J. Parasitol., 90(3), 2004, pp. 653–654
q American Society of Parasitologists 2004
Serological Survey of
Toxoplasma gondii Infection Among Slaughtered Pigs in
Northwestern Taiwan
Chia-Kwung Fan, Kua-Eyre Su*, and Yu-Jen Tsai†,Department of Parasitology, College of Medicine, Taipei Medical University, No. 250
Wu-Hsin Street, Taipei 110, Taiwan, Republic of China; *Department of Parasitology, College of Medicine, National Taiwan University, Taipei, Taiwan, Republic of China; †Taipei Municipal Institute for Animal Health, Taipei, Taiwan, Republic of China.e-mail: tedfan@tmu.edu.tw
TABLEI. Seroprevalence of Toxoplasma antibody among pigs slaugh-tered in northwestern Taiwan using the latex agglutination (LA) test during 1998. Sex/LA titer Male* Tested Positive (%) Female* Tested Positive (%) Total Tested Positive (%) 1:32 1:64 1:128 1:256 1:512 1:1,024 70 — — — — — — 20 (28.6) 4 (20.0) 7 (35.0) 6 (30.0) 2 (10.0) 1 (5.0) 0 (0.0) 41 — — — — — — 12 (29.7) 2 (16.7) 3 (25.0) 3 (25.0) 4 (33.3) 0 (0.0) 0 (0.0) 111 — — — — — — 32 (28.8) 6 (18.8) 10 (31.2) 9 (28.1) 6 (18.8) 1 (3.1) 0 (0.0) * No significant difference was observed (P. 0.05).
ABSTRACT: A serological survey of Toxoplasma gondii infection among slaughtered pigs in the largest slaughterhouse located in Taoyuan County of northwestern Taiwan was conducted using the latex agglu-tination (LA) test during 1998. The overall seroprevalence of T. gondii infection was 28.8% (32/111) with LA titers of 1:32 (6, 18.8%), 1:64 (10, 31.2%), 1:128 (9, 28.1%), 1:256 (6, 18.8%), and 1:512 (1, 3.1%). No significant difference (P. 0.05) in seroprevalence between male (28.6%, 20/70) and female (29.7%, 12/41) slaughtered pigs was ob-served. A decreasing trend in the seroprevalence among slaughtered pigs examined in the same slaughterhouse was observed because of a lower seroprevalence (P, 0.05) than that (44.4%, 128/288) previously reported about 10 yr ago using the LA test. Nevertheless, it is important to avoid eating raw or undercooked pork in order to prevent the acqui-sition of T. gondii infection among people in Taiwan.
Infection by the protozoan parasite Toxoplasma gondii is prevalent in animals and humans worldwide. Humans become infected with T. gondii usually by ingesting oocysts in food and water contaminated by cat feces or by consuming tissue cysts in undercooked meat (Dubey and Beattie, 1988). Pork is considered to be the most important meat source of T. gondii infection in the United States (Dubey, 1986). Two outbreaks of acute toxoplasmosis involving 8 adult patients in Korea were linked to eating uncooked pork (Choi et al., 1997). Most infections are asymptomatic, but in some persons it eventually becomes symptom-atic. Thus, for example, toxoplasmic encephalitis is a major disease in acquired immunodeficiency syndrome patients. Prenatal infection may also occur, resulting in newborns with congenital toxoplasmosis (Jones et al., 2001). In the United States, the T. gondii–associated annual eco-nomic public health burden purportedly exceeds $400 million (Roberts et al., 1994).
In our previous studies, the seroprevalence of latent T. gondii infec-tion among Taiwanese, especially of mountain aboriginal populainfec-tions, was not low, with a range of 2.7–26.7% using the latex agglutination (LA) test (Fan et al., 1998, 2001, 2002). In addition, data from ques-tionnaires concerning risk factors of acquiring T. gondii infection showed that the consumption habits of frequently eating raw or under-cooked pork were considered the chief factor contributing to T. gondii infection among these people (Fan et al., 2001, 2002). Infective tissue cysts have been found repeatedly in commercial cuts of pork of both experimentally and naturally infected pigs (Dubey, 1988). In Taiwan,
studies concerning T. gondii infection in pigs are rather rare. Kundin et al. (1972) were the first to investigate the T. gondii infection in slaugh-tered pigs and indicated the low prevalence (1%, 10/999) in Taiwan. Lee et al. (1975) reported, in contrast, that the prevalence in slaughtered pigs was high (52%, 26/50) in southern Taiwan. In the 1980s, a large-scale survey of pig toxoplasmosis was conducted in 8 counties of Tai-wan using the LA test, and it was found that the overall seroprevalence of latent T. gondii infection in slaughtered pigs was 27.7% (1,073/ 3,880), of which the highest seroprevalence (44.4%, 128/288) was re-corded in the largest slaughterhouse located in Tauyuan County of northwestern Taiwan (Chang et al., 1991). Since then, information con-cerning the status of pig toxoplasmosis in Taiwan is not available. The purpose of this study was to reexamine the prevalence of antibodies to T. gondii in pigs in the largest slaughterhouse located in Tauyuan Coun-ty of northwestern Taiwan.
Between January and October 1998, 111 blood samples were ran-domly collected from pigs having a slaughter weight of 110 kg in a slaughterhouse located in Taoyuan County. Sera were separated by cen-trifugation and kept at270 C until analysis. A commercial LA test kit (Eiken Chemical Co., Tokyo, Japan) was used to test serum anti–T. gondii antibodies (Chang et al., 1991; Gajadhar et al., 1998; Kim et al., 2002). The reactions were performed using a 96-well U-bottom poly-styrene microplate at 2 dilutions, i.e., 1:16 to 1:1,024. To each well was then added 25ml of T. gondii antigen–coated latex particles suspension, which was incubated overnight at room temperature. An agglutination titer$1:32, i.e., 1:32 to 1:1,024, was considered positive. For calcula-tion of the significant differences in seroprevalence between male and female slaughtered pigs, a chi-square test was used and a P value,0.05 was considered significant. Although several studies have indicated that the LA test is not the best serological tool to detect latent T. gondii infection in pigs (Dubey et al., 1995), we still used the LA test because of its moderate agreements with the dye test and modified agglutination test (Dubey et al., 1995).
Of the 111 serum samples tested, 32 (28.8%) were found to be pos-itive with LA titers of 1:32 (6, 18.8%), 1:64 (10, 31.2%), 1:128 (9, 28.1%), 1:256 (6, 18.8%), and 1:512 (1, 3.1%) (Table I). No significant difference (P. 0.05) was observed in the seroprevalence between male (28.6%, 20/70) and female (29.7%, 12/41) pigs in this study (Table I). The results of this study were higher than those of pigs reported in China (10.4%) (Lin et al., 1990), Indonesia (6.3%) (Inoue et al., 2001), Japan (9.1%) (Horio et al., 2001), the Netherlands (1.8%) (van Knapen et al., 1995), and the United States (2.2%) (Dubey et al., 1995). Nev-ertheless, a decreasing trend in the seroprevalence among pigs examined in the same slaughterhouse was observed because of the lower sero-prevalence (P, 0.05) than that (44.4%, 128/288) previously reported about 10-yr ago using the LA test (Chang et al., 1991). The probable reasons for the decreasing seroprevalence are believed to be an im-proved zoohygienic situation on farms, good sanitation, and the exclu-sive use of commercially marketed feeds. However, it is important for people in Taiwan to avoid eating raw or undercooked pork in order to prevent the acquisition of T. gondii infection.
LITERATURE CITED
CHANG, G. N., S. S. TSAI, M. KUO,ANDJ. P. DUBEY. 1991. Epidemiology of swine toxoplasmosis in Taiwan. Southeast Asian Journal of Tropical Medicine and Public Health 22: 111–114.
CHOI, W. Y., H. W. NAM, N. H. KWAK, W. HUH, Y. R. KIM, M. W. KANG, S. Y. CHO,ANDJ. P. DUBEY. 1997. Foodborne outbreaks of human toxoplasmosis. Journal of Infectious Diseases 175: 1280–1282.
DUBEY, J. P. 1986. Toxoplasmosis. Journal of the American Veterinary Medical Association 189: 166–170.
———. 1988. Long-term persistence of Toxoplasma gondii in tissues of pigs inoculated with T. gondii oocysts and effect of freezing on viability of tissue cysts in pork. American Journal of Veterinary Research 49: 910–913.
———,ANDC. P. BEATTIE. 1988. Toxoplasmosis of animals and man. CRC Press, Boca Raton, Florida, 59 p.
———, P. THULLIEZ, R. M. WEIGEL, C. D. ANDREWS, P. LIND,ANDE. C. POWELL. 1995. Sensitivity and specificity of various serologic tests for detection of Toxoplasma gondii infection in naturally in-fected sows. American Journal of Veterinary Research 56: 1030– 1036.
FAN, C. K., C. W. LIAO, T. C. KAO, J. L. LU, ANDK. E. SU. 2001. Toxoplasma gondii infection: Relationship between seroprevalence and risk factors among inhabitants in two offshore islands from Taiwan. Acta Medica Okayama 55: 301–308.
———, K. E. SU, W. C. CHUNG, Y. J. TSAI, H. Y. CHIOU, C. F. LIN, C. T. SU, M. C. TSAI,ANDP. H. CHAO. 1998. Seroprevalence of Toxo-plasma gondii antibodies among Atayal aboriginal people and their hunting dogs in northeastern Taiwan. Japanese Journal of Medical Science and Biology 54: 35–42.
———, ———, G. H. WU,ANDH. Y. CHIOU. 2002. Seroepidemiology of Toxoplasma gondii infection among two mountain aboriginal populations and Southeast Asian laborers in Taiwan. Journal of Parasitology 88: 411–414.
GAJADHAR, A. A., J. J. ARAMINI, G. TIFFIN,ANDJ. R. BISAILLON. 1998. Prevalence of Toxoplasma gondii in Canadian market-age pigs. Journal of Parasitology 84: 759–763.
HORIO, M., K. NAKAMURA,ANDM. SHIMADA. 2001. Risk of Toxoplasma
gondii infection in slaughterhouse workers in Kitakyushu City. Journal of Uoeh 23: 233–243.
INOUE, I., C. S. LEOW, D. HUSIN, K. MATSUO,ANDP. DARMANI. 2001. A survey of Toxoplasma gondii antibodies in pigs in Indonesia. Southeast Asian Journal of Tropical Medicine and Public Health 32: 38–40.
JONES, J. L., D. KRUSZON-MORAN, M. WILSON, G. MCQUILLAN, T. NAVIN, AND J. B. MCAULEY. 2001. Toxoplasma gondii infection in the United States: Seroprevalence and risk factors. American Journal of Epidemiology 154: 357–365.
KIM, J. H., J. K. LEE, E. K. HWANG,ANDD. Y. KIM. 2002. Prevalence of antibodies to Neospora caninum in Korean native beef cattle. Journal of Veterinary Medical Science 64: 941–943.
KUNDIN, W. D., W. F. CHEN, J. H. CROSS, AND G. S. IRVING. 1972. Isolation of Toxoplasma during unsuccessful attempts to isolate Rickettsiae from swine and rodents in Taiwan. Chinese Journal of Microbiology 5: 118–121.
LEE, S., S. CHEN, K. LIU, S. LIN,ANDT. SUZUKI. 1975. Toxoplasmosis in Taiwan. 5. Detection of Toxoplasma cysts from swine lymphno-des and its correlation with titer of indirect hemagglutination test. Journal of the Formosan Medical Association 74: 82–85. LIN, S., Z. C. LING, B. C. ZENG,ANDY. H. YANG. 1990. Prevalence of
Toxoplasma gondii infection in man and animals in Guangdong, Peoples Republic of China. Veterinary Parasitology 34: 357–360. ROBERTS, T., K. D. MURRELL, ANDS. MARKS. 1994. Economic losses
caused by foodborne parasitic diseases. Parasitology Today 10: 419–423.
VANKNAPEN, F., A. F. KREMERS, J. H. FRANCHIMONT,ANDU. NARUCKA. 1995. Prevalence of antibodies to Toxoplasma gondii in cattle and swine in the Netherlands: Towards an integrated control of live-stock production. Veterinary Quarterly 17: 87–91.
J. Parasitol., 90(3), 2004, pp. 654–657
q American Society of Parasitologists 2004
Infectivity of Microsporidia Spores Stored in Seawater at Environmental Temperatures
R. Fayer,Environmental Microbial Safety Laboratory, Agricultural Research Service, United States Department of Agriculture, 10300 Baltimore
Avenue, Beltsville, Maryland 20705-2350;e-mail: rfayer@anri.barc.usda.gov
ABSTRACT: To determine how long spores of Encephalitozoon cuniculi, E. hellem, and E. intestinalis remain viable in seawater at environmental temperatures, culture-derived spores were stored in 10, 20, and 30 ppt artificial seawater at 10 and 20 C. At intervals of 1, 2, 4, 8, and 12 wk, spores were tested for infectivity in monolayer cultures of Madin Darby bovine kidney cells. Spores of E. hellem appeared the most robust, some remaining infectious in 30 ppt seawater at 10 C for 12 wk and in 30 ppt seawater at 20 C for 2 wk. Those of E. intestinalis were slightly less robust, remaining infectious in 30 ppt seawater at 10 and 20 C for 1 and 2 wk, respectively. Spores of E. cuniculi remained infectious in 10 ppt seawater at 10 and 20 C for 2 wk but not at higher salinities. These findings indicate that the spores of the 3 species of Encephali-tozoon vary in their ability to remain viable when exposed to a con-servative range of salinities and temperatures found in nature but, based strictly on salinity and temperature, can potentially remain infectious long enough to become widely dispersed in estuarine and coastal waters. Fourteen species of microsporidia have been reported to infect hu-mans (Kotler and Orenstein, 1999; Cali and Takvorian, 2003). Of these, Encephalitozoon cuniculi, E. hellem, E. intestinalis, and Enterocytozoon bieneusi are zoonotic, infecting domesticated animals (Deplazes et al., 1996; Mansfield et al., 1997; Breitenmoser et al., 1999; Mathis et al., 1999; Rinder et al., 2000; Buckholt et al., 2002; Fayer, Santin, and Trout, 2003) and wildlife (Hersteinsson et al., 1993; Mathis et al., 1996; Thomas et al., 1997; Sulaiman et al., 2003). Encephalitozoon cuniculi has been identified in wild and pet rabbits, wild rats and mice, dogs, cats, foxes, mink, and a variety of monkeys (Bryan and Schwartz, 1999; Deplazes et al., 2000). Encephalitozoon hellem and E. hellem–like
mi-crosporidia have been found in psittacine birds, budgerigar chicks, a wild yellow-streaked lory (Bryan and Schwartz, 1999; Deplazes et al., 2000), and have experimentally infected domesticated chickens (Fayer, Santin, Palmer et al., 2003). Encephalitozoon intestinalis spores have been reported from feces of farm animals in Mexico (dog, pig, cow, goat) (Bornay-Llinares et al., 1998). Although the actual routes of trans-mission are not known, it is possible that the infectious spore stage in urine or feces can contaminate surface waters used for recreation or drinking water (Sparfel et al., 1997; Dowd et al., 1998; Cotte et al., 1999; Fournier et al., 2000). Microscopic and molecular detection of spores in surface waters and circumstantial evidence of waterborne transmission has been reviewed by Bryan and Schwartz (1999). Under experimental conditions, spores of E. cuniculi, E. hellem, and E. intes-tinalis stored in water at environmental temperatures ranging from 10 to 30 C remained infectious long enough to become widely dispersed if exposed to similar conditions in the environment (Li et al., 2003). For example, at 10 C, spores of E. intestinalis were still infectious after 12 mo, whereas those of E. hellem and E. cuniculi were infectious for 9 and 3 mo, respectively. At 30 C, the former 2 species were infectious for 3 wk and 1 mo, respectively, and the latter species for 1 wk. Little is known of how long microsporidians remain infectious in seawater. On the basis of artificially induced filament extrusion from spores of the microsporidian fish parasite Loma salmonae, a decrease was found after storage in seawater, suggesting that spores lost viability (Shaw et al., 2000). Using a similar experimental design as that of Li et al. (2003), the present study was conducted to determine the effect of sa-linity and temperature on longevity of spores of zoonotic species of Encephalitozoon that might be found in estuaries and coastal marine
TABLEI. Viability of Encephalitozoon hellem spores held in 0, 10, 20, and 30 ppt of seawater at 10 and 20 C for 1, 2, 4, 8, and 12 wk, as determined by observation of clusters of proliferating intracellular parasites in Madin Darby bovine kidney cells. (0, 0 clusters; 1, 1–10 clusters; 2, 11–100 clusters; 3, 1011 clusters.) TP* Positive control 5 C 0 ppt, 10 C 10 ppt, 10 C 20 ppt, 10 C 30 ppt, 10 C 0 ppt, 20 C 10 ppt, 20 C 20 ppt, 20 C 30 ppt, 20 C 1 2 4 8 12 3, 2, 3† (TP5 0) 3, 3, 2 (TP5 12) 2, 2, 2 2, 2, 2 2, 1, 1 1, 1, 1 1, 1, 1 2, 2, 2 2, 2, 2 2, 1, 1 1, 1, 0 2, 1, 1 2, 1, 1 2, 2, 2 1, 1, 1 1, 0, 0 1, 1, 0 2, 2, 2 2, 2, 2 2, 1, 1 1, 1, 1 1, 0, 0 2, 2, 1 2, 2, 1 1, 1, 1 1, 1, 0 1, 1, 0 2, 1, 1 1, 1, 1 1, 1, 0 1, 0, 0 0, 0, 0 2, 1, 1 1, 0, 0 1, 0, 0 0, 0, 0 0, 0, 0 1, 1, 1 1, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 * TP, time period (number of weeks spores were held before in vitro viability testing).
† Number of clusters of proliferating parasites within each of 3 wells of MDBK cells.
TABLEII. Viability of Encephalitozoon intestinalis spores held in 0, 10, 20, and 30 ppt of seawater at 10 and 20 C for 1, 2, 4, 8, and 12 weeks, as determined by observation of clusters of proliferating intracellular parasites in Madin Darby bovine kidney cells. (0, 0 clusters; 1, 1–10 clusters; 2, 11–100 clusters; 3, 1011 clusters.) TP* Positive control 0 ppt, 10 C 10 ppt, 10 C 20 ppt, 10 C 30 ppt, 10 C 0 ppt, 20 C 10 ppt, 20 C 20 ppt, 20 C 30 ppt, 20 C 1 2 4 8 12 3, 2, 2† (TP5 0) 2, 2, 2 (TP5 12) 2, 2, 2 2, 2, 1 1, 1, 1 1, 1, 1 1, 1, 1 2, 2, 1 2, 1, 1 1, 1, 1 2, 1, 0 1, 0, 0 2, 1, 1 1, 1, 1 1, 0, 0 1, 1, 0 1, 0, 0 1, 1, 1 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 2, 1, 1 2, 1, 1 1, 1, 1 1, 1, 1 1, 1, 1 2, 2, 1 1, 1, 0 1, 0, 0 1, 0, 0 1, 1, 0 1, 0, 0 1, 0, 0 1, 1, 0 0, 0, 0 0, 0, 0 1, 0, 0 1, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 * TP, time period (number of weeks spores were held before in vitro viability testing).
† Number of clusters of proliferating parasites within each of 3 wells of MDBK cells.
TABLEIII. Viability of Encephalitozoon cuniculi spores held in 0, 10, 20, and 30 ppt of seawater at 10 and 20 C for 1, 2, 4, 8, and 12 weeks, as determined by observation of clusters of proliferating intracellular parasites in Madin Darby bovine kidney cells. (0, 0 clusters; 1, 1–10 clusters; 2, 11–100 clusters; 3, 1011 clusters.) TP* Positive control 0 ppt, 10 C 10 ppt, 10 C 20 ppt, 10 C 30 ppt, 10 C 0 ppt, 20 C 10 ppt, 20 C 20 ppt, 20 C 30 ppt, 20 C 1 2 4 8 12 2, 2, 2† (TP5 0) 2, 2, 2 (TP5 12) 2, 2, 2 1, 1, 1 1, 1, 1 1, 1, 0 1, 0, 0 1, 1, 1 1, 1, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 1, 1, 1 1, 1, 1 1, 1, 0 0, 0, 0 0, 0, 0 1, 1, 0 1, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 0, 0, 0 * TP, time period (number of weeks spores were held before in vitro viability testing).
† Number of clusters of proliferating parasites within each of 3 wells of MDBK cells. waters used for recreation and shellfish harvesting. Knowledge of the effect of salinity and temperature on infectivity of microsporidia in sea-water is necessary for evaluating the risk of sea-waterborne contamination. Many isolates of E. cuniculi, E. hellem, and E. intestinalis have been propagated in vitro in many types of cells (Visvesvara, 2002). In the present study, spores were obtained and propagated as described pre-viously (Li et al., 2003). Briefly, E. cuniculi and E. intestinalis were propagated in monolayer cultures of Madin Darby bovine kidney (MDBK) cells, and E. hellem was propagated in human lung fibroblasts (WI-38). MDBK cells were cultured in Dulbecco modified Eagle me-dium supplemented with 1% nonessential amino acids, 2% N-2-hy-droxythylpiperazine-N9-2-ethane-sulfonic acid, 5% fetal calf serum (FCS), and 1% penicillin–streptomycin in a 5% CO2atmosphere at 35 C. WI-38 cells were similarly cultured, but in minimum essential me-dium with 10% FCS, as well as 1%L-glutamine, and 1% sodium py-ruvate. Spores harvested from culture supernatant by centrifuging at 1,500 g for 15 min were resuspended in deionized water, stained with calcofluor white (Becton Dickinson Microbiology Systems, Sparks, Maryland), pipetted into a well of a Teflon-coated 3-well glass micro-scope slide (Cel-Line, Erie Scientific, Portsmouth, New Hampshire), and counted with the aid of an epifluorescence microscope. For each species, morphologic features and staining intensity appeared uniform,
indicating that the forms examined were spores. Spores of each species were pipetted at 1.53 105spores per tube into 40 microcentrifuge tubes, centrifuged, and the pellets resuspended in artificial seawater at con-centrations of 10, 20, and 30 ppt salinity or in deionized water (Tables I–III). Seawater was constituted from Forty Fathoms Crystal Sea Marine Mix (Marine Enterprises International, Inc., Baltimore, Maryland) dis-solved in deionized water. Spores suspended in deionized water served as controls. Tubes were capped and held in either of 2 circulating water baths at 10 and 20 C that were monitored for temperature twice daily, except on weekends. At 1, 2, 4, 8, and 12 wk, 1 tube for each species of microsporidia at each concentration of salinity was removed from each water bath, spores were aspirated from that tube, and 5 3 104 spores were inoculated into each of the 3 wells of an 8-well Lab-Tek chamber slide (Nalge Nunc Intl., Naperville, Illinois), each well con-taining a monolayer of MDBK cells. After 4 days incubation at 35 C in a 5% CO2atmosphere, the culture medium was decanted, wells were flooded with 100% methanol for 30 min, and slides were air dried. After removing the plastic frame and silicon gasket that formed the wells, each slide was stained by the quick-hot gram-chromotrope method (Moura et al., 1997), a coverslip was affixed, and the entire area of cells within each well was examined by brightfield microscopy. Spores were considered viable and infectious on finding intracellular clusters of
pro-liferating microsporidia. The number of clusters in each well was count-ed, and the numbers 0, 1, 2, and 3 were assigned to represent counts of 0, 1–10, 11–100, and 101 or more clusters per well, respectively (Tables I–III). At the onset (time 0) and termination (12 wk) of the study, 53 104spores of each species held as positive controls in de-ionized water at 5 C were also pipetted into each of 3 wells containing MDBK cell monolayers, then processed and examined in the same man-ner as wells that received spores stored in artificial seawater.
For spores of all 3 species held in deionized water at 10 and 20 C compared with those held in deionized water at 5 C for 12 wk, infec-tivity decreased both with elevated temperatures and length of storage time (Tables I–III). At all time periods for all species at both 10 and 20 C (with 2 exceptions), spores held in seawater were less infectious than those held in deionized water (Tables I–III), indicating a negative effect on infectivity from elevated salinity alone. The degree to which spores were affected was species dependent. Some spores of E. hellem remained infectious at 30 ppt at 10 and 20 C for 12 and 2 wk, respec-tively, fewer spores of E. intestinalis remained infectious at 30 ppt at 10 and 20 C for 1 and 2 wk, and spores of E. cuniculi remained infec-tious at only 10 ppt at 10 and 20 C for 2 wk.
Spores of microsporidia have been detected in a variety of surface waters (Avery and Undeen, 1987; Dowd et al., 1998), and water as a source of human infections has been implied from epidemiological data (Cotte et al., 1999), but information is lacking on the presence of and survival in seawater of microsporidia infectious to humans and other mammals. General interest in survival of microsporidian spores dates back nearly 90 yr, with most efforts to determine the effects of time versus temperature on the viability of spores of Nosema apis, a micro-sporidian parasite of bees, held in water at various temperatures (White, 1919; Revell, 1960; Kramer, 1970; Bailey, 1972; Vavra and Maddox, 1976; Malone et al., 2001) or the mammalian microsporidia E. hellem and E. intestinalis (Kucerova-Pospisilova et al., 1999; Li et al., 2003) and E. cuniculi (Shadduck and Polley, 1978; Waller, 1979; Koudela et al., 1999; Kucerova-Pospisilova et al., 1999; Li et al., 2003) held in water or culture medium. The only study to determine the effect of seawater on spores of microsporidia was that of Shaw et al. (2000), who examined the microsporidian fish parasite L. salmonae.
Infection with all microsporidia begins when the polar filament ex-truded from the spore forms a tube through which the sporoplasm passes into a host cell (Vavra and Larsson, 1999). This process of germination can be artificially induced. However, it is technique dependent, and polar filaments can fail to extrude from spores that are potentially in-fectious or can extrude from spores that lack infectivity. Shaw et al. (2000) examined the germination rate of L. salmonae and found that it decreased from 51 to 0% after 100 days storage at 4 C, suggesting that spores lost viability, although after 95 days, infectivity for fish appeared not to be diminished. Germination was induced in spores of E. intes-tinalis, E. hellem, and E. cuniculi stored in culture medium at 4 C for 48 mo, and the microscopic appearance of intact versus recently ger-minated spores versus those that had lost extruded polar filaments was reported (Kucerova-Pospisilova et al., 1999). When spores of E. cunic-uli, E. hellem, and E. intestinalis stored in deionized water at elevated temperatures for 2, 8, and 10 mo were examined by DIC microscopy and chromotrope-stained spores were examined by brightfield micros-copy, no extruded filament was detected despite the fact that other spores stored under the same conditions were infectious to cultured mammalian cells (Li et al., 2003). These findings suggested that factors other than extrusion of the filament were involved in the loss of infec-tivity (Li et al., 2003). On the basis of those findings, spores in the present study were not examined for polar filament extrusion but were considered infectious based solely on their ability to actually invade and multiply within cultured mammalian cells.
The present study has demonstrated that as the temperature of storage in deionized water increased from 5 to 20 C, infectivity of microspo-ridian spores decreased and as salinity increased from 0 to 30 ppt, infectivity of microsporidian spores decreased. At the highest level of salinity (30 ppt) at both 10 and 20 C, spores of E. hellem were more robust, i.e., remained infectious longer or more were infectious for lon-ger periods, than those of E. intestinalis, which were more robust than those of E. cuniculi, indicating species differences with respect to the effects of salinity. These findings suggest that spores of E. hellem and E. intestinalis could potentially remain infectious in estuarine and ocean waters for weeks and those of E. cuniculi could remain infectious in
low-salinity estuarine waters for weeks, which is sufficient to infect humans and marine mammals or to contaminate shellfish.
The technical assistance of Robert Palmer is gratefully acknowl-edged.
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q American Society of Parasitologists 2004
Ticks (Acari: Ixodidae) Parasitizing Wild Carnivores in Phu Khieo Wildlife Sanctuary,
Thailand
L. I. Grassman, Jr., N. Sarataphan*, M. E. Tewes, N. J. Silvy†, and T. Nakanakrat‡,Feline Research Program, Caesar Kleberg Wildlife
Research Institute, 700 University Boulevard, MSC 218, Texas A&M University–Kingsville, Kingsville, Texas 78363; *Parasitology Section, National Institute of Animal Health, Department of Livestock Development, Chatuchak, Bangkok 10900, Thailand; †Department of Wildlife and Fisheries Sciences, 210 Nagle Hall, Texas A&M University, College Station, Texas 77840; and ‡Phu Khieo Wildlife Sanctuary, P.O. Box 3, Chum Phrae, Khon Kaen 40130, Thailand.e-mail: kslig01@tamuk.edu
ABSTRACT: Ixodid ticks were collected and identified from 8 wild car-nivore species in Phu Khieo Wildlife Sanctuary, northeastern Thailand. Six tick species belonging to 4 genera were recovered and identified from 132 individuals. These included Amblyomma testudinarium (n5 36), Haemaphysalis asiatica (n5 58), H. hystricis (n 5 31), H. semer-mis (n5 3), Rhipicephalus haemaphysaloides (n 5 3), and Ixodes gran-ulatus (n5 1). Leopard cats (Prionailurus bengalensis) (n 5 19) were infested with 4 tick species, whereas yellow-throated marten (Martes flavigula) (n 5 4), clouded leopard (Neofelis nebulosa) (n 5 2), and dhole (Cuon alpinus) (n5 1) were infested with 3 tick species, Asiatic golden cat (Catopuma temmincki) (n5 2) with 2 species, and marbled cat (Pardofelis marmorata), binturong (Arctictis binturong), and large Indian civet (Viverra zibetha) each infested with 1 species. This infor-mation contributes to the knowledge available on the ectoparasites of wild carnivores in Southeast Asia.
The collection and identification of ectoparasites from wild carni-vores in North America and Europe is well documented in the literature, including host species such as raccoon (Procyon lotor) (Sonenshine and Stout, 1971; Rhodes and Norment, 1979; Whitaker and Goff, 1979;
Brillhart et al., 1994; Pung et al., 1994), striped skunk (Mephitis me-phitis) (Durden and Richardson, 2003), coyote (Canis latrans) (Eads, 1948; Pence et al., 1981), red fox (Vulpes vulpes) (Aubert, 1975; Tou-toungi et al., 1991), and river otter (Lutra canadensis) (Eley, 1977). Tick research in Southeast Asia has mainly covered tick identification, distribution, and disease transmission (Toumanoff, 1944; Petney and Keirans, 1994, 1995, 1996; Voltzit and Keirans, 2002), a checklist of Thai ticks (Tanskul et al., 1983), and a tick survey of Malaysian car-nivores and other mammals (Hoogstraal and Wassef, 1984). Except for research on the endoparasites of Thai wild cats (Patton and Rabinowitz, 1994), parasitological research on wild carnivores in Thailand remains largely unstudied. Our objective was to add information about the tick– host relationships of carnivores from Southeast Asia.
Ticks were collected as part of an ecological study of carnivores in northeastern Thailand (Grassman, 2004). Situated in Chaiyaphum Prov-ince (16859–168359N, 1018209–1018559E), Phu Khieo Wildlife Sanctuary (PKWS) is a large, 1,560-km2 evergreen forest dominated by an ap-proximately 1,000-m elevation plateau (Anonymous, 2000). Carnivores were livetrapped from October 1998 to October 2002. Captured carni-vores were anesthetized for physical examination and to attach a radio
TABLEI. Wild carnivore hosts and ticks from Phu Khieo Wildlife Sanc-tuary, Thailand, October 1998 to October 2002.
Host No. of carni-vores No. of tick species Tick species* and stage† Prionailurus bengalensis Martes flavigula Neofelis nebulosa Cuon alpinus Catopuma temmincki Pardofelis marmorata Arctictis binturong Viverra zibetha 19 4 2 1 2 1 1 1 4 3 3 3 2 1 1 1 1LN, 2NA, 5A, 6A 1N, 3LNA, 4A 2A, 3A, 4A 1NA, 2N, 4A 1N, 2A 3N 4A 1LN
* 1, Amblyomma testudinarium; 2, Haemaphysalis asiatica; 3, H. hystricis; 4, Rhipicephalus haemaphysaloides; 5, H. semermis; and 6, Ixodes granulatus. † L, larvae; N, nymphs; A, adults.
collar. During examination, attached and unattached ticks were removed by forceps and placed in plastic vials containing 70% ethanol. Tick specimens were stored at room temperature#4 yr before identification. Parasites were identified by the Parasitology Section of the National Institute of Animal Health, Bangkok, Thailand. Ticks were examined by microscope and identified according to the methods of Yamaguti et al. (1971a, 1971b), Tanskul and Inlao (1989), and Walker et al. (2000). Eight carnivore species (31 individuals) were captured and examined for ticks. Hosts examined included clouded leopard (Neofelis nebulosa) (n5 2), Asiatic golden cat (Catopuma temmincki) (n 5 2), marbled cat (Pardofelis marmorata) (n5 1), leopard cat (Prionailurus bengalensis) (n5 19), dhole (Cuon alpinus) (n 5 1), yellow-throated marten (Martes flavigula) (n5 4), binturong (Arctictis binturong) (n 5 1), and large Indian civet (Viverra zibetha) (n5 1). All ticks (n 5 132) were iden-tified and classified into 6 species: Amblyomma testudinarium (n5 36), Haemaphysalis asiatica (n5 58), H. hystricis (n 5 31), H. semermis (n5 3), Rhipicephalus haemaphysaloides (n 5 3), and Ixodes granu-latus (n5 1) (Table I). Haemaphysalis asiatica was identified the most frequently (43.6%), followed by A. testudinarium (27.1%); however, A. testudinarium was found on more hosts (5) than H. asiatica (4).
No record exists of ixodid ticks parasitizing clouded leopard and marbled cat. In this study, we found H. asiatica, H. hystricis, and R. haemaphysaloides infesting clouded leopards and H. hystricis from a marbled cat. Leopard cats were parasitized by A. testudinarium, H. asia-tica, H. semermis, and I. granulatus. Similarly, ticks parasitizing leopard cats from Thailand recorded by Tanskul et al. (1983) included H. asia-tica, I. granulatus, and I. ovatus. Other parasite records of leopard cats from Nepal included I. ovatus (Hoogstraal et al., 1973) and I. acutitar-sus (Clifford et al., 1975). The occurrence of H. asiatica and A. testu-dinarium identified from the Asiatic golden cat is similar to ticks found on a golden cat in Laos. Robbins et al. (1997) identified H. asiatica, R. haemaphysaloides, and Amblyomma sp. from a single Asiatic golden cat.
Tanskul et al. (1983) recorded H. bispinosa and H. koningsbergeri from a Thai binturong and H. asiatica and I. ovatus from a large Indian civet. We identified R. haemaphysaloides from a binturong and A. tes-tudinarium from a large Indian civet. One record of tick parasitism on dhole from Nepal was identified as I. ovatus (Hoogstraal et al., 1973). Three tick species were identified from a dhole in PKWS, i.e., A. tes-tudinarium, H. asiatica, and R. haemaphysaloides. Two records exist for the yellow-throated marten. Clifford et al. (1975) identified I. tanuki, and Mitchell (1979) identified R. haemaphysaloides and I. tanuki from this mustelid in Nepal. We identified 3 species of ticks from yellow-throated martens, i.e., A. testudinarium, H. hystricis, and R. haema-physaloides.
This study contributed new information on the host–tick fauna of Southeast Asia. Additional research is needed on host–tick identifica-tion, distribuidentifica-tion, and disease agent transmission.
Voucher specimens of ticks were deposited at the Ohio State Museum of Biological Diversity (Columbus, Ohio). Accession numbers: (Hae-maphysalis asiatica) OSAL 003360 (4 specimens), (H. hystricis) OSAL
003361 (4 specimens), (H. semermis) OSAL 003362 (2 specimens), (Rhipicephalus haemaphysaloides) OSAL 003363 (2 specimens), (Ixo-des granulatus) OSAL 003364 (1 specimen), and (Amblyomma testu-dinarium) OSAL 003365 (4 specimens).
This study was supported by the Bosack and Kruger Foundation through the Cat Action Treasury. Support also was provided by the Caesar Kleberg Wildlife Research Institute at Texas A&M University– Kingsville, Sierra Endangered Cat Haven, Hexagon Farm, Parco Faun-istica La Torbiera, Columbus Zoo, Point Defiance Zoo, and Mountain View Farms Conservation Breeding Centre. Research permission was granted by the National Research Council of Thailand (#0004.3/0301) and Royal Forest Department of Thailand. This project was part of the Joint Ph.D. Program between Texas A&M University–Kingsville and Texas A&M University, College Station. Research was approved by the TAMUK Institutional Animal Care and Use Committee (#2003-8-12). This is publication #04-101 of the Caesar Kleberg Wildlife Research Institute. We thank Dr. Hans Klompen of the Museum of Biological Diversity, Ohio Sate University, for cataloging our tick specimens.
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q American Society of Parasitologists 2004
First Report of the Giant Kidney Worm (Dioctophyme renale) in a Harbor Seal
(Phoca vitulina)
V. Hoffman*, T. J. Nolan, and R. Schoelkopf†,Department of Pathobiology, University of Pennsylvania School of Veterinary Medicine, 3800
Spruce Street, Philadelphia, Pennsylvania 19104; *Present address: ORS, VRP Building 28A, RM 106, 28 Library Drive, MSC 5230, Bethesda, Maryland 20892-5230; †Marine Mammal Stranding Center, P.O. Box 713, Brigantine, New Jersey 08203.e-mail: parasit@vet.upenn.edu
FIGURE1. Histological section of the mass found in the peritoneal cavity of the harbor seal. Viable (A) and degenerate (B) Dioctophyme renale ova are surrounded by neutrophils and macrophages. Hematox-ylin and eosin. Bar5 20 mm.
ABSTRACT: A male harbor seal (Phoca vitulina) was found moribund on the coast of New Jersey in January of 2003 and died a few hours later in the Marine Mammal Stranding Center. On necropsy, a single female Dioctophyme renale was recovered from the peritoneal cavity, and a tissue mass was found adjacent to the pelvic urethra and urinary bladder. Within this tissue mass were found D. renale ova. This is the first report of this nematode in the harbor seal and in a North American marine mammal.
An 18.9-kg male harbor seal (Phoca vitulina) was found stranded at Barnegat Lighthouse State Park in New Jersey (latitude 39.76, longitude 74.10) in January 2003. At time of capture the animal had labored breathing and what appeared to be blood was present on the abdominal skin. The animal was treated at the Marine Mammal Stranding Center in Brigantine, New Jersey with 500 ml electrolyte solution, vitamins, and Baytril. The seal was dewormed with 92 mg of levamisole about 6.5 hr before death. The seal was inactive and shivering. The animal died approximately 7 hr after arriving at the stranding center.
At necropsy the urinary bladder was severely distended. It contained watery red fluid that had a strong odor of ammonia. The mucosa was also reddened with a coarsely roughened surface. The ureters were dis-tended measuring 4 mm in diameter bilaterally, but both kidneys were grossly normal. A firm swelling, measuring 5.63 2.6 3 1.8 cm was present adjacent to the pelvic urethra, extending from the trigone of the urinary bladder to the caudal margin of the pelvic bone. The swelling contained a central cavity measuring 4.43 1.2 3 0.6 cm that contained thick opaque fluid. Aerobic cultures of this fluid were negative. The cavity did not communicate with the urethra. Microscopically the wall of the cavitated structure consisted of dense fibrous connective tissue with scattered small bundles of skeletal muscle near the outer surface. Numerous ova were present within multifocal to coalescing inflamma-tory foci. The ova had a double-contoured shell with a mammillated surface (Fig. 1). They contained central granular hypereosinophilic ma-terial and occasionally a single nucleus. Some ova were pale, staining with disrupted and collapsed shells (Fig. 1). Inflammatory cells con-sisted of large numbers of epitheloid macrophages and neutrophils with fewer multinucleated giant cells, lymphocytes, and plasma cells. The urinary bladder had severe necrotizing inflammation with fibrinoid ne-crosis of blood vessels and numerous surface bacteria. The kidneys had
mild multifocal nonsuppurative interstitial nephritis. A large nematode was found adjacent to the rectum in the retroperitoneal space. Parasite eggs were not present in the kidney or the urinary bladder.
Other necropsy findings included numerous lice on the skin of the dorsum (identified as Echinophthirius horridus), chronic ulcerative der-matitis, and cheilitis. Dilated lymphatic vessels were present on the pleural surfaces of the lung. The stomach contained a 2-cm-long white nematode identified as an anisakine. Additional histologic findings in-cluded mild multifocal acute encephalitis of the cerebellum and me-dulla, scattered pulmonary granulomatous and eosinophilic inflamma-tion, and ulcerative gingivitis and dermatitis with intracytoplasmic am-phophilic inclusions in epithelial cells.
The nematode recovered from the peritoneal cavity was a female, brownish-red in color, and measured 22.3 cm long by 4.5 mm wide. The worm was alive when recovered (about 1 day after the death of the seal). The anus was terminal, and the mouth was surrounded by 2 circles of 6 papillae. This led to its identification as Dioctophyme renale.
Eggs recovered from the sediment of the vessel in which the worm was fixed and those seen in histological sections were consistent with this identification (Fig. 1). Ten eggs recovered from the formalin were mea-sured with an ocular micrometer, and their mean size (6SD) was 41.3
6 2.3 mm wide by 68.9 6 3.7 mm long.
At necropsy, dioctophymosis is frequently diagnosed by finding the adult worm, D. renale. In biopsy or cytology material, diagnosis can be definitively made by identifying the mammillated eggs in tissues or urine. Because ova in the seal were found in the wall of the periurethral abscess, it is probable that the worm was residing there after it had migrated into the peritoneal cavity. The intrapelvic lesion could have acted as a functional obstruction leading to the distended and inflamed urinary bladder and hydroureter. The resulting bloody urine lead to the initial observation of bloody fluid on the abdominal skin.
The usual definitive host for D. renale is the mink, although other mustelids and canids are commonly infected. Adult worms are normally found in the right kidney. They are blood-red, and females can measure up to 103 cm long (Measures, 2001). Eggs pass out in the urine, mature to the first stage (within the egg) in about 35 days, after which they are infectious to Lumbriculus variegatus, a freshwater oligochaete. Fish or frogs that ingest the infected oligochaete may act as paratenic hosts. Lesions usually associated with D. renale in mink include destruction of the right kidney. Renal lesions include dilated pelvises, fibrosis, tu-bular atrophy, and chronic inflammation. The worms can survive for years in the natural host without evidence of clinical signs. If the left kidney is also infected or if there is migration into the peritoneal cavity, the animal can die from renal failure or peritonitis, respectively (Mace, 1976; Dyer, 1998).
Immature D. renale have been reported in the body cavities of Cas-pian Seals (Phoca caspica) from the CasCas-pian Sea. (Popov and Taikov, 1985), but this communication is the first report of this parasite in a harbor seal and the first report in a North American marine mammal. Because the life cycle of this parasite takes place in freshwater, the seal must have been infected while feeding in freshwater. Harbor seals are known to venture up freshwater rivers, and some populations stay in freshwater habitats (Baird, 2001).
Cases of D. renale in the peritoneal cavities of its host are not unusual
(Mace, 1976; de Souza Junior and de Padua, 1977; Celerin and McMullen, 1981), however, the eggs laid by such worms cannot pass out of the host to complete the life cycle. It is not known if the eggs recovered from this seal were fertile (unfertilized females will lay eggs). No male was found, and although the seal had been treated with levam-isole 1 day before its death, any dead worms would still be intact. Fertile eggs of D. renale passing in the urine of naturally infected hosts mea-sure 45–47 wide by 73–83 long (Meamea-sures, 2001), slightly larger than the formalin-fixed eggs recovered from this worm.
We wish to acknowledge the help of the staff of the Marine Mammal Stranding Center, the staff of the Pathology Laboratory at the University of Pennsylvania School of Veterinary Medicine, and Dr. Lena Measures for a translated copy of Popov and Taikov’s report.
LITERATURE CITED
BAIRD, R. W. 2001. Status of Harbour Seals, Phoca vitulina, in Canada. Canadian Field-Naturalist 115: 663–675.
CELERIN, A. J.,ANDM. E. MCMULLEN. 1981. Giant kidney worm in a dog. Journal of the American Veterinary Medical Association 179: 245–246.
DESOUZAJUNIOR, F. L.,ANDE. B.DEPADUA. 1977. Dioctophyme renale (Goeze, 1782) (Nematoda, Dioctophymidae) em caes de rua de regiao de Taubate (Sao Paulo, Brasil). Revista de Patologia Tropical 6: 7–10.
DYER, N. W. 1998. Dioctophyma renale in Ranch Mink. Journal of Veterinary Diagnostic Investigation 10: 111–113.
MACE, T. F. 1976. Lesions in mink (Mustela vison) infected with giant kidney worm (Dioctophyme renale). Journal of Wildlife Diseases 12: 88–92.
MEASURES, L. 2001. Dioctophymatosis. In: Parasitic diseases of wild mammals, W. Samuel, M. J. Pybus, and A. A. Kocan (eds.). Iowa State University Press, Ames, Iowa, p. 357–364.
POPOV, V. N.,ANDI. M. TAIKOV. 1985. The discovery of the nematode Dioctophyme renale in the Caspian seal. Vestnik Zoologii 5: 7. [Original title in Russian].
J. Parasitol., 90(3), 2004, pp. 660–663
q American Society of Parasitologists 2004
Localization of a 56-kDa Antigen That is Present in Multiple Developmental Stages of
Neospora caninum
Mark Jenkins, Rodrigo Soares*, Charles Murphy, Andrew Hemphill†, Ryan O’Handley‡, and J. P. Dubey,Animal Parasitic Diseases
Laboratory, Animal and Natural Resources Institute, Agricultural Research Service, United States Department of Agriculture, Beltsville, Maryland 20705; *Departmento de Medicina Veterinaria Preventiva e Saude Animal, Universidade de Sa˜o Paulo, Sa˜o Paulo, SP, Brazil; †Institute of Parasitology, University of Berne, Langgass-Strasse 122, Berne, CH-3012, Switzerland; ‡Atlantic Veterinary College, University of Prince Edward Island, Charlottetown, PEI, C1A 4P3, Canada.e-mail: mjenkins@anri.barc.usda.gov
ABSTRACT: The purpose of the present study was to characterize the intracellular distribution of a native Neospora caninum 56-kDa protein that is recognized by sera from N. caninum–infected dairy cattle. The complementary DNA coding for this protein was expressed in Esche-richia coli as a polyHis fusion protein to which antiserum was prepared and used to localize the antigen in N. caninum tachyzoites and brady-zoites. By sodium dodecylsulfate–polyacrylamide gel electrophoresis and immunoblotting, antirecombinant Nc56 serum recognized a major 56-kDa protein and 2 minor (43 and 39 kDa) proteins of N. caninum tachyzoites. Antiserum to recombinant 56-kDa protein showed this antigen to be pre-sent in both N. caninum tachyzoites and bradyzoites/cysts as detected by immunofluorescence staining. Immunoelectron microscopy revealed the 56-kDa antigen to be present in the apical end of both tachyzoites and bradyzoites and possibly extracellularly secreted by tachyzoites.
Prevention of neosporosis in dairy cattle may rely on drug therapy or vaccination to inhibit reactivation of Neospora caninum during
preg-nancy (Dubey, 1999). Several research groups have identified gene se-quences coding for parasite enzymes on the surface or associated with intracellular organelles. DNA sequences for nucleotide triphosphate hy-drolase (Asai et al., 1998) and a subtilisinlike serine protease (Nc-p65; Louie and Conrad, 1999) have been reported. Gene sequences have also been described for surface antigens (NcSAG1 [Nc-p36], Hemphill, Fuchs et al., 1997; NcSRS2 [Nc-p43], Hemphill, Felleisen et al., 1997) and proteins associated with micronemes (NcMIC2, Lovett et al., 2000; NcMIC3, Naguleswaran et al., 2001; NcMIC1, Keller et al., 2002) and dense granules (NcGRA7, Lally et al., 1997; NcGRA6, Liddell et al., 1998; NcGRA2, Ellis et al., 2000). In the present study, a complemen-tary DNA (cDNA) coding for a recombinant N. caninum antigen that is related to a native 56-kDa protein located in the apical end of N. caninum tachyzoites and bradyzoites is described.
A N. caninum tachyzoite cDNA library prepared in UNIZAP-XR vector (Stratagene, La Jolla, California) was immunoscreened with rab-bit sera specific for a native 56-kDa N. caninum protein. Antinative Nc56 serum was prepared by excising a horizontal strip of nitrocellulose
FIGURE 1. Characterization of Nc56 transcript and protein in Neo-spora caninum tachyzoites. Left panel, Northern blot hybridization of DIG-labeled Nc56 DNA to N. caninum tachyzoite RNA size-fraction-ated on a guanidine thiocyanate agarose gel; kb, RNA molecular size standards. Right panel, immunoblotting analysis of N. caninum tachy-zoite protein probed with antisera to recombinant Nc56 protein.1, re-ducing (12-mercaptoethanol) SDS-PAGE; 2, nonreducing (22 mer-captoethanol) SDS-PAGE; MrS, molecular weight markers.
membrane containing sodium dodecylsulfate–polyacrylamide gel elec-trophoresis (SDS-PAGE)–fractionated whole N. caninum tachyzoite protein corresponding to a 56-kDa protein that was identified by sera from N. caninum–infected cattle. The nitrocellulose strip was minced into small pieces, mixed with ImmunoMax SR adjuvant (Zonagen Inc., The Woodlands, Texas), and injected into 2 female New Zealand white rabbits (Covance, Denver, Pennsylvania) using an 18-gauge needle and 1.0-ml syringe. The rabbits received a booster immunization with the native Nc56-impregnated nitrocellulose membrane at 1-mo postprimary immunization and were bled for serum 2 wk later.
Using an in vivo excision protocol, pBluescript plasmid harboring the Nc56 cDNA was prepared and subjected to dideoxy sequencing using BigDye terminators and analysis on an ABI 377 DNA sequencer (Applied Biosystems, Foster City, California). The cDNA was excised from pBluescript by restriction enzyme digestion and inserted into the pBAD-His expression vector (Invitrogen, Carlsbad, California). DNA sequencing of the recombinant pBAD-His–Nc56 was conducted to en-sure that Nc56 was cloned in-frame with the pBAD-His open-reading frame (ORF). Recombinant polyHis–Nc56 protein was expressed by 0.2% arabinose induction of pBAD-His–Nc56–transformed Escherichia coli DH5 at midlog growth at 37 C. Recombinant protein was extracted under denaturing conditions (3 M urea), and soluble Nc56 protein was purified by NiNTA affinity chromatography following manufacturer’s instructions (Invitrogen). NiNTA-purified Nc56 was emulsified in ImmunoMax SR adjuvant (Zonagen) and used for intramuscular injec-tion into 2 female New Zealand white rabbits (Covance, 1mg/rabbit). The rabbits received a booster immunization with the same amount of purified antigen at 1-mo postprimary immunization and were bled for serum 2 wk later.
Northern blot hybridization was conducted to estimate the size of the Nc56 messenger RNA transcript. Neospora caninum tachyzoite RNA was prepared using Trizol reagent and procedures recommended by the manufacturer (Invitrogen). RNA was suspended in diethyl pyrocarbon-ate–treated H2O. The RNA concentration was estimated by measuring absorbance at optical density 260/280. RNA (20mg) was mixed with deionized formamide and formaldehyde, heat denatured for 5 min at 65 C, chilled on ice, mixed with RNA sample buffer, and electrophoresed in 13 TBE at 7.5 V/cm on 1.2% agarose containing 20 mM guanidine thiocyanate using described procedures (Goda and Minton, 1995). RNA molecular weight markers (New England Biolabs, Beverly, Massachu-setts) were denatured in the same manner and electrophoresed in a sep-arate well to allow for size estimation of the hybridizing RNA band. After electrophoresis, the RNA was transferred overnight to Nylon membrane (Roche, Indianapolis, Indiana) using a Turboblotter (Schlei-cher and Schuell, Keene, New Hampshire) and exposed to 1,200 mJ UV light in a Stratalinker (Stratagene). The Northern blot was treated with blocking reagent (Roche) for 1 hr at 50 C and then probed for 16 hr with digoxigenin (DIG)–labeled Nc56 insert DNA. DIG labeling was performed using Nc56 bacteriophage DNA and the DIG PCR-probe kit following manufacturer’s instructions (Roche). After hybridization, the blots were washed 2 times at room temperature (RT) with 2.03 standard saline citrate (SSC) and 0.1% SDS and 2 times at 50 C with 0.23 SSC and 0.1% SDS. The blots were washed with maleic acid buffer, treated with blocking reagent, and then incubated overnight with a 1:25,000 dilution of alkaline phosphatase–labeled anti-DIG reagent (Roche). The blots were washed 3 times with maleic acid wash buffer, once with alkaline phosphatase buffer, treated with CDP-Star substrate (Roche), and analyzed using EpiChemII Darkroom and LabWorks Software (UVP Inc., Upland, California).
To estimate the size of Nc56-related N. caninum proteins, NC-1 ta-chyzoites were harvested and purified from cell culture using described procedures (Bjerkas et al., 1994). Neospora caninum tachyzoite total protein was extracted as described (Bjerkas et al., 1994), treated with sample buffer with or without 2-mercaptoethanol, size-fractionated by SDS-PAGE, and transferred to Immobilon (Millipore, Bedford, Mas-sachusetts) on a semidry blotter (BioRad, Hercules, California). The blots were treated with phosphate-buffered saline (PBS) containing 2% nonfat dry milk (NFDM) for 1 hr at RT, followed by incubation for 2 hr at RT with a 1:500 dilution of rabbit anti-Nc56 or negative control serum. The blots were then incubated for 1 hr with 10mg/ml biotiny-lated goat anti-rabbit IgG (Sigma Chemical Co., St. Louis, Missouri), for 1 hr with 10mg/ml avidin-peroxidase (Sigma), followed by
perox-idase substrate 4-chloro-1-naphthol (Sigma). The blots were washed 3 times for 5 min per wash between each incubation step.
For immunofluorescence antibody (IFA) staining, N. caninum tachy-zoites were dried onto multiwell slides and then treated for 1 hr with PBS-NFDM to block nonspecific immunoglobulin binding in subse-quent steps. The slides were washed once with PBS, air-dried, incubated for 2 hr with a 1:100 dilution of rabbit antiserum against recombinant Nc56 protein, then incubated with a 1:50 dilution of fluorescein-labeled goat anti-rabbit IgG (H1L chain specific, Sigma) for 1 hr. The slides were washed 3 times with PBS and air-dried between each incubation step. After the last step, the slides were air-dried, overlaid with anti-bleaching mounting medium (Vector Laboratories, Burlingame, Cali-fornia) and a coverslip, and then examined under epifluorescence mi-croscopy at3400 magnification.
To localize Nc56 antigen, in vitro–cultured N. caninum tachyzoites or bradyzoites (Vonlaufen et al., 2002) were suspended in 0.1 M cac-odylate buffer containing 3% paraformaldehyde and 0.5% glutaralde-hyde for 10 min at RT. The fixed tachyzoites were washed 2 times with 0.1 M cacodylate buffer and pelleted by centrifugation. The parasites were dehydrated in a graded ethanol series, infiltrated overnight in LR White hard-grade acrylic resin (London Resin Company, London, U.K.), and cured at 55 C for 24 hr. Thin sections (90 nm thick) were obtained using a Diatome diamond knife on a Reichert/AO Ultracut microtome and collected on 200-mesh nickel grids. The grids were floated on drops of PBS containing 0.1 M glycine and 1% bovine serum albumin for 10 min, washed with PBS, floated on drops of PBS-NFDM-Tw20, and then floated on drops containing a 1:100 dilution of rabbit antiserum in PBS-NFDM-Tw20. The grids were incubated for 2 hr at RT, washed 3 times with PBS-NFDM-Tw20, and floated for 1 hr at RT on drops of a 1:50 dilution of gold particle (10-nm diameter)–labeled goat anti-rabbit IgG (H1L chain specific, Sigma). The grids were washed 3 times with PBS-Tw20, twice with deionized H2O, air-dried, stained with 5% uranyl acetate for 30 min, and examined under a Hi-tachi H500H transmission electron microscope at 75 kV.
Immunoscreening N. caninum tachyzoite cDNA libraries with anti-sera specific for a native 56-kDa N. caninum tachyzoite protein revealed a 2,607-nucleotide (nt) DNA insert (GenBank AY340638). An ORF beginning at nt 3 and terminating at nt 1,004 coding for a predicted
