled and stored at 280 C until use. The antibody activity against N. caninum in the serum was examined by Neospora agglutination test (NAT) according to the method described by Lindsay et al. (2002), using 2 3 107/ml N. caninum tachyzoites fixed with phosphate-buffered saline (PBS) containing 4% paraformaldehyde as the antigen. For the control, sheep serum was obtained before inoculation with 106 N. caninum tachyzoites and on day 49 after inoculation and was inactivated as mentioned above. The sera tested were diluted to a concentration of either 10 or 40% with D-MEM10FBS and then mixed with an equal volume of the N. caninum suspension. Next, either BAE cell or Caco-2 cell monolayers were placed on 15-mm–diameter, round coverslips, challenged with 0.2 ml of the parasite–serum mixture, and then incubated at 37 C. At 24 hr after challenge, the coverslips were thoroughly washed with PBS, fixed with methanol, and stained with Giemsa. The number of intracellular N. caninum was determined in a total of 400 cells and indicated as a number of the parasite per 100 cells. All experiments were performed in triplicate and repeated at least twice. Data from each experiment were evaluated using the Student’s t-test. The level of significance in all analysis was 95%. To examine the intracellular growth of N. caninum in the BAE cell culture, we compared 3H-uracil–incorporation rates between parasiteinfected cells cultured either with human serum or FBS, according to the method described by Innes et al. (1995). Briefly, 2 ml of either diluted serum was mixed with the same volume of the N. caninum suspension, 1 ml of the mixture was added to BAE cells on 3-cmdiameter petri dishes, and this was followed by incubation at 37 C in a CO2 incubator. After 18 hr of incubation, 3H-uracil (Amersham, Tokyo, Japan) was added at 0.37 MBq/ml/well, and incubation was continued for a further 24 hr. Afterward, the BAE cells were dissolved in 100 ml of 1 N NaOH, and then, 1 ml of 20% trichloroacetic acid was added. The precipitates were collected on glass filters, and radioactivity was measured with a b-scintillation counter.

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FIGURE 3. Photomicrographs of scid mice tissues from each group on day 22 after challenge. Neospora caninum clusters were associated with severe degeneration and necrosis of the parenchymal cells and inflammatory reactions (arrow). There were also many necrosis areas and degradation foci. (A) brain; (B) oviduct; (C) lung; (D) skeletal muscle of scid mice administered human serum; (E) brain; (F) oviduct; (G) lung; and (H) skeletal muscle of scid mice administered FBS. Hematoxylin and eosin staining. Bar 5 100 mm.

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The effect of human serum on N. caninum was also examined in an inoculation test in 2 groups of scid mice,1 of them receiving human serum and the other FBS. Two mice in each group were preinoculated intraperitoneally with 0.5 ml/day of either human serum or FBS for 7 days and then challenged with 2 3 105 N. caninum. The survival time and mortality of the mice in each group were monitored until all mice in 1 group were dead. Organ tissue samples from the mice that died during the monitoring period were examined microscopically for the presence of N. caninum. Also, surviving mice at the end of the period were examined for pathological changes in their organs. At necropsy of the scid mice, representative tissue samples (liver, spleen, kidney, heart, lung, alimentary tract, pancreas, adrenal gland, genital organ, peritoneum, skeletal muscle, brain, and spinal cord) were collected and fixed in 10% buffered formalin. These tissues were processed and embedded in paraffin wax by routine procedures. Sections were stained with hematoxylin and eosin. The human serum, FBS, BS, and sheep serum obtained before infection showed no antibody activity (NAT titer ,1:10), whereas serum obtained after infection did show antibody activity (NAT titer 1:160). As shown in Figure 1a, after 24 hr, the number of N. caninum in BAE cells incubated with human serum at a concentration of 10% was no different from that in cells incubated with 10% FBS, BS, or sheep serum obtained before infection. For incubation with BS, the number of intracellular tachyzoites was somewhat lower than for incubation with 10% human serum and FBS, and the number obtained with 10% infected sheep serum was significantly lower. The number of N. caninum in Caco-2 cells was somewhat lower than in BAE cells, but there was no significant difference among the sera, except for the infected sheep serum. BAE cells were therefore used for the subsequent experiments. For human serum at a concentration of 40%, the extent of N. caninum invasion was similar to that for FBS (Fig. 1b). Figure 2 shows the results for examination of N. caninum growth using 3H-uracil–incorporation rates, for both infected and noninfected cell cultures incubated with human serum. There was no significant difference in 3H-uracil–incorporation rates between infected cells cultured with human serum and those cultured with FBS. However, the levels of incorporation for human serum tended to be higher than those for FBS. In the in vivo experiments using scid mice, human serum was observed to have no protective effect against N. caninum infection, and mice inoculated with human serum died on day 20 or 22 after challenge. Many N. caninum clusters were observed in the organs and tissues examined; which, listed in decreasing order of cluster frequency, were the brain, oviduct, adrenal gland, lung, stomach, spleen, skeletal muscle, pancreas, and mesenteric lymph nodes (Fig. 3). These N. caninum clusters were associated with severe degeneration/necrosis of the parenchymal cells and inflammatory reactions. There were many necrosis areas and degradation foci. However, in the case of FBS inoculation, there were still mice surviving on day 22 after challenge. There seemed to be fewer lesions in these mice than in those administered human serum. This study indicates that human serum at certain concentrations has no effect on the invasion of N. caninum into host cells or its growth in them. Furthermore, the results of the inoculation test in scid mice administered human serum also demonstrated that it had no protective ability against N. caninum infection. However, the human serum used in these experiments was heat inactivated to avoid any cytotoxic effect on host cells, so there is still a possibility that heat-labile components of serum that were affected by the inactivation could exert a protective action. With the infected sheep serum had NAT titer, there was a marked inhibitory effect on the invasion of N. caninum into BAE cells and Caco-2 cells. This indicates that the anti–N. caninum antibody detected by NAT may play a role in protection against N. caninum. O’Handley et al. (2002) demonstrated that there was an increase in NAT titers during the experimental infection of sheep. In this study, we could not explain the reason why the levels of 3H-uracil incorporation for human serum tended to be higher than those for FBS, and the scid mice administered with FBS showed fewer lesions than those administered human serum. Dubey and Lindsay (1996) reported that most batches of commercial fetal sera have N. caninum antibodies, and binding of non-

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specific immunoglobulins or other proteins in culture media to tachyzoites can be recognized nonspecifically by anti-bovine conjugates. In this regard,1 possibility is that N. caninum antibodies or binding of nonspecific immunoglobulins or other proteins in FBS, which were not detected by NAT, may have an inhibitory effect on the invasion of N. caninum into host cells or its growth in them. Because N. caninum is an obligate intracellular parasite, cell-mediated immunity should play a major role in protection. To determine more clearly whether human serum provides any resistance to N. caninum infection, further study is necessary. This would involve examining resistance effected by natural killer cells, macrophages, and dendritic cells, which participate in natural resistance, and induction of cellmediated immune responses to the parasite. It would also be important to continue searching for N. caninum microscopically in tissue biopsies, cerebrospinal fluid, fetal blood, and amniotic fluid and also to examine for the presence of its DNA in these tissues and body fluids. We are grateful to M. Watarai, Laboratory of Veterinary Microbiology, Obihiro University of Agriculture and Veterinary Medicine for providing Caco-2 cells. This work was supported in part by Grant-inaid for Scientific Research (The 21st Century Center-of-Excellence Program) from the Ministry of Education, Culture, Sports, Science, and Technology, Japan. LITERATURE CITED DUBEY, J. P., AND D. S. LINDSAY. 1996. A review of Neospora caninum and nesporosis. Veterinary Parasitology 67: 1–59. GRAHAM, D. A., V. CALVERT, M. WHYTE, AND J. MARKS. 1999. Absence of serological evidence for human Neospora caninum infection. Veterinary Record 144: 672–673. INNES, E. A., W. R. M. PANTON, J. MARKS, A. J. TREES, J. HOLMDAHL, AND D. BUXTON. 1995. Interferon gamma inhibits the intracellular multiplication of Neospora caninum, as shown by incorporation of 3H uracil. Journal of Comparative Pathology 113: 95–100. KOBAYASHI, Y., M. YAMADA, Y. OMATA, T. KOYAMA, A. SAITO, T. MATSUDA, K. OKUYAMA, S. FUJIMOTO, H. FURUOKA, AND T. MATSUI. 2001. Naturally-occurring Neospora caninum infection in an adult sheep and her twin fetuses. Journal of Parasitology 87: 1434–1436. KOYAMA, T., Y. KOBAYASHI, Y. OMATA, M. YAMADA, H. FURUOKA, R. MAEDA, T. MATSUI, A. SAITO, AND T. MIKAMI. 2001. Isolation of Neospora caninum from the brain of a pregnanat sheep. Jounral of Parasitology 87: 1486–1488. LINDSAY, D. S., J. P. DUBEY, AND R. B. DUNCAN. 1999. Confirmation that the dog is a definitive host for Neospora caninum. Veterinary Parasitology 82: 327–333. ———, S. E. LITTLE, AND W. R. DAVIDSON. 2002. Prevalence of antibodies to Neospora caninum in white-tailed deer, Odocoileus virginianus, from the southeastern United States. Journal of Parasitology 88: 415–417. MCALLISTER, M. M., J. P. DUBEY, D. S. LINDSAY, W. R. JOLLEY, R. A. WILLS, AND A. M. MCGUIRE. 1998. Dogs are definitive hosts of Neospora caninum. International Journal for Parasitology 28: 1473–1482. O’HANDLEY, R., S. LIDDEL, C. PARKER, M. C. JENKINS, AND J. P. DUBEY. 2002. Experimental infection of sheep with Neospora caninum oocysts. Journal of Parasitology 88: 1120–1123. SUNDERMANN, C. A., AND B. H. ESTRIDGE. 1999. Growth of and competition between Neospora caninum and Toxoplasma gondii in vitro. International Journal for Parasitology 29: 1725–1732. TANAKA, T., Y. OMATA, M. NARISAWA, A. SAITO, K. SHIMAZAKI, I. IGARASHI, H. HIRUMI, AND N. SUZUKI. 1997. Growth inhibitory effect of bovine lactoferrin on Toxoplasma gondii tachyzoites in murine macrophages: Role of radical oxygen and inorganic nitrogen oxide in Toxoplasma growth-inhibitory activity. Veterinary Parasitology 68: 27–33. TRANAS, J., R. A. HEINZEN, L. M. WEISS, AND M. M. MCALLISTER. 1999. Serological evidence of human infection with the protozoan Neospora caninum. Clinical and Diagnostic Laboratory Immunology 6: 765–767.