Comparative Parasitology 68(2) 2001 - Peru State College

Volume 68 July 2001 Number 2 

Formerly the 

Journal of the Helminthological Society of Washington 

A semiannual journal of research devoted to 

Helminthology and all branches of Parasitology 

CONTENTS 

CRISCIONE, C. D., AND W. F. FONT. Development and Specificity of Oochoristica 

javaensis (Eucestoda: Cyclophyllidea: Anoplocephalidae: Linstowiinae) . 149 

CRISCIONE, C. D., AND W. F. FONT. Artifactual and Natural Variation of Oochoristica 

javaensis: Statistical Evaluation of In Situ Fixation 156 

BOLEK, M. G., AND J. R. COGGTNS. Seasonal Occurrence and Community Structure of 

Helminth Parasites in Green Frogs, Rana damitans melanota, from Southeastern 

Wisconsin, U. S.A „ 164 

FORRESTER, D. J., G. W. FOSTER, AND J. E. THUL. Blood Parasites of the Ring-Necked 

Duck (Aythya collaris) on Its Wintering Range in Florida, U.S.A 173 

CARKENO, R. A., L. A. DURDEN, D. R. BROOKS, A. ABRAMS, AND E. P. HOBERG. 

Parelaphostrongylus tennis (Nematoda: Protostrongylidae) and Other Parasites of 

White-Tailed Deer (Odocoileus virginianus) in Costa Rica 177 

JOY, J. E., AND R. B. TUCKER. Cepedietta michiganensis (Protozoa) and Batracholandros 

magnavulvaris (Nematoda) from Plethodontid Salamanders in West Virginia, 

U.S.A x 185 

AGUIRRE-MACEDO, M. L., T. SCHOLZ, D. GoNzALEZ-SoLfs, V. M. VIDAL-MARTINEZ, P. 

POSEL, G. ARJANO-TORRES, S. DUMAILO, AND E. SIU-ESTRADA. Some Adult 

Endohelminths Parasitizing Freshwater Fishes from the Atlantic Drainages of 

Nicaragua .. 190 

SALGADO-MALDONADO, G., G. CABANAS-CARRANZA, J. M. CASPETA-MANDUJANO, E. 

SOTO-GALERA, E. MAYEN-PENA, D. BRAILOVSKY, AND R. BAEZ-VALE. Helminth 

Parasites of Freshwater Fishes of the Balsas River Drainage Basin of Southwestern 

Mexico 196 

SALGADO-MALDONADO, G., G. CABANAS-CARRANZA, E. SOTO-GALERA, J. M. CASPETA- 

MANDUJANO, R. G. MORENO-NAVARRETE, P. SANCHEZ-NAVA, AND R. AGUILAR- 

AGUILAR. A Checklist of Helminth Parasites of Freshwater Fishes from the 

Lerma- Santiago River Basin, Mexico . 204 

CURRAN, S. S., R. M. OVERSTREET, T. T. DANG, AND T. L. NGUYEN. Singhiatrema vietnamensis 

sp. n. (Digenea: Ommatobrephidae) and Szidatia taiwanensis (Fischthal 

and Kuntz, 1975) comb. n. (Digenea: Cyathocotylidae) from Colubrid Snakes in 

Vietnam . ; 219 

KUZMIN, Y., V V TKACH, AND S. D. SNYDER. Rhabdias ambystomae sp. n. (Nematoda: 

Rhabdiasidae) from the North American Spotted Salamander Ambystoma maculaturn 

(Amphibia: Ambystomatidae) 228 

(Continued on Outside Back Cover) 

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EDITORIAL BOARD 

2001 

WALTER A BOEGER 

WILLIAM F. FONT 

DONALD FORRESTER 

J. RALPH LICHTENFELS 

JOHN S. MACKIEWICZ 

BRENT NICKOL 

WILLIS A. REID, JR. AND JANET W. REID, EDITORS 

2002 

DANIEL R. BROOKS 

HIDEO HASEGAWA 

SHERMAN S. HENDRIX 

JAMES E. JOY 

DAVID MARCOGLIESE 

DANTE S. ZARLENGA 

© The Helminthological Society of Washington 2001 

ISSN 1525-2647 

2003 

ROY C. ANDERSON 

DIANE P. BARTON 

LYNN K. CARTA 

RALPH P. ECKERLIN 

FUAD M. NAHHAS 

DANNY B. PENCE 

This paper meets the requirements of ANSI/NISO Z39.48-1992 (Permanence of Paper). 


Comp. Parasitol. 

68(2), 2001, pp. 149-155 

Development and Specificity of Oochoristica javaensis (Eucestoda: 

Cyclophyllidea: Anoplocephalidae: Linstowiinae) 

CHARLES D. CRISCIONE' AND WILLIAM F. FONT2 

Department of Biological Sciences, Southeastern Louisiana University, Hammond, Louisiana 70402, U.S.A. 

(2 e-mail: wffont@selu.edu) 

ABSTRACT: Because assumptions of strict host specificity and geographic isolation apparently have been used 

as criteria in determining species of Oochoristica, studies were conducted to address the effects that these 

assumptions could have on resolving the taxonomy of Oochoristica. Experimental infections of native fence 

lizards, Sceloporus undulatus imdulatus, and Indo-Pacific geckos, Hemidactylus garnotii, demonstrated that the 

exotic tapeworm Oochoristica javaensis lacked host specificity. In addition, tapeworms with gravid proglottids, 

a stage of development that has not been previously reported for any species of Oochoristica, were obtained 

from both hosts. Evidence against the assumption of geographic isolation stems from the fact that lizard species 

known to harbor Oochoristica spp. have been introduced beyond their native ranges, and in some cases, these 

introductions predate the species descriptions. Lack of support for either assumption indicates a need for more 

rigorous analyses and experimentation to determine species of Oochoristica. 

KEY WORDS: Oochoristica javaensis, Cestoda, Hemidactylus turcicus, Hemidactylus garnotii, Sceloporus undulatus 

undulatus, Mediterranean geckos, Indo-Pacific geckos, fence lizards, development, host specificity, biogeography, 

taxonomy. 

Approximately 80 species have been described 

in the cosmopolitan genus Oochoristica 

Liihe, 1898 (Bursey and Goldberg, 1996a, b; 

Bursey et al., 1996, 1997; Brooks et al., 1999). 

These anoplocephalid cestodes predominantly 

parasitize lizards, but also snakes, turtles, and a 

few marsupials (Schmidt, 1986; Beveridge, 

1994). Development has been examined for 

Oochoristica vacuolata Hickman, 1954, Oochoristica 

osheroffi Meggitt, 1934, and Oochoristica 

anolis Hardwood, 1932 (Hickman, 1963; 

Widmer and Olsen, 1967; Conn, 1985). Although 

larval and adult development has been 

examined for 3 species of Oochoristica, only 

Conn (1985) tried to determine host specificity 

experimentally. In his experiments, however, he 

was unable to infect wall lizards, Podarcis muralis 

(Laurenti, 1768) and mice, Mm musculus 

Linnaeus, 1758. Curiously, no other attempts 

have been made to determine the specificity of 

a species of Oochoristica. This is unfortunate in 

that, as Brooks et al. (1999) pointed out, one of 

the major criteria used in resolving the taxonomy 

of Oochoristica has been the assumption of 

a high degree of host specificity exhibited by 

species in this genus. They also mentioned re- 

1 Corresponding author. Current address: Department 

of Zoology, 3029 Cordley Hall, Oregon State 

University, Corvallis, Oregon 97331, U.S.A. (e-mail: 

crischar@bcc.orst.edu). 

149 

striction to particular geographic regions as a 

criterion that has ostensibly been used in the past 

to identify species of Oochoristica. 

In a survey of helminths of the Mediterranean 

gecko, Hemidactylus turcicus (Linnaeus, 1758), 

from Louisiana, U.S.A., a species of Oochoristica 

was recovered (C. D. Criscione, unpublished 

data); however, there were difficulties in 

identifying this species. These problems were 

associated in part with the assumptions of strict 

host specificity and geographic isolation. It became 

apparent that in addition to the lack of 

specificity experiments, introduced and native 

host distributions were often ignored when identifying 

species of Oochoristica. Neither assumption 

has been properly addressed, and in order 

to validate the identification of any species of 

Oochoristica, these assumptions should be tested. 

In light of these problems with the taxonomy 

of Oochoristica, the primary objective of this 

study was to examine the development of Oochoristica 

javaensis Kennedy, Killick, and Beverley-Burton, 

1982, and to test the assumption 

of specificity via experimental infections. In addition, 

we comment on the assumption of geographic 

isolation. 

Materials and Methods 

To minimize variation among hosts, gravid proglottids 

of O. javaensis were obtained from 15 worms recovered 

from the small intestine of a single female 


150 COMPARATIVE PARASITOLOGY, 68(2), JULY 2001 

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CRISCIONE AND FONT—DEVELOPMENT OF OOCHORISTICA JAVAENSIS 151 

Table 2. Measurements of Oochoristica javaensis obtained from experimentally infected Hemidactylus 

garnotii (HEGA), and Sceloporus u. undulatus (SCUN) for day 105 postexposure; measurements in (xm 

unless noted otherwise. 

Variable 

Total 

Proglottid number 

Neck 

Scolex 

Sucker 

Immature proglottid 

Genital pore position|| 

Mature proglottid 

Cirrus sac 

Ovary 

Vitellaria 

Testis 

Testes number 

Gravid proglottid 

Oncosphere 

Hook 

L± (mm) 

W 

L (mm) 

W 

L 

W 

L 

W 

L 

W 

L 

W 

L 

W 

L 

W 

L 

W 

L 

W 

L (mm) 

W 

L 

L 

Sample 

size* 

5 

5 

5 

5 

5 

5 

5 

5 

15 

15 

15 

15 

15 

15 

15 

15 

15 

15 

15 

15 

15 

15 

15 

15 

15 

15 

15 

HEGA 

n = 5f 

SCUN 

Sample 

size n = 1 

61.6-86.9 (67.7 ± 48.3)§ 1 54.5 

109-144 (128 ± 5.7) 

1 140 

166-229 (191 ± 10.7) 1 221 

1.16-1.79 (1.52 ± 0.12) 1 1.12 

90-191 (152 ± 17.7) 1 218 

78-277 (140 ± 36.3) 1 164 

35-90 (61.8 ± 9.5) 1 82 

47-82 (66.4 ± 6.4) 1 105 

482-561 (520 ± 6.6) 

3 379-403 (390 ± 7.1) 

300-387 (348 ± 7) 

3 427-466 (443 ± 11.9) 

0.24-0.28 (0.27 ± 0.004) 3 0.24-0.28 (0.26 ± 0.01) 

593-695 (643 ± 9.1) 3 403-411 (406 ± 2.7) 

616-790 (712 ± 12.4) 3 648-695 (679 ± 15.7) 

43-51 (46.7 ± 0.47) 3 47-51 (49.7 ± 1.3) 

137-164 (146 ± 1.99) 3 109-121 (117 ± 4.0) 

265-351 (314 ± 6.0) 

3 168-211 (91 ± 12.6) 

195-265 (230 ± 5.1) 

3 133-187 (159 ± 15.7) 

137-191 (169 ± 4.5) 3 86-105 (96.3 ± 5.6) 

82-144 (118 ± 4.3) 3 82-101 (89.7 ±5.8) 

39_47 (43.5 ± 0.77) 3 35-39 (36.3 ± 1.3) 

35-55 (46.2 ± 1.3) 3 39-43(41.7 ± 1.3) 

23-35 (30.8 ± 0.78) 3 21-30 (26.3 ± 2.7) 

571-749 (629 ± 12.6) 3 433-473 (453 ± 11.6) 

1.50-2.28 (1.92 ± 0.05) 3 1.48-1.52 (1.49 ± 0.01) 

20-34 (25.9 ± 1.04) 3 22 (22 ± 0.0) 

16-28 (21.5 ± 0.79) 3 18-20(19.3 ± 0.67) 

10-12 (11.6 ± 0.21) 3 12 (12 ± 0.0) 

* Sample size refers to the total number of each character that was measured. 

t Number of tapeworms measured. 

± L = length, W = width. 

§ Range followed by mean ± 1 SE in parentheses. 

|| Genital pore position was calculated as a ratio of the position along the length of the mature proglottid from the anterior end 

(length to the center of the genital pore -^ length of proglottid). 

mediate host for the genus Oochoristica. Oochoristica 

javaensis became established in 7 individuals 

of the experimental definitive hosts. 

Successful infection occurred in only 1 of 16 H. 

turciciis; this Mediterranean gecko was examined 

on day 1 PE and had an intensity of 4. 

Oochoristica javaensis was recovered from 3 of 

10 H. garnotii on days 7, 28, and 105 PE and 

had intensities of 1, 7, and 6, respectively. Three 

of 5 S. u. undulatus were infected with 2, 3, and 

1 tapeworms on days 10, 30, and 105 PE, respectively. 

The 10 A. carolinensis and 5 R. 

sphenocephala were negative for infections. 

Measurements for specimens from experimental 

infections are in Tables 1 and 2. 

Proglottid formation did not occur prior to 

day 28 PE (Figs. 1 and 2). Terminal proglottids 

of tapeworms recovered on day 28 PE from H. 

garnotii had developing ovaries, testes, vitellaria, 

and cirrus sacs (Fig. 3). A median terminal 

excretory pore was present in each terminal proglottid, 

and in 1 worm there was a developed 

genital atrium. Terminal proglottid length ranged 

from 514 to 822 (mean = 659, SE = 38.7, n = 

7) and width from 174 to 277 (mean = 216, SE 

= 12.4, n — 7). Two of the worms from S. u. 

undulatus examined on day 30 PE also had developing 

reproductive organs (Fig. 4). One measured 

442 X 158 (L X W); the other had been 

torn. The third tapeworm recovered from day 30 

PE had been damaged in the mounting process; 

however, prior examination had shown that no 

more than 15 proglottids had formed, sexual primoridia 

had just begun to develop, and total 




length did not exceed 1.2 mm. By day 105 PE, 

development of O. javaensis progressed to strobilas 

with gravid proglottids in both H. garnotii 

(Fig. 5) and 5. u. undulatus (Fig. 6). Although 

it was possible that tapeworms from day 105 PE 

were natural infections because experimental definitive 

hosts were not laboratory-raised, prior 

fecal examinations were negative for all hosts 

used in experiments. 

Discussion 

Development of Oochoristica javaensis 

Prior to the present study, the most developed 

stage experimentally obtained for a species of 

Oochoristica was a terminal mature proglottid 

(Conn, 1985). Our experimental infections with 

O. javaensis were successful in obtaining gravid 

specimens. Susceptible definitive hosts for O. javaensis 

included H. turcicus, H. garnotii, and 5. 

u. undulatus; however, the fact that only 1 of 16 

control hosts, H. turcicus, became infected suggests 

that exposure techniques may have been 

flawed. Possible problems may have been the 

temperature at which experimental hosts were 

housed or the inoculation method. When dealing 

with small hosts, the stomach tube may not be 

the best method, and another, such as placing 

metacestodes in gel capsules, may prove to be 

more efficient. Despite the scarcity of infections, 

however, the developing worms obtained from 

H. garnotii and S. u. undulatus indicated a lack 

of specificity for O. javaensis. 

On day 105 PE, Conn (1985) recovered a single 

specimen of O. anolis from A. carolinensis 

that had mature proglottids with fully formed 

male and female reproductive systems. The terminal 

proglottid, however, still had a median excretory 

pore, suggesting that this specimen had 

not yet shed any proglottids. Specimens of O. 

javaensis from H. garnotii on day 28 PE also 

had median excretory pores in the terminal proglottids. 

Although most of the specimens that we 

recovered from day 28 PE did not have fully 

developed reproductive organs, their stage of development 

had greatly surpassed the develop- 


mental stage of O. anolis from day 28 PE (Conn, 

1985). Oochoristica anolis from green anoles on 

day 28 PE had just begun to form proglottids 

with genital anlages and had a maximum total 

length of 3.25 mm (Conn, 1985). Widmer and 

Olsen (1967) reported immature O. osheroffi 

with a maximum total length of 4.14 mm on day 

28 PE in the prairie rattlesnake, Crotalus viridis 

Rafinesque, 1818, but the mean total worm 

length from our study for day 28 PE was 8.32 

mm (Table 1). These comparisons suggest a 

more rapid development in the definitive host 

for O. javaensis than for O. anolis or O. osheroffi. 

However, small sample sizes in our study 

and infection techniques differing from previous 

life cycle studies of Oochoristica spp. prevented 

definitive comparison of developmental patterns 

among species. Likewise, the low number of infections 

precluded examination of host-induced 

variation for O. javaensis. Although development 

in S. u. undulatus appeared slightly slower 

than in H. garnotii up to day 30 PE (Table 1), 

measurements of gravid specimens from H. garnotii 

and S. u. undulatus on day 105 PE (Table 

2) may suggest plasticity for some characters. 

Host specificity and geographic isolation 

Results from our study experimentally demonstrated 

for the first time that a single species 

of Oochoristica can infect more than 1 species 

of host. Oochoristica javaensis infected hosts 

representing 2 unrelated lizard families, Phrynosomatidae 

for S. u. undulatus and Gekkonidae 

for H. garnotii (Estes et al., 1988; Pough et al., 

1998). It is not known if the ecology of either 

S. it. undulatus or H. garnotii would predispose 

natural populations of these hosts to the establishment 

of O. javaensis. Development in different 

hosts demonstrated via our laboratory experiments, 

however, raises questions with regard 

to the use of host specificity as a taxonomic criterion 

for species of Oochoristica. Specificity in 

the laboratory and in the field should be examined 

for other species of Oochoristica in light of 

these results for 2 reasons. First, lack of speci- 

Figures 1-6. Development of Oochoristica javaensis from Hemidactylus garnotii (HEGA) and Sceloporus 

u. undulatus (SCUN) at different days postexposure. Photomicrographs were taken with differential 

interference contrast. Bars = 200 jjim. 1. Day 7 in HEGA. 2. Day 10 in SCUN. 3, 4. Terminal proglottids 

from day 28 in HEGA and day 30 in SCUN, respectively. 5, 6. Mature proglottids from day 105 in HEGA 

and SCUN, respectively. CS = cirrus sac, GA = genital atrium, O = ovary, T = testis, and V = vitellaria. 



ficity means that tapeworms of the same species 

will be exposed to different environments in different 

species of hosts, thus presenting opportunities 

for host-induced variation. If variation is 

induced, then this may affect the current morphometrically 

based taxonomy of species of 

Oochoristica. Second, a better understanding of 

the degree to which species of Oochoristica can 

switch hosts is imperative in light of the conservation 

implications associated with introduced 

organisms and their parasites (see Barton, 

1997). 

Introduced lizards will have consequences not 

only for conservation, but also for parasite taxonomy. 

If in the past, species of Oochoristica 

have been transmitted with their exotic lizard 

hosts, then current assumptions of biogeographic 

isolation may be incorrect. This is not to say that 

there was never a biogeographic pattern that paralleled 

Oochoristica speciation, but especially 

because of anthropogenic effects, species of 

Oochoristica may have colonized new areas before 

many of them were ever described (see Bursey 

et al. [1996] for a list of authority dates). 

This possibility exists because records of some 

introduced geckos, i.e., H. turcicus in Florida 

(Stejneger, 1922) and Hemidactylus mabouia 

(Moreau de Jonnes, 1818) in South America and 

the Caribbean (Kluge, 1969), predate many 

Oochoristica species descriptions. 

It is interesting to note that several authors 

(Bursey and Goldberg, 1996b; Bursey et al., 

1996; Brooks et al., 1999) listed O. vanzolinii as 

a Neotropical species of Oochoristica from Brazil 

without regard to the fact that it was described 

from the introduced house gecko, H. mabouia 

(Rego and Oliveira Rodrigues, 1965). It 

has been hypothesized that H. mabouia naturally 

colonized the New World via rafting or was 

transported during the slave trades over 400 yr 

ago (Kluge, 1969). In either case, H. mabouia 

colonized the New World from Africa, thus presenting 

the opportunity for parasite transport. It 

is also interesting to note that in describing O. 

javaensis, Kennedy et al. (1982) listed O. vanzolinii 

as the species that most resembled their 

specimens. 

Brooks et al. (1999) stated that both assumptions, 

specificity and geographic isolation, were 

not evidence for new species, and suggested that 

in describing new species many morphological 

characters should be provided. We strongly support 

their contention; however, the full extent to 

Copyright © 2011, The Helminthological Society of Washington 

which certain features are plastic is still unknown 

for this genus. As indicated by Criscione 

and Font (2001), proglottid morphology of O. 

javaensis exhibited a high degree of plasticity 

that may have resulted from a crowding effect 

(Read, 1951). Providing more characters may alleviate 

some problems, but it will not solve the 

underlying difficulties associated with the taxonomy 

of Oochoristica. The morphologically 

based taxonomy of Oochoristica will only be 

validated upon experimentation establishing the 

variation of characters within the genus and/or 

the use of molecular data. 

Acknowledgments 

We express our gratitude to Dr. Richard Seigel, 

Caroline Kennedy, and Tom Lorenz for collecting 

the fence lizards and leopard frogs, and 

to Amanda Vincent and Jonathan Willis for their 

help with experiments and care of lizards. 

Thanks are also extended to Dr. Murray Kennedy 

and Dr. Bruce Conn for loaning us specimens 

of Oochoristica from their private collections, 

and to Judith Price at the Canadian Museum 

of Nature (CMNPA) and Pat Pilitt, Dr. Eric 

Hoberg, and Dr. J. Ralph Lichtenfels at the 

USNPC for their assistance and loan of specimens. 

Literature Cited 

Barton, D. P. 1997. Introduced animals and their parasites: 

the cane toad, Bufo marinus, in Australia. 

Australian Journal of Ecology 22:316-324. 

Beveridge, I. 1994. Family Anoplocephalidae Cholodkovsky, 

1902. Pages 315-366 in L. F. Khalil, A. 

Jones, and R. A. Bray, eds. Keys to the Cestode 

Parasites of Vertebrates. Commonwealth Agricultural 

Bureau, Wallingford, U.K. 

Brooks, D. R., G. Perez-Ponce de Leon, and L. 

Garcia-Prieto. 1999. Two new species of Oochoristica 

Liihe, 1898 (Eucestoda: Cyclophyllidea: 

Anoplocephalidae: Linstowiinae) parasitic in 

Ctenosaura spp. (Iguanidae) from Costa Rica and 

Mexico. Journal of Parasitology 85:893-897. 

Bursey, C. R., and S. R. Goldberg. 1996a. Oochoristica 

macallisteri sp. n. (Cyclophyllidea: Linstowiidae) 

from the side-blotched lizard, Ufa stansburiana 

(Sauria: Phrynosomatidae), from California, 

USA. Folia Parasitologica 43:293-296. 

, and . 1996b. Oochoristica maccoyi n. 

sp. (Cestoda: Linstowiidae) from Anolis gingivinus 

(Sauria: Polychortidae) collected in Anguilla, 

Lesser Antilles. Caribbean Journal of Science 32: 

390-394. 

-, and D. N. Woolery. 1996. Oochoristica 

piankai sp. n. (Cestoda: Linstowiidae) and 

other helminths of Moloch horridus (Sauria:

Agamidae) from Australia. Journal of the Helminthological 

Society of Washington 63:215-221. 

-, C. T. McAllister, and P. S. Freed. 1997. 

Oochoristica jonnesi sp. n. (Cyclophyllidea: Linstowiidae) 

from the house gecko, Hemidactylus 

mabouia (Sauria: Gekkonidae), from Cameroon. 


64:55-58. 

Conn, D. B. 1985. Life cycle and postembryonic development 

of Oochoristica anolis (Cyclophyllidea: 

Linstowiidae). Journal of Parasitology 71:10— 

16. 

Criscione, C. D., and W. F. Font. 2001. Artifactual 

and natural variation of Oochoristica javaensis'. 

statistical evaluation of in situ fixation. Comparative 

Parasitology 68:156-163. 

Estes, R., K. De Queiroz, and J. A. Gauthier. 1988. 

Phylogenetic relationships within Squamata. Pages 

119-281 in R. Estes and G. Pregill, eds. Phylogenetic 

Relationships of the Lizard Families, Essays 

Commemorating Charles L. Camp. Stanford 

University Press, Stanford, California, U.S.A. 

Hicknian, J. L. 1963. The biology of Oochoristica 

vacuolata Hickman (Cestoda). Papers and Proceedings 

of the Royal Society of Tasmania 97:81- 

104. 

Kennedy, M. J., L. M. Killick, and M. Beverley- 

Burton. 1982. Oochoristica javaensis n. sp. (Eu- 


Editors' Acknowledgments 

cestoda: Linstowiidae) from Gehyra mutilata and 

other gekkonid lizards (Lacertilia: Gekkonidae) 

from Java, Indonesia. Canadian Journal of Zoology 

60:2459-2463. 

Kluge, A. G. 1969. The evolution and geographical 

origin of the New World Hemidactylus mabouiabrookii 

complex (Gekkonidae, Sauria). Miscellaneous 

Publications, Museum of Zoology, University 

of Michigan 138:1-78. 

Pough, F. H., R. M. Andrews, J. E. Cadle, M. L. 

Crump, A. H. Savitzky, and K. D. Wells. 1998. 

Herpetology. Prentice-Hall, Upper Saddle River, 

New Jersey, U.S.A. 577 pp. 

Read, C. P. 1951. The "crowding effect" in tapeworm 

infections. Journal of Parasitology 37:174-178. 

Rego, A. A., and H. Oliveira Rodrigues. 1965. Sobre 

duas Oochoristica parasitas de lacertilios (Cestoda, 

Cyclophillidea). Revista Brasileira de Biologia 

25:59-65. 

Schmidt, G. D. 1986. Handbook of Tapeworm Identification. 

CRC Press, Boca Raton, Florida, U.S.A. 

675 pp. 

Stejneger, L. 1922. Two new geckos to the fauna of 

the United States. Copeia 1922:56. 

Widmer, E. A., and O. W. Olsen. 1967. The life 

history of Oochoristica osheroffi Meggitt, 1934 

(Cyclophyllidea: Anoplocephalidae). Journal of 

Parasitology 53:343-349. 

We acknowledge, with thanks, the following persons for providing valuable help and insights in reviewing 

manuscripts for Comparative Parasitology: Roy Anderson, Prema Arasu, Carter Atkinson, Scott 

Baird, James Baldwin, Diane Barton, Ian Beveridge, Reginald Blaylock, Walter Boeger, Rodney Bray, 

Daniel Brooks, Michael Burt, Goran Bylund, Charles Bursey, Ronald Campbell, Melanie Chapman, 

Anindo Choudhury, James Coggins, William Coil, David Cone, Bruce Conn, Thomas Cribb, John Cross, 

Lawrence Curtis, Murray Dailey, William Davidson, Emmet Dennis, Marie-Claude Durette-Desset, Ralph 

Eckerlin, Burton Endo, Alan Fedynich, Stephen Feist, Jacqueline Fernandez, David Fitch, Donald Forrester, 

Scott Gardner, Lynda Gibbons, David Gibson, Tim Goater, Stephen Goldberg, Robert Goldstein, Hideo 

Hasegawa, Richard Heard, Sherman Hendrix, John Hnida, Eric Hoberg, John Holmes, Patrick Hudson, 

David Huffman, Jean-Pierre Hugot, Kym Jacobson, Francisco Jimenez-Ruiz, James Joy, Michael Kent, 

Mike Kinsella, Greg Klassen, Ronald Ko, Delane Kritsky, Kevin Lafferty, Murray Lankester, Ralph Lichtenfels, 

Eugene Lyons, John Mackiewicz, David Marcogliese, Jean Mariaux, Chris McAllister, Donald 

McAlpine, Frantisek Moravec, Janice Moore, Patrick Muzzall, Fuad Nahhas, Robin Overstreet, Raphael 

Payne, Thomas Platt, Robert Poulin, Annie Prestwood, Robert Rausch, Amilcar Rego, Mark Rigby, John 

Riley, Klaus Rohde, Benjamin Sacks, William Samuel, Gerhard Schad, Thomas Scholz, Maria Sepiilveda, 

Scott Seville, Takeshi Shimazu, John Sullivan, Stephen Taft, Dennis Thoney, Tellervo Valtonen, William 

Wardle, Patricia Wilber, and Darwin Wittrock. 




Artifactual and Natural Variation of Oochoristica javaensis: 

Statistical Evaluation of In Situ Fixation 

CHARLES D. CRISCIONE' AND WILLIAM F. FONT2 

Department of Biological Sciences, Southeastern Louisiana University, Hammond, Louisiana 70402, U.S.A. 

(e-mail: 2wffont@selu.edu) 

ABSTRACT: Lack of knowledge of the extent of natural morphological variation can undermine proper taxonomic 

decisions. Confounding this problem is artifactual variation that arises from improper fixation techniques. For 

the morphologically based taxonomy of the cestode genus Oochoristica, little information exists on the plasticity 

of important taxonomic characters. In addition, paratypes of several species of Oochoristica are highly contracted 

and contorted. These paratypes were recovered from preserved hosts; thus, the tapeworms were killed and fixed 

inside the host (in situ fixation). Experiments demonstrated that in situ fixation of Oochoristica javaensis results 

in highly contracted specimens, and statistical comparisons between relaxed and in situ-fixed tapeworms revealed 

significant differences for proglottid measurements. Natural variation for the paratypes recovered from preserved 

hosts is likely misrepresented by the artificial variation induced by in situ fixation. Lastly, data from natural 

infections suggested that proglottid characters of O. javaensis are plastic and may be subject to crowding effects. 

KEY WORDS: Oochoristica javaensis, Cestoda, Hemidactylus turcicus, Mediterranean gecko, fixation techniques, 

crowding effects, morphological variation, taxonomy. 

The taxonomy of the cestode genus Oochoristica 

Liihe, 1898, has been based solely on morphology 

without knowledge of the extent of natural 

intraspecific morphological variation. Parasite 

morphological variation may be the result of 

genetic determinants, host-induced effects, parasite 

intensity effects, or external habitat influences. 

Stunkard (1957) and Haley (1962) discussed 

the importance of environmental and 

host-induced variation for the systematics of helminth 

parasites, citing such factors as different 

host species, host age, host diet, or infection intensity 

as causes of variation. They also emphasized 

the need to assess experimentally the stability 

of taxonomic characters when identifying 

a species. 

In addition to the lack of knowledge on intraspecific 

variation, natural variation of some species 

of Oochoristica may be masked by artificial 

morphological variation induced by fixation 

techniques. Several paratype specimens of 

Oochoristica examined from the U.S. National 

Parasite Collection (USNPC), Beltsville, Maryland, 

U.S.A. were highly contracted and contorted. 

Examination of the respective species descriptions 

revealed that these paratypes (listed 

below) were obtained from formalin-fixed hosts. 


of Zoology, 3029 Cordley Hall, Oregon State 

University, Corvallis, Oregon 97331, U.S.A. (e-mail: 

crischar@bcc.orst.edu). 

156 


That is, they were removed from host specimens 

deposited in museum collections without regard 

to the effects of host fixation on internal parasites. 

Bakke (1988) and others have qualitatively 

illustrated the distorting effects of improper fixation 

techniques on the morphology of soft-bodied 

helminths, but comparisons of fixation techniques 

have not been tested statistically to examine 

for quantitative differences in the measurements 

of important taxonomic characters. 

The purpose of our report was to provide a 

quantitative assessment of the artifactual morphological 

variation induced by killing and fixing 

tapeworms within a host, i.e., in situ fixation. 

In order to address the effects that improper fixation 

methods may have on the morphologically 

based taxonomy of Oochoristica, statistical 

comparisons of in situ-fixed tapeworms to ones 

that were collected alive and killed in a relaxed 

state were conducted with specimens of Oochoristica 

javaensis Kennedy, Killick, and Beverley-Burton, 

1982. In addition, we provide data 

regarding the effects of intensity on O. javaensis 

morphology. 


Fixation experiments 

To mimic lizard fixation techniques used for museum 

collections, 15 Mediterranean geckos, Hemidactylus 

turcicus (Linnaeus, 1758), were collected from the 

campus of Louisiana State University (LSU), Baton 

Rouge, Louisiana, U.S.A. (30°24.92'N; 91010.81'W), 

where they were known to have a high prevalence of

infection with O. javaensis (C. D. Criscione, unpublished 

data). Geckos were killed using an overdose of 

ether and immediately fixed via subcutaneous, oral 

cavity, and body cavity injections of unheated 10% 

formalin. Oral cavity injections insured that the tapeworms 

were killed immediately. After 6 days in 10% 

formalin, geckos were soaked in water for 24 hr to 

remove the formalin. Geckos were then transferred to 

70% ethanol for storage until dissection 4 days later. 

In situ-fixed tapeworms recovered upon necropsy were 

stored in 70% ethanol, stained in Semichon's acetocarmine, 

dehydrated in ethanol, cleared in xylene, and 

mounted in Canada balsam. Comparisons were made 

with relaxed tapeworms that were killed with hot water 

(90°C) and fixed and stored in alcohol-formalin-acetic 

acid solution (AFA). Relaxed worms were obtained in 

a helminth survey of//, turcicus from LSU in the summer 

of 1998 (C. D. Criscione, unpublished data); staining 

and mounting techniques were the same as for the 

in situ-fixed specimens. 

Quantitative analyses included measurements of mature 

proglottids from 5 relaxed and 5 in situ-fixed specimens 

of O. javaensis. Three mature proglottids, located 

just anterior to the first proglottid displaying evidence 

of egg production, were selected from each individual. 

Length and width were measured for each 

mature proglottid and for the ovary, vitellaria, and 1 

testis within each proglottid. One testis was randomly 

selected from each proglottid, ovary length was measured 

for the ovary lobe opposite the genital atrium, 

and ovary width was measured across both lobes. Although 

multiple testes are present within a single proglottid, 

only 1 was chosen in order to facilitate the use 

of appropriate statistical tests. In order to test for in 

situ-fixation effects, a nested ANOVA design was used 

to control the pseudoreplication of measuring 3 proglottids 

from 1 tapeworm. Tapeworms nested within 

type of fixation constituted the experimental units, i.e., 

true replicates, with the error term being the proglottids, 

i.e., pseudoreplicates, nested within individual 

tapeworms. Principal components analysis (PCA) with 

Varimax rotation was used as a data reduction technique 

and to examine latent relationships among the 

variables. A variable was considered to load on a factor 

if its correlation to the factor was >|0.5| (Hair et 

al., 1999). The resulting factors with their standardized 

factor scores were then tested for differences between 

relaxed and in situ-fixed tapeworms in the nested design. 

Statistical significance was determined at P < 

0.05. 

Analysis of natural morphological variation 

The analysis of intensity effects on the morphology 

of O. javaensis included 5 tapeworms from each of 3 

Mediterranean geckos that were naturally infected with 

15, 28, and 64 tapeworms. Criteria and morphological 

characters used for measurements were the same as 

those used in the fixation experiments. Experimental 

design using factor scores from a PCA with Varimax 

rotation was also the same, in that a nested ANOVA 

was used to test for intensity effects. A priori contrasts 

of 15 versus 28 and 28 versus 64 were computed. In 

order to conduct this analysis, the 5 tapeworms from 

CRISCIONE AND FONT—VARIATION OF OOCHORISTICA JAVAENSIS 157 

each intensity level were treated as true replicates, 

when in fact they were pseudoreplicates. 

In addition, 10 relaxed tapeworms that were killed 

with hot water (90°C) were selected to provide measurements 

representative of O. javaensis recovered in 

a helminth survey of H. turcicus (C. D. Criscione, unpublished 

data). This representative data set included 

specimens from geckos with intensities ranging from 

1 to 64, and had at least 1 tapeworm from each of 5 

collection locations in southeastern Louisiana, U.S.A. 

[Bayou Segnette State Park in Westwego (29°53.18'N; 

90°09.80'W); Fairview-Riverside State Park in Madisonville 

(30°24.55'N; 90°08.41'W); a residential 

neighborhood in Metairie (30°00.76'N; 90°08.90'W); 

Southeastern Louisiana University in Hammond 

(30°30.67'N; 90°27.98'W); and LSU]. PCA with Varimax 

rotation was applied to this data set to examine 

for latent relationships among the same variables used 

in the fixation and intensity analyses. 

Specimens examined 

Museum specimens examined from the USNPC and 

the Canadian Museum of Nature (CMNPA), Ottawa, 

Ontario, Canada included the following: O. javaensis, 

2 paratypes (CMNPA nos. 1982-0693, 1982-0695); 

Oochoristica anolis Hardwood, 1932, 1 voucher 

(USNPC no. 75748) and the holotype (USNPC no. 

30898); Oochoristica bezyi Bursey and Goldberg, 

1992, 2 paratypes (USNPC no. 81874); Oochoristica 

bresslaui Fuhrmann, 1927, 1 voucher (USNPC no. 

89087); Oochoristica chinensis Jensen, Schmidt, and 

Kuntz, 1983, 2 paratypes (USNPC no. 077168); Oochoristica 

islandensis Bursey and Goldberg, 1992, 1 

paratype (USNPC no. 82225); Oochoristica macallisteri 

Bursey and Goldberg, 1996, 1 voucher (USNPC 

no. 89267) and 2 paratypes (USNPC no. 86196); Oochoristica 

mccoyi Bursey and Goldberg, 1996, 2 

vouchers (USNPC nos. 85403, 85408) and 1 paratype 

(USNPC no. 86343); Oochoristica novaezealandae 

Schmidt and Allison, 1985, 5 paratypes (USNPC no. 

78407); Oochoristica osheroffi Meggitt, 1934, 1 

voucher (USNPC no. 80433); Oochoristica parvula 

(Stunkard, 1938), 3 vouchers (USNPC no. 84397); 

Oochoristica piankai Bursey, Goldberg, and Woolery, 

1996, 1 voucher (USNPC no. 88189) and 1 paratype 

(USNPC no. 84589); Oochoristica scelopori Voge and 

Fox, 1950, 2 vouchers (USNPC nos. 84234, 87529). 

We deposited voucher specimens of O. javaensis from 

H. turcicus in the USNPC (nos. 90344-90348). 

Results 

For the fixation analysis, PCA revealed 3 latent 

variables from the 8 measured (Table 1). 

Examination of the variable factor loadings revealed 

that factor 1 consisted of all the vertical 

measurements, while factors 2 and 3 were characterized 

by horizontal measures. Factors 1, 2, 

and 3 were renamed vertical, horizontal-1, and 

horizontal-2, respectively. All 3 factors showed 

significant worm-to-worm variation within each 

type of fixation (Fvcrtical = 9.20, FhorizontaM = 



Table 1. Oochoristica javaensis: variable factor loadings, factor eigenvalues, and percent total variance 

accounted for by each factor from the Varimax rotated correlation matrix of the fixation data set. 

Testis length 

Ovary length 

Vitellaria length 

Proglottid length 

Ovary width 

Proglottid width 

Testis width 

Vitellaria width 

Eigenvalues 

Percent of total variance explained by the factor 

* Bold print shows loadings where variable loaded onto factor. 

42.20, Fhorizontal_2 = 5.31, df = 8, 20, P < 0.001); 

thus, the fixation main effect was tested with the 

mean-square values of the subgroups, tapeworms 

nested within fixation method. The vertical 

factor showed a significant effect between in 

situ-fixed and relaxed tapeworms (Fig. 1) (F]>8 = 

11.927, P = 0.009); however, neither horizontal 

factor was significant. Table 2 provides the raw 

measurements of the variables used in the analysis. 

PCA revealed that the 8 variables used in the 

intensity data set constituted only 1 factor (Table 

•3 0 

CJ 

1 -1 

-2 

In situ Relaxed 

Form of Fixation 

Figure 1. Plot of the mean factor scores and 

95% confidence intervals for the vertical factor of 

relaxed and in situ-fixed specimens of Oochoristica 

javaensis. 

Factor 1 

(vertical) 

0.917* 

0.875 

0.863 

0.805 

-0.203 

-0.236 

-0.092 

0.463 

3.320 

41.495 

Varimax rotated loading matrix 


Factor 2 

(horizontal- 1) 

-0.112 

-0.176 

-0.237 

-0.497 

0.912 

0.869 

0.469 

0.177 

2.184 

27.296 

Factor 3 

(horiz.ontal-2) 

0.209 

0.209 

0.262 

0.023 

-0.082 

-0.096 

-0.835 

0.793 

1.498 

18.723 

3), thus showing that all 8 characters from relaxed 

specimens varied together. Even though 

there was significant worm-to-worm variation 

for this factor (F12 30 = 12.62, P < 0.001), there 

was still a significant intensity effect (F2 12 = 

13.42, P < 0.001) (Fig. 2). The a priori contrast 

between 15 and 28 was significant (F, 12 = 

10.58, P = 0.007), but 28 versus 64 was not. 

Figure 3A-C displays mature proglottid variation 

for tapeworms recovered from intensities of 

15, 28, and 64, and Table 4 gives the raw measurements 

of the variables. Measurements of 10 

O. javaensis tapeworms from Louisiana are given 

in Table 5. Based on results from the fixation 

experiments, only tapeworms exhibiting little to 

no wrinkling (i.e., contraction) were used to provide 

the representative measurements of O. javaensis 

collected in our survey. The fact that the 

8 variables in the representative data emerged as 

only 1 factor from the PCA (Table 3) again demonstrated 

that all 8 characters from relaxed specimens 

varied together. 

Discussion 

Variation resulting from different fixation 

techniques or conditions only confounds taxonomic 

problems in which the range of natural 

morphological variation is not known. Experimental 

data revealed 2 quantitative problems 

with using in situ-fixed tapeworms. The first was 

that a significant reduction in the vertical factor 

without a significant change in horizontal factors 

produced an accordion effect. High vertical factor 

scores were determined by large length measurements 

of the proglottid and its ovary, vitel-


Table 2. Measurements of mature proglottids from specimens of Oochoristica javaensis fixed in a relaxed 

state and specimens fixed in situ; all measurements are in jxm. 



Ovary width 

Ovary length 



Testis width 

Testis length 

Sample size* 

15 

15 

15 

15 

15 

15 

15 

15 

Relaxed 

n = 5t 

482-648 (589 ± 15.8)± 

356-640 (493 ± 25.8) 

246-35 1 (288 ± 8.06) 

152-238 (201 ± 6.63) 

125-195 (161 ± 5.11) 

90-137 (104 ± 3.83) 

31-43 (38.7 ± 0.83) 

27-47 (39.5 ± 1.23) 


t Number of tapeworms used in measurements. 

:j: Range followed by mean ± 1 SE in parentheses. 

laria, and testes (Table 2); thus, relaxed tapeworms 

had a higher mean factor score (Fig. 1). 

Contraction from in situ fixation resulted in mature 

proglottids wider than long (Fig. 3D-F), but 

completely relaxed tapeworms from hot-fixed 

specimens yielded mature proglottids longer 

than wide (Fig. 3A-C). 

The second quantitative effect pertained to the 

correlative relationships among the mature proglottid 

characters and was revealed by the PCA 

itself. If only relaxed specimens are incorporated 

into the PCA, i.e., the intensity and representative 

data sets, all 8 characters vary together as 

Table 3. Oochoristica javaensis: variable factor 

loadings, factor eigenvalues, and percent total variance 

accounted for by each factor from the correlation 

matrix of the intensity data set and representative 

data set. 

Ovary width 


Ovary length 



Testis width 

Testis length 


Eigenvalues 

Percentage of total variance 

explained by the factor 

Loading matrices ;_ 

for the O 

Intensity 

data set 

Factor 1 

0.924* 

0.921 

0.916 

0.893 

0.866 

0.841 

0.753 

0.751 

5.929 

74.116 

Representative 

Q 

data set "g 

Factor 1 W 

r~ 

0.909 a 

0.854 ^ 

0.814 ^ 

0.851 ^ 

0.784 

0.825 

0.749 

0.443 

4.996 

62.445 

Bold print shows loadings where variable loaded onto factor. 

In situ-fixed 

n = 5 

403-845 (635 

213-490 (312 

261-355 (310 

109-183 (144 

113-148 (130 

43-94 (70.5 

35-47 (43.5 

20-35 (27.1 

± 34.1) 

± 23.0) 

± 7.01) 

± 5.69) 

± 2.86) 

± 4.11) 

± 0.95) 

± 1.07) 

1 factor (Table 3); but when in situ-fixed tapeworms 

are incorporated into the PCA, i.e., the 

fixation data set, vertical measurements become 

independent of horizontal measurements (Table 

1). Contraction of the in situ-fixed tapeworms 

altered the correlative nature of the 8 variables 

and divided 1 factor into 3 factors. 

Contraction of helminth parasites resulting 

from improper fixation has been documented 

many times in the parasite literature (Bakke, 

1988); however, our study may be the first to 

quantify the effects of different forms of fixation 

and to analyze the data statistically. The empirical 

evidence provided in the current study not 

only supported the conclusions of Bakke (1988) 

1.5 

0.5 

-0.5 

-1.5 

15 28 64 

Intensity of Infection 

Figure 2. Plot of the mean factor scores and 

95% confidence intervals for tapeworms collected 

at intensities of 15, 28, and 64. 



Figure 3. Mature proglottid variation of Oochoristica javaensis from Hemidactylus turcicus. Photomicrographs 

were taken with differential interference contrast. A-C. Natural variation of specimens from 

intensities of 15, 28, and 64, respectively; all were fixed in a relaxed state. D-F. Artificial variation showing 

contraction that resulted from in situ fixation. Bars = 200 u.m. CS = cirrus sac, GA = genital atrium, O 

= ovary, T = testis, and V = vitellaria. 

Table 4. Mature proglottid measurements of Oochoristica javaensis from Hemidactylus turcicus with intensities 

of 15, 28, and 64; all measurements are in |xm. 

Level of 

intensity 



Ovary width 

Ovary length 



Testis width 

Testis length 

Sample 

size* 

15 

15 

15 

15 

15 

15 

15 

15 

n 

15 

= 5t 

506-616 (560 ± 7.06):j: 

545-75 1 (624 ± 14.5) 

238-293 (268 ± 3.89) 

156-226 (189 ± 4.48) 

129-203 (161 ± 5.78) 

74-137 (105 ± 4.95) 

39-47 (42.7 ± 0.61) 

39-51 (43 d: 

0.87) 

28 

n = 5 

387-450 (409 

395-553 (470 

195-254 (225 

125-179 (160 

86-133 (117 

70-113 (90.3 


t Number of tapeworms used in measurements. 

± Range followed by mean ± 1 SE in parentheses. 

± 5) 

± 14.7) 

± 4.85) 

± 3.83) 

± 3.37) 

± 3.3) 

35-43 (39 ± 0.78) 

31-47 (39 ± 1.03) 


n 

269-545 

419-545 

121-281 

78-183 

59-117 

43-101 

27-43 

27-47 

64 

= 5 

(387 

(463 

(198 

(126 

± 31.7) 

± 8.7) 

± 14.2) 

± 7.9) 

(90.4 ± 5.19) 

(73.5 ± 4.91) 

(35 ± 1.29) 

(36.6 ± 1.4)


Table 5. Measurements of Oochoristica javaensis from naturally infected Hemidactylus turcicus in southeastern 

Louisiana; measurements in jxin unless noted otherwise. 

Variable 

Total 

Proglottid number 

Neck 

Scolex 

Sucker 

Immature proglottid 

Genital pore position* 

Mature proglottid 

Cirrus sac 

Ovary 

Vitellaria 

Testis 

Testes number 

Gravid proglottid 

Oncosphere 

Hook 

Li (mm) 

W 

L (mm) 

W 

L 

W 

L 

W 

L 

W 

L 

W 

L 

W 

L 

W 

L 

W 

L 

W 

L (mm) 

W 

L 

L 

Sample size* 

10 

10 

10 

10 

10 

10 

10 

10 

30 

30 

30 

30 

30 

30 

30 

30 

30 

30 

30 

30 

30 

30 

30 

30 

30 

30 

30 

n = 10t 

22.2-105 (53.4 ± 7.4)§,|| 

86-164 (131 ± 7.8)|| 

158-237 (205 ± 8.1)|| 

1.12-1.58 (1.38 ± 0.05)|| 

148-246 (195 ± 9.1)|| 

98-183 (140 ± 8.9) 

51-90 (74.1 ± 3.9)|| 

62-117 (89 ± 5.3)|| 

261-506 (408 ± 14.5) 

237-371 (297 ± 7.6) 

0.24-0.33 (0.28 ± 0.004)|| 

277-648 (490 ± 19.5)|| 

395-751 (509 ± 17.8)|| 

43-55 (47.7 ± 0.58) 

86-144 (116 ± 2.8)|| 

133-390 (268 ± 11.8)|| 

78-316 (184 ± 0.6)|| 

62-269 (144 ± 8.9)|| 

43-221 (106 ± 6.6)|| 

27-51 (40.2 ± 0.98)|| 

31-47 (40.9 ± 0.9) 

17-46 (26.6 ± 1.4)|| 

158-650 (492 ± 29.4)|| 

0.85-1.99 (1.26 ± 0.05)|| 

20-34 (25.3 ± 0.69)|| 

18-28 (23.2 ± 0.52)|| 

8-12 (11.5 ± 0.18)|| 


t Number of tapeworms measured. 

± L = length, W = width. 

§ Range followed by mean ± 1 SE in parentheses. 

|| Indicates that the range of the character extends values reported in the original description (Kennedy et al., 1982). 

# Genital pore position was calculated as a ratio of the position along the length of the mature proglottid from the anterior 

end (length to the center of the genital pore + length of proglottid). 

but also quantitatively demonstrated the misleading 

representation of morphological characters 

used in the taxonomy of Oochoristica resulting 

from in situ fixation. We believe that our 

more rigorous analysis is important because, despite 

the elegant studies of Bakke (1988) and 

previous workers, the practice of describing improperly 

fixed specimens is still widespread 

among parasite taxonomists. 

In addition to the quantitative changes resulting 

from in situ fixation, 2 qualitative effects further 

demonstrated the inappropriateness of using 

in situ-fixed specimens in species descriptions. 

Distortion of the scolex (Fig. 4A, B) and proglottids 

(Fig. 4C, D) prevented accurate measurements 

of these characters. This is not to say 

that every scolex and mature proglottid fixed in 

situ will be rendered useless for species identi- 

fication, but the number of appropriate characters 

available for analysis will be greatly reduced. 

As seen in Figure 4A-D, one would have 

difficulty in finding a true scolex width, and contortion 

in the strobila would limit the number of 

proglottids suitable for examination. 

Species descriptions based on contracted or 

disfigured specimens will misrepresent the true 

natural variation by decreasing the means of vertical 

characters and artificially inflating the dispersion 

of measurements if used in conjunction 

with relaxed specimens. Such may be the case 

with several paratype (O. bezyi, O. islandensis, 

O. macallisteri, O. mccoyi, O. piankai} and 

voucher (O. mccoyi, O. par~vula, O. piankai, O. 

scelopori) specimens examined in our study. 

These specimens may represent true species, but 

their reported natural variation is more than like- 



Figure 4. Comparisons of relaxed and in situ-fixed specimens of Oochoristicajavaensis. A-D. Distorting 

effects of in situ fixation on scolices and proglottids. E-H. Relaxed specimens. Bars = 200 (xm for A, B, 

E, and F (differential interference contrast). Bars = 500 (Jim for C, D, G, and H (brightfield). 

ly masked within the artificial variation induced 

by in situ fixation. Characters that do not reflect 

their natural variation should not be used to describe 

species. Oochoristica spp. recovered from 

fixed museum hosts may provide historical 

abundance data, but identification of these specimens 

should be made with extreme caution, and 

ideally, in conjunction with specimens fixed appropriately. 

Intensity effects were examined because a 

crowding effect has been documented as a cause 

of variation in the size of tapeworms (Read, 

1951), and because of the occurrence of different 

intensities in naturally infected H. turcicus. 

PCA for the intensity data set produced 1 factor 

in which all 8 variables loaded high (Table 3); 

therefore, individual proglottids with large measurement 

values received high factor scores. Although 

not quantified, there was no apparent 

crowding effect observed for natural infections 

with intensities between 1 and 15. Tapeworms 

from an intensity of 15 had significantly greater 

factor scores than specimens from 28 and 64 

(Fig. 2), thus indicating that crowding reduces 

the size of the respective morphological characters 

(Table 4) in O. javaensis. Brooks and 


Mayes (1976) reported similar crowding effects 

for O. bivitellobata recovered from the prairie 

racerunner, Cnemidophorus sexlineatus Lowe, 

1966. Their results and ours, however, should be 

considered only preliminary for 2 reasons. First, 

data were obtained from natural infections; thus 

other factors that induce variation were not controlled. 

Second, the intensity levels were not 

replicated. Ideally, one would wish to sample 

tapeworms from multiple hosts harboring all 

possible intensity levels. Both reports, however, 

suggest that morphological characters for species 

of Oochoristica can be variable and may be 

subject to intensity levels. 

Measurements of the 10 0. javaensis specimens 

given in Table 5 extend the ranges of several 

characters provided in the original description 

of O. javaensis (Kennedy et al., 1982). 

These measurements were based on specimens 

with little or no contraction and are provided to 

give a representation of O. javaensis collected 

from H. turcicus in southeastern Louisiana. 

Means of several characters (Table 5) do not 

match those provided by Kennedy et al. (1982); 

however, based on the lack of host specificity 

displayed in laboratory experiments (Criscione

and Font, 2001), the indication of plasticity in 

morphological characters (Table 4), and the examination 

of O. javaensis paratypes, it was determined 

that the specimens from H. turcicus in 

southeastern Louisiana were O. javaensis. 

In summary, statistical analyses demonstrated 

that measurements of in situ-fixed tapeworms, 

i.e., specimens recovered from preserved hosts, 

distorted the true natural variation of O. javaensis. 

The intraspecific variation of several species 

of Oochoristica may be misrepresented because 

they were described from highly contracted, in 

situ-fixed specimens. Additionally, morphological 

characters used in the taxonomy of Oochoristica 

have not been examined for their stability 

when exposed to different environmental 

or host-induced conditions. Our analyses indicated 

that proglottid morphology was highly 

variable and that this plasticity may have resulted 

from crowding. 


We extend our thanks to Dr. Murray Kennedy 

and Dr. Bruce Conn for loaning us specimens of 

Oochoristica from their private collections, and to 

Judith Price at the CMNPA and Pat Pilitt, Dr. Eric 

Hoberg, and Dr. Ralph Lichtenfels at the USNPC 

for their assistance and loan of specimens. 


Notice of Dues Increase 


Bakke, T. A. 1988. Morphology of adult Phyllodistomum 

umblae (Fabricius) (Platyhelminthes, Gorgoderidae): 

the effect of preparation, killing and 

fixation procedures. Zoologica Scripta 17:1-13. 

Brooks, D. R., and M. A. Mayes. 1976. Morphological 

variation in natural infections of Oochoristica 

bivitellobata Loewen, 1940 (Cestoidea: Anoplocephalidae). 

Transactions of the Nebraska Academy 

of Sciences 3:20-21. 

Criscione, C. D., and W. F. Font. 2001. Development 

and specificity of Oochoristica javaensis (Eucestoda: 

Cyclophyllidea: Anoplocephalidae: Linstowiinae). 

Comparative Parasitology 68:149-155. 

Hair, J. F., Jr., R. E. Anderson, R. L. Tatham, and 

W. C. Black. 1999. Multivariate Data Analysis, 

5th ed. Prentice-Hall, Upper Saddle River, New 

Jersey, U.S.A. 730 pp. 

Haley, A. J. 1962. Role of host relationships in the 

systematics of helminth parasites. Journal of Parasitology 

48:671-678. 

Kennedy, M. J., L. M. Killick, and M. Beverley- 

Burton. 1982. Oochoristica javaensis n. sp. (Eucestoda: 

Linstowiidae) from Gehyra mutilata and 

other gekkonid lizards (Lacertilia: Gekkonidae) 

from Java, Indonesia. Canadian Journal of Zoology 

60:2459-2463. 

Read, C. P. 1951. The "crowding effect" in tapeworm 

infections. Journal of Parasitology 37:174-178. 

Stunkard, H. W. 1957. Intraspecific variation in parasitic 

flatworms. Systematic Zoology 6:7-18. 

At the January 2001 meeting of the Helminthological Society of Washington, the Society voted to 

increase the membership dues to US$30.00 for individual U.S. members and to US$33.00 for individual 

foreign members. Institution subscription rates will increase as follows: US$55.00 (U.S.A.), US$57.00 

(Canada and Mexico), US$60.00 (all other countries). This is the first dues increase in six years and is 

necessitated by increased management costs to the Society. The increases will take effect in January, 2002. 




Seasonal Occurrence and Community Structure of Helminth 

Parasites in Green Frogs, Rana clamitans melanota, from 

Southeastern Wisconsin, U.S.A. 

MATTHEW G. BOLEK' AND JAMES R. COGGINS 

Department of Biological Sciences, University of Wisconsin-Milwaukee, Milwaukee, Wisconsin 53201, 

U.S.A. (e-mail: coggins@csd.uwm.edu) 

ABSTRACT: From April to October 1996, 75 green frogs, Rana clamitans melanota, were collected from Waukesha 

County, Wisconsin, U.S.A. and examined for helminth parasites. Seventy-one (94%) of 75 green frogs 

were infected with 1 or more helminth species. The component community consisted of 12 species: 5 trematodes, 

2 cestodes, and 5 nematodes. Approximately 2,790 (72%) trematodes, 447 (11.5%) cestodes, and 636 (16.5%) 

nematodes were found. A significant correlation existed between wet weight and helminth species richness. 

Helminth populations and communities were seasonally variable and/or did not show significant differences 

during the year. Haematoloechus varioplexus showed seasonal variation in size during the year that was related 

to recruitment period. The helminth fauna of green frogs was depauperate and dominated by indirect-life-cycle 

parasites. Host diet and aquatic habitat were important in the transmission dynamics of these species. Host size, 

sex, and time of collection were also important factors in structuring helminth communities of green frogs and 

may mask any simple explanations. 

KEY WORDS: Rana clamitans, Haematoloechus varioplexus, Halipegus eccentricus, Glyptheltnins quieta, Cosmocercoides 

sp., Oswaldocruzia pipiens, Waltonella sp., Mesocestoides sp., metacercariae, Trematoda, Nematoda, 

Cestoda, Amphibia, seasonal study, Wisconsin, U.S.A. 

Studies by Kennedy et al. (1986) on freshwater 

fish, birds, and a mammal have developed 

predictions in determining helminth community 

structure, particularly that ectotherm and endotherm 

helminth communities are fundamentally 

different: the former are species poor and noninteractive, 

while the latter are diverse and interactive. 

A review by Aho (1990) indicated that 

helminth communities of amphibians are highly 

variable, depauperate, and noninteractive in 

structure, but there is a need to examine and 

reexamine more species from different locations. 

To date there are few studies that utilized helminth 

community measures in amphibian hosts 

(Goater et al., 1987; Aho, 1990; Muzzall, 1991a, 

b; Goldberg et al., 1995; Barton, 1996; Yoder 

and Coggins, 1996; McAlpine, 1997; Bolek and 

Coggins, 2000). A number of helminth surveys 

of green frogs, Rana clamitans melanota Rafinesque, 

1820, have been published (Campbell, 

1968; Williams and Taft, 1980; Coggins and Sajdak, 

1982; McAlpine and Burt, 1998), but there 

are few studies on the helminth infracommunity 

and component community structure of this spe- 

1 Corresponding author. Current address: School of 

Biological Sciences, University of Nebraska-Lincoln, 

Lincoln, Nebraska 68588, U.S.A. (e-mail: mbolek@ 

unlserve.unl.edu). 

164 


cies (Muzzall, 199la; McAlpine, 1997), and 

none that incorporated a seasonal component. 

Green frogs are large, semiaquatic frogs inhabiting 

freshwater ponds, lakes, swamps, and 

slow-moving streams in North America. They 

spend most of their time around the water's 

edge. They occur from Newfoundland to western 

Ontario, and south to eastern Oklahoma, 

southern Illinois, northern Georgia, and eastern 

North Carolina. In Wisconsin, these frogs overwinter 

buried in the mud and are active from 

early April through October (Vogt, 1981). Green 

frogs are largely sit-and-wait gape-limited predators, 

feeding on any accessible prey of appropriate 

size, including aerial, aquatic, and terrestrial 

invertebrates, primarily insects (Scale, 

1987; Werner et al., 1995). Here we report on 

the seasonal helminth community structure of 

green frogs from southeastern Wisconsin. Specifically, 

we were interested in how host habitat, 

age and/or size, diet, sex, and seasonality were 

important in determining helminth populations 

and communities of green frogs. 


A total of 75 green frogs, R. clamitans melanota, 

was collected from April to October 1996 at a small 

spring-fed permanent pond located at the Carroll College 

field station in Waukesha County, Wisconsin,

U.S.A. (42°59'N; 88°21'W). Ten to 15 frogs were collected 

monthly around the periphery of the pond by a 

dip-net. Animals were placed in plastic containers, 

transported to the laboratory, stored at 4°C, and euthanized 

in MS 222 (ethyl m-aminoben/.oate methane sulfonic 

acid) within 72 hr of capture. Snout-vent length 

(SVL) and wet weight (WW) were recorded for each 

individual. Frogs were individually toe-clipped and 

frozen. At necropsy, the digestive tracts, limbs and 

body wall musculature, and internal organs were examined 

for helminth parasites. Each organ was placed 

individually in a petri dish and examined under a stereomicroscope. 

The body cavities were rinsed with distilled 

water into a petri dish and the contents examined. 

All individuals were sexed by gonad inspection during 

necropsy. Worms were removed and fixed in alcohol— 

formaldehyde-acetic acid or formalin. Trematodes and 

cestodes were stained with acetocarmine, dehydrated 

in a graded ethanol series, cleared in xylene, and 

mounted in Canada balsam. Nematodes were dehydrated 

to 70% ethanol, cleared in glycerol, and identified 

as temporary mounts. Prevalence, mean intensity, 

and abundance are according to Bush et al. (1997). 

Mean intensity was not calculated for the unidentified 

kidney metacercariae because they could not be counted 

accurately, and overall abundance was reported as 

an estimate of encysted metacercariae counted on the 

surface of the kidneys. Mean helminth species richness 

is the sum of helminth species per individual frog, including 

noninfected individuals, divided by the total 

sample size. All values are reported as the mean ± 1 

standard deviation. Undigested stomach contents were 

identified to class or order following Borror et al. 

(1989). Stomach contents are reported as a percent = 

the number of prey items in a given class or order, 

divided by the total number of prey items recovered 

X 100. Voucher specimens have been deposited in the 

H. W. Manter Helminth Collection, University of Nebraska, 

Lincoln, Nebraska, U.S.A. (accession numbers 

HWML 15354, Haematoloechus varioplexus Stafford, 

1902; 15355, Glypthelmins quieta Stafford, 1900; 

15356, kidney metacercariae; 15357, Mesocestoides 

sp.; 15358, diplostomid metacercariae; 15359, Halipegus 

eccentricus Thomas, 1939; 15360, unidentified 

adult tapeworm; 15361, Cosmocercoides sp.; 15362, 

unidentified larval nematode; 15363, unidentified species 

of Waltonella Schacher, 1974; 15364, Oswaldocruzia 

pipiens Walton, 1929). 

The chi-square test for independence was calculated 

to compare differences in prevalence among host sex. 

Yates' adjustment for continuity was used when sample 

sizes were low. A single-factor, independent-measures 

ANOVA and Scheffe's posthoc test were used to 

compare among seasonal differences in mean intensity 

and mean helminth species richness. When variances 

were heteroscedastic, the Kruskal-Wallis test and the 

Kolmogorov—Smirnov two-sample test were used. Student's 

?-test was used to compare differences in mean 

intensity and mean helminth species richness between 

sex of hosts. Approximate Mests were calculated when 

variances were heteroscedastic (Sokal and Rohlf, 

1981). Pearson's correlation was used to determine relationships 

among host SVL and WW and abundance 

of helminth parasites, excluding larval platyhelminths. 

BOLEK AND COGGINS—HELMINTH COMMUNITIES IN GREEN FROGS 165 

Pearson's correlation was calculated for host SVL and 

WW and helminth species richness per individual frog. 

Because WW gave a stronger correlation than SVL in 

each case, it is the only parameter reported. Because 

of low sample size during certain collection periods, 

data were pooled on a bimonthly basis to form samples 

of 15—20 frogs per season. Larval helminths were not 

included in the seasonal analysis, because they can 

also accumulate throughout the amphibian's life and 

thus mask monthly recruitment dynamics in adult 

frogs. 

Results 

A total of 75 adult green frogs, 43 males and 

32 females, was collected during April through 

October 1996. No significant difference existed 

in the number of male and female frogs collected 

throughout the year (x2 = 7.01, P > 0.05). 

The overall means of SVL and WW of green 

frogs were 68.8 ± 10.8 mm (range 39.8-89.4 

mm) and 35.8 ± 15.5 g (5.4-75.6 g), respectively. 

There was no significant difference in 

mean SVL (t = 0.10, P > 0.05) or mean WW 

(t = 0.24, P > 0.05) in male and female frogs. 

Stomach content analyses of green frogs revealed 

a broad range of aerial, terrestrial, and 

aquatic invertebrates. Sixteen different groups of 

invertebrates were recovered from stomach contents 

of green frogs, with coleopterans, gastropods, 

and diplopodans making up the largest 

percentage. 

Seventy-one (94%) of 75 R. clamitans melanota 

were infected with helminth parasites. The 

component community consisted of 12 species 

(5 trematodes, 2 cestodes, and 5 nematodes). Of 

these, 8 have indirect life cycles, 1 has a direct 

life cycle, and the life cycles of 3 are unknown. 

Overall mean helminth abundance, excluding 

larval platyhelminths, was 16.5 ± 38 with most 

frog infracommunities having 10 or fewer 

worms. In terms of abundance, digeneans dominated 

adult helminth communities (61.5% of total 

adult helminths). Prevalence ranged from 

80% for kidney metacercariae to 1.3% for an 

unidentified adult cestode, a filarid nematode of 

the genus Waltonella, and an unidentified encysted 

nematode. Values for overall prevalence, 

mean intensity, mean abundance, and total number 

of helminths recovered are summarized in 

Table 1. 

Statistically significant differences in prevalence 

or mean intensity existed between male 

and female frogs for H. varioplexus, H. eccentricus, 

kidney metacercariae, and unidentified 

species of Mesocestoides Valunt, 1863 and Cos- 



Table 1. Prevalence, mean intensity (MI), mean abundance (MA), and total helminths found in 75 specimens 

of Rana clamitans melanota in Wisconsin. 

Trematoda 

Species 

Haematoloechus varioplexus 

Halipegus eccentricus 

Glypthelmins quieta 

Unidentified metacercariae:i: 

Diplostomid metacercariae* 

Cestoda 

Unidentified adult cestode 

Mesocestoides sp.* 

Nematoda 

Oswaldocruzia pipiens 

Cosmoccrcoides sp. 

Waltonclla sp.t 

Larval nematode 

Encysted nematode 

* Underestimate. 

t New host record. 

± Not counted. 

Prevalence: MI ± 1 SD 

Number of 

worms 

number (%) (range) MA ± 1 SD recovered 

33 (44) 

17 (22.6) 

11 (14.6) 

60 (80) 

12 (16.0) 

1 (1.3) 

14 (18.6) 

21 (28) 

19 (25) 

1 (1.3) 

9 (12) 

1 (1.3) 

5.3 ± 7 

d-30) 

1.9 ± 1.7 

(1-8) 

35.5 ± 34.7 

(1-110) 

NC:i: 

35 ± 41 

(1-100) 

1 

31.9 ± 22 

(12-86) 

14.7 ± 23 

(1-90) 

3.3 ± 4.3 

(1-19) 

4 

25.9 ± 61 

(1-200) 

1 

mocercoides Harwood, 1930 (Table 2). Both H. 

varioplexus, a lung trematode, and H. eccentricus, 

a trematode of the eustachian tubes, had significantly 

higher mean intensities in male frogs, 

while the kidney metacercariae occurred at a 

higher prevalence in males. Female frogs had 

significantly higher mean intensities of Mesocestoides 

sp. and Cosmocercoides sp. 

Mean helminth species richness was 2.68 ± 

1.29 species per frog. Infections with multiple 

species were common, with 0, 1,2, 3, 4, and 5 

species occurring in 4, 8, 23, 20, 13, and 7 frogs, 

respectively. No statistically significant differences 

in mean helminth species richness were 

found between male (2.76 ± 1.01) and female 

frogs (2.56 ± 1.52, t = 0.66, P > 0.05). A nonsignificant 

positive correlation was found between 

overall helminth abundance, excluding 

larval platyhelminths, and WW (/• = 0.04, P > 

0.05). Nonsignificant relationships were also observed 

for most helminth species: unidentified 

adult cestode (r = -0.01, P > 0.05), H. vario- 

2.3 ± 5.4 

0.4 ± 1.1 

5.2 ± 18 

NC 

5.7 ± 20.7 

0.01 ± 0.1 

5.9 ± 15.3 

4.1 ± 13.7 

0.8 ± 2.6 

0.05 ± 0.5 

3.5 ± 23 

0.01 

± 0.1 

176 

33 

391 

> 1,770 

420 

1 

446 

310 

62 

4 

259 


1 

Lungs 

Location 

Eustachian tubes 

Small intestine 

Kidneys, body cavity 

Leg muscles 


Leg muscles, lungs 

Small intestine, stomach 

Large intestine, small intestine 

Body cavity 

Large intestine 


plexus (r = 0.10, P > 0.05), H. eccentricus (r 

= 0.19, P > 0.05), G. quieta (r = -0.02, P > 

0.05), O. pipiens (r = -0.12, P > 0.05), unidentified 

larval nematode (r = -0.04, P > 

0.05), Waltonella sp. (r = 0.20, P > 0.05), and 

encysted nematodes (r = -0.19, P > 0.05). The 

nematode Cosmocercoides sp. had a significant 

positive correlation with WW (/• = 0.31, P < 

0.01). A significant positive Pearson's correlation 

also existed between species richness and 

WW (r = 0.31, P < 0.01). However, correlations 

between frog WW and species richness 

were not significant in May—June (/• = 0.31, P 

> 0.05), July-August (r = 0.01, P > 0.05), and 

September-October (/- = 0.29, P > 0.05) but 

were significant for the April collection (r = 

0.60, P < 0.02). 

The trematodes H. varioplexus and H. eccentricus 

occurred throughout the year, with highest 

prevalences observed during the fall (September-October) 

collection, 65% and 30%, respectively. 

The intestinal trematode, G. quieta, was

Table 2. Prevalence (Pr) and 

Rana clamitans melanota. 

Trematoda 

Species 

Haematoloechus varioplexus 

Halipegns eccentricus 


Unidentified metacercariae* 

Diplostomid metacercariae* 

Cestoda 

Unidentified adult cestode 

Mesocestoides sp.* 

Nematoda 

Oswaldocruzia pipiens 

Cosmocercoides sp. 

Waltonella sp. 

Larval nematode 

Encysted nematode 

* Underestimate. 

± Not counted. 


mean intensity (MI) of helminth parasites in male and female green frogs, 

Measure of 

parasitism 

Pr 

MI ± 

Pr 

MI ± 

PI- 

MI ± 

Pr 

MI ± 

Pr 

MI ± 

Pr 

MI ± 

Pr 

MI ± 

Pr 

MI ± 

Pr 

MI ± 

Pr 

MI ± 

Pr 

MI ± 

PI- 

MI ± 

1 SD 

1 SD 

1 SD 

1 SD 

1 SD 

1 SD 

1 SD 

1 SD 

1 SD 

1 SD 

1 SD 

1 SD 

first observed during midspring (May-June) 

with a prevalence of 5%. Prevalence for this 

species reached its maximum (30%) during summer 

(July-August) and decreased during the fall 

collection (20%). Seasonal mean intensity of 

adult platyhelminths followed similar patterns as 

prevalence, but no significant differences existed 

for any of the adult platyhelminths recovered, H. 

eccentricus (adjusted H = 2.02, P > 0.05), H. 

varioplexus (F = 0.34, P > 0.05), or G. quieta 

(t = 0.43, P > 0.05). 

Although prevalence and intensity of H. varioplexus 

did not vary significantly throughout 

the collection period, mean length of worms did 

(Fig. 1). Greatest mean length of worms (4.1 

mm) was recorded in early spring (April), when 

all individuals were gravid adults, and reached 

a minimum during midspring (1.84 mm), when 

immature individuals were common. Statistically 

significant differences in mean length were 

observed for April and May—June collections, 

Males 

N = 43 

46.5 (20/43) 

7 ± 8.6 

23.3 (10/43) 

2.3 ± 2.1 

1 1 .6 (5/43) 

21.8 ± 27.9 

90.7 (39/43) 

NC:!: 

18.6 (8/43) 

45.3 ± 46.1 

2.3 (1/43) 

1 

23.3 (10/43) 

24.9 ± 16.2 

27.9 (12/43) 

16.6 ± 25.8 

20.9 (9/43) 

2+1.7 

0 (0/43) 

0 

9.3 (4/43) 

11.5 ± 11.2 

0 (0/43) 

0 

Females 

N = 32 

40.6 ( 1 3/32) 

2.8 ± 2.2 

21.9 (7/32) 

1.4 ± 0.5 

18.8 (6/32) 

47 ± 38.1 

65.6 (21/32) 

NC 

12.5 (4/32) 

14.5 ± 23.7 

0 (0/32) 

0 

12.5 (4/32) 

49.3 ± 24.8 

28.1 (9/32) 

12.2 ± 19.6 

31.3 (10/32) 

4.4 ± 5.6 

3.1 (1/32) 

4 

18.8 (6/32) 

35.5 ± 80.6 

3.1 (1/32) 

1 

Statistic 

X2 = 0.24 

f',= 3.07 

X2 = 0.02 

f',= 2.71 

X2 = 0.46 

t = 1 .35 

X2 = 5.72 

X2odj = 0.16 

t = 1.23 

X2ailJ = 0.02 

X2adj = 0.78 

t = 6.69 

x2 = o.oo 

/ = 0.44 

X2 = LOO 

r',= 3.81 

X2ailj = 0.02 

X2:,Ji = 1-67 

f',= 1.67 

X2a(lj = 0.02 

P 

>0.05 

0.05 

0.05 

>0.05 

0.05 

>0.05 

>().05 

>0.05 

0.05 

>0.05 

>0.05 

0.05 

>0.05 

>0.05 

>0.05 

April and July-August collections, May-June 

and September—October collections, and July- 

August and September-October collections (F = 

11.8, P < 0.05, single-factor, independent-measure 

ANOVA; P < 0.05 for all pair-wise comparisons, 

Scheffe's test). 

The nematodes Waltonella sp. and an unidentified 

encysted larva were recovered infrequently 

as single infections during midspring and fall 

collections, respectively. Prevalence and intensity 

values for O. pipiens, Cosmocercoides sp., 

and unidentified larval nematodes were low and/ 

or erratic over the 7-mo period. The nematodes 

O. pipiens, Cosmocercoides sp., and unidentified 

larval nematodes were first observed during 

midspring and persisted until fall, with prevalence 

being highest in summer for O. pipiens 

(62%) and midspring and summer for Cosmocercoides 

sp. (40%) and larval nematodes 

(20%). However, only O. pipiens exhibited statistically 

significant differences (adjusted H = 


168 COMPARATIVE PARASITOLOGY, 68(2), JULY 2001 

7 -r 

N = 20 N = 27 N = 28 N = 79 

Apr May-Jun Jul- Aug Sep-Oct 

Date 

Figure 1. Mean lengths of Haematoloechus varioplexus 

from Rana clamitans melanota. N = number 

of worms measured from all frogs recovered in 

each sampling period. 

9.45, P < 0.05) in mean intensity. The two-sample 

Kolmogorov-Smirnov test revealed significant 

differences in mean intensities during the 

May-June (27 ± 30) and July-August (9.8 ± 

17) collections. 

Mean helminth species richness fluctuated 

seasonally (Fig. 2) and was lowest (1.53) during 

early spring and highest (3.1) during the summer 

collections. Statistically significant differences 

in mean helminth species richness were observed 

for April and May—June collections, 

April and July—August collections, and April 

and September-October collections (F = 6.01, 

P < 0.05, single-factor, independent-measure 

ANOVA; P < 0.05 for all pairwise comparisons, 

Scheffe's test). No statistically significant differences 

in frog WW were observed during the 

year (F = 0.37, P > 0.05). 

Discussion 

Wisconsin green frogs had high overall helminth 

prevalence, with parasite infracommunities 

being dominated by indirect life-cycle parasites. 

Of the identified parasites, only 1 direct 

life-cycle nematode, O. pipiens, was present, 

with most helminth species displaying a prevalence 

below 50% and/or low mean intensities 

below 30. 

Most green frogs contained identifiable stom- 

5-r 


= 15 N = 20 N = 20 N = 20 

May-Jim Jul-Aug Sep-Oct 

Date 

Figure 2. Mean helminth species richness of 

Rana clamitans melanota during April, May-June, 

July-August, and September-October 1996. N = 

number of frogs collected on each date. 

ach contents containing predominantly beetles, 

gastropods, and diplopods. In total, 16 different 

groups of terrestrial, aerial, and aquatic invertebrates 

comprised their stomach contents. 

These results appear similar to those of other 

investigators (Hamilton, 1948; Stewart and Sandison, 

1972; McAlpine and Dilworth, 1989; 

Werner et al., 1995). Hamilton (1948) found that 

the principal foods of green frogs collected in 

New York, U.S.A. consisted of beetles, flies, and 

grasshoppers, with a total of 15 different items 

recovered from adult frogs and 20 different prey 

items recovered from various sized individuals. 

The most common helminth recovered was an 

unidentified kidney metacercaria. This larval 

trematode had an overall prevalence of 80% and 

mean intensity of over 30 worms per frog. Four 

other digeneans were recovered from green 

frogs: a diplostomid metacercaria encysted in 

the musculature, and 3 adult trematodes: H. varioplexus, 

H. eccentricus, and G. quieta. Frogs 

become infected with H. varioplexus, a lung 

trematode, and H. eccentricus, a eustachian tube 

trematode, by eating infected odonates (Krull, 

1931; Dronen, 1975, 1978; Wetzel and Esch, 

1996). Glypthelmins quieta, a trematode of the 

small intestine, is acquired when frogs ingest 

prey such as tadpoles, frogs, and/or shed skin 

infected with metacercariae (Prudhoe and Bray, 

1982). Therefore, diet was important in the

transmission dynamics of these 3 trematode species 

in this study. 

Haematoloechus varioplexus and H. eccentricus 

were recovered from frogs throughout the 

year, increasing, although not significantly, in 

both prevalence and mean intensity during the 

fall collection. Recently, Wetzel and Esch (1997) 

have shown that the life span of H. eccentricus 

may be variable, with trematodes capable of maturing 

in as little as 1 wk and being lost the 

following week. Because of the small number of 

these flukes recovered in our study, little can be 

said about their recruitment throughout the year. 

Krull (1931) estimated that the life-span of Haematoloechus 

medioplexus Stafford, 1902, averaged 

1 yr, while studies by Kennedy (1980) on 

species of Haematoloechus have shown that 

trematodes can reach full length in only 60 days. 

The size differences observed for H. varioplexus 

during the year (Fig. 1) may be significant in 

understanding recruitment of this species. The 

seasonal variation in length of H. varioplexus 

suggests that adult worms are lost during early 

spring and that new infections begin during 

midspring and continue throughout the year. 

These results are similar to those of Ward 

(1909), who observed lung flukes of Rana pipiens 

Schreber, 1782, being lost during breeding, 

and recruitment occurring throughout the year. 

Two cestode species were recovered from 

green frogs during this study, the larval tetrathyridium 

of Mesocestoides sp. and a single 

specimen of an adult cestode that could not be 

identified because the scolex was lost. The complete 

life cycles of Mesocestoides spp. are currently 

unknown; however, a number of mammals, 

amphibians, and reptiles are known to 

serve as second intermediate hosts, while carnivorous 

mammals serve as definitive hosts. The 

tetrathyridian stage has been reported from a variety 

of mammals and reptiles but is rare in amphibians 

(McAllister and Conn, 1990). The life 

cycles of these 2 species of cestode are unknown, 

although frog diet may be important in 

their transmission dynamics. 

Nematodes in the genus Waltonella typically 

are found in the body cavity of species of Rana. 

Adult worms release microfilaria into the bloodstream, 

and mosquitoes serve as vectors, infecting 

frogs while feeding (Witenberg and Gerichter, 

1944). The only report of filarial worms in 

R. clamitans is of microfilaria recovered from 1 

frog in Ontario, Canada (Barta and Desser, 


1984). Therefore, R. clamitans melanota is apparently 

a new host record for Waltonella sp. 

(Esslinger, 1986; Baker, 1987). Waltonella 

americana was previously reported and described 

in Wisconsin leopard frogs by Walton 

(1929). 

The nematode Cosmocercoides sp. was recovered 

from the large intestine of green frogs. 

Confusion exists in the literature on the identification 

of species of Cosmocercoides in amphibians 

and reptiles (Baker, 1987; Vanderburgh 

and Anderson, 1987). The major difference in 

species identification is the number of rosette 

papillae per subventral row in males, with male 

Cosmocercoides dukae Holl, 1928 (gastropod 

parasite) having 9—21 rosette papillae, averaging 

13-14, and C. variabilis (amphibian parasite) 

having 15-25, averaging 20 or 21. Because of 

this overlap and the presence of only 5 damaged 

males out of 62 nematodes recovered, species 

identification was not possible. Interestingly, no 

worms were found in the lungs or body cavity 

of any green frogs, and Cosmocercoides sp. occurred 

in frogs in months when gastropods were 

commonly found in the stomach contents. We 

suspect that specimens of Cosmocercoides sp. 

recovered are C. dukae, although this cannot be 

confirmed. 

Differential infection between host sex and 

prevalence or mean intensity was observed for 

a number of helminth species. Male frogs had a 

significantly higher prevalence of kidney metacercariae 

and significantly higher mean intensities 

of H. varioplexus and H. eccentricus than 

female frogs. Male green frogs are territorial 

during the breeding season and defend their 

aquatic breeding site from potential competitors 

(Martof, 1953; Oldham, 1967). Thus, unlike the 

females, they remain in the water for longer periods 

of time and may be exposed to cercariae 

of the kidney trematode for longer periods. Because 

males remain in a relatively small area of 

the pond during the breeding season, they may 

occur in a microhabitat conducive to becoming 

infected with digeneans. Recently, Wetzel and 

Esch (1997), in a seasonal study of Halipegus 

occidualis Stafford, 1905 and H. eccentricus in 

green frogs, suggested that certain areas of a 

pond may be "hot spots" for infection with digenetic 

trematodes. Therefore, male frogs in 

these "hot spots" may feed more often on 

emerging odonates containing metacercariae of 

species of Haematoloechus and Halipegus, ex- 



plaining the higher mean intensities of these 

trematodes observed in male frogs (Wetzel and 

Esch, 1996). 

Female frogs had significantly greater mean 

intensities of Cosrnocercoides sp. and Mesocestoides 

sp. than males. Although Cosmocercoides 

sp. could not be identified to species, 

both C. variabilis and C. dukae occur in terrestrial 

habitats. Female frogs spend more time 

on the ground and have a higher probability 

of encountering these nematodes in a terrestrial 

habitat, either by skin-penetrating C. variabilis 

or by feeding on terrestrial mollusks, 

hosts for C. dukae. Unfortunately, nothing can 

be stated about the transmission dynamics of 

Mesocestoides sp., and no conclusions can be 

drawn from this difference. The observed differences 

in host sex are probably due to ecological 

differences in their habitat preference 

throughout the year. 

Significant positive relationships between 

WW and species richness were observed in 

green frogs. In this study, frogs in the later 

collections had greater species richness than in 

early collections (Fig. 2); therefore, time of 

exposure was more important in developing 

richer helminth communities than was frog 

weight during the May-June, July-August, 

and September—October collections. This is 

supported by the results showing significant 

differences in species richness over time and 

nonsignificant correlations between WW and 

species richness. Observations linking higher 

species richness with larger host size have 

been reported in green frogs and other species 

of Rana by Muzzall (199 la) and Me Alpine 

(1997). These investigators suggested that older 

individuals may have a longer exposure 

time and possess more surface area for colonization 

by skin-penetrating nematodes and 

digenean metacercariae. Also, larger frogs 

possess a greater gape size and may feed on 

larger, and a wider number, of intermediate 

hosts than smaller individuals. As in their 

studies, our data also support the island size 

hypothesis, which predicts that larger host individuals 

should support higher species richness 

than smaller individuals (Holmes and 

Price, 1986). McAlpine (1997) also stated that 

aspects of host ecology, such as diet and habitat, 

and parasite transmission may confound 

any simple relationship between the diversity 

of helminth communities and size of hosts. 


Data from the present study suggest that time 

of transmission may also have a similar confounding 

effect. 

The depauperate helminth community structure 

of Carroll College green frogs was similar 

to the community structure of green frogs examined 

from Michigan, U.S.A. and New 

Brunswick, Canada by Muzzall (199la) and 

McAlpine (1997), respectively. Of the 120 

frogs examined from Michigan, 108 (90%) 

were infected with a total of 13 species of helminths 

(8 trematodes, 1 cestode, and 4 nematodes) 

while 164 of 234 (70.1%) green frogs 

examined from Canada were infected with 18 

species of helminths (10 trematodes, 3 cestodes, 

and 5 nematodes). As in our study, in 

terms of abundance, digeneans dominated the 

adult helminth communities of frogs in Michigan 

(96.5%) and New Brunswick (60.8%), indicating 

that diet and a semiaquatic habitat 

were also important in structuring helminth 

communities dominated by indirect-life-cycle 

parasites at those locations. Similarly, in both 

of those studies, adult frogs had higher species 

richness than young-of-the-year individuals, 

indicating that size and/or age were also important 

in acquiring new species of helminths 

into the infracommunity of these hosts. Data 

from our study and the Michigan and New 

Brunswick frogs suggest that although helminth 

species composition, richness, and prevalence 

may be variable depending on collection 

site and the ecological factors influencing 

variation in life history traits in local populations 

at those locations, green frog helminth 

communities are dominated by digenetic trematodes 

acquired in a semiaquatic habitat and/ 

or through the frogs' diet. 


We thank Dr. Susan Lewis, Department of Biology, 

Can-oll College for allowing access to the 

field station and permission to collect frogs, and 

Melissa Ewert and Luke Bolek for help in collecting 

frogs. We also thank 2 anonymous reviewers 

and the editors Drs. Willis A. Reid, Jr. 

and Janet W Reid for improvements on an earlier 

draft of the manuscript. 


Aho, J. M. 1990. Helminth communities of amphibians 

and reptiles: comparative approaches to understanding 

patterns and processes. Pages 157-

196 in G. W. Esch, A. O. Bush, and J. W. Aho, 

eds. Parasite Communities: Patterns and Processes. 

Chapman and Hall, New York, New York, 

U.S.A. 335 pp. 

Baker, M. R. 1987. Synopsis of the Nematoda parasitic 

in amphibians and reptiles. Memorial University 

of Newfoundland Occasional Papers in Biology 

11:1-325. 

Barta, J. R., and S. S. Desser. 1984. Blood parasites 

of amphibians from Algonquin Park, Ontario. 

Journal of Wildlife Diseases 20:180-189. 

Barton, D. P. 1996. Helminth infracommunities in Litoria 

genimaculata (Amphibia: Anura) from 

Birthday Creek, an upland rainforest stream in 

northern Queensland, Australia. International 

Journal for Parasitology 26:381-385. 

Bolek, M. G., and J. R. Coggins. 2000. Seasonal occurrence 

and community structure of helminth 

parasites from the eastern American toad, Bufo 

americanus americanus, from southeastern Wisconsin, 

U.S.A. Comparative Parasitology 67:202- 

209. 

Borror, D. J., C. A. Triplehorn, and N. F. Johnson. 

1989. An Introduction to the Study of Insects, 6th 

ed. Harcourt Brace College Publishers, Philadelphia, 

Pennsylvania, U.S.A. 875 pp. 

Bush, A. O., K. D. Lafferty, J. M. Lotz, and A. W. 

Shostak. 1997. Parasitology meets ecology on its 

own terms: Margolis et al. revisited. Journal of 


Campbell, R. A. 1968. A comparative study of the 

parasites of certain Salientia from Pocahontas 

State Park, Virginia. Virginia Journal of Science 

19:13-29. 

Coggins, J. R., and R. A. Sajdak. 1982. A survey of 

helminth parasites in the salamanders and certain 

anurans from Wisconsin. Proceedings of the Helminthological 


Dronen, N. O. 1975. Studies on the life cycle of Haematoloechus 

coloradensis Cort, 1915 (Digenea: 

Plagiorchiidae), with emphasis on host susceptibility 

to infection. Journal of Parasitology 61:657- 

660. 

. 1978. Host-parasite population dynamics of 

Haematoloechus coloradensis Cort, 1915 (Digenea: 

Plagiorchiidae). American Midland Naturalist 

99:330-349. 

Esslinger, J. H. 1986. Redescription of Foleyellid.es 

striatus (Ochoterena and Caballero, 1932) (Nematoda: 

Filarioidea) from a Mexican frog, Rana 

montezumae, with reinstatement of the genus Foleyellides 

Caballero, 1935. Proceedings of the Helminthological 


Goater, T. M., G. W. Esch, and A. O. Bush. 1987. 

Helminth parasites of sympatric salamanders: ecological 

concepts at the infracommunity, component, 

and compound community levels. American 

Midland Naturalist 118:289-299. 

Goldberg, S. R., C. R. Bursey, and I. Ramos. 1995. 

The component parasite community of three sympatric 

toad species, Bufo cognatus, Bufo debilis 

(Bufonidae), and Spea multiplicata (Pelobatidae) 

from New Mexico. Journal of the Helminthological 

Society of Washington 62:57—61. 


Hamilton, W. J., Jr. 1948. The food and feeding behavior 

of the green frog, Rana clamitans Latreille, 

in New York State. Copeia 1948:203-207. 

Holmes, J. C., and P. W. Price. 1986. Communities 

of parasites. Pages 187-213 in J. Kikkawa and D. 

J. Anderson, eds. Community Ecology: Pattern 

and Process. Blackwell Scientific Publications, 

Boston, Massachusetts, U.S.A. 432 pp. 

Kennedy, M. J. 1980. Host-induced variations in Haematoloechus 

buttensis (Trematoda: Haematoloechidae). 

Canadian Journal of Zoology 58:427- 

442. 

Kennedy, R. C., A. O. Bush, and J. M. Aho. 1986. 

Patterns in helminth communities: why are birds 

and fish different? Parasitology 93:205-215. 

Krull, W. H. 1931. Life history studies on two frog 

lung flukes, Pneumonoeces mediopiexus and 

Pneumobites parviplexus. Transactions of the 

American Microscopical Society 50:215—277. 

Martof, B. S. 1953. Territoriality in the green frog, 

Rana clamitans. Ecology 34:165-167. 

McAllister, C. T., and D. B. Conn. 1990. Occurrence 

of tetrathyridia of Mesocestoides sp. (Cestoidea: 

Cyclophyllidea) in North American anurans (Amphibia). 


McAlpine, D. F. 1997. Helminth communities in bullfrogs 

(Rana catesbeiand), green frogs (Rana 

clamitans), and leopard frogs (Rana pipiens) from 

New Brunswick, Canada. Canadian Journal of Zoology 

75:1883-1890. 

, and M. B. Burt. 1998. Helminths of bullfrogs, 

Rana catesbeiana, green frogs, R. clamitans, 

and leopard frogs, R. pipiens in New Brunswick. 

Canadian Field-Naturalist 112:50-68. 

-, and T. G. Dilworth. 1989. Microhabitat and 

prey size among three species of Rana (Anura: 

Ranidae) sympatric in eastern Canada. Canadian 

Journal of Zoology 67:2244-2252. 

Muzzall, P. M. 1991 a. Helminth infracommunities of 

the frogs Rana catesbeiana and Rana clamitans 

from Turkey Marsh, Michigan. Journal of Parasitology 

77:366-371. 

. 1991b. Helminth communities of the newt, 

Notophthalrnus viridescens, from Turkey Marsh, 

Michigan. Journal of Parasitology 77:87-91. 

Oldham, R. S. 1967. Orienting mechanisms of the 

green frog, Rana clamitans. Ecology 48:477-491. 

Prudhoe, O. B. E., and R. A. Bray. 1982. Platyhelminth 

parasites of the Amphibia. British Museum 

(Natural History)/Oxford University Press, London, 

U.K. 217 pp. 

Scale, D. B. 1987. Amphibia. Pages 467-552 in T. J. 

Pandian and F. J. Vanberg, eds. Animal Energetics. 

Vol. 2. Academic Press, San Diego, California, 

U.S.A. 631 pp. 

Sokal, R. R., and J. F. Rohlf. 1981. Biometry, 2nd 

ed. W. H. Freeman and Company, New York, New 

York, U.S.A. 859 pp. 

Stewart, M. M., and P. Sandison. 1972. Comparative 

food habits of sympatric mink frogs, bullfrogs and 

green frogs. Journal of Herpetology 6:241-244. 

Vanderburgh, D. J., and R. C. Anderson. 1987. The 

relationship between nematodes of the genus Cosmocercoides 

Wilkie, 1930 (Nematoda: Cosmocer- 



coidea) in toads (Bufo americanus) and slugs 

(Deroceras laeve). Canadian Journal of Zoology 

65:1650-1661. 

Vogt, R. C. 1981. Natural History of Amphibians and 

Reptiles of Wisconsin. The Milwaukee Public 

Museum and Friends of the Museum, Inc., Milwaukee, 

Wisconsin, U.S.A. 205 pp. 

Walton, A. C. 1929. Studies on some nematodes of 

North American frogs. I. Journal of Parasitology 

15:227-249. 

Ward, H. B. 1909. The influence of hibernation and 

migration on animal parasites. Proceedings of the 

7th International Zoological Congress (Boston). 

12pp. 

Werner, E. E., G. A. Wellborn, and M. A. McPeek. 

1995. Diet composition in postmetamorphic bullfrogs 

and green frogs: implications for interspecific 

predation and competition. Journal of Herpetology 

29:600-607. 

Wetzel, E. J., and G. W. Esch. 1996. Influence of 

odonate intermediate host ecology on the infection 

dynamics of Halipegus spp., Haematoloechus longiplexus, 

and Haematoloechus complexus (Trematoda: 

Digenea). Journal of the Helminthological 


-, and . 1997. Infrapopulation dynamics 

of Halipegus occidualls and Halipegus eccentricus 

(Digenea: Hemiuridae): temporal changes 

within individual hosts. Journal of Parasitology 

83:1019-1024. 

Williams, D. D., and S. J. Taft. 1980. Helminths of 

anurans from NW Wisconsin. Proceedings of the 

Helminthological Society of Washington 47:278. 

Witenberg, G., and C. Gerichter. 1944. The mor 

phology and life history of Foleyella duboisis 

with remarks on allied filarids of amphibians. 

Journal of Parasitology 30:245-256. 

Yoder, H. R., and J. R. Coggins. 1996. Helminth 

communities in the northern spring peeper, Pseudacris 

c. crucifer Wied, and the wood frog, Rana 

sylvatica Le Conte, from southeastern Wisconsin. 


63:211-214. 

Announcement of the 

FOURTH INTERNATIONAL CONGRESS OF NEMATOLOGY 

at the 

Tenbel Resort, Tenerife, Canary Islands, Spain 

June 8-13, 2002 

Sponsor: The International Federation of Nematology Societies (IFNS). 

Scientific Program: 4 full days of scientific sessions with an opening plenary session, symposia, colloquia, 

discussion sessions and offered papers arranged in 4 poster sessions. The themes will include the systematics, 

molecular biology, genomics, genetics, management, ecology, and physiology of parasitic, entomophagous, 

and free-living nematodes. Abstracts of offered papers will be accepted between December 

1, 2001 and March 1, 2002. 

Accommodations: The Tenbel resort is a large hotel complex in an extensive tropical garden setting. A 

range of accommodations will be available to Congress participants. Special interest and scenic tours will 

be arranged during and after the FICN. Details of available accommodations and tourism opportunities 

will be posted on the IFNS web site at http:\\www.ifns.org. 

Registration: Registrations for the FICN will be accepted after December 1, 2001. Registration forms 

and details regarding regular, student, spouse, and accompanying person registrations are available from 

the IFNS web site at www.ifns.org, or from Dr. Maria Arias, FICN Local Arrangements Chair, CSIC 

Centre de Ciencias Medioambiantales, Serrano 115 DPDO, Madrid 28006, Spain. 




Blood Parasites of the Ring-Necked Duck (Ay thy a collaris} on Its 

Wintering Range in Florida, U.S.A. 

DONALD J. FORRESTER, 1


Voucher specimens have been deposited in the International 

Reference Centre for Avian Haematozoa at 

the Queensland Museum, Brisbane, Australia (accession 

nos. G462509-G462640) and the U.S. National 

Parasite Collection in Beltsville, Maryland, U.S.A. (accession 

nos. 88248-88254). Prevalence data for each 

parasite were evaluated by using PROC FREQ for 2tailed 

Fisher's exact tests with regard to locality, gender, 

and age (SAS Institute, 1989). Significance was 

taken at P < 0.05. Terminology used was according to 

Bush et al. (1997). 

Results 

Five species of blood parasites were identified, 

2 protozoans (Haemoproteus nettionis 

(Johnson and Cleland, 1909) and Leucocytozoon 

simondi Mathis and Leger, 1910) and microfilariae 

of 3 nematodes (Splendidofilaria fallisensis 

(Anderson, 1954) and 2 unidentified species). 

The microfilariae of 5. fallisensis (n = 1,096) 

were 75.1 (41-130) in length and 4.9 (4.1-6.0) 

in width, with a rounded anterior, and a tapering 

tail that ended bluntly. Sheaths were lacking. 

Microfilariae of 2 unidentified filaroid species 

were observed in 36 ring-necked ducks. Microfilariae 

of Species I were long and slender with 

a bluntly rounded anterior and an overall uniform 

body width that tapered slightly to a rounded 

posterior. These microfilariae (n = 94) were 

150 (110-190) in length and 5.8 (5.0-6.0) in 

width. They had a well-defined sheath that could 

be seen at 1 or both ends of the body. The cephalic 

space was 4.8 (3-7, n = 10). Fixed points 

(n — 10), expressed as percentages of body 

length, were as follows: excretory cell 36.4 (35— 

38), inner body 64.9 (62-67), anal pore 86.6 

(83-90). Microfilariae of Species II (n = 287) 

were similar in appearance to Species I, except 

they were 150 (120-210) in length and 5.8 (5.0- 

6.0) in width and lacked a sheath. The cephalic 

space was 5.2 (3-6, n = 10). Fixed points (n = 

10) were as follows: excretory cell 36.5 (35-39), 

inner body 64.7 (63-67), anal pore 86.6 (85- 

90). 

The overall prevalences were H. nettionis 

(5.3%), L. simondi (9.2%), S. fallisensis 

(39.2%), filaroid Species I (6.0%), and filaroid 

Species II (13.8%). Of the sample, 97 ducks 

were infected with 1 species of parasite, 43 with 

2, and 8 with 3. The most common multiple infection 

was a combination of L. simondi and S. 

fallisensis (n = 15), followed by combinations 

of the 2 unidentified filaroids (n = 10). 

There were no significant differences in prevalences 

of the 5 species of parasites among the 


collection periods, and therefore data for all 

years were combined. Prevalences of H. nettionis 

in Regions 1 (7%) and 2 (7%) were significantly 

higher (P = 0.03) than in Region 3 (0%). 

The prevalence of L. simondi was significantly 

higher (P = 0.0005) in Region 1 (19%) than in 

Regions 2 (8%) and 3 (1%). Microfilariae of 

Species II were more prevalent (P = 0.016) in 

Regions 2 (14%) and 3 (22%) than in Region 1 

(6%). There were no regional differences in the 

prevalences of the microfilariae of S. fallisensis 

and Species I. 

Microfilariae of S. fallisensis, Species I, and 

Species II were more prevalent in female ducks 

(46%, 9%, and 23%) than in males (33%, 3%, 

and 5%) (P = 0.028, 0.042, and 0.00002). The 

prevalences of H. nettionis and L. simondi were 

similar. 

Leucocytozoon simondi was more prevalent 

(P = 0.021) in juveniles (16%) than in adults 

(6%). On the other hand, filaroid Species I and 

II were both more prevalent (P = 0.029 and 

0.001) in adults (8% and 18%) than in juveniles 

(1% and 4%). There were no age differences in 

the prevalences of H. nettionis and S. fallisensis. 

Discussion 

Although this is the first report of H. nettionis 

and L. simondi from ring-necked ducks in Florida, 

both have been identified in ring-necked 

ducks overwintering in Texas, U.S.A. (Loven et 

al., 1980). Prevalences in the 20 ducks examined 

in Texas were similar to those of our Florida 

birds, i.e., 5% for H. nettionis and 10% for L. 

simondi. No microfilariae were reported in the 

Texas study. Unidentified microfilariae were 

seen in 1 of 6 ring-necked ducks examined in 

Maine, U.S.A.; the authors stated that their microfilariae 

were 45—65 long and 4-6 wide with 

a short, narrow, and slightly twisted tail (Nelson 

and Gashwiler, 1941). In a study of the blood 

parasites of 178 ring-necked ducks in the Maritime 

Provinces of Canada (New Brunswick, 

Nova Scotia, and Prince Edward Island), the 

prevalence of H. nettionis was higher (10%), and 

of L. simondi was lower (5%), than those in our 

birds from Florida (Bennett et al., 1975). No microfilariae 

were reported in the Maritime study. 

In another study, Bennett and Inder (1972) 

found microfilariae in 1 of 10 ring-necked ducks 

from Newfoundland, Canada, but provided no 

descriptions or measurements. 

Anderson's (1956) description of the micro-

filariae of S. fallisensis was similar to our specimens, 

with the exception of ranges of the 

lengths and the lack of sheaths; however, he noted 

that the sheaths of S. fallisensis were extremely 

delicate, and he was unable to see them 

in most specimens stained with Giemsa. Anderson 

(1956) also described a microfilaria from a 

European teal (Anas crecca Linnaeus, 1758) that 

he called Type D. His Type D microfilariae were 

similar to those of 5. fallisensis except for the 

length (range = 110-138) and the lack of a 

sheath. The microfilariae in our ring-necked 

ducks that we are calling 5. fallisensis could actually 

represent 2 species. However, it is possible 

that because we measured twice as many microfilariae 

as did Anderson (1956), that we have 

determined that the range of lengths for the microfilariae 

of this species is more extensive than 

previously recognized. Therefore, we are calling 

those microfilariae that fell in the range of 41- 

130 in length, but otherwise conformed to Anderson's 

(1956) description, S. fallisensis. The 

high number of combined infections of S. fallisensis 

and L. simondi in the same bird (n = 15) 

was probably due to the fact that both parasites 

utilize the same species of simuliid blackflies as 

vectors (Fallis et al., 1951; Anderson, 1968). 

Of the 36 ducks that had unidentified microfilariae, 

2 had Species I only, while 17 had Species 

II only. Seventeen of 36 ducks had both 

Species I and II microfilariae. The fact that Species 

I usually occurred with Species II and only 

twice by itself and that the range of lengths and 

several fixed points were almost equal may support 

the idea that these are variations of a single 

species. In many ways (morphologic and metric) 

our microfilariae resemble those of Chandlerella 

bus hi, described by Bartlett and Anderson 

(1987) from American coots (Fulica americana 

Gmelin, 1789) in Manitoba, Canada. They were 

not able to see the sheaths on microfilariae of C. 

bushi in blood films made from fresh heart blood 

and stained with Giemsa. However, they were 

able to see the sheaths on Giemsa-stained blood 

films of heart blood taken from thawed carcasses 

or specimens teased from lung tissue. The lack 

of visible sheaths of microfilariae in our ringnecked 

duck blood films might be because they 

were made from fresh heart blood. The identification 

of the microfilariae from ring-necked 

ducks will have to await the discovery of adult 

worms and further study and comparison of their 

FORRESTER ET AL.—BLOOD PARASITES OF RING-NECKED DUCKS 175 

intrauterine microfilariae with those from the 

blood. 

The regional differences in the prevalences of 

H. nettionis and L. simondi may have been a 

reflection of the location of the breeding grounds 

and flyways used by different subpopulations of 

ring-necked ducks. Because transmission of the 

blood parasites of waterfowl does not occur in 

Florida (Thul et al., 1980; Thul and O'Brien, 

1990; Forrester et al., 1994), the ducks must become 

infected either on the breeding grounds or 

during migration. The types and numbers of arthropod 

vectors found on various breeding 

grounds might differ and thereby influence the 

acquisition of these blood parasites in various 

segments of the North American population. 

Ring-necked ducks that overwinter in Florida 

are known to breed during the summer months 

in various prairie provinces of Canada across to 

Ontario and the eastern U.S.A. (Bellrose, 1976). 

Some ducks that breed in the more western regions 

of Canada migrate eastward and then 

move southward. Others migrate southward and 

pass through Wisconsin, Indiana, Tennessee, and 

Georgia. Most of the ring-necked ducks in Florida 

originate from Ontario, Manitoba, and the 

District of Mackenzie (Bellrose, 1976). The lower 

prevalence of infections of L. simondi in 

adults may be due to age-related immunity. Reasons 

for the gender differences in prevalences 

are unknown. 


We thank H. F. Percival, T C. Hines, C. W. 

Jeske, and A. R. Woodward for assistance in collecting 

ducks, R. C. Littell for helping with statistical 

analyses, and R. C. Anderson, E. C. Greiner, 

and M. G. Spalding for reviewing the manuscript 

and offering useful suggestions. This research 

was supported in part by the Florida Fish 

and Wildlife Conservation Commission and is a 

contribution of Federal Aid to Wildlife Restoration, 

Florida Pitman—Robertson Project W-41. 

This is Florida Agricultural Experiment Station 

Journal Series No. R-07749. 


Anderson, R. C. 1956. Ornithofilaria fallisensis n. sp. 

(Nematoda: Filarioidea) from the domestic duck 

with descriptions of microfilariae in waterfowl. 

Canadian Journal of Zoology 32:125-137. 

. 1968. The simuliid vectors of Splendidofilaria 

fallisensis of ducks. Canadian Journal of Zoology 

46:610-611. 



Bartlett, C. M., and R. C. Anderson. 1987. Chandlerella 

bushi n. sp. and Splendidofilaria caperata 

Hibler, 1964 (Nematoda: Filarioidea) from Fulica 

americana (Gruiformes: Rallidae) in Manitoba, 

Canada. Canadian Journal of Zoology 65:2799- 

2802. 

Bellrose, F. C. 1976. Ducks, Geese & Swans of North 

America. Stackpole Books, Harrisburg, Pennsylvania, 

U.S.A. 543 pp. 

Bennett, G. F., and J. G. Inder. 1972. Blood parasites 

of game birds from insular Newfoundland. Canadian 


, A. D. Smith, W. Whitman, and M. Cameron. 

1975. Hematozoa of the Anatidae of the Atlantic 

Flyway. II. The Maritime Provinces of Canada. 


-, M. Whiteway, and C. B. Woodworth-Lynas. 

1982. Host-parasite catalogue of the avian 

Haematozoa. Memorial University of Newfoundland 

Occasional Papers in Biology 5:1-243. 

Bishop, M. A., and G. F. Bennett. 1992. Host-parasite 

catalogue of the avian Haematozoa, Supplement 

1, and Bibliography of the avian blood-inhabiting 

Haematozoa, Supplement 2. Memorial 

University of Newfoundland Occasional Papers in 

Biology 15:1-244. 





Fallis, A. M., D. M. Davies, and M. A. Vickers. 

1951. Life history of Leucocytozoon simondi 

Mathis and Leger in natural and experimental infections 

and blood changes produced in the avian 

host. Canadian Journal of Zoology 29:305-328. 

New Book Available 

Forrester, D. J., J. M. Kinsella, J. W. Mertins, R. 

D. Price, and R. E. Turnbull. 1994. Parasitic helminths 

and arthropods of fulvous whistling-ducks 

(Dendrocygna bicolor) in southern Florida. Journal 

of the Helminthological Society of Washington 

61:84-88. 

Loven, J. S., E. G. Bolen, and B. W. Cain. 1980. 

Blood parasitemia in a South Texas wintering waterfowl 

population. Journal of Wildlife Diseases 

16:25-28. 

Martin, E. M. 1996. Tables of 1981-1990 average 

U.S. waterfowl harvest by species and county. 

Unpublished data from the Division of Migratory 

Bird Management, U.S. Fish and Wildlife Service, 

Laurel, Maryland, U.S.A. 

Nelson, E. C., and J. S. Gashwiler. 1941. Blood parasites 

of some Maine waterfowl. Journal of Wildlife 

Management 5:199-205. 

Robertson, W. B., and G. E. Woolfenden. 1992. 

Florida Bird Species: An Annotated List. Florida 

Ornithological Society Special Publication Number 

6, Gainesville, Florida, U.S.A. 260 pp. 

SAS Institute, Inc. 1989. SAS/STAT User's Guide, 

Version 6, 4th ed., Vol. 1. SAS Institute, Inc., 

Gary, North Carolina, U.S.A. 943 pp. 

Thul, J. E., D. J. Forrester, and E. C. Greiner. 1980. 

Hematozoa of Wood Ducks (Aix sponsd) in the 

Atlantic Flyway. Journal of Wildlife Diseases 16: 

383-389. 

, and T. O'Brien. 1990. Wood duck hematozoan 

parasites as biological tags: Development of 

a population assessment model. Pages 323-334 in 

Proceedings of the 1988 North American Wood 

Duck Symposium, St. Louis, Missouri, U.S.A. 

Metazoan Parasites in the Neotropics: A Systematic and Ecological Perspective. Editors Guillermo 

Salgado-Maldonado, Alfonso N. Garcia-Aldrete, and Victor M. Vidal-Martinez. 2000. Published by the 

Universidad Nacional Autonoma de Mexico (UNAM) to celebrate the 70th Anniversary of the Institute 

de Biologia. 310 pp. ISBN 968-36-8827-6. 6%" X 8Vi" soft cover. Cost: US$20.00 or MXP$200.00 per 

copy plus shipping and handling. Available from Institute de Biologia, UNAM, Secretaria Tecnica, Apartado 

Postal 70-153, CP 04510 Mexico D.F., Mexico or on-line at the internet web-page http:// 

www.ibiologia.unam.mx. Payment by check only, made payable to "Institute de Biologia, UNAM". This 

volume is a compilation of 11 original contributions by an internationally diverse group of 18 researchers 

addressing a broad range of parasite systematics and biogeographical issues in South and Central American 

vertebrates. 




Parelaphostrongylus tennis (Nematoda: Protostrongylidae) and Other 

Parasites of White-Tailed Deer (Odocoileus virginianus} in Costa Rica 

RAMON A. CARRENO,1'5 LANCE A. DuRDEN,2 DANIEL R. BROOKS,3 ARTHUR ABRAMS,4 AND 

ERIC P. HOBERG4 

1 Department of Environmental Biology, University of Guelph, Guelph, Ontario, Canada NIG 2W1 (e-mail: 

rcarreno@uoguelph.ca), . 

2 Institute of Arthropodology and Parasitology, Georgia Southern University, Statesboro, Georgia 30460, 

U.S.A. (e-mail: Idurden@gsvms2.cc.gasou.edu), 

3 Department of Zoology, University of Toronto, Toronto, Ontario, Canada M5S 3G5 (e-mail: dbrooks@ 

zoo.utoronto.ca), and 

4 Biosystematics and National Parasite Collection Unit, USDA, Agricultural Research Service, BARC East 

1180, Beltsville, Maryland 20715, U.S.A. 

ABSTRACT: Parasites were collected from 2 female white-tailed deer (Odocoileus virginianus) in the Area de 

Conservacion Guanacaste, Costa Rica, in early June 1999. Both deer were parasitized by the ticks Amblyomma 

parvum and Haemaphysalis juxtakochi as well as the hippoboscid fly, Lipoptena mazamae. One deer also hosted 

the ticks Boophilus microplus, Ixodes qffinis, and Anocentor nitens. Both deer were infected by larvae of the 

nasopharyngeal botfly Cephenemyia jellisoni, and the helminths Eucyathostomum webbi, Gongylonema pulchrum, 

Parelaphostrongylus tennis, and Paramphistomum liorchis, whereas Setaria yehi, an undescribed species 

of Ashworthius, and Onchocerca cervipedis occurred in single hosts. A cysticercus of Taenia oinissa was found 

encapsulated in the lung parenchyma of 1 host. This is the first report of these endoparasites from Central 

America. 

KEY WORDS: Ashworthius sp., biodiversity, ticks, Boophilus microplus, Gongylonema pulchrum, Haemaphysalis 

juxtakochi, Ixodes affinis, Odocoileus virginianus, white-tailed deer, helminth parasites, Parelaphostrongylus 

tenuis, cysticercus, Costa Rica. 

The white-tailed deer Odocoileus virginianus 

(Zimmermann, 1780) has a widespread Nearctic 

and Neotropical range, extending from southern 

Canada and the United States through Mexico 

and Central America to Bolivia, the Guianas, 

and northern Brazil (Reid, 1997). The subspecies 

described from Costa Rica, Odocoileus virginianus 

truei Merriam, 1898, ranges from the 

southeastern edge of Mexico to northeastern 

Panama (Whitehead, 1972; Mendez, 1984). The 

parasite fauna of O. virginianus and other cervids 

is well documented in North America 

(Walker and Becklund, 1970; Davidson et al., 

1981). However, very little information is available 

on parasites of cervids in the southern parts 

of their range, including Central America. This 

is significant because white-tailed deer are hosts 

to several serious pathogens and parasites of cervids 

and other animals, including the tick Ixodes 

scapularis Say, 1821, which is the main North 

American vector of the agent of Lyme borreli- 

5 Corresponding author. Present address: Department 

of Nematology, University of California, Davis, California 

95616, U.S.A. (e-mail: racarreno@ 

ucdavis.edu). 

177 

osis. Additionally, one of the most important 

parasites in O. virginianus is Parelaphostrongylus 

tenuis (Dougherty, 1945), the meningeal 

worm. This species is not pathogenic in O. virginianus, 

but when snails infected with its larvae 

are ingested by other ruminants such as moose 

(Alces alces (Linnaeus, 1758)), fallow deer 

(Dama dama (Linnaeus, 1758)), reindeer (Rangifer 

tarandus (Linnaeus, 1758)), and llamas 

(Lama spp.), severe neurologic disease can result 

from adult worms in the brain and central 

nervous system (Anderson, 1964, 1970; Nettles 

et al., 1977; Krogdahl et al., 1987; Rickard et 

al., 1994). 

The following report is part of a biodiversity 

inventory of eukaryotic parasites of vertebrates 

in the Area de Conservacion Guanacaste (ACG) 

in northwestern Costa Rica. 


We collected 2 adult female O. virginianus within 

the ACG, Guanacaste, Costa Rica (10°57'N; 85°48'W) 

in early June 1999. Ectoparasites were collected within 

1 hr postmortem. Internal organs were then removed, 

following procedures suggested by Nettles (1981), and 

examined for endoparasites. In addition to onsite ex- 



amination, several aliquots of abomasal and intestinal 

contents were fixed and later searched for parasites. 

Contents from the abomasum and small intestine were 

suspended in 2 liter of water. Duplicate aliquots of 200 

ml were removed, fixed in 5% formalin, and subsequently 

examined using a stereomicroscope. Digeneans 

were fixed in hot 10% formalin and stored in 70% 

ethanol. Ectoparasites and nematodes were stored in 

70% ethanol without fixation in formalin. Prior to examination, 

nematodes were cleared in either glycerine 

or phenol-alcohol. Tissues examined for sarcocysts 

were embedded in paraffin, sectioned, and stained with 

both hematoxylin and eosin and periodic acid-Schiff 

stains for diagnosis. Amphistome digeneans were prepared 

using the same methods and identified using descriptions 

by Kennedy et al. (1985) and Sey (1991). 

Voucher tick specimens are deposited in the U.S. National 

Tick Collection (USNTC), Institute of Arthropodology 

and Parasitology, Georgia Southern University, 

Statesboro, Georgia, U.S.A. Voucher specimens 

of other parasites are deposited in the U.S. National 

Parasite Collection (USNPC), United States Department 

of Agriculture, Beltsville, Maryland, U.S.A. Accession 

numbers are listed in Table 1. 

Results 

A total of 18 parasite species was found in 

the 2 deer (Table 1). Both deer hosted the hippoboscid 

louse fly Lipoptena mazamae and the 

ixodid ticks Amblyomma parvum and Haemaphysalis 

juxtakochi. We also found second- and 

third-stage larvae of Cephenemyia jellisoni 

(identified using keys in Bennett and Sabrosky 

[1962]) in the nasal sinuses of both deer. One 

deer also hosted the ticks Anocentor nitens, Boophilus 

microplus, and Ixodes affinis. 

Both deer hosted the digenean trematode Paramphistomum 

liorchis in their rumens. They also 

were infected by 3 species of nematodes: Gongylonema 

pulchrum in the submucosa of the 

esophagus, Eucyathostomum webbi in the large 

intestine, and Parelaphostrongylus tennis in the 

inner surface of the dura and from cranial sinuses 

and nerves. Meristic data for the latter 

species (Table 2) did not differ from those reported 

for P. tennis in O. virginianus from North 

America (Anderson, 1963; Carreno and Lankester, 

1993), although the esophageal lengths of 

the 2 males were greater than those reported in 

North American specimens. 

A cysticercus of Taenia omissa was found encapsulated 

in fibrous connective tissue in the 

lung parenchyma. Identification of the cysticercus 

was based on structure and measurements of 

rostellar hooks and the occurrence in deer; morphology 

of the intact cysticercus was consistent 

with observations by Rausch (1981). 

Two specimens of Setaria yehi were collected 

from 1 deer, 1 in the rectum, and the other in 

the posterior region of the body cavity. From the 

abomasal intestinal aliquots, few nematodes in 

the abomasa and none in the small intestines 

were found. A female specimen of Mazamastrongylus 

sp. occurred in the abomasum of 1 

deer. It could not be identified to species based 

on the diagnostic features within the genus. 

Hoberg (1996) demonstrated polymorphism in 

vulvar anatomy, and no males were found. In 

the abomasum of the second deer, specimens of 

an undescribed species of Ashworthius were 

found. 

Protozoan infections were not obvious in 

these animals (blood smears and fecal examinations 

were negative), except for the presence 

of unidentified sarcocysts in the hind leg muscles 

of 1 host. 

Discussion 

Parasites in white-tailed deer 


This report constitutes the first data on parasites 

of O. virginianus in Costa Rica. The ranges 

of C. jellisoni, P. liorchis, E. webbi, O. cervipedis, 

and, most importantly, P. tennis are now 

extended south of North America. These range 

extensions suggest that the parasites may also be 

present in Mexico, other parts of Central America, 

and perhaps South America, coinciding with 

the distribution of this cervid. 

There is little information on the parasites of 

cervids south of the United States. Captive O. 

virginianus from the Yucatan Peninsula have 

been reported as hosts of nematodes (Haemonchus 

sp., Cooperia spp., Isospora spp., Eimeria 

spp., Trichuris spp., Strongyloides spp.) and cestodes 

(Moniezia spp.), based on examination of 

fecal samples and fecal culture of nematode L3 

larvae (Montes-Perez et al., 1998). Several parasites 

have been listed from O. virginianus from 

Mexico and Central America, including arthropods, 

Cephenemyia sp., L. mazamae, A. nitens, 

H. juxtakochi, I. affinis, and nematodes, Haemonchus 

conforms (Rudolphi, 1802) Cobb, 

1898, and Gongylonema pulchrum, as well as 

several other species not found in the present 

study (Mendez, 1984). In South America, O. virginianus 

has been reported as a host for the 

nematodes H. contortus, Setaria sp., Oesophagostomum 

asperum Railliet and Henry, 1913, 

and Mecistocirrus sp., as well as the arthropods

CARRENO ET AL.—DEER PARASITES IN COSTA RICA 179 

Table 1. Parasites recovered from Odocoileus virginianus in Guanacaste, Costa Rica. 

Protozoa 

Sarcocystis sp. 

Parasite Deer Deer 2 Accession number 

Arthropoda: Acari 

Arnblyomma parvum Aragao, 1908 

Anocentor nitens (Neumann, 1897) 

Boophilus microplus (Canestrini, 1887) 

Haemaphysalis juxtakochi (Cooley, 1946) 

Ixodes qffinis Neumann, 1899 

Amblyomma sp. (immature stages) 

Arthropoda: Diptera 

Lipoptena mazamae Rondani, 1878 

Cephenemyia jellisoni Townsend, 1941 

Trematoda 

Paramphistomum liorchis Fischoeder, 

1901 

Cestoda 

Taenia omissa Liihe, 1910 

Nematoda 

Setaria yehi (Desset, 1966) 

Ashworthius sp. 

Gongylonenm pulchmm Molin, 1857 

Eucyathostomum webbi Pursglove, 1976 

Parelaphostrongylus tennis (Dougherty, 

1945) 

Mazamastrongylus sp. 

Onchocerca cervipedis Wehr and Dikmans, 

1935 

+ USNPC 90056 

+ + RML 122837, RML 122838 

+ RML 122838 

+ - RML 122838 

+ + RML 122837, RML 122838 

+ - RML 122838 

+ + RML 122837, RML 122838 

+ USNPC 90062, USNPC 90063 

10 USNPC 90053, USNPC 90054 

+ (> 1,000) + (


Table 2. Morphometric measurements for P. tennis 

recovered from O. virginianus in Guanacaste, Costa 

Rica; measurements are given in microns (u.m) unless 

otherwise noted. 

Males 

Length (mm) 

Width (posterior to esophagus) 

Esophagus length 

Nerve ring from anterior 

Excretory pore from anterior 

Gubernaculum length 

Crura length 

Spicules 

Spicule branch 

Females 

Length (mm) 

Width (posterior to esophagus) 

Nerve ring from anterior 

Excretory pore from anterior 

Vulva from posterior 

Anus from posterior 

N Mean Range 

1 

2 

2 

2 

1 

3 

3 

6 

2 

1 

2 

2 

2 

8 

8 

47.00 

170.00 

807.50 

120.00 

110.00 

117.00 

36.17 

225.00 

7 1 .50 

105.50 

210.00 

105.50 

108.50 

185.50 

56.96 

— 

140-200 

795-820 

107-133 

— 

110-131 

25.7-44.0 

211-236 

70-73 

— 

190-230 

102-109 

106-111 

165-214 

45.5-79.0 

gated geographically and seasonally. For example, 

in Alabama, Durden et al. (1991) recovered 

4 species of ticks from 537 O. virginianus examined 

during winter (November-January), but 

only 2 of these species (/. scapularis and Dermacentor 

albipictus (Packard, 1869)) were common. 

In Alberta, Samuel et al. (1980) recorded 

just 1 species of tick (D. albipictus) from 148 

O. virginianus examined in all months, whereas 

Smith (1977) recorded 7 tick species from this 

host from 12 southeastern states, again with just 

2 species (/. scapularis and Amblyomma americanum 

(Linnaeus, 1758)) being common. In 

southern Texas, Samuel and Trainer (1970) recovered 

6 species of ticks from 404 O. virginianus 

examined, with 3 of these species (A. 

americanum, Amblyomma inornatum Banks, 

1909, and Amblyomma maculatum Koch, 1844) 

being prevalent. 

The known geographical range of A. parvutn 

extends from Mexico to Argentina. Throughout 

this range it parasitizes a wide variety of mammals 

(Fairchild et al., 1966; Jones et al., 1972). 

In Panama, Fairchild et al. (1966) recorded A. 

parvum from white-tailed deer, cattle, domestic 

cats, sloth, human, anteater (Tamandua sp.), and 

cotton rats (Sigmodon spp.). There is an unpublished 

USNTC record of this tick from a horse 

in Costa Rica. 

The tropical horse tick, A. nitens, is a pest of 


equines in the neotropics and is the main vector 

of Babesia equi (Laveran, 1901), the protozoan 

that causes equine piroplasmosis (Strickland et 

al., 1976). In adjoining Panama, Fairchild et al. 

(1966) reported this tick from horses, cattle, and 

deer. The USNTC contains 8 collections of A. 

nitens from Costa Rica: 6 from horses, 1 from 

a domestic cat, and 1 from a rabbit (Sylvilagus 

sp.). 

The southern cattle tick, B. microplus, was 

formerly widespread throughout the New World 

as a major pest of domestic cattle, where it 

caused tremendous economic damage through 

its role as a vector of Babesia bigemina (Smith 

and Kilborne, 1893), an agent of bovine piroplasmosis 

(=Texas cattle fever) (Strickland et 

al., 1976; Bram and George, 2000). This tick 

also is a vector of Anaplasma marginale Theiler, 

1910 and Babesia bovis (Babes, 1888) Starcovici, 

1893. In Costa Rica, Hermans et al. (1994) 

reported high infection rates for these 3 hemoparasites 

in B. microplus and high seroprevalences 

to them in cattle. In Panama, Fairchild et 

al. (1966) reported this tick from cattle, horses, 

pigs, and dogs, with single collections from a 

goat and a deer. The USNTC contains 23 collections 

of B. microplus from Costa Rica: 15 

from cattle, 2 from horses, 2 from vegetation, 

and 1 each from a human, a tapir (Tapirus sp.), 

a gray fox (Urocyon cinereoargenteus (Schreber, 

1775)), and a fringe-lipped bat (Trachops 

cirrhosus (Spix, 1823)). 

Deer are the preferred hosts of H. juxtakochi, 

which is widely distributed from Mexico to Argentina 

(Jones et al., 1972). However, this tick 

has occasionally also been collected from rodents, 

humans, tapirs, coatimundis (Nasua sp.), 

peccaries (Tayassu spp.), porcupines (Coendou 

spp.), and lagomorphs (Fairchild et al., 1966; 

Jones et al., 1972). The USNTC contains 1 other 

Costa Rican collection of H. juxtakochi, also 

from a white-tailed deer. 

Ixodes affinis has a disjunct geographical distribution, 

with 1 focus in the southeastern 

U.S.A. (coastal Florida, Georgia, and South Carolina), 

and the other focus extending from Mexico 

to Brazil (Fairchild et al., 1966; Durden and 

Keirans, 1996). Because it is a member of the 

Ixodes ricinus complex, several members of 

which are vectors of the Lyme disease spirochete, 

Borrelia burgdorferi Johnson, Schmid, 

Hyde et al., 1984, this tick could be an enzootic 

vector of this zoonotic pathogen. It parasitizes a

ange of host species, but most collections, especially 

of adults, are from deer and larger carnivores 

(Fairchild et al., 1966; Durden and Keirans, 

1996). The USNTC includes 5 additional 

Costa Rican collections of /. qffinis: 2 from ocelots 

(Leopardus pardalis (Linnaeus, 1758)), and 

1 each from a human, a horse, and a long-tailed 

weasel (Mustela frenata Lichtenstein, 1831). 

Digeneans 

Brokx (1984) and Mendez (1984) reported 

amphistome digeneans, Cotylophoron sp. and 

Paramphistomum cervi (Zeder, 1790), respectively, 

in the stomach of O. virginianus. Unfortunately 

no voucher specimens exist from those 

accounts, so we cannot confirm their identifications. 

The only amphistomes we found were P. 

liorchis, the species most commonly reported 

from O. virginianus in North America (Kennedy 

et al., 1985). 

Cestodes 

Taenia omissa has a broad geographic distribution 

in the Western Hemisphere, coinciding 

with the range of the cougar, Puma concolor 

(Linnaeus, 1771), and deer intermediate hosts 

including Odocoileus and Mazama in North and 

South America (Rausch, 1981; Rausch et al., 

1983). Consistent with the current study, cysticerci 

generally are found in the thoracic cavity, 

including the lungs and pericardium (Forrester 

and Rausch, 1990). Although cysticerci have 

been reported in brocket deer (Mazama cf. gouazoubira 

(Fischer, 1814)) from eastern Colombia 

(Rausch, 1981), there are apparently no prior records 

from Central America. Prevalence and intensity 

of infection in deer may be influenced by 

differences in population density of cougars 

across the range of this parasite—host assemblage 

(Forrester and Rausch, 1990). 

Nematodes 

This is the first record of P. tennis south of 

the United States. The presence of elaphostrongyline 

nematodes in cervids is a major concern 

in the translocation of these animals in wildlife 

projects and the game ranching industry (Lankester 

and Fong, 1989; Samuel et al., 1992; 

Miller and Thorne, 1993; Davidson et al., 1996). 

Parelaphostrongylus tennis is of great concern 

in future wildlife management and conservation 

practices in Central America. An overall decline 

of O. virginianus populations in Mexico and 


Central America due to overhunting and habitat 

loss (Mendez, 1984) raises the possibility of reintroducing 

deer to areas where they have been 

extirpated. The effects of P. tennis on the only 

other Central American cervid, the brocket deer 

(Mazama americana (Erxleben, 1777)), are unknown. 

As P. tennis is highly pathogenic in 

most cervids other than O. virginianus, it may 

also be pathogenic in Mazama spp. Until the 

pathogenic significance (if any) of P. tennis to 

M. americana has been determined, the translocation 

of both it and Central American O. virginianus 

may be problematic in areas inhabited 

by other cervids that may be susceptible to parelaphostrongylosis. 

The translocation of infected O. virginianus 

to areas in which P. tennis is absent may result 

in the establishment of the parasite in other areas. 

The importation of deer from Pennsylvania 

to an island off the Georgia coast may have resulted 

in the establishment of P. tennis in an area 

outside its normal range (Davidson et al., 1996). 

Similarly, the translocation of reindeer (Rangifer 

tarandus) from Norway to Newfoundland has 

led to the establishment of Elaphostrongylus 

rangiferi Mitskevitch, 1960, another pathogenic 

species, in this region of North America (Lankester 

and Fong, 1989). Other cervids such as 

red deer (Cervus elaphus Linnaeus, 1758) and 

moose (Alces alces (Linnaeus, 1758)) may also 

harbor Elaphostrongylus species, and North 

American elk have been shown to have potential 

for surviving infection with and passing larvae 

of P. tennis (Samuel et al., 1992). These studies 

indicate a need for reliable diagnosis of P. tennis 

in ungulates and the need to determine the full 

distribution of this parasite. This information can 

help to prevent the spread of the parasite to uninfected 

host populations. Cervids of western 

North America are of particular concern, as P. 

tennis has not been recorded in western states 

and provinces (Miller and Thorne, 1993). 

The presence of P. tennis in Central American 

deer is also of evolutionary significance. As yet, 

we do not know if M. americana or any of the 

South American cervids such as Pudu spp. and 

Ozotoceros spp. are hosts for protostrongylids. 

Phylogenetic analysis of the family Protostrongylidae 

and comparison with host distribution, 

however, indicates that cervids are the basal 

hosts of these parasites (Carreno and Hoberg, 

1999). Discovery of new or already described 

protostrongyles in these hosts will contribute 



further to an understanding of the evolution of 

the Protostrongylidae. 

The presence of an undescribed species of 

Ashworthius is also of interest. Species of this 

genus of the Haemonchiinae have never been 

reported from hosts in the Western Hemisphere, 

although both Haernonchus and Mecistocirrus 

are known in Central and South America. In Africa 

and Eurasia, species of Ashworthius have 

been reported from cervids and members of 2 

subfamilies of the Bovidae, but not species of 

Bos (Pike, 1969; Drozdz et al., 1998). The presence 

of Ashworthius sp. in a wild cervid in the 

Western Hemisphere may have been the result 

of introduction and colonization from bovine 

hosts following European settlement since the 

1500s (Hoberg, 1997). The host distribution for 

this genus and its apparent absence in domestic 

bovids, however, suggest otherwise. Alternatively, 

the distribution of Ashworthius spp. in Central 

America may be relatively archaic, reflecting 

an extended history with endemic cervids. Historical 

reconstruction of the biogeography of 

Ashworthius in the New World is currently hampered 

by the paucity of survey data regarding 

parasitism in wild ruminants in Central and 

South America. Additionally, it is important to 

note that superficially some species of Ashworthius 

may resemble and be confused with Haemonchus 

spp. 

Conclusions 

The purpose of parasite inventories is 2-fold. 

First, the information obtained from parasite 

faunas can contribute valuable integrative information 

on our knowledge of the biosphere that 

serves as an indicator of biodiversity (Brooks 

and Hoberg, 2000). Secondly, in the case of the 

white-tailed deer, it is important to develop an 

understanding of the distribution of potential 

pathogens of wildlife. Many wildlife pathogens 

pose a serious threat to global biodiversity and 

also include several zoonoses (Daszak et al., 

2000), and it is thus necessary to assess carefully 

the global distribution of potentially pathogenic 

parasites (Hoberg, 1997). The results of this 

study are an important contribution to biodiversity 

initiatives in Costa Rica. They have important 

implications in conservation projects in the 

region, as some parasites of white-tailed deer, 

such as P. tennis, may be pathogenic in other, 

less common endemic hosts such as M. arneri- 


The uncertainty of species identifications in 

other reports, such as the amphistome digeneans 

(Brokx, 1984; Mendez, 1984), demonstrates the 

need for deposition of suitable voucher specimens 

in documenting parasite fauna. Are we 

dealing with a widespread and relatively uniform 

parasite fauna of white-tailed deer throughout 

their range, or one that is highly localized 

depending on habitat? A lack of voucher specimens 

to confirm identifications, and a lack of 

sufficient expert taxonomists available to provide 

those identifications (the taxonomic impediment: 

Brooks and Hoberg, 2000), prevents us 

from making such assessments. And such assessments, 

in turn, are critical for management 

policy, including game farming, sport hunting, 

and the interface between wildlands and agroscape. 


We thank the scientific and technical staff of 

the ACG for support of this study, in particular 

Sigifredo Marin, Roger Blanco, Alejandro Masis, 

Maria Marta Chavarria, Felipe Chavarria, 

Guillermo Jimenez, Carolina Cano, Elda Araya, 

Fredy Quesada, Dunia Garcia, Roberto Espinoza, 

Elba Lopez, and Petrona Rios. Special thanks 

to Dr. Dan Janzen, technical adviser to the ACG, 

and to Calixto Moraga for their timely and accurate 

assistance. We are also grateful to Dr. 

Murray Lankester, Lakehead University, Thunder 

Bay, Canada, for advice on necropsy and 

collection procedures. Funds for this study were 

provided in part by an operating grant from the 

Natural Sciences and Engineering Research 

Council (NSERC) of Canada to D.R.B. Study of 

tick specimens was funded in part by National 

Institutes of Health grant Al 40729 to L.A.D. 


Anderson, R. C. 1963. The incidence, development, 

and experimental transmission of Pneiunostrongylus 

tenuis Dougherty (Metastrongyloidea: Protostrongylidae) 

of the meninges of the white-tailed 

deer (Odocoileus virginianus borealis) in Ontario. 


. 1964. Neurologic disease in moose infected 

experimentally with Pneiimostrongyliis tennis 

from white-tailed deer. Pathologia Veterinaria 1: 

289-322. 

-. 1970. Neurologic disease in reindeer (Rangifer 

tarandus tarandus) introduced into Ontario. 


Bennett, G. F., and C. W. Sabrosky. 1962. The Ne 

arctic species of the genus Cephenemyla (Diptera,

Oestridae). Canadian Journal of Zoology 40:431- 

448. 

Bram, R. A., and J. E. George. 2000. Introduction 

of nonindigenous arthropod pests of animals. 

Journal of Medical Entomology 37:1-8. 

Brokx, P. A. 1984. South America. Pages 525-546 in 

L. K. Halls, ed. White-Tailed Deer: Ecology and 

Management. Wildlife Management Institute. 

Stackpole Books, Harrisburg, Pennsylvania, 

U.S.A. 870 pp. 

Brooks, D. R., and E. P. Hoberg. 2000. Triage for 

the biosphere: the need and rationale for taxonomic 

inventories and phylogenetic studies of parasites. 

Comparative Parasitology 67:1—25. 

Capelle, K. J. 1971. Myiasis. Pages 279-305 in J. W. 

Davis and R. C. Anderson, eds. Parasitic Diseases 

of Wild Mammals. Iowa State University Press, 

Ames, Iowa, U.S.A. 

Carreno, R. A., and E. P. Hoberg. 1999. Evolutionary 

relationships among the Protostrongylidae 

(Nematoda: Metastrongyloidea) as inferred from 

morphological characters, with consideration of 

parasite—host coevolution. Journal of Parasitology 

85:638-648. 

, and M. W. Lankester. 1993. Additional information 

on the morphology of the Elaphostrongylinae 

(Nematoda: Protostrongylidae) of North 

American Cervidae. Canadian Journal of Zoology 

71:592-600. 

Daszak, P., A. A. Cunningham, and A. D. Hyatt. 

2000. Emerging infectious diseases of wildlife— 

threats to biodiversity and human health. Science 

287:443-449. 

Davidson, W. R., G. L. Doster, and R. C. Freeman. 

1996. Parelaphostrongylus tennis on Wassaw Island, 

Georgia: a result of translocating whitetailed 

deer. Journal of Wildlife Diseases 32:701- 

703. 

, F. A. Hayes, V. F. Nettles, and F. E. Kellogg 

(eds.). 1981. Diseases and Parasites of White- 

Tailed Deer. Miscellaneous Publication No. 7, Tall 

Timbers Research Station, Tallahassee, Florida, 

U.S.A. 458 pp. 

Drozdz, J., A. W. Demiaszkiewicz, and J. Lachowicz. 

1998. Ashworthius sidemi (Nematoda, Trichostrongylidae) 

a new parasite of the European bison 

Bison bonasus (L.) and the question of independence 

of A. gagarini. Acta Parasitologica 43:75- 

80. 

Durden, L. A., and J. E. Keirans. 1996. Nymphs of 

the genus Ixodes (Acari: Ixodidae) of the United 

States: taxonomy, identification key, distribution, 

hosts, and medical/veterinary importance. Thomas 

Say Publications in Entomology: Monographs, 

Entomological Society of America, Lanham, 

Maryland, U.S.A. 95 pp. 

, S. Luckhart, G. R. Mullen, and S. Smith. 

1991. Tick infestations of white-tailed deer in Alabama. 


Fairchild, G. B., G. M. Kohls, and V. J. Tipton. 

1966. The ticks of Panama (Acarina: Ixodoidea). 

Pages 167-219 in R. L. Wenzel and V. J. Tipton, 

eds. Ectoparasites of Panama. Field Museum of 

Natural History, Chicago, Illinois, U.S.A. 


Forrester, D. J., and R. L. Rausch. 1990. Cysticerci 

(Cestoda: Taeniidae) from white-tailed deer, Odocoileus 

virginianus, in southern Florida. Journal of 


Hermans, P., R. H. Dwinger, G. M. Buening, and 

M. V. Herrero. 1994. Seasonal incidence and hemoparasite 

infection rates of ixodid ticks (Acari, 

Ixodidae) detached from cattle in Costa Rica. Revista 

de Biologia Tropical 42:623-632. 

Hoberg, E. P. 1996. Emended description of Mazamastrongylus 

peruvianus (Nematoda: Trichostrongylidae), 

with comments on the relationships of 

the genera Mazamastrongylus and Spiculopteragia. 


. 1997. Parasite biodiversity and emerging 

pathogens: a role for systematics in limiting impacts 

on genetic resources. Pages 71-83 in K. E. 

Hoagland and A. Y. Rossman, eds. Global Genetic 

Resources: Access, Ownership and Intellectual 

Property Rights. Association of Systematics Collections, 

Washington, D.C., U.S.A. 

Jones, E. K., C. M. Clifford, J. E. Keirans, and G. 

M. Kohls. 1972. The ticks of Venezuela (Acarina: 

Ixodoidea) with a key to the species of Amblyornma 

in the western hemisphere. Brigham Young 

University Science Bulletin, Biological Series 

17(4): 1-40. 

Kennedy, M. J., M. W. Lankester, and J. B. Snider. 

1985. Paramphistomum cervi and Paramphistomum 

liorchis (Digenea: Paramphistomatidae) in 

moose, Alces alces, from Ontario, Canada. Canadian 


Krogdahl, D. W., J. P. Thilstead, and S. K. Olsen. 

1987. Ataxia and hypermetria caused by Parelaphostrongylus 

tennis infection in llamas. Journal 

of the American Veterinary Medical Association 

190:191-193. 

Lankester, M. W., and D. Fong. 1989. Distribution 

of elaphostrongyline nematodes (Metastrongyloidea: 

Protostrongylidae) in Cervidae and possible 

effects of moving Rangifer spp. into and within 

North America. Alces 25:133-145. 

Maa, T. C. 1963. Genera and species of Hippoboscidae 

(Diptera): types, synonymy, habitats and natural 

groupings. Pacific Insects Monograph 6:1- 

186. 

Mendez, E. 1984. Mexico and Central America. Pages 

513-524 in L. K. Halls, ed. White-Tailed Deer: 

Ecology and Management. Wildlife Management 

Institute. Stackpole Books, Harrisburg, Pennsylvania, 

U.S.A. 870 pp. 

Miller, M. W., and E. T. Thorne. 1993. Captive cervids 

as potential sources of disease for North 

America's wild cervid populations: avenues, implications, 

and preventive management. Transactions 

of the North American Wildlife and Natural 

Resources Conference 58:460-467. 

Montes-Perez, R. C., R. I. Rodriguez-Vivas, J. F. de 

J. Torres-Acosta, and L. G. Ek-Pech. 1998. Seguimiento 

anual de la parasitosis gastrointestinal 

de venados cola blanca Odocoileus virginianus 

(Artiodactyla: Cervidae) en cautiverio en Yucatan, 

Mexico. Revista de Biologia Tropical 46:821-827. 

Nettles, V. F. 1981. Necropsy procedures. Pages 6-16 



in W. R. Davidson, F. A. Hayes, V. F. Nettles, and 

F. E. Kellogg, eds. Diseases and Parasites of 

White-Tailed Deer. Miscellaneous Publication no. 

7, Tall Timbers Research Station. Southeastern 

Cooperative Wildlife Disease Study, Athens, 

Georgia, U.S.A. 458 pp. 

, A. K. Prestwood, and R. D. Smith. 1977. 

Cerebrospinal parelaphostrongylosis in fallow 

deer. Journal of Wildlife Diseases 13:440-444. 

Pike, A. 1969. A revision of the genus Ashworthius 

Le Roux, 1930 (Nematoda: Trichostrongylidae). 

Journal of Helminthology 43:135-144. 

Rausch, R. L. 1981. Morphological and biological 

characteristics of Taenia rileyi Loewen, 1929 

(Cestoda: Taeniidae). Canadian Journal of Zoology 

59:653-666. 

, C. Maser, and E. P. Hoberg. 1983. Gastrointestinal 

helminths of the cougar, Felis concolor 

L., in northeastern Oregon. Journal of Wildlife 

Diseases 19:14-19. 

Reid, F. A. 1997. A Field Guide to the Mammals of 

Central America and Southeast Mexico. Oxford 

University Press, Oxford, U.K. 334 pp. 

Rickard, L. G., B. B. Smith, E. J. Gentz, A. A. 

Frank, E. G. Pearson, L. L. Walker, and M. J. 

Pybus. 1994. Experimentally induced meningeal 

worm (Parelaphostrongylus tennis) infection in 

the llama (Lama glama): clinical evaluation and 

implications for parasite translocation. Journal of 

Zoo and Wildlife Medicine 25:390-402. 

Samuel, W. M., E. R. Grinnell, and A. J. Kennedy. 

1980. Ectoparasites (Mallophaga, Anoplura, Acari) 

on mule deer, Odocoileus hemionus, and whitetailed 

deer, Odocoileus virginianus, of Alberta, 

Canada. Journal of Medical Entomology 17:15- 

17. 

, M. J. Pybus, D. A. Welch, and C. J. Wilke. 

1992. Elk as a potential host for meningeal worm: 


implications for translocation. Journal of Wildlife 

Management 56:629-639. 

-, and D. O. Trainer. 1970. Amblyomma (Ac- 

arina: Ixodidae) on white-tailed deer, Odocoileus 

virginianus (Zimmermann), from South Texas 

with implications for theileriasis. Journal of Medical 

Entomology 7:567-574. 

Sey, O. 1991. CRC Handbook of the Zoology of Amphistomes. 

CRC Press Inc., Boca Raton, Florida, 

U.S.A. 480 pp. 

Smith, J. S. 1977. A survey of ticks infesting whitetailed 

deer in 12 southeastern states. M.S. Thesis, 

University of Georgia, Athens, Georgia, U.S.A. 61 

pp. 

Strickland, R. K., R. R. Gerrish, J. L. Hourrigan, 

and G. O. Schubert. 1976. Ticks of Veterinary 

Importance. USD A, APHIS Handbook No. 485. 

U.S. Government Printing Office, Washington. 

D.C., U.S.A. 122 pp. 

, , and J. S. Smith. 1981. Arthropods. 

Pages 363-389 in W. R. Davidson, F. A. Hayes, 

V. F. Nettles, and F. E. Kellogg, eds. Diseases and 

Parasites of White-Tailed deer. Miscellaneous 

Publication no. 7, Tall Timbers Research Station. 

Southeastern Cooperative Wildlife Disease Study, 

Athens, Georgia, U.S.A. 458 pp. 

Tonn, R. J., G. M. Kohls, and K. Arnold. 1963. 

Ectoparasites of birds and mammals of Costa 

Rica. 2. Ticks. Revista de Biologia Tropical 11: 

217-220. 

Walker, M. L., and W. W. Becklund. 1970. Checklist 

of the internal and external parasites of deer, 

Odocoileus hemionus and O. virginianus, in the 

United States and Canada. Special Publication No. 

1, Index Catalogue of Medical and Veterinary Zoology, 

U.S. Government Printing Office, Washington, 

D.C., U.S.A. 45 pp. 

Whitehead, G. K. 1972. Deer of the World. Constable 

and Company, London, U.K. 194 pp.



Cepedietta michiganensis (Protozoa) and Batracholandros 

magnavulvaris (Nematoda) from Plethodontid Salamanders in West 

Virginia, U.S.A. 

JAMES E. JOY' AND ROBERT B. TUCKER 

Department of Biological Sciences, Marshall University, Huntington, West Virginia 25755, 

U.S.A. (e-mail: joy@marshall.edu) 

ABSTRACT: The gastrointestinal tracts of 38 plethodontid salamanders (25 Plethodon punctatus and 13 Plethodon 

wehrlei), collected at high elevation sites in Pendleton and Randolph counties, West Virginia, U.S.A., were 

examined for parasites in 1996. Sixty percent of P. punctatus and 23% of P. wehrlei were infected by the ciliate 

Cepedietta michiganensis, while prevalences of the nematode Batracholandros magnavulvaris were 52% and 

30% for P. punctatus and P. wehrlei, respectively. Mean intensities were 3.4 nematodes per infected host for 

P. punctatus and 4.0 for P. wehrlei. Only 2 of 61 B. magnavulvaris collected were males. This is the first report 

of parasites from these plethodontid species and the first record of C. michiganensis from West Virginia. 

KEY WORDS: Batracholandros magnavulvaris, Cepedietta michiganensis, Plethodon punctatus, Plethodon 

wehrlei, Ciliata, Nematoda, West Virginia, U.S.A. 

The Cow Knob salamander, Plethodon punctatus 

Highton, 1972, is a large plethodontid salamander 

known only from higher elevations 

(>730 m) of the Shenandoah and Great North 

mountains in Augusta, Rockingham, and Shenandoah 

counties of Virginia (Buhlmann et al., 

1988; Conant and Collins, 1991). The range of 

this rare species in West Virginia is restricted to 

higher elevations (>730 m) of Hardy and Pendleton 

counties in the eastern panhandle. All 25 

P. punctatus examined for this study were collected 

on Shenandoah Mountain in Pendleton 

County, West Virginia, from June through August 

1996, under a permit granted by the West 

Virginia Division of Natural Resources 

(WVDNR) and written permission from the U.S. 

Fish and Wildlife Service. 

Wehrle's salamander, Plethodon wehrlei 

Fowler and Dunn, 1917, is considered a near 

sibling of P. punctatus, and has been recorded 

from a wide range of elevations in 28 of West 

Virginia's 55 counties (Green and Pauley, 1987). 

The geographic range of P. wehrlei extends 

from southwestern New York to northwestern 

North Carolina (Conant and Collins, 1991). All 

13 P. wehrlei individuals used in this study were 

collected from Shaver's Mountain in Randolph 

County, West Virginia, from May through August 

1996, under a permit from the WVDNR. 

The original purpose of collecting these plethodontid 

species was to obtain reproductive and 

Corresponding author. 

185 

ecological data for use in forest and wildlife 

management plans. Because there are no published 

reports of parasites from either species, 

these collections also offered the opportunity to 

examine them for parasites. 


All salamanders were anesthetized in Chloretone® 

within 48 hr of collection. Snout-to-vent lengths (SVL) 

were measured with vernier calipers to the nearest 0.1 

mm. Salamanders were killed by decapitation, sexed, 

and the small and large intestines were removed for 

examination. The SVL for P. punctatus (ft/mean in mm 

± 1 SD) was 18/63.8 ± 6.9 for males and 7/65.3 ± 

7.9 for females. Because the difference in mean SVL 

for males versus females was not significant (£0.05,23 = 

0.469), individuals of both host sexes were combined 

to calculate prevalence of ciliate infection and prevalence 

and mean intensity of nematode infection. The 

SVL for P. wehrlei (/z/mean in mm ± 1 SD) was 6/ 

59.8 ± 9.3 for males and 7/59.9 ± 11.7 for females. 

Again the difference in mean SVL for males versus 

females was not significant (?oo5,ii = 0.017), and the 

data for both host sexes were combined for calculations 

of prevalence and mean intensity. 

During necropsy it was evident that both salamander 

species harbored astomatous ciliates and nematodes. 

Whole mounts of these ciliates and nematodes were 

prepared by staining in Semichon's acetic carmine, dehydrating 

in an ethanol series, clearing in xylene, and 

mounting in Permount®. Voucher specimens have been 

deposited in the U.S. National Parasite Collection, 

Beltsville, Maryland 20705, U.S.A., under accession 

numbers USNPC 89838 (Cepedietta michiganensis) 

and USNPC 89839 (Batracholandros magnavulvaris). 

Results 

The astomatous ciliate, C. michiganensis 

(Woodhead, 1928) Corliss, de Puytorac, and 



Table 1. Published reports of amphibian hosts harboring Cepedietta michiganensis, with prevalences (P) 

by locality; host scientific names, taxonomic authorities, and dates after Frost (1985). 

Host common name and species 

Arnbystoma jeffersonianum (Green, 1 827) (Jefferson salamander) 

Ambystoma opacum (Gravenhorst, 1807) (marbled salamander) 

Desmognathus fitscus (Green, 1818) (northern dusky 

salamander) 

Desmognathus phoca (Matthes, 1855) (seal salamander) 

Eurycea bislineata (Green, 1818) (northern two-lined 

salamander) 

Eurycea cirrigera (Green, 1830) (southern two-lined 

salamander) 

Eurycea guttolineata (three-lined salamander):!: 

Hemidactylium scutatum (Temminck and Schlegel, 

1838) (four-toed salamander) 

Plethodon albagula Grobman, 1944 (western slimy salamander) 

Plethodon cinereus (Green, 1818) (red-backed salamander) 

Plethodon fourchensis Duncan and Highton, 1979 

(Fourche Mountain salamander) 

Plethodon glutinosus (Green, 1818) (northern slimy salamander) 

Plethodon jordani Blatchley, 1901 (Jordan's salamander) 

Plethodon ouachitae Dunn and Heinze, 1933 (Rich 

Mountain salamander) 

Plethodon punctatus Highton, 1972 (Cow Knob salamander) 

Plethodon wehrlei Fowler and Dunn, 1917 (Wehrle's 

salamander) 

Pseudotriton moiitanus Baird, 1849 (midland mud salamander) 

Rana clamitans Latreille, 1801 (southern green frog) 

Rana sylvatica LeConte, 1825 (wood frog) 

P(%) 

1/NI* 

7 

0/12| 

6 

9 

6 

18 

16 

0/2 It 

70 

1/NI* 

9 

18 

33 

5 

1 l/7t 

41-72§ 

20 

14 

0-49 

48 

60 

23 

33 

— 

20 

Michigan 

North Carolina 


Locality 



New Hampshire 




Michigan 

Reference 

Woodhead, 1928 

Rankin, 1937a 

Rankin, 1937a 

Rankin, 1937a 

Rankin, 1937a 

Muzzall et al., 1997 

Mann, 1932 

Mann, 1932 

Rankin, 1937a 

Woodhead, 1928 

Arkansas McAllister et al., 1993 

Ohio 

New Hampshire 

Michigan 

Arkansas 



Tennessee 

Arkansas 

Great Smoky Mts. of Tennessee- 


Tennessee 

Arkansas 

West Virginia 

West Virginia 


Ohio 

Ohio 

Hazard, 1937 

Muzzall et al., 1997 

Muzzall, 1990 

Winter et al., 1986 

Mann, 1932 

Rankin, 1937a 

Powders, 1970 


Powders, 1967 

Powders, 1970 


Present study 

Present study 

Rankin, 1937a 

Odlaug, 1954 

Hazard, 1937 

* Single individual infected/sample size not included. 

t Prevalence in Durham, North Carolina, U.S.A. area of the central Piedmont/prevalence in the mountains. 

:!: Eurycea guttolineata, the three-lined salamander, is considered a subspecies of the long-tailed salamander, E. longicauda 

(Green, 1818). 

§ Prevalences inversely related to altitude and higher in fall months. 

Lorn, 1965, was found in 60% (15/25) and 23% 

(3/13) of P. punctatus and P. wehrlei, respectively 

(Table 1). This ciliate species was primarily 

aggregated in the duodenum of both host 

species, either free in the lumen or attached to 

the intestinal epithelium. Cepedietta michiganensis 

individuals were often so numerous that 

they appeared to occlude the duodenum; however, 

because there was no gross distention of 

this organ it was unlikely that any blockage of 


food materials actually occurred. No damage to 

cells of the host's intestinal epithelium was evident. 

There were a few instances where small 

numbers of ciliates were found in the middle and 

posterior small intestine, large intestine, or gall 

bladder of P. punctatus, as well. 

The oxyurid nematode, Batracholandros 

magnavulvaris (Schad, 1960) Fetter and Quentin, 

1976, was found in the large intestine of 

52% (13/25) P. punctatus and 31% (4/13) P.

JOY AND TUCKER—SALAMANDER PARASITES 187 

Table 2. Published reports of hosts harboring Batracholandros magnavulvaris, with prevalences (P) and 

mean intensities (x) by locality; host scientific names, taxonomic authorities, and dates after Frost (1985). 

Host common name and species 

Aneides flavipunctatus (Strauch, 1870) (black salamander) 

Aneides aeneus Cope and Packard, 1881 (green salamander) 

Desmognathus brimleyorum Stejneger, 1895 (Ouachita 

dusky salamander) 

Desmognathus fuscus (Green, 1818) (northern dusky salamander) 

Desmognathus monticola Dunn, 1916 (seal salamander) 

Desmognathus ochrophaeus Cope, 1959 (Allegheny 

Mountain dusky salamander) 

Desmognathus phoca (Matthes, 1855) (seal salamander) 

Desmognathus quadrimaculatus Holbrook, 1840 (blackbellied 

salamander) 

Eurycea bislineata (Green, 1818) (northern two-lined 

salamander) 

Eurycea guttolineata (three-lined salamander) 

Eurycea lucifuga Rafinesque, 1822 (cave salamander) 

Leiirognathus marmoratus Moore, 1 899 (shovel-nosed 

salamander) 

Notophthalmus viridescens (Rafinesque, 1820) (red-spotted 

newt, red eft) 

Notophthalmus viridescens (red-spotted newt, adult) 

Plethodon caddoensis Pope and Pope, 1951 (Caddo 


Plethodon cinereus (Green, 1818) (red-backed salamander) 

Plethodon fourchensis Duncan and Highton, 1 979 

(Fourche Mountain salamander) 

Plethodon glutinosus (Green, 1818) (northern slimy salamander) 

Plethodon ouachitae Dunn and Heinze, 1933 (Rich 


Plethodon punctatus Highton, 1 972 (Cow Knob salamander) 

Plethodon serratus Grobman, 1944 (southern red-backed 

salamander) 

Plethodon wehrlei Fowler and Dunn, 1917 (Wehrle's salamander) 

Plethodon yonahlossee Dunn, 1917 ( Yonahlossee salamander) 

P (%) 

50 

24 

77 

30 

48 

— 

10 

6 

27 

— 

48 

60 

50 

23 

14 

40 

14 

25 

15 

7 

31 

27 

11 

2 

0/64± 

100 

6 

100 

8 

12 

0/2* 

50 

28 

— 

9 

33 

0/3$ 

14 

52 

22 

31 

33 

A' 

3 

5 

3 


wehrlei individuals (Table 2). Mean intensities 

(±1 SD) were 3.4 (±1.9) and 4.0 (±4.0) for P. 

punctatus and P. wehrlei, respectively. Of the 61 

B. magnavulvaris individuals collected, only 2 

(both in P. wehrlei) were males. 

Discussion 

Prevalences of 60% and 23% for C. michiganensis 

in P. punctatus and P. wehrlei from 

West Virginia fall within ranges cited by previous 

investigators for this species (Table 1). Finding 

C. michiganensis species throughout the intestinal 

tract, with heavy aggregations in the duodenum, 

is consistent with the observations of 

Winter et al. (1986), who noted that C. michiganensis 

could be found throughout the intestine 

of Plethodon ouachitae Dunn and Heinze, 1933, 

but was concentrated in the anterior third of this 

organ. There was a single case of C. michiganensis 

infection in the gallbladder of P. punctatus, 

but ciliates were not attached to the epithelium 

of the gall bladder. Winter et al. (1986) 

also reported C. michiganensis from the gall 

bladder of P. ouachitae, adding that the ciliates 

were not attached to the epithelium. Three other 

reports mentioned occurrence of this ciliate in 

the host's gall bladder: Muzzall (1990) for Plethodon 

cinereus (Green, 1818); McAllister et al. 

(1993) for Plethodon albagula Grobman, 1944; 

and Muzzall et al. (1997) for Eurycea bislineata 

(Green, 1818) and P. cinereus. Previous reports 

of amphibian hosts harboring species of C. michiganensis 

are summarized in Table 1. 

Prevalences of 52% and 31% recorded in the 

present study for B. magnavulvaris in P. punctatus 

and P. wehrlei, respectively, are not unusual. 

This nematode species exhibits little host 

specificity and is found in widely varying prevalences 

(McAllister et al., 1995), an observation 

supported by the findings of other investigators 

summarized in Table 2. McAllister et al. (1995) 

also reported that prevalence for B. magnavulvaris 

in Desmognathus brimleyorum Stejneger, 

1895 varies seasonally, being highest (34%) in 

mid-March versus only 17% for late May. While 

we had relatively few salamanders in our sample, 

all B. magnavulvaris individuals were collected 

from 13 of 21 P. punctatus in June and 

July. Similarly, 4 of the 9 P. wehrlei examined 

in May through July were infected. None of the 

8 plethodontids (4 P. punctatus and 4 P. wehrlei) 

sampled in August were infected, suggesting 

that the variations in prevalence by season noted 


by McAllister et al. (1995) may be a normal 

pattern. Mean intensities of infection at 3.4 and 

4.0 for P. punctatus and P. wehrlei, respectively, 

were relatively high compared with previous reports 

(Table 2). Only 2 B. magnavulvaris individuals 

collected in the present study were 

males. This heavily female-biased sex ratio for 

B. magnavulvaris is similar to the observations 

of previous investigators (Dyer et al., 1980; 

Muzzall, 1990; Joy et al., 1993; McAllister et 

al., 1995). 


We are grateful to William Tolin, U.S. Fish 

and Wildlife Service, for granting the requisite 

collection permit for P. punctatus, and to Thomas 

Pauley for reviewing this manuscript. 


Buhlmann, K. A., C. A. Pague, J. C. Mitchell, and 

R. B. Glascow. 1988. Forestry operations and terrestrial 

salamanders: techniques in a study of the 

Cow Knob salamander, Plethodon punctatus. Pages 

38-44 in R. C. Szaro, K. E. Severson, and D. 

R. Patton, eds. Management of Amphibians, Reptiles 

and Mammals in North America. USDA Forest 

Service, Rocky Mountain Forest and Range 

Experiment Station, Fort Collins, Colorado, 

U.S.A. Technical Report RM-166. 

Bursey, C. R., and D. R. Schibli. 1995. A comparison 

of the helminth fauna of two Plethodon cinereus 

populations. Journal of the Helminthological Society 

of Washington 62:232-236. 

Conant, R., and J. T. Collins. 1991. A Field Guide 

to Reptiles and Amphibians of Eastern and Central 

North America. Houghton Mifflin Company, 

Boston, Massachusetts, U.S.A. 450 pp. 

Dunbar, J. R., and J. D. Moore. 1979. Correlations 

of host specificity with host habitat in helminths 

parasitizing the plethodontids of Washington 

County, Tennessee. Journal of the Tennessee 

Academy of Science 54:106-109. 

Dyer, W. G. 1991. Helminth parasites of amphibians 

from Illinois and adjacent midwestern states. 

Transactions of the Illinois State Academy of Science 

84:1-19. 

, R. A. Brandon, and R. L. Price. 1980. Gas 

trointestinal helminths in relation to sex and age 

of Desmognathus fuxcus (Green, 1818) from Illinois. 

Proceedings of the Helminthological Society 


-, and S. B. Peck. 1975. Gastrointestinal parasites 

of the cave salamander, Eurycea lucifuga Rafinesque, 

from the southeastern United States. Canadian 


Ernst, E. M. 1974. The parasites of the red-backed 

salamander, Plethodon cinereus. Bulletin of the 

Maryland Herpetological Society 10:108-1 14. 

Fischthal, J. H. 1955. Ecology of worm parasites in

south-central New York salamanders. American 

Midland Naturalist 53:176-183. 

Frost, D. R. 1985. Amphibian Species of the World: 

A Taxonomic and Geographic Reference. Allen 

Press, Inc. and The Association of Systematics 

Collections, Lawrence, Kansas, U.S.A. 732 pp. 

Goater, T. M., G. W. Esch, and A. O. Bush. 1987. 

Helminth parasites of sympatric salamanders: ecological 

concepts at infracommunity, component 

and compound community levels. American Midland 

Naturalist 118:289-300. 

Green, N. B., and T. K. Pauley. 1987. Amphibians 

and Reptiles in West Virginia. University of Pittsburgh 

Press, Pittsburgh, Pennsylvania, U.S.A. 241 

pp. 

Hazard, F. O. 1937. Two new host records for the 

protozoan Haptophyra michiganensis Woodhead. 


Joy, J. E., T. K. Pauley, and M. L. Little. 1993. 

Prevalence and intensity of Thelandros magncivulvaris 

and Omeia papillocaucia (Nematoda) in 

two species of desmognathine salamanders from 

West Virginia. Journal of the Helminthological 


Lehmann, D. L. 1954. Some helminths of West Coast 

urodeles. Journal of Parasitology 40:231. 

Mann, D. R. 1932. The ecology of some North Carolina 

salamanders with special reference to their 

parasites. M.A. Thesis, Duke University, Durham, 

North Carolina, U.S.A. 52 pp. 

McAllister, C. T., C. R. Bursey, S. J. Upton, S. E. 

Trauth, and D. B. Conn. 1995. Parasites of' Desmognathus 

briinleyorum (Caudata: Plethodontidae) 

from the Ouachita Mountains of Arkansas 

and Oklahoma. Journal of the Helminthological 


, S. J. Upton, and S. E. Trauth. 1993. Endoparasites 

of western slimy salamanders, Plethodon 

albagula (Caudata: Plethodontidae), from 

JOY AND TUCKER—SALAMANDER PARASITES 189 

Arkansas. Journal of the Helminthological Society 


Muzzall, P. M. 1990. Endoparasites of the red-backed 

salamander, Plethodon c. cinereus, from southwestern 

Michigan. Journal of the Helminthological 


, C. R. Peebles, and T. M. Burton. 1997. Endoparasites 

of plethodontid salamanders from Paradise 

Brook, New Hampshire. Journal of Parasitology 

83:1193-1195. 

Odlaug, T. O. 1954. Parasites of some Ohio Amphibia. 

Ohio Journal of Science 54:126-128. 

Powders, V. N. 1967. Altitudinal distribution of the 

astomatous ciliate Cepedietta michiganensis 

(Woodhead) in a new host, Plethodon jordani 

Blatchley. Transactions of the American Microscopical 

Society 86:336—338. 

. 1970. Altitudinal distribution of the protozoan 

Cepedietta michiganensis in the salamanders 

Plethodon glutinosus and Plethodon jordani in 

eastern Tennessee. American Midland Naturalist 

83:393-403. 

Rankin, J. S., Jr. 1937a. An ecological study of parasites 

of some North Carolina salamanders. Ecological 

Monographs 7:169-270. 

. 1937b. New helminths from North Carolina 

salamanders. Journal of Parasitology 23:29-42. 

Schad, G. A. 1963. Thelandros magnavidvaris (Rankin, 

1937) Schad, 1960 (Nematoda: Oxyuroidea) 

from the green salamander, Aneides aeneus. Canadian 


Walton, A. C. 1940. Some nematodes from Tennessee 

Amphibia. Journal of the Tennessee Academy of 

Science 15:402-405. 

Winter, E. A., W. M. Zawada, and A. A. Johnson. 

1986. Comparison of the symbiotic fauna of the 

family Plethodontidae in the Ouachita Mountains 

of western Arkansas. Proceedings of the Arkansas 

Academy of Sciences 40:82-85. 

Woodhead, A. F. 1928. Haptophyra michiganensis sp. 

nov., a protozoan parasite of the four-toed salamander. 





Some Adult Endohelminths Parasitizing Freshwater Fishes from the 

Atlantic Drainages of Nicaragua 

M. LEOPOLDINA AGUIRRE-MACEDO, M TOMAS ScHOLZ,1-2 DAVID GONZALEZ-SOUS,' 

VICTOR M. VIDAL-MARTINEZ,' PETR PosEL,3 GREGORY ARJONA-TORRES,' 

SVETLANA DUMAILO,3 AND EDUARDO SlU-ESTRADA3 

1 Laboratorio de Parasitologfa, Centre de Investigacion y Estudios Avanzados del Institute Politecnico 

Nacional (CINVESTAV-IPN), Unidad Merida, Carretera Antigua a Progreso Km 6, A.P. 73 Cordemex, C.P. 

97310, Merida, Yucatan, Mexico (e-mail: leo@mda.cinvestav.mx), 

2 Institute of Parasitology, Academy of Sciences of the Czech Republic, Branisovska 31, 370 05 Ceske 

Budejovice, Czech Republic (e-mail: tscholz@paru.cas.cz), and 

3 Bluefields Indian and Caribbean University, P.O. Box 88, Bluefields, Nicaragua (e-mail: bicu@ibw.com.ni) 

ABSTRACT: Adults of 12 endoparasitic helminths were recorded from 8 freshwater fish species from the South 

Atlantic Autonomous Region, Nicaragua: 8 digeneans Crassicutis cichlasomae, Magnivitellinum simplex, Oligogonotylus 

manteri, Prosthenhystera obesa, Saccocoelioides sogandaresi, Saccocoelioides sp. 1, Saccocoelioides 

sp. 2, and Allocreadiidae gen. sp. ("Crepidostomum" sp.); 3 nematodes Procamallanus (Spirocamallanus) 

rebecae, Procamallanus (Spirocamallanus) neocaballeroi, and Rhabdochona kidderi kidderi; and 1 acanthocephalan 

Neoechinorhynchus golvani. Comparative measurements among 5. sogandaresi from Poecilia velifera, 

Saccocoelioides sp. 1 from Cichlasoma maculicaiida, and Saccocoelioides sp. 2 from Astyanax fasciatus, as well 

as drawings of the 2 latter species are given for future reference. All but C. cichlasomae and O. manteri are 

reported from Nicaragua for the first time, and most taxa also represent new geographical records for Central 

America. The majority of species have previously been found in freshwater fishes from southeastern Mexico, 

which indicates a close similarity of the helminth faunas of both regions, in accordance with previous data on 

the larval stages of endohelminths and gill monogeneans. 

KEY WORDS: Helminths, parasites, Digenea, Nematoda, Acanthocephala, freshwater fishes, Nicaragua. 

During a short visit by 4 of the authors 

(M.L.A.M., T.S., V.M.V.M., and G.A.T.) to Nicaragua 

in March 1999, brackish and freshwater 

fishes from the Autonomous Region of the 

South Atlantic were examined for helminth parasites. 

Because information on helminths parasitizing 

freshwater fishes in Nicaragua is limited 

to the report by Watson (1976), in which several 

species of trematodes from Lake Nicaragua were 

reported, a list of adult endohelminths is provided, 

with morphological data on some taxa. The 

results of a survey of larval stages of endohelminths 

found in the same fish hosts and ancyrocephaline 

monogeneans from the gills of cichlids, 

have already been published (Aguirre-Macedo 

et al., 2001; Vidal-Martinez, Scholz, and 

Aguirre-Macedo, 2001). 


A total of 56 fish of the following 8 species was 

examined: tetra Astyanax fasciatus (Cuvier, 1819) (8 

specimens examined) (family Characidae); pastel cichlid 

Amphilophus alfari (Meek, 1907) (3); convict cichlid 

4 Corresponding author. 

190 


Archocentrus nigrofasciatus (Giinther, 1869) (3); blackbelt 

cichlid Cichlasoma maculicaiida (Regan, 1805) 

(12); jaguar cichlid Cichlasoma managuense (Giinther, 

1867) (13); butterfly cichlid Herotilapia multispinosa 

(Giinther, 1867) (8) (Cichlidae); molly Poecilia velifera 

(Regan, 1814) (5) (Poeciliidae); and long-whiskered catfish 

Rhamdia nicaraguensis (Giinther, 1864) (4) (Pimelodidae). 

Fish were collected by hook and line and 

throw nets from 7 localities of the Atlantic drainages of 

Nicaragua in the Autonomous Region of the South Atlantic 

(Region Autonoma del Atlantico Sur—RAAS): 

Torsuani River (11°47'06"N; 83°52'38"W); Mahogany 

River (12°03'22"N; 83°59'07"W); Cano Negro Stream 

(12°00'55"N; 84°01'10"W), in Bluefields City; Walpatara 

Bridge (12°00'14"N; 83°45'58"W); Loonku Creek 

(11°59'05"N; 83°46'48"W); Cano Marandn Stream 

(12°00'10"N; 83°46'39"W); and Puente Chino 

(12°00'30"N; 83°46'13"W). The number offish sampled 

in individual localities and map locations are given in 

Aguirre-Macedo et al. (2001). 

Fish were transported alive to the laboratory of the 

Bluefields Indian and Caribbean University (BICU), 

where they were examined by routine helminthological 

procedures outlined by Vidal-Martinez, Aguirre-Macedo 

et al. (2001). All helminths were studied in fresh 

preparations and counted in situ. Adult helminths were 

considered those with fully developed reproductive organs 

regardless of the presence of eggs. Eventually, 

some digeneans were fixed with a glycerin-ammonium 

picrate (GAP) mixture following the methodology out-

AGUIRRE-MACEDO ET AL.—ENDOHELMINTHS FROM NICARAGUAN FISHES 191 

Table 1. Some endoparasitic helminths collected from Nicaraguan freshwater fishes. 

Digcnea 

Helminth species 

Crassicutis cichlasomae Manter, 1936 

Magnivitellinum simplex Kloss, 1966 

Oligogonotylus manteri Watson, 1976 

IProsthenhystera obesa (Diesing, 1850) 

Saccocoelioides sogandaresi Lumsden, 

1964 

Saccocoelioides sp. 1 


Allocreadiidae gen. sp. 

Nematoda 

Procamallanus (Spirocamallanus) rebecae 

Andrade-Salas, Pineda-Lopez, and 

Osorio-Sarabia, 1994 

Procamallanus (Spirocamallanus) neocaballeroi 

(Caballero-Deloya, 1977) 

Rhabdochona kidderi kidderi Pearsc, 

1936 

Acanthocephala 

Neoechinorhynchus golvani Salgado-Maldonado, 

1978 

Hosts 

(no. infected/examined) 

Cichlasoma maculicauda (5/7) 

Astyanax fasciatus ( 1 /4) 

(1/2) 

C. maculicauda (2/7) 

Cichlasoma managuense (1/2) 

A. fasciatus (1/2) 

Poet-ilia velifera (1/1) 

C. maculicauda (1/3) 

(6/7) 

A. fasciatus (2-4) 

A. fasciatus ( 1 /4) 

(1/2) 

Intensity 

range 

1-8 

1 

6 

3-6 

1 

1 

1 

17 

4-42 

1-3 

1 

4 

Amphilophus alfari (1/3) 29 

C. maculicauda (21 \) 1—2 

Herotilapia rnultispinosa (3/8) 1—3 

A. fasciatus (1/9) 3 

C. maculicauda (1/11) 1 

C. maculicauda (1/11) 1 

A. alfari (1/1) 

C. managuense (1/2) 

C. managuense (2/4) 

H. rnultispinosa (1/4) 

H. rnultispinosa (1/3) 

lined by Ergens (1969). Measurements are given in 

micrometers. Drawings were made with the aid of a 

drawing tube. Reference specimens were deposited in 

the Coleccion Nacional de Helmintos (CNHE), Mexico 

City, Mexico, and the Laboratory of Parasitology, 

CINVESTAV-IPN (CHCM), Merida, Mexico. 

Results 

A total of 12 helminth species was found. 

Data on the hosts, localities, and infection range 

are provided in Table 1. All but 1 species 

(IProsthenhystera obesa) were located in the intestine; 

IP. obesa inhabited the gall blader. 

Among the species found, 3 trematodes were 

not identified to species level: 2 species of Saccocoelioides 

and a trematode of the subfamily 

Allocreadiinae (Allocreadiidae). Measurements 

of the first 2 species (Figs. 1-4), together with 

those of a congeneric species {Saccocoelioides 

sogandaresi) from Poecilia velifera are presented 

in Table 2. 

1 

1 

1-8 

3 

2 

Locality 

Cano Maranon 

Torsuani River 

Loonku Creek 


Puente Chino 

Loonku Creek 

Cano Maranon 


Cano Maranon 



Loonku Creek 


Loonku Creek 

Deposit 

accession nos. 

CNHE/CHCM 

4193/ 

4 1 95/ 

4196/ 

4194/ 

/383 

/384 

/380 

4142/ 

/393, 393-1 

Mahogany River 4143/ 

Torsuani River /396 


Loonku Creek 

Puente Chino 

Cano Negro 

Loonku Creek 

Puente Chino 

41911 

Discussion 

The number of species of adult endohelminths 

recorded was lower than that of larval stages, in 

particular metacercariae of digeneans, found in 

the same fish hosts (Aguirre-Macedo et al., 

2001). Adult trematodes represented the dominant 

group (8 species) in this study, whereas 

nematodes were fewer (only 3 species). Only 1 

acanthocephalan occurred in fishes examined. 

No adult cestodes were found, even in the pimelodid 

catfish Rhamdia nicaraguensis, but 

only 4 fish of this species were examined. Thus, 

it is probable that more fish and localities need 

to be sampled to find adult cestodes, especially 

considering the low prevalence (




Table 2. Measurements of species of Saccocoelioides from Nicaraguan freshwater fishes (n = number of 

specimens measured). 

Host 

Body shape 

Body length 

Maximum width 

Oral sucker 

Ventral sucker 

Sucker ratio 

Position of acctabulum 

Prepharynx 

Pharynx 

Oral sucker/pharynx ratio 

Extent of ceca 

Tcstis 

Hermaphroditic sac 

Ovary 

Extent of vitellarium 

Eggs 

Saccocoelioides 

sogandaresi 

(n = 1) 

Poecilia velifera* 

Widely oval 

995 

310 

100 X 122 

123 X 130 

1.14:1 

48% of body length 

38 

90 X 98 

1.18:1 

Anterior line of testis 

102 X 82 

150 X 76 

— 

— 

— 

Specimens fixed with GAP under pressure. 

Ponce de Leon, 1996) from fish of the genus 

Rhamdia in the Yucatan Peninsula (see Scholz 

et al., 1996). 

In South America, cestodes appear to be the 

dominant component of the fauna of endohelminths 

in freshwater fishes, in terms of the number 

of species and genera (Thatcher, 1991; Rego 

et al., 1999). These cestodes belong almost exclusively 

to the order Proteocephalidea, and they 

occur most frequently in siluriform fishes, including 

pimelodids (de Chambrier and Vaucher, 

1999; Rego, 2000). 

The endohelminth fauna of fishes from the 

Atlantic coastal drainages of Nicaragua closely 

resembles that of southeastern Mexico. Similar 

to the larval stages of endohelminths (Aguirre- 

Macedo et al., 2001), a majority of species 

found occur in congeneric fish hosts from the 

Yucatan Peninsula (Moravec et al., 1995; Scholz 

et al., 1995, 1996; Salgado-Maldonado et al., 

1997; Scholz and Vargas-Vazquez, 1998; Vidal- 

Martinez, Aguirre-Macedo et al., 2001). This 

similarity indicates close relationships between 


(n = 3) 

Astyanax fasciatus 

Elongate, with tapering ends 

1,070-1,210 

290-320 

90-112 X 100-120 

105-115 X 113-125 

0.83-1.01:1 

33-35% 

45-50 

70-78 X 63-69 

1.35-1.61:1 

About midline of testis 

274-290 X 140-173 

245-280 X 143-160 

98-105 X 80-104 

Far postacetabular 

73-75 X 46-50 


(« = 15) 

Cichlasoma maculicauda 

Oval to elongate 

470-680 

150-335 

67-105 X 75-125 

56-120 X 50-125 

0.63-1.30:1 

31-39% 

36-72 

50-82 X 47-75 

1.01-1.46:1 

About midline to % of testis 

72-175 X 58-145 

77-162 X 62-135 

37-87 X 35-87 

About midline of acetabulum 

67-81 X 36-47 

the helminth faunas of freshwater fishes in Central 

America and southeastern Mexico, in accordance 

with the analysis of Vidal-Martinez and 

Kennedy (2000) and the general biogeography 

of the neotropics (Briggs, 1984). Vidal-Martinez, 

Scholz and Aguirre-Macedo (2001) also 

found a marked resemblance between gill monogeneans 

of cichlids from Nicaragua and those 

from Yucatan. 

Three species of trematodes, Magnivitellinum 

simplex, IP. obesa, and S. sogandaresi, all nematodes, 

and the acanthocephalan Neoechinorhynchus 

golvani, previously found in North and 

South America (Travassos et al., 1969; Thatcher, 

1991; Perez-Ponce de Leon et al., 1996; Salgado-Maldonado 

et al., 1997; Moravec, 1998), are 

reported from Central America for the first time. 

With the exception of the trematodes Oligogonotylus 

manteri and Crassicutis cichlasomae reported 

from Lake Nicaragua by Watson (1976), 

all species also represent new geographical records 

from Nicaragua. This reflects the shortage 

Figures 1—4. 1, 2. Saccocoelioides sp. 1 from Cichlasoma maculicauda. 1. Total view from the ventral 

aspect. 2. Tegumental spines at pharyngeal level. 3, 4. Saccocoelioides sp. 2 from Astyanax fasciatus. 3. 

Total view ventral from the ventral aspect. 4. Tegumental spines at pharyngeal level. Scale bars in millimeters. 



of data on helminths of freshwater fishes in this 

country and in Central America in general. 

Three species of the genus Saccocoelioid.es 

Szidat, 1954, were found in this study, differing 

from each other in the size and shape of the 

body, their spination, relative size of the pharynx 

and hermaphroditic sac, extent of the vitellaria, 

egg size, etc. However, 2 of them (Figs. 

1-4) remain unidentified to species level because 

they differ from all hitherto described taxa 

(see Szidat, 1954; Lunaschi, 1984). They are not 

described as new species because of the unsatisfactorily 

resolved taxonomy of the genus, with 

numerous taxa having been inadequately described. 

To provide data for subsequent species 

identification, measurements of all 3 species are 

provided in Table 2. 

The allocreadiid trematode found in Astyanax 

fasciatus most resembles in its morphology the 

species Crepidostomum platense Szidat, 1954, 

and Crepidostomum stenopteri Mane-Garzon 

and Gascon, 1973, described from the intestine 

of several pimelodid catfishes from Argentina 

and from the characid fish "dentudo transparente" 

Charax (=Asiphonichthys) stenopterus 

(Cope, 1894) from Uruguay, respectively (see 

Szidat, 1954; Mane-Garzon and Gascon, 1973). 

However, there are marked differences among 

the present specimens and both species of Crepidostomum 

in the extent of the vitelline follicles, 

position of the ventral sucker, and in the 

shape and relative positions of the testes. It is, 

therefore, probable that the trematode from Nicaragua 

represents at least a new species. Nevertheless, 

it is not described formally in this paper 

because it differs, as do both species from 

South America, in its morphology from members 

of Holarctic species of the genus Crepidostomum 

Braun, 1900, and may even represent a 

different genus. 

The present report is the second on adults of 

helminth parasites from Nicaragua, following 

that of Watson (1976). It is evident that much 

more data on the fish parasites from larger samples 

of fish hosts must be obtained for better 

understanding of their species composition and 

relationships to those from other areas of the 

Neotropical region. 


The authors are indebted to Dr. Anindo 

Choudhury, National Wildlife Health Center, 

Madison, Wisconsin, U.S.A., for valuable advice 


and helpful suggestions. Thanks are also due to 

the students of the School of Marine Biology of 

BICU for help in collecting fish, and to the authorities 

of this university for enabling us to use 

its facilities. A short visit by M.L.A.M. and 

V.M.V.M. to the Czech Republic in August 1999 

was supported by the Institute Mexicano de 

Cooperacion Internacional de la Secretaria de 

Relaciones Exteriores, Mexico. 


Aguirre-Macedo, M. L., T. Scholz, D. Gonzalez-Solis, 

V. M. Vidal-Martinez, P. Posel, G. Arjona- 

Torres, E. Siu-Estrada, and S. Dumailo. 2001. 

Larval helminths parasitizing freshwater fishes 

from the Atlantic coast of Nicaragua. Comparative 

Parasitology 68:42-51. 

Briggs, J. C. 1984. Freshwater fishes and biogeography 

of Central America and the Antilles. Systematic 

Zoology 33:428-435. 

de Chambrier, A., and C. Vaucher. 1999. Proteocephalidae 

et Monticelliidae (Eucestoda: Proteocephalidea) 

parasites des poissons d'eau douce au 

Paraguay, avec descriptions d'un genre nouveau 

et de dix especes nouvelles. Revue Suisse de 

Zoologie 106:165-240. 

Ergens, R. 1969. The suitability of ammonium-picrate 

in preparing slides of lower Monogenoidea. Folia 

Parasitologica 16:320. 

Lunaschi, L. I. 1984. Helmintos parasitos de peces de 

agua dulce de la Argentina I. Tres nuevas especies 

del genero Saccocoelioides Szidat, 1954 (Trematoda 

Haploporidae). Neotropica 30(83):31-42. 

Mane-Garzon, F., and A. Gascon. 1973. Digenea de 

peces de agua dulce del Uruguay, I. Una nueva 

especie del genero Crepidostomum Braum, 1900 

del intestine de Asiphonichthys stenopterus. Revista 

de Biologia del Uruguay 1:11-14. 

Moravec, F. 1998. Nematodes of Freshwater Fishes 

of the Neotropical Region. Academia, Prague, 

Czech Republic. 464 pp. 

, C. Vivas-Rodriguez, T. Scholz, J. Vargas- 

Vazquez, E. Mendoza-Franco, and D. Gonzalez-SoIis. 

1995. Nematodes parasitic in fishes of 

cenotes (=sinkholes) of the Peninsula of Yucatan, 

Mexico. Part 1. Adults. Folia Parasitologica 42: 

115-129. 

Perez-Ponce de Leon, G., L. Garcia-Prieto, D. Osorio-Sarabia, 

and V. Leon-Regagnon. 1996. Listados 

Faunisticos de Mexico. VI. Helmintos Parasitos 

de Peces de Aguas Continentales de Mexico. 

Institute de Biologia, Universidad Nacional 

Autonoma de Mexico, Mexico, D.F., Mexico. 100 

pp. 

Rego, A. A. 2000. Cestode parasites of neotropical 

teleost freshwater fishes. Pages 135-154 in A. N. 

Garcfa-Aldrete, G. Salgado-Maldonado, and V. M. 

Vidal-Martmez, eds. Metazoan Parasites in the 

Neotropics: Ecological, Taxonomic and Evolutionary 

Perspectives. Commemorative Volume of 

the 70th Anniversary of the Institute de Biologia,


Universidad Nacional Autonoma de Mexico. 

Mexico, D.F., Mexico. 

J. C. Chubb, and G. C. Pavanelli. 1999. 

Cestodes in South American freshwater teleost 

fishes: keys to the genera and brief descriptions. 

Revista Brasileira de Zoologia 16:299-367. 

Salgado-Maldonado, G., R. Pineda-Lopez, V. M. 

Vidal-Martinez, and C. R. Kennedy. 1997. A 

checklist of metazoan parasites of cichlid fish 

from Mexico. Journal of the Helminthological Society 


Scholz, T., and J. Vargas-Vazquez. 1998. Trematodes 

parasitizing fishes of the Rio Hondo River 

and freshwater lakes of Quintana Roo, Mexico. 


65:91-95. 

, , F. Moravec, C. Vivas-Rodriguez, 

and E. F. Mendoza-Franco. 1995. Cenotes (sinkholes) 

of the Yucatan Peninsula of Yucatan, Mexico, 

as habitat of adult trematodes of fish. Folia 

Parasitologica 42:37-47. 

, , , , and . 1996. 

Cestoda and Acanthocephala of fishes from cenotes 

( = sinkholes) of Yucatan, Mexico. Folia Parasitologica 

43:141-152. 

Szidat, L. 1954. Trematodos nuevos de peces de agua 

dulce de la Republica Argentina y un intento para 

alcarar su caracter marino. Revista del Institute de 

Investigacion de las Ciencias Naturales y Museo 

Argentine de Ciencias Naturales "Bernardino Rivadavia" 

3(1): 1-85. 

Thatcher, V. E. 1991. Amazon fish parasites. Amazoniana 

11:263-572. 

Travassos, L., J. F. Teixeira de Freitas, and A. 

Kohn. 1969. Trematodeos do Brasil. Memorias do 

Institute Oswaldo Cruz 67:1-886. 

Vidal-Martinez, V. M., M. L. Aguirre-Macedo, T. 

Scholz, D. Gonzalez-Solis, and E. F. Mendoza- 

Franco. 2001. Atlas of the Helminth Parasites of 

Cichlid Fish of Mexico. Academia, Prague, Czech 

Republic. 165 pp. 

, and C. R. Kennedy. 2000. Zoogeographical 

determinants of the helminth fauna composition 

of neotropical cichlid fish. Pages 227-290 in A. 

N. Garcfa-Aldrete, G. Salgado-Maldonado, and V. 

M. Vidal-Martinez, eds. Metazoan Parasites in the 

Neotropics: Ecological, Taxonomic and Evolutionary 

Perspectives. Commemorative Volume of 

the 70th Anniversary of the Institute de Biologia, 

Universidad Nacional Autonoma de Mexico. 

Mexico, D.F., Mexico. 

T. Scholz, and M. L. Aguirre-Macedo. 

2001. Dactylogyridae of cichlid fishes from Nicaragua, 

Central America, with descriptions of 

Gussevia herotilapiac and three new species of 

Sciadicleithrum (Monogenea: Ancyrocephalinae). 


Watson, D. E. 1976. Digenea of fishes from Lake Nicaragua. 

Pages 251-260 in T. B. Thorson, ed. Investigations 

of the Ichthyofauna of Nicaraguan 

Lakes. School of Life Sciences, University of Nebraska, 

Lincoln, Nebraska, U.S.A. 

Relocation of the Onderstepoort Helminthological Collection 

We have been notified that the Onderstepoort Helminthological Collection has moved and been renamed, 

and is now under the curatorship of Professor J. Boomker, Department of Veterinary Tropical Diseases, 

and Dr. E. van den Berg, Plant Protection Unit, University of Pretoria, Private Bag X04, Onderstepoort, 

0110 South Africa. The collection is now known as the National Collection of Animal Helminths and is 

fully accessible. Prospective lenders, or those seeking further information, can notify Professor Boomker 

at the above address, phone: +27-12-529-8166, fax: +27-12-529-8312, or e-mail: jboomker@op.up.ac.za. 




Helminth Parasites of Freshwater Fishes of the Balsas River 

Drainage Basin of Southwestern Mexico 

GUILLERMO SALGADO-MALDONADO,1'4 GUILLERMINA CABANAS-CARRANZA,1 

JUAN M. CASPETA-MANDUJANO,2 EDUARDO SoTO-GALERA,3 ELIZABETH MAYEN-PENA,' 

DAVID BRAILOVSKY,' AND RAFAEL BAEZ-VALE' 

1 Institute de Biologia, Universidad Nacional Autonoma de Mexico, Apartado Postal 70-153, CP 04510, 

Mexico (e-mail: gsalgado@mail.ibiologia.unam.mx), 

2 Centre de Investigaciones Biologicas, Universidad Autonoma del Estado de Morelos, Av. Universidad 1001, 

CP 62210, Cuernavaca, Morelos, Mexico, and 

3 Laboratorio de Ictiologfa y Limnologia, Escuela Nacional de Ciencias Biologicas, Institute Politecnico 

Nacional, Carpio y Plan de Ayala, Santo Tomas, Mexico, D.F., Mexico 

ABSTRACT: This study presents the results of the first survey of the helminth parasites in fishes in the Balsas 

River drainage, southwestern Mexico. A total of 25 species of helminth parasites in 13 freshwater fish species 

(n = 1,045) was collected between December 1995 and September 1998. The most prevalent and widespread 

helminth species was the Asian tapeworm Bothriocephalus acheilognathi. Two features characterize the helminth 

fauna of the Balsas River basin fishes: (1) a predominance of nematode and trematode species coupled with a 

scarcity of monogeneans and acanthocephalans; and (2) all helminths found had previously been reported from 

other regions of Mexico; therefore the composition of the helminth fauna of the fishes of the Balsas River 

drainage is not very distinct from that of fishes from other previously studied freshwater basins in Mexico. 

KEY WORDS: Monogenea, Digenea, Cestoda, Nematoda, Acanthocephala, freshwater fish, Balsas River, southwestern 

Mexico, survey. 

The Balsas River basin is the largest river 

drainage in southwestern Mexico. The Balsas 

River has its source at about 3,660 m altitude in 

the Sierra Madre del Sur. It flows generally eastwest 

through the states of Tlaxcala, Puebla, 

Guerrero, and Michoacan and receives several 

major inflows from the states of Oaxaca, Mexico, 

Morelos, and Jalisco before emptying into 

the Pacific Ocean. This river has a fish fauna 

composed of 37 species of 26 genera in 10 families. 

In addition to native fishes, exotic species 

such as Asian cyprinids (carps) and African 

cichlids (tilapias) have been introduced into 

many areas. 

Little information exists about the occurrence 

of helminth parasites of fishes from the Balsas 

River (Osorio-Sarabia, 1982, 1984; Salgado- 

Maldonado et al., 1998; Caspeta-Mandujano and 

Moravec, 2000; Caspeta-Mandujano et al., 2000; 

Moravec, 2000; Moravec et al., 2000), and the 

present report is the first survey of the helminth 

parasites of fishes of this drainage system. The 

aim of this work is to report the survey results, 

and the distribution and intensity data for these 

helminth parasites. 


196 



A total of 1,045 fish was collected from 28 sites, 

mostly rivers, in the Balsas River basin between December 

1995 and September 1998 (Table 1, Fig. 1). 

Fish at each site were captured by electrofishing or 

by gill nets. Live fish were brought to the laboratory 

and examined within 48 hr after capture using standard 

procedures. The following fishes were examined (* indicates 

species endemic to the Balsas River basin): Cyprinidae—*Hybopis 

boucardi (Giinther, 1868) (Balsas 

shiner, n= 111); Characidae—Astyanax fasciatus (Cuvier, 

1819) (Mexican tetra, n = 166); Ictaluridae—*Ictalurus 

balsanus (Jordan and Snyder, 1899) (Balsas 

catfish, n = 1); Ictalurus punctatus (Rafinesque, 1818) 

(channel catfish, n = 2); Goodeidae—Goodca atripinnis 

Jordan, 1880 (blackfin goodea, n = 6); *Ilyodon 

whitei (Meek, 1904) (Balsas splitfin, n = 59); Poeciliidae—Heterandria 

bimaculatiis (Heckel, 1848) (spottail 

killifish, n = 88), Poecilia reticulata Peters, 1860 

(guppy, n — 20), Poecilia sphenops Valenciennes in 

Cuvier and Valenciennes, 1846 (Mexican molly, n = 

261), Poeciliopsis gracilis (Heckel, 1848) (porthole 

livebearer, n = 156), Poeciliopsis infans (Woolman, 

1894) (Lerma livebearer, n = 20); Cichlidae—*Cichlasoma 

istlanum (Jordan and Snyder, 1899) (redside 

cichlid, n = 32), Cichlasoma nigrofasciatum (Giinther, 

1867) (convict cichlid, n = 123). Fish sample sizes per 

site are given in Table 1. 

All helminths recovered from each fish were counted. 

Digeneans (adults and metacercariae), cestodes, 

and nematodes were fixed in hot 10% neutral formalin. 

Acanthocephalans were placed in distilled water and 

refrigerated overnight (6-12 hr) to evert the proboscis,

Table 1. Helminths found in freshwater fishes from the Balsas River basin, Mexico. 

^Locality (no 

total no. of helm 

Species Site Hosts 

Monogenea 

Gyrodactylus sp. Gills 

P. gracilis PL (2/18; 7; 0.4 ± 1.1 [3-4]) 

P. infans VJ (1/20; 1) 

A.fasciatus PL (3/13; 12; 0.9 ± 1.8 [3-5]), 

[1-2]), PT (5/10; 30; 3.0 ± 3 

Urocleidoides cf. costaricensis (Price Gills 

and Bussing, 1967) Kritsky and 

Leiby, 1972 

/. whitei CH (1/22; 3) 

P. sphenops AR (1/7; 1), CH (2/43; 2; 0.05 

P. gracilis AR (9/11; 17; 1.5 ± 1.1 [1-4]) 

A. fasciatus CU (1/11; 1) 

P. reticulata AC (3/20; 3; 0.2 ± 0.4 [1-1]) 

P. sphenops HA (2/15; 16; 1.1 ± 3.9 [1-15] 

H. boucardi JU (1/8; 1), IX (5/10; 18; 1.8 ± 

Trematoda 

Saccocoelioides sogandaresi Lums- Intestine 

den, 1961 

Magnivitellinum simplex Kloss, 1966 Intestine 

Diplostomum cf. compaction (Lutz, Eyes, brain, body cavity 

1928) 

Posthodiplostomum minimum Muscle, liver, eyes, 

(MacCallum, 1921) mesentery, body cavity 

G. atripinnis JU (6/6; 801; 134.0 ± 37.5 [83 

H. bimacidata HJ (1/25; 1) 

P. sphenops HA (5/15; 39; 2.6 ± 6.0 [1-22] 

1), AM (1/16; 1); OT (1/2; 1 

P. infans VJ (19/20; 307; 15.4 ± 14.3 [3 

C. istlanum AM (1/4; 2) 

C. nigrofasciatum CO (5/44; 5; 0.1 ± 0.3 [1-1]), 

AM (8/20; 25; 1.2 ± 2.3 [1-9]) 

H. boucardi CU (5/14; 10; 0.7 ± 1.3 [1-4]) 

A.fasciatus PL (1/13; 1), HJ (1/5; 1) 

P. sphenops CO (1/10; 1), HJ (10/40; 14; 0. 

P. gracilis PL (1/18; 2), HJ (2/3; 3; 1.0 ± 

C. istlanum AM (3/4; 157; 39.2 ± 53.3 [16 

C. nigrofasciatum CH (17/22; 125; 5.7 ± 5.7 [1-2 

16.6 ± 13.1 [1-55]), AM (18 

A. fasciatus AM (4/26; 5; 0.2 ± 0.5 [1-2]) 

Uvulifer sp. Skin, fins 

A. fasciatus AM (2/26; 5; 0.2 ± 0.7 [2-3]) 

/. whitei CH (18/22; 1926; 87.5 ± 97.4 

P. sphenops CH (6/43; 28; 0.6 ± 2.3 [1-13] 

P. gracilis CH (3/15; 4: 0.3 ± 0.6 [1-2]), 

C. nigrofasciatum CH (2/22; 2; 0.1 ± 0.3 [1-1]), 

Clinostomum complanatum (Rudol- Fins, opercula, body 

phi, 1814) cavity 

Centrocestus formosanus (Nishigori, Gills 

1924) 


Table 1. Continued. 

'•'Locality (no. 

total no. of helmi 

Species Site Hosts 

H. boucardi CU (15/19; 103; 5.4 ± 10.8 [1-4 

[1-6]), IX (1/10; 1), PT (7/7; 

(5/15; 8; 0.5 ± 1.0 [1-4]), AM 


H. bimacuiata CO ((1/13; 1), HJ (5/25; 7; 0.3 ± 

P. reticulata AC (1/20; 1) 

P. sphenops PT (1/5; 1), MI (1/15; 2), XO (3 

0.4 ± 0.9 [1-3]), PL (2/13; 2; 

5; 1) 

P. gracilis CO (2/19; 2; 0.1 ± 0.3 [1-1]), T 

C. istlamim TE (1/11; 2) 

C. nigrofasciatum HJ (3/21; 5; 0.2 ± 0.7 [1-3]) 

P. sphenops HA ((2/15; 2; 0.1 ± 0.4 [1-1]), T 

XO (3/16; 6; 0.4 ± 0.8 [2-2]) 

P. gracilis TE (9/16; 15; 0.9 ± 1.1 [1-3]) 


P. gracilis TE (1/16; 1) 

C. istlamim TE (3/11; 11; 1.0 ± 2.6 [1-9]) 

Intestine 

Cestoda 

Bothriocephalus archeilognathi Yamaguti, 

1934 

Body cavity, mesentery, 

liver 

Glossocercus auritus (Rudolphi, 

1819) Bona, 1994 

P. sphenops TE (2/6; 18; 3.0 ± 6.0 [3-15]) 

P. gracilis TE (4/16; 19; 1.2 ± 2.4 [2-8]) 

Parvitaenia cochlearii Coil, 1955 Liver 

Parvitaenia macropeos (Wedl, 1855) Liver 

Baer and Bona, 1960 

Valipora minuta (Coil, 1950) Baer Liver, gall bladder 

and Bona, 1960 

A. fasciatus PL (1/13; 5) 

P. sphenops CH (7/43; 42; 1.0 ± 2.7 [1-10]), 

± 4.5 [1-13]), HJ (5/40; 116; 

C. nigrofasciatum CH (1/22; 1) 

H. boucardi JU (5/8; 15; 1.9 ± 2.1 ([1-6]), C 

1.7 [2-4]), RP (6/7; 80; 11.4 ± 

0.3 ± 0.5 [1-1]), MI (7/10; 34 

C. istlamim CO (1/1; 4), AM (4/4; 143; 35.8 

C. nigrofasciatum AR (8/16; 35; 2.2 ± 3.6 [1-14]), 

1.0 [1-6]), HJ (17/21; 99; 4.7 

C. atripinnis JU (5/6; 33; 5.5 ± 4.9 [1-13]) 

Nematoda 

Capillaria cyprinodonticola Huffman Intestine, 

and Bullock, 1973 liver 

Rhabdochona canademis Moravec Intestine 

and Arai, 1971 

Rhabdochona kidderi Pearse, 1936 Intestine 

A. fasciatus CO (2/13; 2; 0.1 ± 0.4 [0-1]), P 

0.2 ± 0.4 [1-1]), PT (1/10; 1) 

Rhabdochona lichtenfelsi Sanchez- Intestine 

Alvarez et al., 1998 

Rhabdochona mexicana Caspeta- Intestine 

Mandujano et al., 2000 


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MEXICO ,-•* ! \TLAXCAl_A 

Figure 1. The Balsas River drainage basin of southwestern Mexico, showing the fish collection sites. 

Double circles indicate sites where no infected fish were collected. 

then fixed in hot 10% formalin. Digeneans, cestodes, 

and acanthocephalans were stained with Mayer's paracarmine 

or Ehrlich's hematoxylin, dehydrated through 

a graded alcohol series, cleared in methyl salicylate, 

and whole-mounted. Nematodes were cleared with 

glycerine for light microscopy and stored in 70% ethanol. 

Voucher specimens of all taxa have been deposited 

in the Coleccion Nacional de Helmintos, Institute 

de Biologfa, Universidad Nacional Autonoma de Mexico. 

Infection parameters utilized are those proposed 

by Margolis et al. (1982), that is, prevalence (% infected) 

and abundance of infection (number of parasites 

per examined fish), expressed as mean ± standard 

deviation, followed by the range of intensity. 

Results 

The parasites encountered, their hosts, collection 

locations, infection sites, and prevalence, 

abundance, and range of intensity of helminth 

species are summarized in Table 1. 

Only 18 of the 1,045 host fish examined harbored 

monogeneans. Eight Gyrodactylus sp. 

were collected from 2 P. gracilis and 1 P. infans. 

Fifteen A. fasciatus were found to harbor 

66 Urocleidoides cf. costaricensis. 

Only 36 (3.4%) of the necropsied fish (3 species) 

were parasite-free. These fish included both 

ictalurid species, and 33 /. whitei from CR. Fish 

at 8 of the 28 collection sites sampled were not 


infected at all: HU, PE, CI, PA, XA, TL, AH, 

and CR (Fig. 1). 

The most prevalent and widespread helminth 

parasite was the cestode Bothriocephalus acheilognathi 

that was recorded in 8 of the 13 Balsas 

River fish species, at infection intensities 

from 1 to 46. 

Discussion 

Data from this survey provide further evidence 

to support Moravec's (1998) contention 

that nematodes represent a significant component 

of helminth faunas in tropical freshwater 

fishes. They also corroborate the statement of 

Salgado-Maldonado and Kennedy (1997) that 

richness in digenean species is a characteristic 

of these helminth communities. In contrast, 

acanthocephalans were found to be very rare in 

the Balsas River survey, supporting the claim of 

Salgado-Maldonado et al. (1992) that adult 

acanthocephalans are generally very rare parasites 

in Mexican freshwater fishes. Adult cestodes 

are not common parasites in Mexican 

freshwater fishes; however, this survey found 4 

metacestode species. Previous surveys from 

most other geographical areas in Mexico (Pine-

da-Lopez et al., 1985; Leon, 1992; Jimenez- 

Garcia, 1994; Salgado-Maldonado et al., 1997) 

did not reveal a rich fauna of cestodes (but see 

Scholz et al., 1996). Monogeneans have only exceptionally 

been reported from freshwater fishes 

in Mexico (Lamothe-Argumedo, 1981), but a 

number of species have been found recently, in 

particular in southeastern Mexico (Kritsky et al., 

1994, 2000; Mendoza-Franco et al., 1997, 

1999). 

Most of the parasites recorded in this survey 

are shared with freshwater fishes inhabiting other 

Mexican drainage basins (see Pineda-Lopez 

et al., 1985; Jimenez-Garcia, 1994; Moravec, 

Vivas-Rodriguez, Scholz, Vargas-Vazquez, 

Mendoza-Franco, and Gonzalez-Soli's, 1995; 

Moravec, Vivas-Rodriguez, Scholz, Vargas-Vazquez, 

Mendoza-Franco, Schmitter-Soto, and 

Gonzalez-Soli's, 1995; Scholz et al., 1995, 1996; 

Salgado-Maldonado et al., 1997; Moravec, 

1998; Moravec et al., 2000; Scholz and Vargas- 

Vazquez, 1998; Scholz and Salgado-Maldonado, 

2000). Six adult species are of neotropical origin: 

Urocleidoid.es cf. costaricensis, M. simplex, 

R. kidderi, R. lichtenfelsi, R. mexicana, and N. 

golvani. Saccocoelioides sogandaresi, Rhabdochona 

canadensis, and C. cyprinodonticola have 

been recorded in various freshwater fishes in 

Canada and southern North America (Lumsden, 

1963; Moravec and Aral, 1971; Moravec, 1998). 

Twelve of 25 helminth species recorded during 

this survey were larval forms that utilized 

small freshwater fishes as intermediate hosts. All 

these allogenic species are widespread taxa, with 

wide distributions within Mexico and broad host 

specificity. Thus, they can be regarded as an 

ecological component of the fish parasite communities 

in the Balsas River basin. The metacercariae 

of C. complanatum, P. minimum, and 

Diplostomum cf. compactum, as well as the larvae 

of nematodes Eustrongylides sp., Contracaecum 

sp., and Acuariidae gen. sp. have been 

commonly recorded in cichlids, poeciliids, characiids, 

pimelodids, and other fish families from 

southern Mexico (Pineda-Lopez, 1985; Pineda- 

Lopez et al., 1985; Osorio-Sarabia et al., 1987; 

Jimenez-Garcia, 1994; Moravec, Vivas-Rodriguez, 

Scholz, Vargas-Vazquez, Mendoza-Franco, 

Schmitter-Soto, and Gonzalez-Solis, 1995; 

Scholz et al., 1995; Salgado-Maldonado et al., 

1997). They have also been reported in atherinids, 

goodeids, and other fish families from the 

Lerma Santiago River basin in the highland pla- 

SALGADO-MALDONADO ET AL.—HELMINTHS OF MEXICAN FISHES 201 

teau of central Mexico (Osorio-Sarabia et al., 

1986; Salgado-Maldonado and Osorio-Sarabia, 

1987; Leon, 1992; Peresbarbosa et al., 1994). 

All these helminth species are widely distributed 

in North America, and some are worldwide 

(Hoffman, 1967; Yamaguti, 1971; Gibson, 

1996). 

Some of the helminths found have been introduced 

to Mexico with exotic fish or other animals. 

The Asian fish tapeworm (B. acheilognathi) 

has been disseminated globally in association 

with Asian cyprinids (grass and common 

carp) introduced to several countries for use in 

aquaculture (Salgado-Maldonado et al., 1986). 

This tapeworm has broad host specificity and 

now occurs in more than 15 freshwater fish species 

in Mexico (Garcia and Osorio-Sarabia, 

1991). We found B. acheilognathi widely distributed 

within the Balsas River basin, parasitizing 

8 fish species, mainly poeciliids. 

Another example is the heterophyid trematode 

C. formosanus that was introduced into Mexico 

most probably with the imported thiarid snail 

Melanoides tuberculata (Miiller, 1774) serving 

as the first intermediate host. This trematode has 

rapidly spread to an extensive area, including 

central Mexico and both the Atlantic and Pacific 

coasts, apparently aided by the previous expansion 

of M. tuberculata within Mexico. The 

metacercariae of C. formosanus are encysted in 

the gills of a wide spectrum of native fishes including 

members of Atherinidae, Cichlidae, Cyprinidae, 

Eleotridae, Goodeidae, Ictaluridae, and 

Poeciliidae (see Scholz and Salgado-Maldonado, 

2000). The adults are parasites in piscivorous 

birds and mammals. An increasing number of 

recent records of C. formosanus in numerous 

new hosts and regions, including the Balsas River 

drainage, suggests that this helminth is continuing 

to expand its distribution (Scholz and 

Salgado-Maldonado, 2000). 

Too few studies have been undertaken to draw 

conclusions about the zoogeographic characteristics 

of the helminth communities in the freshwater 

fish species of the Balsas River basin. 

However, two general statements can be made 

about these faunas. The first is that nematode 

and trematode species predominate, with only a 

few monogeneans and acanthocephalans being 

present. Second, all helminths found had previously 

been reported from other regions of Mexico; 

therefore the taxonomic composition of the 

helminth fauna of the fishes of the Balsas River 



drainage is not very distinct from that seen in 

other previously studied freshwater basins in 

Mexico. 


This study was supported by project no. 

400355-5-27668N from the Consejo Nacional 

para la Ciencia y la Tecnologia (CONACyT), 

Mexico and by the Comision Nacional para el 

Conocimiento y Uso de la Biodiversidad (CON- 

ABIO), Mexico, project nos. H007 and L051. 

We are indebted to Drs. Frantisek Moravec and 

Tomas Scholz for identification of nematodes 

and cestodes. Thanks are also due Aitzane Delgado-Yoshino, 

Erick Alcantara, Alejandra Hernandez-Rodriguez, 

Nancy Minerva Lopez-Flores, 

Isabel Cristina Caneda-Guzman, Norman 

Mercado-Silva, and Felipe Villegas-Marquez for 

their assistance in the field and laboratory. 


Caspeta-Mandujano, J. M., and F. Moravec. 2000. 

Two new intestinal nematodes of Profundulus labialis 

(Pisces, Cyprinodontidae) from fresh waters 

in Mexico. Acta Parasitologica 45:332-339. 

, , M. A. Delgado-Yoshino, and G. 

Salgado-Maldonado. 2000. Seasonal variations 

in the occurrence and maturation of the nematode 

Rhabdochona kidderi in Cichlasoma nigrofasciatum 

of the Amacuzac River, Mexico. Helminthologia 

37:29-33. 

Garcia, P. L., and D. Osorio-Sarabia. 1991. Distribucion 

actual de Bothriocephalus acheilognathi 

en Mexico. Anales del Institute de Biologia, Universidad 

Nacional Autonoma de Mexico, Serie 

Zoologia 62:523-526. 

Gibson, D. I. 1996. Trematoda. In L. Margolis and Z. 

Kabata, eds. Guide to the Parasites of Fishes of 

Canada. Part IV. Canadian Special Publications of 

Fisheries and Aquatic Sciences 124. 373 pp. 

Hoffman, G. L. 1967. Parasites of North American 

Freshwater Fishes. University of California Press, 

Berkeley, California, U.S.A. 486 pp. 

Jimenez-Garcia, M. I. 1994. Fauna helmintologica de 

Cichlasoma fenestratum (Pisces: Cichlidae) del 

lago de Catemaco, Veracruz, Mexico. Anales del 

Institute de Biologia, Universidad Nacional Autonoma 

de Mexico, Serie Zoologia 64:75-78. 

Kritsky, D. C., E. F. Mendoza-Franco, and T. 

Scholz. 2000. Neotropical Monogenoidea. 36. 

Dactylogyrids from the Gills of Rhamdia guatemalensis 

(Siluriformes: Pimelodidae) from Cenotes 

of the Yucatan Peninsula, Mexico, with Proposal 

of Ameloblastella gen. n. and Aphanoblastella 

gen. n. (Dactylogyridae: Ancyrocephalinae). 

Comparative Parasitology 67:76-84. 

, V. M. Vidal-Martinez, and R. Rodriguez- 

Canul. 1994. Neotropical Monogenoidea 19. Dactylogyridae 

of cichlids (Perciformes) from the Yucatan 

Peninsula, with descriptions of three new 


species of Sciadicleithriim Kritsky, Thatcher and 

Boeger, 1989. Journal of the Helminthological Society 


Lamothe-Argumedo, R. 1981. Monogeneos parasitos 

de peces. VIII. Descripcion de una nueva especie 

del genero Octomacrum Mueller, 1934 (Monogenea: 

Discocotylidae). Anales del Instituto de Biologia, 

Universidad Nacional Autonoma de Mexico, 

Serie Zoologia 51:51—60. 

Leon, R. V. 1992. Fauna helmintologica de algunos 

vertebrados acuaticos de la cienega de Lerma, Estado 

de Mexico. Anales del Instituto de Biologia, 

Universidad Nacional Autonoma de Mexico, Serie 

Zoologia 63:151-153. 

Lumsden, R. D. 1963. Saccocoelioides sogandaresi 

sp. n., a new haploporid trematode from the sailfin 

molly Mollienisia latipinna Le Sueur in Texas. 


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. 

Mendoza-Franco, E. F., T. Scholz, and V. M. Vidal- 

Martinez. 1997. Sciadicleithriim meekii sp. n. 

(Monogenea: Ancyrocephalinae) from the gills of 

Cichlasoma meeki (Pisces: Cichlidae) from cenotes 

(=sinkholes) of the Yucatan Peninsula, Mexico. 

Folia Parasitologica 44:205-208. 

, , C. Vivas-Rodriguez, and J. Vargas- 

Vazquez. 1999. Monogeneans of freshwater fishes 

from cenotes (sinkholes) of the Yucatan Peninsula, 

Mexico. Folia Parasitologica 46:267-273. 


of the Neotropical Region. Academia, Prague, 

Czech Republic. 464 pp. 

—. 2000. Systematic status of Laurotravassoxyuris 

bravoae Osorio-Sarabia, 1984 (Nematoda: 

Pharyngodonidae) [=Atractis bravoae (Osorio- 

Sarabia, 1984) n. comb.: Cosmocercidae]. Systematic 


, and H. P. Arai. 1971. The North and Central 

American species of Rhabdochona Railliet, 1916 

(Nematoda: Rhabdochonidae) of fishes, including 

Rhabdochona canadensis sp. nov. Journal of the 

Fisheries Research Board of Canada 28:1645- 

1662. 

, G. Salgado-Maldonado, and J. M. Caspeta- 

Mandujano. 2000. Rhabdochona mexicana sp. n. 

(Nematoda: Rhabdochonidae) from the intestine 

of characid fishes in Mexico. Folia Parasitologica 

47:211-215. 

, C. Vivas-Rodriguez, T. Scholz, J. Vargas- 

Vazquez, E. Mendoza-Franco, and D. Gonzalez-Solis. 

1995. Nematodes parasitic in fishes of 

cenotes (=sinkholes) of the Peninsula of Yucatan, 

Mexico. Part 1. Adults. Folia Parasitologica 42: 

115-129. 

, , , , , J- J- 

Schmitter-Soto, and D. Gonzalez-Solis. 1995. 

Nematodes parasitic in fishes of cenotes ( = sinkholes) 

of the Peninsula of Yucatan, Mexico. Part 

2. Larvae. Folia Parasitologica 42:199-210. 

Osorio-Sarabia, D. 1982. Descripcion de una nueva

especie del genero Goezici Zeder, 1800 (Nematoda: 

Goeziidae) en peces de agua dulce de Mexico. 

Anales del Instituto de Biologfa, Universidad Nacional 

Autonoma de Mexico, Serie Zoologia 52: 

71-87. 

. 1984. Descripcion de una especie nueva del 

genero Laurotravassoxyuris Vigueras, 1938 

(Nematoda: Syphaciidae) en peces de agua dulce 

de Mexico. Anales del Instituto de Biologfa, Universidad 


Zoologia 54:23-33. 

, G. Perez, and G. Salgado-Maldoiiado. 

1986. Helmintos de peces del lago de Patzcuaro, 

Michoacan I: Helmintos de Chirostoma estor el 

"pescado bianco." Taxonomfa. Anales del Instituto 

de Biologfa, Universidad Nacional Autonoma 

de Mexico, Serie Zoologfa 57:61-92. 

-, R. Pineda-Lopez, and G. Salgado-Maldonado. 

1987. Fauna helmintologica de peces dulceacufcolas 

de Tabasco. Estudio preliminar. Universidad 

y Ciencia 4:5-31. 

Peresbarbosa, R. E., G. Perez, and L. Garcia-Prieto. 

1994. Helmintos parasites de tres especies de 

peces (Goodeidae) del lago de Patzcuaro, Michoacan. 

Anales del Instituto de Biologfa, Universidad 


Zoologfa 65:201-204. 

Pineda-Lopez, R. 1985. Infeccion por metacercarias 

(Platyhelminthes: Trematoda) en peces de agua 

dulce de Tabasco. Universidad y Ciencia 2:47-60. 

, V. M. Carballo-Cruz, M. G. Fucugauchi, 

and L. Garcia-Magana. 1985. Metazoarios parasitos 

de peces de importancia comercial en la region 

de Los Rfos, Tabasco, Mexico. Pages 197- 

270 in Usumacinta: Investigacion Cientffica en la 

Cuenca del Usumacinta. Gobierno del Estado de 

Tabasco, Mexico. 

Salgado-Maldonado, G., G. Cabanas-Carranza, 

and J. M. Caspeta-Mandujano. 1998. Crcptotrema 

agonostomi n. sp. (Trematoda: Allocreadiidae) 

from the intestine of freshwater fish of Mexico. 


, S. Guillen-Hernandez, and D. Osorio-Sarabia. 

1986. Presencia de Bothriocephalus acheilognathi 

Yamaguti, 1934 (Ccstoda: Bothriocephal- 


idae) en peces de Patzcuaro, Michoacan, Mexico. 


Autonoma de Mexico, Serie Zoologfa 57: 

213-218. 

, M. I. Jimenez-Garcia, and V. Leon-Regagnon. 

1992. Presence of Octospiniferoides chandleri 

Bullock, 1957 in Heterandria bimaculata 

from Catemaco Veracruz, and considerations 

about the acanthocephalans of freshwater fishes of 

Mexico. Memorias do Instituto Oswaldo Cruz 

87(supplement 1):239-240. 

, and C. R. Kennedy. 1997. Richness and similarity 

of helminth communities in the tropical 

cichlid fish Cichlasoma umphthalmus from the 

Yucatan Peninsula, Mexico. Parasitology 114: 

581-590. 

, and D. Osorio-Sarabia. 1987. Helmintos de 

algunos peces del lago de Patzcuaro. Ciencia y 

Desarrollo 13:41-57. 

, R. Pineda-Lopez, V. M. Vidal-Martinez, 

and C. R. Kennedy. 1997. A checklist of metazoan 

parasites of cichlid fish from Mexico. Journal 


64:195-207. 

Scholz, T., and G. Salgado-Maldonado. 2000. The 

introduction and dispersal of Centrocestus formosanus 

(Nishigori, 1924) (Digenea: Heterophyidae) 

in Mexico: a review. American Midland Naturalist 

143:185-200. 

, and J. Vargas-Vazquez. 1998. Trematodes 

from fishes of the Rio Hondo River and freshwater 

lakes of Quintana Roo, Mexico. Journal of the 

Helminthological Society of Washington 65:91- 

95. 

, , F. Moravec, C. Vivas-Rodriguez, 

and E. Mendoza-Franco. 1995. Metacercariae of 

trematodes of fishes from cenotes ( = sinkholes) of 

the Yucatan Peninsula, Mexico. Folia Parasitologica 

42:173-192. 

, , , , and . 1996. 

Cestoda and Acanthocephala of fishes from cenotes 

( = sinkholes) of Yucatan, Mexico. Folia Parasitologica 

43:141-152. 

Yamaguti, S. 1971. Synopsis of Digenetic Vertebrates 

of Vertebrates. Vol. I. Keigaku Publishing Company, 

Tokyo, Japan. 1,074 pp. 




A Checklist of Helminth Parasites of Freshwater Fishes from the 

Lerma-Santiago River Basin, Mexico 

GUILLERMO SALGADO-MALDONADO,1-4 GUILLERMINA CABANAS-CARRANZA,1 EOUARDO SOTO- 

GALERA,2 JUAN M. CASPETA-MANDUJANO,3 R. GRISELDA MORENO-NAVARRETE,1 

PETRA SANCHEZ-NAVA,' AND ROGELIO AGUILAR-AGUILAR' 

1 Institute de Biologia, Universidad Nacional Autonoma de Mexico, Apartado Postal 70-153, CP 04510, 

Mexico, D.F., Mexico (e-mail: gsalgado@mail.ibiologia.unam.mx), 

2 Laboratorio de Ictiologfa y Limnologfa, Escuela Nacional de Ciencias Biologicas, Institute Politecnico 

Nacional, Carpio y Plan de Ayala, Santo Totnas, CP 11340, Mexico D.F., Mexico, and 

3 Centre de Investigaciones Biologicas, Universidad Autonoma del Estado de Morelos, Avenida Universidad 

1001, CP 62210, Cuernavaca, Morelos, Mexico 

ABSTRACT: A checklist based on previously published records and original data is presented for the helminth 

parasites reported from 33 freshwater fish species from the Lerma-Santiago river basin, west-central Mexico. 

The checklist contains 43 helminth species, 6 (14%) of which are endemic to the basin. Fourteen of the 43 are 

allogenic species, mostly Nearctic in origin. Three species are anthropogenically introduced colonizers, of which 

the Asian fish tapeworm Bothriocephalus acheilognathi is the most widely distributed species in the basin. The 

checklist includes 75 new host records, and records of 12 localities where no previous surveys had been conducted. 

KEY WORDS: Digenea, Monogenea, Cestoda, Nematoda, Acanthocephala, freshwater fishes, Lerma-Santiago 

river basin, west-central Mexico, survey. 

At least 375 freshwater fish species, of which 

approximately 60% are endemic, occur in Mexico, 

and over 500 species occur if those living 

in estuaries and coastal lagoons are included 

(Miller, 1982; Espinosa-Perez, 1993). The Lerma-Santiago 

river basin in west-central Mexico 

has the highest percentage of endemism of any 

major river basin in Mexico, with 30 of its 42 

(72%) fish species found nowhere else (Espinosa-Perez, 

1993; Soto-Galera et al., 1998). 

This river basin (Fig. 1) drains much of westcentral 

Mexico and consists of 2 major rivers, the 

Lerma and the Santiago. The Lerma River basin 

is the most important hydrologic system of the 

Mexican Central Highland Plateau. It originates in 

the State of Mexico at an elevation of 3,000 m and 

flows for 700 km through the states of Queretaro, 

Guanajuato, Michoacan, and Jalisco before emptying 

into Chapala Lake at 1,500 m elevation. The 

Santiago River drains from Chapala Lake, flowing 

through the state of Jalisco to the Pacific Ocean. 

The fish fauna of the Lerma-Santiago river basin 

has long been studied by ichthyologists (Diaz- 

Pardo et al., 1993; Soto-Galera et al., 1998). No 

regional survey of the parasite fauna of these fish 

has been published, and the literature is scattered 


204 


in taxonomic papers (Flores-Barroeta, 1953; Lamothe-Argumedo, 

1970, 1981, 1988; Lamothe-Argumedo 

and Cruz-Reyes, 1972; Osorio-Sarabia et 

al., 1986; Salgado-Maldonado et al., 1986; Salgado-Maldonado 

and Osorio-Sarabia, 1987; Alarcon, 

1988; Alarcon and Castro-Aguirre, 1988; Garcfa- 

Prieto et al., 1988; Garcfa-Prieto and Osorio-Sarabia, 

1991; Leon-Regagnon, 1992; Peresbarbosa- 

Rojas et al., 1994; Perez-Ponce de Leon et al., 

1994; Espinosa-Huerta et al., 1996; Mendoza- 

Garfias et al., 1996; Astudillo-Ramos and Soto- 

Galera, 1997; Pineda-Lopez and Gonzalez, 1997; 

Sanchez-Alvarez et al., 1998; Guzman-Cornejo 

and Garcfa-Prieto, 1999; Caspeta-Mandujano et 

al., 1999; Moravec et al., 2000, 2001; Scholz and 

Salgado-Maldonado, 2000, 2001). This paper 

compiles the extant information on the helminth 

parasites of freshwater fishes in the Lerma-Santiago 

river basin and includes original data derived 

from our own research. The species referred to in 

theses and scientific meetings do not constitute formal 

publications and are consequently not considered 

herein. This checklist should facilitate future 

research on the ecology, zoogeography, and biodiversity 

of this important river basin. 


As part of an ongoing parasitological investigation into 

the helminth fauna of the freshwater fishes of Mexico, a

JALISCO 


Chap ' 

LAKE'CHAPALA 

Figure 1. The Lerma-Santiago River drainage basin of west-central Mexico, showing the fish collection 

sites. Locality codes as in Table 1. 

review of the literature dealing with freshwater fish helminth 

parasites in the entire Lerma-Santiago river basin 

was made. In addition, a total of 1,177 fish of 18 species 

(Table 1), from 11 localities in the Lerma-Santiago river 

basin (Table 2, Fig. 1) was examined for the presence of 

helminths from January to October 1997 and from January 

to March 1998. 

At each site, fish were captured using electrofishing or 

gill nets. The numbers of fish examined at each locality 

and collection data are given in the parasite-host list (Table 

3). After capture, the fish were taken live to the laboratory 

and examined within 48 hr using standard procedures. 

Briefly, all the external surfaces, viscera, and 

musculature of each fish host were examined under a 

stereomicroscope, and all the helminths encountered in 

each fish were counted. Digeneans (adults and larvae), 

cestodes, and nematodes were fixed in hot 4% neutral 

formalin. Acanthocephalans were placed in distilled water, 

refrigerated overnight (6—12 hr) to evert the proboscis, 

and then fixed in hot 10% formalin. Digeneans, cestodes, 

and acanthocephalans were stained with Mayer's paracarmine 

or Ehrlich's hematoxylin, dehydrated using a 

graded alcohol series, cleared in methyl salicylate, and 

whole mounted. Nematodes were cleared with glycerine 

for light microscopy and stored in 70% ethanol. Voucher 

specimens of all taxa have been deposited in the National 

Helminth Collection (Coleccion Nacional de Helmintos 

[CNHE]), Institute of Biology, National Autonomous 

University of Mexico (UNAM), Mexico City. Infection 

parameters utilized are those proposed by Margolis et al. 

(1982), that is, prevalence (% infected) and mean intensity 

of infection (number of parasites per examined fish). 

Voucher specimens of the following species, found in 

fish from the Lerma-Santiago river basin and deposited 

in the CNHE, were examined: Posthodiplostomum minimum 

(MacCallum, 1921) (nos. 001253, 001476, 

001748); Bothriocephalus acheilognathi Yamaguti, 1934 

(no. 000434); Proteocephalus pusillus Ward, 1910 (nos. 

000383-000386); Proteocephalus sp. (no. 000425); Ligula 

intestinalis (Linnaeus, 1758) [nos. 448(F) and 

449(F)], and Contracaeciim sp. [nos. 002508(F) and 

002253(F)]. 

Results 

A host-parasite checklist is presented herein 

as Table 3. In this study, 43 helminth species are 

reported from 33 species of freshwater fishes of 

the Lerma-Santiago river basin, west-central 

Mexico. Six (14%) of the 43 species are endemic 

to the basin: A. mexicanum, M. bravoae, O. 

mexicanum, R. lichtenfelsi, Spinitectus sp., and 

B. nayaritensis. Fourteen are allogenic species 

that mature in, and are transported by, birds: C. 


206 COMPARATIVE PARASITOLOOY, 68(2), JULY 2001 

Table 1. Fish species from the Lerma-Santiago river basin of west-central Mexico that were examined 

for helminths in 1997 and 1998. 

Fish species Common name 

Sample 

size (n) 

Cyprinidae 

*Algansea tincella (Valenciennes in Cuvier and Valenciennes, 1844) Spottail chub 17 

jCyprinux carpio Linnaeus, 1758 Common carp 45 

'•'"Notropis sallei (Giinther, 1868) Azteca chub 37 

*Yuriria alta (Jordan, 1880) Lerma chub 49 

Goodeidae 

''Girardinichthys multiradiatiis (Meek, 1904) Darkedged splitfin 503 

'•'•Goodea atripinnis Jordan, 1880 Blackfin goodea 143 

*Xcnotoca variants (Bean, 1887) Jeweled splitfin 56 

Poeciliidae 

Poecilia sphenops Valenciennes //; Cuvier and Valenciennes, 1846 Mexican molly 23 

*Poeciliopxis infans (Woolman, 1894) Lerma livebearer 16 

Poeciliopsis sp. 13 

Atherinidae 

*Atherinella crystallina (Jordan and Culver ;'/; Jordan, 1895) Blackfin silverside 48 

*Chirostoma humboldtianitm (Valenciennes in Cuvier and Valenciennes, 

1835) Shortfin silverside 46 

*Chirostoma jordani: Woolman, 1894 Mesa silverside 64 

*Chirostoma labarcae Meek, 1902 Sharpnose silverside 3 

*Chirostoma riojai Solorzano and Lopez, 1966 Toluca silverside 78 

Cichlidae 

Cichlasoma beani (Jordan, 1889) Sinaloan cichlid 32 

Centrarchidae 

t Lepomis macrochirus Rafinesque, 1819 Bluegill 2 

Gobiidac 

Awaous tajasica (Lichtenstein, 1822) River goby 2 

* Species endemic to the Lerma-Santiago river basin. 

t Species introduced to the Lerma-Santiago river basin. 

complanatum, Diplostomum sp., P. minimum, C. 

formosanus, L. intestinalis, C. cf. ralli, P. caballeroi, 

P. cf. urseus, P. cochlearii, V. campylane 

ristrota, V. mutabills, Eustrongylides sp., 

Contracaecum sp., and P. brevis. Three species 

are recent, anthropogenically introduced colonizers: 

C. formosanus, P. tornentosa, and B. acheilognathi, 

which is the most widely distributed 

species in the basin. Twelve of the 33 fish species 

examined have not previously been surveyed 

for parasites, and present data expand the 

spectrum of fish hosts, to the effect that the list 

provides 75 new records for hosts and locations. 

Discussion 

Only 8 fish species have been examined in 

sufficient numbers to enable evaluation of helminth 

community composition and structure: Algansea 

lacustris, Chirostoma estor, C. attenu- 


atutn, Goodea atripinnis, Alloophorus robustus, 

Allotoca diazi, Micropterus salmoides, and Cyprinus 

carpio. Patzcuaro Lake has been systematically 

sampled, while other localities have only 

been sampled occasionally and with few fish examined. 

From large areas of the basin no data 

on fish parasites exist at all. There is also limited 

information on the parasites of fish in rivers and 

other water bodies. Parasitological knowledge 

for the Lerma-Santiago river basin is fragmentary, 

as many studies did not record all the helminth 

species because they were prepared for 

taxonomic ends, such as the description of a single 

species. As a result, most of the research in 

the basin merely indicates where data are most 

needed. 

A notable aspect of the present data is a highly 

characteristic endemic helminth component in 

the Lerma-Santiago river basin. Of the 43 re-


Table 2. Codes and features of the localities sampled or reported in the literature from which hosts were 

collected. 

Code 

Bata 

Bizn 

Chap 

Chic 

Coin 

Cons 

Cuit 

Igna 

Lagu 

Lerm 

Patz 

Rami 

Rsan 

Sala 

Taza 

Tila 

Trin 

Viet 

Zira 

Locality name 

Presa El Batan 

Presa La Biznaga 

Lago de Chapala 

Lago de Chicnahuapan 

("Almoloya del Rio") 

Presa Cointzio 

Presa Constitucion de 1917 

Lago de Cuitzeo 

Presa Ignacio Allende 

La Lagunilla 

Cienega de Lerma 

Lago de Patzcuaro 

Presa Ignacio Ramirez 

Rio Santiago (Aguamilpa) 

Lago de Salazar 

Las Tazas 

Santiago Tilapa, Laguna de 

Guadalupe Victoria 

Trinidad Fabela 

Villa Victoria 

Lago de Zirahuen 

Habitat 

type 

AR* 

AR 

NL 

NL 

AR 

AR 

NL 

AR 

WL 

WL 

NL 

AR 

RI 

NL 

AR 

NL 

AR 

AR 

NL 

State (coordinates) 

Queretaro (20°13'13"N; 100°24'39"W) 

Guanajuato (21°25'30"N; 100°52'52.7"W) 

Jalisco (20°08'-20°22'N; 102°42'-103°25'W) 

=|: AR = Artificial reservoir; NL = natural lake; WL = wetland; RI = river. 

corded helminth species, 6 (14%) are endemic 

to the basin: the digeneans A. mexicanum and 

M. bravoae, from atherinids and the goodeid G. 

multiradiatus, respectively; the monogenean O. 

mexicanum, a parasite of the cyprinid A. lacustris', 

and the nematodes R. lichtenfelsi, from the 

goodeids A. robustus, A. diazi, and G. atripinnis, 

and a species of Spinitectus, previously referred 

to as S. carolini, from the atherinids C. attenuatum 

and C. estor. Additionally, the nematode 

species B. nayaritensis, a parasite of C. beani in 

the Santiago River, may be endemic to this basin, 

because there is no other record of this species 

in Mexico (Moravec, 1998), and cichlids 

are the best studied fish family from a parasitological 

point of view (Salgado-Maldonado et 

al., 1997; Vidal-Martfnez and Kennedy, 2000). 

It is thought that the present hydrological configuration 

of the Lerma-Santiago river basin was 

created during the Pliocene Age by orogenic activity 

that isolated it from the ocean (Barbour, 

1973; Echelle and Echelle, 1984). The fish fauna 

of the basin consists of the descendants of marine 

ancestors that invaded the freshwater bodies, 

as well as Nearctic components such as cyprinids. 

It is assumed that by at least 5 million yr 

ago the fish species in the basin had established 

themselves, evolving and diversifying from their 

Estado de Mexico (19°11'N; 99°30'W) 

Michoacan (19°36'46"N; 101°17'58"W) 

Queretaro (20°25'00"N; 100°05'00"W) 

Guanajuato-Michoacan (20°04'34"- 1 9°53'25"N; 101° 1 9'34"-l 00°50'20"W) 

Guanajuato (20°55'N; 100°50'W) 

Estado de Mexico (19°08'30"N; 99°30'12"W) 


Michoacan (19°41'-19°32'N; 101°27'-101°53'W) 


Nayarit (21°46'42"N; 104°55'36"W) 


Estado de Mexico (not located) 

Estado de Mexico (19°1 1'15"N; 99°23'56"W) 

Estado de Mexico (19°48'N; 99°46'W) 


Michoacan (19°21'14"-19°29'32"N; 101°30'33"-101°46'15"W) 

original marine ancestors. The parasite fauna 

must also have evolved and diversified during 

this period of isolation, the current assemblage 

of endemic helminth species being the product 

of these evolutionary processes. To the extent to 

which the fish species adapted to these environments 

and speciated within them, so did their 

helminth communities, with some being lost and 

others developing in the new hosts. In other 

words, both the fish of the Lerma-Santiago river 

basin, and their parasites developed in isolation. 

The fish parasite fauna of this basin is also 

enriched through colonization by allogenic species 

transported by birds. As a result, the fish 

helminth communities in the basin have an 

abundant (14 of the total 43 species) component 

of allogenic species that mature in, and are 

transported by, birds: C. complanatum, Diplostomum 

sp., P. minimum, C. formosanus, L. intestinalis, 

C. cf. ralli, P. caballeroi, P. cf. urseus, 

P. cochlearii, V. campylancristrota, V. mutabilis, 

Eustrongylid.es sp., Contracaecum sp., 

and P. brevis, most of which occur throughout 

the American continent or are cosmopolitan. 

Many factors may have favored this colonization. 

They include the small size of the fish in 

this basin, their gregarious habits, their shallow 

water habitat, their status in the food web, and 


Table 3. Parasite-host list of helminths collected from fish of the Lerma-Santiago river basin of west-c 

Number 

of 

hosts 

exam- Preva 

ined mean in 

Locality 

Host/infection site(s)* 

Parasite 

Adult Trematoda 

Family Allocreadiidae Stossich, 1903 

AUocreadium mexicanurn Osorio-Sarabia, Perez and Salgado-Maldonado, 

1986 

6% 

4% 

23% 

13% 

24% 

"Low 

40% 

216 

195 

48 

8 

209 

64 

5 

Patz 

Patz 

Rsan 

Tila 

Patz 

Lagu 

Viet 

C. estorll 

C. attenuatum/l 

A. crystallina/I 

C. riojaill 

M. salmoides/Pc, I 

G. midtiradiatusl\I 

Crepidostomum cooperi Hopkins, 1931 

Margotrema bravoae Lamothe-Argumedo, 1970 

9 

Chap 

/. dugesill 

Family Gorgoderidae (Looss, 1901) 

Phyllodistomum lacustris (Loewen, 1929) 

6% 

17 

Igna 

A. tincella/l 

Larval Trematoda 

Family Cryptogonimidae Ciurea, 1933 

Cryptogonimidae gen. sp. 

Family Proterodiplostomidae Dubois, 1936 

Proterodiplostomurn sp. 

6% 

67% 

45% 

20% 

15% 

33% 

3% 

20% 

61% 

30% 

3% 

28% 

36% 

17 

15 

94 

50 

75 

3 

31 

5 

18 

20 

29 

46 

14 

Igna 

Rami 

Chic 

Lagu 

Rami 

Sala 

Trin 

Viet 

Bizn 

Igna 

Trin 

Viet 

Rami 

A. tincella/Bc 

N. sallei/Bc 

G. multiradiatus/Bc 

/Be 

/Be 

/Be 

/Be 

/Be 

G. atripinnis/Bc 

/Be 

/Be 

C. humboIdtianum/L, 

C. riojai/Bc 

90% 

9 

9 

Family Clinostomidae Liihe, 1901 

Clinostomum complanatum (Rudolphi, 1814) 

30 

41 

31 

Cuit 

Patz 

Patz 

A. robustusll 

/L, M 

A. diazi/L, M 



Number 

of 

hosts 

exam- Preva 

ined mean i 

Locality 


Parasite 

13% 

0.6% 

5% 

27% 

30 

178 

22 

41 

Cuit 

Patz 

Igna 

Cuit 

G. atripinnisll 

/L 

/L 

X. variatusH 

Family Diplostomidae Poirier, 1886 

Diplostomum sp. 

10% 

79% 

9% 

11% 

7% 

14% 

30% 

5% 

216 

38 

22 

9 

30 

42 

30 

390 

6 

30 

41 

31 

9 

30 

178 

35 

41 

30 

195 

42 

216 

30 

30 

17 

15 

1 

8 

10 

118 

13 

Patz 

Bizn 

Rsan 

Igna 

Cuit 

Zira 

Cuit 

Patz 

Lerm 

Cuit 

Patz 

Patz 

Lerm 

Cuit 

Patz 

Patz 

Cuit 

Patz 

Patz 

Zira 

Patz 

Cuit 

Cuit 

Igna 

Rami 

Lagu 

Chic 

Igna 

Chic 

Rami 

C. estor/B 

C. jor'danifM. 

P. sphenops/Bc 

Y. alta/M 


C. attenuatum/1 

C. jordanill 

A. lacustris/M 

N. sallei/1 

A. robustusll 

/L, M, Mu, E 

A. diazi/L, M, Mu, E 

G. multiradiatusH 


/L, Mu 

/L, M, Mu, E 

X. variants!! 

C. attenuatum/1^, M, Mu 

/L, M, Mu, E, B 

/L, M, Mu 

C. estor/L, Mu, E, B 

C. jordani/1 

O. aureusll 

A. tincella/M 

N. sallei/M 

/M 

/M 

Y. altall, L, M 

G. multiradiatus/M 

/M 

Diplostomum (Tylodelphys) sp. 

Posthodiplostomum minimum (MacCallum, 1921) Dubois, 

1936 

93% 

87% 

62% 

80% 

100% 

98% 

81% 

95% 

67% 

7% 

82% 

7% 

100% 

22% 

80% 

22% 

8% 



Preval 

mean in 

Number 

of 

hosts 

examined 

Locality 


Parasite 

60% 

55% 

25% 

57% 

5% 

100% 

100% 

48% 

52% 

100% 

26% 

8% 

25 

22 

4 

35 

22 

9 

2 

46 

23 

2 

23 

13 

Bizn 

Igna 

Trin 

Igna 

Rsan 

Igna 

Rami 

Viet 

Igna 

Igna 

Rami 

Tila 

G. atripinnis/M 

/L, M 

/M 

X. variatusfL, M 

P. sphenops/M 

P. infans/L, M 

/M 

C. humboldtianumfL 

C. jordani/L, M 

C. labarcaefL, M 

C. riojaifL, M 

/L, M 

? 

31 

Patz 

A. diazill 

Family Plagiorchiidae Liihe, 1901 

Ochetosorna sp. 

Family Heterophyidae Odhner, 1914 

Centrocestus formosanus (Nishigori, 1924) 

18% 

50% 

27% 

100% 

20% 

62% 

38% 

50% 

17 

14 

11 

1 

5 

13 

48 

2 

Igna 

Igna 

Igna 

Rsan 

Igna 

Rsan 

Rsan 

Rsan 

A. tincella/G 

Y. alta/G 

G. atripinnis/G 

P. sphenops/G 

P. infans/G 

Poeciliopsis sp./G 

A. crystallina/G 

L. macrochirus/G 

L2% 

25 

Rsan 

C. beani/G 

Monogenea 

Family Dactylogyridae Bychowsky, 1933 

Sciadicleithrum sp. 

28% 

23% 

46 

22 

Chic 

Rsan 

G. multiradiatuslG 

P. sphenopslG 

Family Gyrodactylidae Cobbold, 1864 

Gyrodactylus elegans Nordmann, 1832 

Gyrodactyhis sp. 

62% 

390 

Patz 

A. lacustrislG 

Family Discocotylidae Price, 1936 

Octomacrum mexicanum Lamothe-Argumedo, 1981 



Number 

of 

hosts 

exam- Preva 

ined mean i 

Locality 


Parasite 

0.3% 

390 

Patz 

A. lacustns/l 

Adult Cestoda 

Order Caryophyllidea van Beneden in Carus, 1863 

Caryophyllidea gen. sp. 

9 

Family Bothriocephalidae Blanchard, 1849 

Bothriocephalus acheilognathi Yamaguti, 1934 

5% 

13% 

390 

6 

178 

5 

41 

31 

9 

41 

9 

36 

30 

195 

42 

216 

234 

9 

9 

25 

43 

209 

40 

17 

15 

17 

3 

43 

2 

63 

50 

75 

Chap 

Patz 

Lerm 

Patz 

Lerm 

Patz 

Patz 

Lerm 

Bata 

Chap 

Cons 

Patz 

Patz 

Zira 

Patz 

Coin 

Chap 

Patz 

Bata 

Cons 

Patz 

Cons 

Igna 

Rami 

Igna 

Rami 

Rami 

Trin 

Chic 

Lagu 

Rami 

A. mbescensll 

A. lacustrisll 

N. sallei/I 

C. carpioll 

l\ robustus/l 

12% 

8% 

13% 

7% 

24% 

2% 

2% 

24% 

40% 

l% 

8% 

6% 

13% 

24% 

33% 

2% 

100% 

3% 

26% 

3% 

A. diazifl 

G. multiradiatusll 


/I 

X. variatusll 


/I 

/I 

C. estor/l 

C. humboldtianumll 

C. ocotlanell 

C. grandocule/l 

Cliirostoma sp./I 

/I 

M. salmoidesll 

O. niloticiis/l 

A. tincella/l 

N. sallei/l 

Y. alta/l 

/I 

C. carpioll 

11 

G. miiltiracliatus/l 

l\ 



_| H * < -l 

O O O O O O 

=: £ £ 5: =: 5 

a! a


Number 

of 

hosts 

exam- Prev 

ined mean 

Locality 


Parasite 

8 

3% 

22% 

3 

2% 

2% 

33% 

10 

4% 

9% 

3% 

4% 

24 

38 

23 

38 

48 

46 

3 

20 

50 

75 

31 

25 

Igan 

Bizn 

Igna 

Bizn 

Rsan 

Viet 

Rami 

Rami 

Lagu 

Rami 

Trin 

Rsan 

X. variatus/M 

C. jordani/M, L 

C. jordani/L 

C. jordanifL 

A. crystallinafL 

C. humboldtianum/Gb 

C. jordani/Gb 

C. riojai/Gb 

G. miiltiradiatus/Gb 

Paradilepis caballeroi Rysavy and Macko, 1973 

Paradilepis cf. urceus (Wedl, 1855) 

Paradilepis sp. 

Parvitaenia cochlearii Coil, 1955 

Valipora campy lancristrota (Wedl, 1855) 

C. beani/L, Gb 

Valipora mutabilis Linton, 1927 

0.5% 

195 

Patz 


Order Cyclophyllidea 

Cyclophyllidea gen. sp. 

Adult Nematoda 

Family Capillariidae Neveau-Lemaire, 1936 

Pseudocapillaria tomentosa (Dujardin, 1843) 

5% 

10 

0.5% 

2% 

1 

7% 

8% 

5% 

33% 

20 

178 

195 

216 

110 

43 

75 

184 

3 

Patz 

Patz 

Patz 

Patz 

Patz 

Patz 

Patz 

Patz 

Rami 

A. robustiis/l 

G. atripinnis/I 

C. attenuatum/I 

C. estorfl 

/I 

C. carpio/l 

N. sallei/l 

(1986) but Moravec et al 

5% 

4% 

20 

25 

Igna 

Bizn 

Remarks: This species was originally described as Capillaria patzcuarensis by Osorio-Sarabia et al. 

from Europe with cyprinids. 

Capillariidae gen. sp. 

G. atripinnis/I 

/I 

14% 

7 

Rsan 

C. beani/I 

Family Cucullanidae Cobbold, 1864 

Dichelyne mexicanus Caspeta-Mandujano, Moravec and 

Salgado-Maldonado, 1999 



Number 

of 

hosts 

exam- Preva 

ined mean in 

* Locality 

Parasite Host/infection site(s) 

0.5% 

390 

Patz 

Family Philometridae Baylis and Daubney, 1926 

Philometridae gen. sp. A. lacustrisfBc 

7 

7 

7 

40% 

8% 

7 

44% 

360 

41 

31 

20 

178 

35 

25 

Cuit 

Patz 

Patz 

Cuit 

Patz 

Patz 

Rsan 

Family Rhabdochonidae Travassos, Artigas, and Pereira, 1928 

Rhabdochona lichtenfelsi Sanchez-Alvarez, Garcia, and A. robustusfl 

Perez, 1998 /I 

A. diazi/i 

G. atripinnis/l 

n 

n 

Beaninema nayaritense Caspeta-Mandujano, Moravec, and C. beani/l 

Salgado-Maldonado, 2000 

10% 

14% 

43% 

12% 

30 

195 

42 

216 

Patz 

Patz 

Zira 

Patz 

Family Cystidicolidae Skrjabin, 1846 

Spinitectus sp. C. attenuatum/l 

11 

l\ estor/I 

Remarks: The nematodes were originally reported as Spinitectus carolini Holl, 1928, but they in fact belong to a separate spec 

of the Czech Republic, personal communication). 

Larval Nematodes 

Family Dioctophymatidae Railliet, 1915 

? 

2% 

2% 

13% 

1% 

10% 

41 

178 

195 

30 

209 

10 

Patz 

Patz 

Patz 

Patz 

Patz 

Bizn 

A. robustus/M, Be 

G. atripinnis/Mu 

C. attenuatum/M 

n 

M. salmoidesfMu 

G. atripinnis/Mu 

Eustrongylides sp. 

Family Anisakidae Railliet and Henry, 1912 

Contracaeciim sp. 

0.3% 

6% 

13% 

1 1 % 

5% 

31% 

390 

17 

15 

9 

22 

35 

Patz 

Igna 

Rami 

Igna 

Igna 

Igna 

A. lacustris/l 

A. tincella/M 

N. sallei/M 

Y. alta /M 

G. atripinnis/M, Be 

X. variatus/M 


COMPARA1


Number 

of 

hosts 

exam- Prev 

ined mean 

Locality 


Parasite 

119 

3 

17 

12 

50% 

9 

38 

23 

25 

2 

Igna 

Bizn 

Igna 

Rsan 

Rsan 

P. infansIM 

C. joi'dani/M 

/M 

C. beani/L,, M 

A. tajasica/M 

5% 

0.8% 

2% 

Patz 

Patz 

Patz 

Patz 

Patz 

Patz 

Patz 

Patz 

Rami 

Rami 

Bizn 

Trin 

Igna 

A. robustiisfL 

A. laciistris/l 

C. carpioll 

A. ?-obnstns/M, I 

A. diazi/M, I 

G. atripinnis/M, I 

/I 

M. salmoidesfl 

N. salleifl 

G. miiltiradiatiis/I 

G. atripinnisfl 

fl 

X. variatus/l 

Family Gnathostomatidae Railliet, 1895 

Gnathostoma sp. 

Spiroxys sp. 

1 

l% 

33 

15 

6 

3 

5 

20 

390 

184 

41 

31 

35 

178 

209 

3 

13 

18 

29 

21 

20% 

25 

Rsan 

C. beanifl 

Acanthocephala Adult 

Family Neoechinorhynchidae Ward, 1953 

Neoechinorhynchus golvani Salgado-Maldonado, 1978 

Acanthocephala Larvae 

Family Polymorphidae Meyer, 1931 

Polymorphic brevis Van Cleave, 1916 

0.3% 

2% 

3% 

4% 

8% 

3% 

6% 

390 

184 

41 

31 

178 

195 

216 

209 

35 

Patz 

Patz 

Patz 

Patz 

Patz 

Patz 

Patz 

Patz 

Igna 

A. lacustris/M 

C. carpio/L., M 

A. robustus/M, Mu 

A. diazi/M, Mu 

G. atripinnis/L,, M 

C. attenuatum/L., M 

C. estor/L, M 

M. salmoides/L,, M 

X. variatus/L, 

* B = brain; Be = body cavity; E = eyes; G = gills; Gb = gall bladder; I = intestine; L = liver; M = mesentery; Mu = 



their situation along the annual migratory routes 

of Nearctic birds. Additionally, the low number 

of helminth species in the basin may have readily 

allowed invasion of these communities by 

allogenic species. 

Three helminth species on the list are recent, 

anthropogenically introduced colonizers. The 

first is the cestode B. acheilognathi, which is the 

most widely distributed species in the basin and 

is found in 22 host species. The second is the 

heterophyid trematode C. formosanus. The actual 

distribution of this helminth within the basin 

has not been evaluated, because the intermediate 

host, the thiarid snail Melanoides tuberculata 

Miiller, 1774, has established along riverbanks 

and in riverbeds, where few fish have been sampled. 

Both these species were introduced recently 

into Mexico; the cestode together with 

Asian carp (Salgado-Maldonado et al., 1986), 

and the trematode most probably with the intermediate 

snail host (Scholz and Salgado-Maldonado, 

2000). The third species is the capillariid 

nematode P. tomentosa, reported from atherinids 

and goodeids, as well as from cultured carp, C. 

carpio, from Mexico, where it was probably introduced 

along with its fish host from Europe 

(Moravec, 1998; Moravec et al., 2001). 

The proportions among the helminth groups 

that constitute the communities in the fish of the 

Lerma-Santiago river basin are also distinctive. 

The dominance in species numbers of nematodes 

and trematodes (principally metacercariae) 

is a pattern characteristic of the fish helminth 

communities of southeastern Mexico (Scholz et 

al., 1995; Salgado-Maldonado and Kennedy, 

1997; Scholz and Vargas-Vazquez, 1998) and 

the Balsas River basin in central Mexico (Salgado-Maldonado 

et al., 2001). However, data in 

the present study show that cestodes, both adults 

and metacestodes, are almost as important in the 

Lerma-Santiago river basin in terms of numbers 

as the nematodes and trematodes. Most cestodes 

found, such as V. campylancristrota, occur 

throughout the American continent or are cosmopolitan 

(see Scholz and Salgado-Maldonado, 

2001). The presence of 5 monogenean species 

in the basin is also notable, as it is a higher 

number than recorded in other drainages in central 

Mexico. However, the monogenean fauna of 

freshwater fishes in southeastern Mexico is even 

richer (Kritsky et al., 1994, 2000; Mendoza- 

Franco et al., 1997, 1999, 2000). 

It is still not possible to form conclusions 


about the zoogeographic characteristics of the 

fish helminth parasite communities in the Lerma-Santiago 

river basin, as very few studies 

have been done. However, the data that do exist 

suggest that the proportion of endemic parasites 

is high, and thus very distinctive, as compared 

for example to the lack of endemic species 

among the helminth parasites of fishes from the 

Balsas River drainage (Salgado-Maldonado et 

al., 2001). The helminth communities were 

probably initially poor, and have been invaded 

by allogenic, Nearctic species transported by 

birds that have enriched these multispecific assemblages. 

Research into fish helminth parasites in the 

Lerma-Santiago river basin has been restricted to 

descriptions of some species, and more detailed 

studies have been carried out only in Patzcuaro 

Lake. Obviously, more complete inventories of 

the fish parasites in this basin are urgently required. 

Almost 7% of the fish species that originally 

inhabited the basin are extinct, and an additional 

23% are classified as endangered or vulnerable 

because of population decline associated 

with continuous habitat degradation and introduction 

of competing and predatory species that are 

added to the natural predation pressures in these 

ecosystems (Soto-Galera et al., 1998). 


This study was supported by project no. 

27668N from the Consejo Nacional de Ciencia 

y Tecnologia (CONACyT), Mexico, and by project 

no. H007 from the Comision Nacional para 

el Estudio y Uso de la Biodiversidad (CONA- 

BIO), Mexico. We are indebted to Dr. Frantisek 

Moravec for confirmation of identification of 

nematodes and Dr. Tomas Scholz for identification 

of cestodes. We also thank Nancy Minerva 

Lopez-Flores, Isabel Jimenez-Garcia, Cris Caneda-Guzman, 

Rafael Baez-Vale, Norman Mercado-Silva, 

and Felipe Villegas-Marquez for 

their assistance in the field and laboratory. 


Alarcon, G. C. 1988. Diagnostico e identificacion de 

una parasitosis helmmtica en Carassius carassius 

en un centre piscicola. Revista Latinoamericana 

de Microbiologia 30:297. 

, and J. L. Castro-Aguirre. 1988. Tratamiento 

experimental con Mebendazol para bothriocefalosis 

en Carassius carassius. Revista Latinoamericana 

de Microbiologfa 30:298. 

Astudillo-Ramos, L., and E. Soto-Galera. 1997. Es-

tudio helmintologico de Chirostoma humboldtianum 

y Girardinichthys multiradiatus capturados en 

el Lerma. Zoologia Informa 35:53-59. 

Barbour, C. D. 1973. A biogeographical history of 

Chirostoma (Pisces: Atherinidae): a species flock 

from the Mexican Plateau. Copeia 1973:533-556. 

Caspeta-Mandujano, J. M., F. Moravec, and G. Salgado-Maldonado. 

1999. Observations on cucullanid 

nematodes from freshwater fishes in Mexico, 

including Dichelyne mexicanus sp. n. Folia Parasitologica 

46:289-295. 

, , and . 2001. Two new species 

of rhabdochonids (Nematoda: Rhabdochonidae) 

from freshwater fishes in Mexico, with a description 

of a new genus. Journal of Parasitology 87: 

139-143. 

Diaz-Pardo, E., M. A. Godinez-Rodriguez, E. Lopez-Lopez, 

and E. Soto-Galera. 1993. Ecologia 

de los peces de la cuenca del rio Lerma, Mexico. 

Anales de la Escuela Nacional de Ciencias Biologicas 

39:103-127. 

Echelle, A. A., and A. F. Echelle. 1984. Evolutionary 

genetics of a "species flock": atherinid fishes on 

the Mesa Central of Mexico. Pages 93-110 in A. 

A. Echelle and I. Kornfield, eds. Evolution of Fish 

Species Flocks. University of Maine at Orono 

Press, Orono, Maine, U.S.A. 

Espinosa-Huerta, E., L. Garcia-Prieto, and G. 

Perez. 1996. Helminth community structure of 

Chirostoma attemiatiim (Osteichthyes: Atherinidae) 

in two Mexican lakes. Southwestern Naturalist 

41:288-292. 

Espinosa-Perez, H. 1993. Riqueza y diversidad de peces. 

Ciencias, Mexico 7:77-84. 

Flores-Barroeta, L. 1953. Cestodos de vertebrados I. 

Ciencia 13:31-36. 

Garcia-Prieto, L., H. Mejia, and G. Perez. 1988. 

Hallazgo del plerocercoide de Ligula intestinalis 

(Cestoda) en algunos peces dulceacuicolas de 

Mexico. Anales del Institute de Biologia, Universidad 


Zoologia 58:887-888. 

, and D. Osorio-Sarabia. 1991. Distribucion 

actual de Bothriocephalus acheilognathi en Mexico. 

Anales del Institute de Biologia, Universidad 

Nacional Autonoma de Mexico, Serie Zoologia 

62:523-526. 

Guzman-Cornejo, M. C., and L. Garcia-Prieto. 

1999. Trematodiasis en algunos peces del lago de 

Cuitzeo, Michoacan, Mexico. Revista de Biologia 

Tropical 47:593=596. 

Kritsky, D. C., E. F. Mendoza-Franco, and T. 

Scholz. 2000. Neotropical Monogenoidea. 36. 

Dactylogyrids from the gills of Rhamdia guatemalcnsis 

(Siluriformes: Pimelodidae) from cenotes 

of the Yucatan Peninsula, Mexico, with proposal 

of Ameloblastella gen. n. and Aphanoblastella 

gen. n. (Dactylogyridae: Ancyrocephalinae). 

Comparative Parasitology 67:76-84. 

, V. M. Vidal-Martinez, and R. Rodriguez- 

Canul. 1994. Neotropical Monogenoidea 19. Dactylogyridae 

of cichlids (Perciformes) from the Yucatan 

Peninsula, with descriptions of three new 

species of Sciadicleithrum Kritsky, Thatcher, and 


Boeger, 1989. Journal of the Helminthological Society 


Lamothe-Argumedo, R. 1970. Trematodos de peces 

VI. Margotrema bravoae gen. nov. sp. nov. 

(Trematoda: Allocreadiidae) parasito de Lermichthys 

multiradiatus Meek. Anales del Institute de 

Biologia, Universidad Nacional Autonoma de 

Mexico, Serie Zoologia 41:87-92. 

. 1981. Monogeneos parasitos de peces VIII. 

Descripcion de una nueva especie del genero Octomacrum 

Miiller, 1934 (Monogenea: Discocotylidae). 

Anales del Institute de Biologia, Universidad 

Nacional Autonoma de Mexico, Serie Zoologia 

51:56-60. 

. 1988. Trematodos de peces VIII. Primer registro 

de Phyllodistomum lacustri (Loewen, 1924) 

parasito de Ictalurus dugesi en Mexico. Anales 

del Institute de Biologia, Universidad Nacional 

Autonoma de Mexico, Serie Zoologia 58:487- 

496. 

-, and A. Cruz-Reyes. 1972. Hallazgo de Ligula 

intestinalis (Goeze, 1782) Gmelin, 1790 en 

Lermichthys multiradiatus (Meek) (Pisces: Goodeidae). 

Revista de la Sociedad Mexicana de Historia 

Natural 33:99-100. 

Leon-Regagnon, V. 1992. Fauna helmintologica de 

algunos vertebrados acuaticos de la Cienega de 

Lerma, Mexico. Anales del Institute de Biologia, 

Universidad Nacional Autonoma de Mexico, Serie 

Zoologia 63:151-153. 

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. 

Mendoza-Franco, E. F., T. Scholz, and V. M. Vidal- 

Martinez. 1997. Sciadicleithrum meekii sp. n. 

(Monogenea: Ancyrocephalinae) from the gills of 

Cichlasoma meeki (Pisces: Cichlidae) from cenotes 

( = sinkholes) of the Yucatan Peninsula, Mexico. 

Folia Parasitologica 44:205-208. 

, , C. Vivas-Rodriguez, and J. Vargas- 

Vazquez. 1999. Monogeneans of freshwater fishes 

from cenotes (sinkholes) of the Yucatan Peninsula, 

Mexico. Folia Parasitologica 46:267-273. 

-, V. M. Vidal-Martinez, M. L. Aguirre-Macedo, 

R. Rodriguez-Canul, and T. Scholz. 2000. 

Species of Sciadicleithrum (Dactylogyridae: Ancyrocephalinae) 

of cichlid fishes from southeastern 

Mexico and Guatemala: new morphological 

data and host and geographical records. Comparative 


Mendoza-Garfias, B., L. Garcia-Prieto, and G. 

Perez. 1996. Helmintos de la "acumara" Algansea 

lacustris en el lago de Patzcuaro, Michoacan, 

Mexico. Anales del Instituto de Biologia, Universidad 


Zoologia 67:77=88. 

Miller, R. R. 1982. Pisces. Pages 486-581 in S. H. 

Hurlbert and A. Villalobos, eds. Aquatic Biota of 

Mexico, Central America and the West Indies. San 

Diego State University, San Diego, California, 

U.S.A. 




of the Neotropical Region. Academy of Sciences 

of the Czech Republic, Prague, Czech Republic. 

464 pp. 

, R. Aguilar-Aguilar, and G. Salgado-Maldonado. 

2001. Systematic status of Capillaria 

patzcuarensis Osorio-Sarabia, Perez-Ponce de 

Leon et Salgado-Maldonado, 1986 (Nematoda: 

Capillariidae) from freshwater fishes in Mexico. 

Acta Parasitologica 46:8-11. 

-, G. Salgado-Maldonado, and D. Osorio-Sarabia. 

2000. Records of the bird capillariid, Ornithocapillaria 

appendiculata (Freitas, 1933) 

comb, n., from freshwater fishes in Mexico, with 

remarks on Capillaria patzcuarensis Osorio-Sarabia, 

Perez-Ponce de Leon and Salgado-Maldonado, 

1986. Systematic Parasitology 45:53-59. 

Osorio-Sarabia, D., G. Perez, and G. Salgado-Maldonado. 

1986. Helmintos de peces del lago de 

Patzcuaro, Michoacan, I. Helmintos de Chirostoma 

estorel "pescado bianco." Taxonomfa. Anales 

del Instituto de Biologfa, Universidad Nacional 

Autonoma de Mexico, Serie Zoologfa 57:61-92. 

Peresbarbosa-Rojas, R. E., G. Perez, and L. Garcia. 

1994. Helmintos parasites de tres especies de peces 

(Goodeidae) del lago de Patzcuaro, Michoacan. 

Anales del Instituto de Biologia, Universidad 

Nacional Autonoma de Mexico, Serie Zoologfa 

65:201-204. 

Perez-Ponce de Leon, G., B. Mendoza G., and G. 

Pulido F. 1994. Helminths of the charal prieto 

Chirostoma atteniiatum (Osteichthes: Atherinidae), 

from Patzcuaro Lake, Michoacan, Mexico. 


61:139-141. 

Pineda-Lopez, R., and C. Gonzalez. 1997. Bothriocephalus 

acheilognathr. presencia e importancia 

de un invasor asiatico infectando peces de Queretaro. 

Zoologfa Informa 35:5-12. 

Salgado-Maldonado, G., G. Cabanas-Carranza, J. 

M. Caspeta-Mandujano, E. Soto-Galera, E. 

Mayen-Pena, D. Brailovsky, and R. Baez-Vale. 

2001. Helminth parasites of freshwater fishes of 

the Balsas River drainage, southwestern Mexico. 


, S. Guillen-Hernandez, and D. Osorio-Sarabia. 

1986. Presencia de Bothriocephalus acheilognathi 

Yamaguti, 1934 (Cestoda: Bothriocephalidae) 

en peces de Patzcuaro, Michoacan, Mexico. 


Autonoma de Mexico, Serie Zoologfa 57: 

213-218. 


, and C. R. Kennedy. 1997. Richness and similarity 

of helminth communities in the tropical 

cichlid fish Cichlasoma iirophthalmus from the 

Yucatan Peninsula, Mexico. Parasitology 1 14: 

581-590. 

, and D. Osorio-Sarabia. 1987. Helmintos de 

algunos peces del lago de Patzcuaro. Ciencia y 

Desarrollo 113:41-57. 

-, R. Pineda-Lopez, V. M. Vidal-Martinez, 

and C. R. Kennedy. 1997. A checklist of mctazoan 

parasites of cichlid fish from Mexico. Journal 


64:195-207. 

Sanchez-Alvarez, A., L. Garcia-Prieto, and G. 

Perez. 1998. A new species of Rhahdochona Railliet, 

1916 (Nematoda: Rhabdochonidae) from endemic 

goodeids (Cyprinodontiformes) from two 

Mexican lakes. Journal of Parasitology 84:840- 

845. 

Scholz, T., and G. Salgado-Maldonado. 2000. The 

introduction and dispersal of Centrocestus forinosanus 

(Nishigori, 1924) (Digenea: Heterophyidae) 

in Mexico: a review. American Midland Naturalist 

143:185-200. 

, and . 2001. Metacestodes of the family 

Dilepididae (Cestoda: Cyclophyllidea) parasitizing 

fishes in Mexico. Systematic Parasitology 

49:23-39. 

, and J. Vargas-Vazquez. 1998. Trematodes 

from fishes of the Rio Hondo River and freshwater 

lakes of Quintana Roo, Mexico. Journal of the 


95. 

-, F. Moravec, C. Vivas-Rodriguez, 

and E. Mendoza-Franco. 1995. Metacercariae of 

trematodes of fishes from cenotes ( = sinkholes) of 

the Yucatan Peninsula, Mexico. Folia Parasitologica 

42:173-192. 

Soto-Galera, E., E. Diaz-Pardo, E. Lopez-Lopez, 

and J. Lyons. 1998. Fish as indicators of environmental 

quality in the Rio Lerma Basin, Mexico. 

Aquatic Ecosystem Health and Management 

1:267-216. 

Vidal-Martinez, V. M., and C. R. Kennedy. 2000. 

Zoogeographical determinants of the helminth 

fauna composition of Neotropical cichlid fish. 

Pages 250-278 in G. Salgado-Maldonado, A. N. 

Garcfa-Aldrete, and V. M. Vidal-Martfnez, eds. 

Metazoan Parasites in the Neotropics: A Systematic 

and Ecological Perspective. Instituto de Biologfa, 

Universidad Nacional Autonoma de Mexico, 

Mexico City, Mexico.



Singhiatrema vietnamensis sp. n. (Digenea: Ommatobrephidae) and 

Szidatia taiwanensis (Fischthal and Kimtz, 1975) comb. n. (Digenea: 

Cyathocotylidae) from Colubrid Snakes in Vietnam 

STEPHEN S. CURRAN,' ROBIN M. OVERSTREET,U DANG TAT THE,2 AND NGUYEN THI LE2 

1 The University of Southern Mississippi, Gulf Coast Research Laboratory, P.O. Box 7000, Ocean Springs, 

Mississippi 39566, U.S.A. (e-mail: robin.overstreet@usm.edu and stephen.curran@usm.edu) and 

2 Institute of Ecology and Biological Resources, Nghia do, Cau Giay, Hanoi, Vietnam (ntminh@ncst.ac.vn) 

ABSTRACT: A new digenean is described and a second species is redescribed from the colubrid rear-fanged 

water snakes Enhydris chinensls (Gray) and Enhydris phunbea (Boie) captured from several regions in Vietnam 

during 1996-1998. Singhiatrema vietnamensis sp. n. (Ommatobrephidae) from the small intestine of both snakes 

is characterized by the extent of the ceca, the position of the vitellaria, the size of the eggs, and the host. Szidatia 

taiwanensis (Fischthal and Kuntz, 1975) comb. n. (Cyathocotylidae) is redescribed from the holotype and specimens 

from the gallbladder of both snakes. The species is transferred from the genus Mesostephanoides primarily 

because it does not have a large cirrus that is spined; it is characterized by the shape of the seminal vesicle, 

length of the ceca, body size relative to the forebody, number of testes in hindbody, egg size, size of tribocytic 

organ, and the infection site in the host. Concerning the classification of Singhiatrema Simha within Echinostomatiformes, 

we consider Singhiatrematinae Simha a junior synonym of Ommatobrephinae Poche. We discuss 

the classification of Gogatea Lutz and Szidatia Dubois and consider Gogatinae Mehra a junior synonym of 

Szidatiinae Dubois. The use of different fixation methods can produce artifacts characteristic at the generic level. 

KEY WORDS: Singhiatrema vietnamensis sp. n., Ommatobrephidae, Szidatia taiwanensis comb, n., Cyathocotylidae, 

fixation artifacts, Enhydris chinensis, Enhydris plumbea, Colubridae, snakes, Vietnam. 

Two unrelated digeneans, a new intestinal ommatobrephid 

and a gallbladder cyathocotylid, 

each infecting 2 species of rear-fanged colubrid 

water snakes (Enhydris chinensis (Gray, 1842) 

and Enhydris plumbea (Boie, 1827)) from Vietnam, 

are herein described. The life history of 

neither parasite is known. The new ommatobrephid 

is most similar to species of Singhiatrema 

Simha, 1954, which are usually found in the intestines 

of water snakes. Exceptions include 

Singhiatrema najai Chattopadhyaya, 1967, from 

the intestine of the Indian cobra Naja naja (Linnaeus, 

1758), and Singhiatrema lali Chakrabarti, 

1967, from the intestine of a freshwater emydid 

turtle (Hardella thurgii (Gray, 1831)) (Chattopadhyaya, 

1967). All prior known species are 

reported from the Indian subcontinent only. 

The gallbladder cyathocotylid that we found 

in Vietnam also occurs in one of the same hosts 

in Taiwan. It exhibits a relationship with Szidatia 

joyeuxi (Hughes, 1929), which infects the intestine 

of the colubrid water snake Matrix maura 

(Linnaeus, 1758) (as Tropidonotus viperinus 

(Sonnini and Latreille, 1802)) in an oasis in Tunis, 

Morocco, and has a cercaria that is shed 

from the freshwater snail Melanopsis sp. and de- 


219 

velops in the leg muscles of the frog Rana rudibunda 

Pallas, 1771 (as Rana esculenta rudibunda; 

see Langeron, 1924; Hughes, 1929; Dubois, 

1938). 


The second and third authors captured a total of 43 

specimens of E. chinensis and 51 specimens of E. 

plumbea in Ha Noi, Nam Ha, Thai Binh, Na Nam, 

Nam Dinh, Hai Duong, and Hai Phong provinces (Red 

River Delta) in Vietnam. They collected digeneans live 

from the intestines and gallbladders of both snakes 

from all the provinces. For the first collections, they 

relaxed the specimens in distilled water, then placed 

them in physiological saline, and finally fixed them in 

a cold 5% formalin solution. However, specimens collected 

later were used for the descriptions. These were 

placed directly into saline, then fixed with near boiling 

water without coverslip pressure, and finally pipetted 

into 5% formalin. Additional unmeasured specimens 

were fixed under pressure for examination of specific 

features. Whole mounts of worms were prepared by 

staining specimens with either carmine or Van 

Cleave's hematoxylin with additional Ehrlich's hematoxylin. 

These were dehydrated, cleared with clove oil, 

and mounted on slides with Canada balsam. Measurements 

are given for the holotype, followed in parentheses 

by the range of measurements of each feature 

derived from specimens heat-fixed without pressure. 

Measurements are given in micrometers unless otherwise 

stated. Specimens were deposited in the United 

States National Parasite Collection (USNPC), Belts- 



ville, Maryland, U.S.A., and the Harold W. Manter 

Laboratory (HWML) of the University of Nebraska 

State Museum, Lincoln, Nebraska, U.S.A. 

Description 

Results 

Ommatobrephidae Poche, 1926 

Singhiatrema vietnamensis sp. n. 

(Figs. 1 and 2) 

Based on 5 specimens: Ommatobrephinae 

Poche, 1926. Body pyriform, elongate, 2.56 mm 

(2.41-2.61 mm) long, 0.76 mm (0.76-0.91 mm) 

wide at maximum width near posterior end of 

body, lacking tegumental spines but possessing 

minute interrupted longitudinal striae. Oral sucker 

with subterminal mouth, 170 (170—207) long, 

189 (189-217) wide. Head crown having single 

row of 22—23 spines 31—56 long, arranged 11— 

12 per side; row interrupted dorsally and ventrally. 

Prepharynx short, less than % length of 

pharynx; pharynx oval, 95 (95-106) long, 78 

(78-106) wide. Esophagus 640 (547-640) long, 

89 (80—105) wide. Ceca extending beyond posterior 

extent of testes. Acetabulum 290 (285- 

329) long, 379 (363-407) wide, lying between 

anterior % and anterior J/2 of body. Ratio of 

widths of oral sucker to acetabulum 1:1.8-2.0. 

Testes 2, weakly (incompletely) lobed, lying 

opposite, located near terminal end of body, with 

anterior margins diverging from each other; left 

testis 292 (285-340) long, 217 (200=234) wide; 

right testis 279 (251-335) long, 234 (195-234) 

wide. Cirrus sac oval, 179 (179-227) long, 102 

(102—110) wide, medial, lying between cecal bifurcation 

and acetabulum, containing large bipartite 

seminal vesicle, short pars prostatica, and 

short muscular ejaculatory duct; pars prostatica 

a short duct surrounded by prostatic cells at anterior 

of sac; external seminal vesicle lacking. 

Ovary pretesticular, spherical, 73 (73-95) in 

diameter, slightly dextral in 4 of 5 specimens, 

slightly sinistral in 1 specimen. Oviduct communicating 

with Laurer's canal, then forming 

ootype; Laurer's canal straight, directed dorsally 

from region of ootype, opening on dorsal surface 

at level of ovary; ootype surrounded by 

Mehlis' gland; Mehlis' gland compact, usually 

larger than ovary, consisting of relatively small 

cells; ootype receiving relatively long common 

vitelline duct from vitelline reservoir, communicating 

with uterus; uterus intercecal, postacetabular, 

with proximal portion a uterine seminal 


receptacle and with distal portion a metraterm; 

metraterm thick walled, approximately length of 

cirrus sac, sinistral to cirrus sac, entering genital 

atrium anteriorly; genital atrium relatively small, 

ventral to anterior portion of cirrus sac in heatkilled 

specimens (directed anteriorly in coldkilled 

specimens when fixed under pressure with 

cirrus sac displaced anteriorly); genital pore 

opening medially or submedially near anterior 

region of cirrus sac in heat-killed specimens 

(near anterior acetabular margin in contracted 

specimens). Vitellarium consisting of 2 lateral 

bands of irregularly shaped follicles; bands lying 

ventral to ceca between posterior level of acetabulum 

and middle of testes, communicating to 

vitelline reservoir by left and right transverse 

main collecting channel; left channel about 195 

long, right channel about 223-280 long; vitelline 

reservoir approximately 45—56 long, 111-195 

wide. Eggs 95-117 long, 61-75 wide, with 

thickened knob at posterior end, with eyespots 

visible in miracidia of some specimens; eyespots 

2, lightly pigmented in developing miracidia, 

darkly pigmented and fused in developed miracidia. 

Excretory vesicle Y-shaped, with triangular 

posterior bladder; main stem slender, medial, 

concealed by overlapping lobes of testes, extending 

anteriorly from bladder and bifurcating 

at anterior margins of testes; arms reaching anteriorly 

to approximate level just posterior to 

pharynx; excretory pore subterminal, opening 

on dorsal surface. 

Taxonomic summary 

TYPE HOST: Enhydris chinensis (Gray, 

1842), rear-fanged water snake or Chinese water 

snake (Colubridae). Other host: Enhydris plumbea 

(Boie, 1827), rear-fanged water snake, rice 

paddy snake, or plumbeous water snake (Colubridae). 

TYPE LOCALITY: Ha Noi Province, Vietnam. 

Other localities: throughout Red River Delta, Vietnam, 

in Nam Ha, Thai Binh, Nam Ha, Nam 

Dinh, Hai Duong, and Hai Phong provinces. 

INFECTION SITE: Small intestine. 

PREVALENCE AND INTENSITY OF INFECTION: Ten 

of 43 specimens of E. chinensis (23%) each 

hosted 1-5 individual worms; 9 of 51 specimens 

of E. plumbea (18%) each hosted 1—3 worms. 

SPECIMENS DEPOSITED: Holotype USNPC No. 

90037; paratypes USNPC No. 90038, HWML 

Nos. 15388 (E. chinensis), 15389 (E. plumbea).

CURRAN ET AL.—TWO VIETNAMESE SNAKE DIGENEANS 221 

Figures 1-6. 1. Ventral view of holotype of Singhiatrema vietnamensis sp. n.; scale bar = 300 u.m. 2. 

Ventral view of specimen of 5. vietnamensis exposed to fresh water and slight pressure prior to and during 

cold fixation; scale bar = 300 (mm. 3. Ventral view of Szidatia taiwanensis; scale bar = 200 \nm. 4. Ventral 

view of S. taiwanensis showing detail of tribocytic organ; scale bar = 200 (Jim. 5. Lateral view of 5. 

taiwanensis; scale bar = 200 (Jim. 6. Ventral view of 5. taiwanensis exposed to fresh water prior to cold 

fixation; scale bar = 275 (xm. 



ETYMOLOGY: This species is named for its 

type locality, Vietnam. 

Remarks 

Singhiatrema vietnamensis is consistent with 

members of Ommatobrephidae because it has a 

preacetabular cirrus sac with an internal seminal 

vesicle; large embryonated eggs, with some containing 

an oculate miracidium; and opposite incompletely 

lobed testes located in the posterior 

region of the body, often with axes diverging 

anteriorly. The species belongs in Ommatobrephinae 

rather than Parorchiinae Lai, 1936, because 

the collar of spines is interrupted dorsally 

and ventrally as opposed to being arranged in a 

continuous row. In addition, the tegument is 

smooth rather than spinous, an external seminal 

vesicle is absent, spines do not occur on the intromittent 

organ, and the specimens are parasites 

in reptiles rather than birds. The 2 genera within 

Ommatobrephinae, Singhiatrema and Ommatobrephus 

Nicoll, 1914, are distinguished by the 

presence of a dorsally interrupted row of collar 

spines in the former and the absence of the collar 

spines in the latter. The presence of a dorsally 

and ventrally interrupted row of 22-23 collar 

spines in 5. vietnamensis enables us to assign 

the worms to the genus Singhiatrema. Singhiatrema 

was originally unveiled to science in an 

oral presentation and abstract at the Forty-first 

Session of the Indian Science Congress (Simha, 

1954). The genus was described in more detail 

by Simha (1958) and placed in Echinostomatidae 

Poche, 1926, by the author without a subfamily 

designation. Singhiatrema singhia Simha, 

1954, from a colubrid ratsnake (Ptyas mucosus 

(Linnaeus, 1758)) from Hyderabad in southern 

India, was designated the type species (Simha, 

1954). Two other species, Singhiatrema longifurca 

Simha, 1958, and Singhiatrema hyderabadensis 

Simha, 1958, parasitize another colubrid 

water snake, the checkered keelback Xenochrophis 

piscator (Schneider, 1799) (as Tropidonotus 

piscator}, from the same locality 

(Simha, 1958). Three other Indian species have 

since been added to the genus. Singhiatrema najai 

parasitizes the Indian cobra (N. najd) in Hyderabad, 

S. lali parasitizes the turtle H. thurgii 

in Lucknow, and Singhiatrema piscatora Dwivedi, 

1968, parasitizes the water snake X. piscator 

in Chhindwara (Chakrabarti, 1967; Chattopadhyaya, 

1967; Dwivedi, 1968). 

Singhiatrema vietnamensis differs from its 


congeners, with the exception of S. lali, by having 

ceca that extend to the posterior region of 

the body and vitelline follicles that lie ventral 

and lateral to the ceca in bands stretching from 

the posterior margin of the acetabulum to the 

midlevel of the testes. Singhiatrema vietnamensis 

differs from S. lali by having larger eggs 

(103-119 |xm long by 45-56 jxm wide vs. 65- 

87 |xm long by 34-39 |xm wide), ceca that reach 

beyond the posterior level of the testes rather 

than to their midlevel, longer collar spines (31- 

56 (Jim rather than 7-9 jxm), and a snake rather 

than a turtle definitive host. Yamaguti (1971) reported 

24 collar spines for S. lali; however, 

Chakrabarti (1967) did not report the number of 

collar spines in the description of S. lali, and the 

illustration of the species did not permit the collar 

spines to be counted. Specimens of S. lali 

were apparently never deposited in a lending 

museum. We report 22-23 collar spines in our 

specimens of 5. vietnamensis. Three of 5 of our 

specimens had 22 collar spines and the remaining 

2 had 23 collar spines. It is possible that 

some specimens could lose spines in life or handling, 

but biological variation probably exists. 

Initially, we obtained 4 specimens of S. vietnamensis 

that had been placed in fresh water 

prior to their fixation with slight pressure in unheated 

formalin (see Fig. 2). The overall shape 

of these specimens, as well as their measurements, 

varied dramatically from those specimens 

fixed with heat and used in the above description. 

Specimens subjected to fresh water and 

pressure had an oval rather than pyriform body 

shape, and their width was greater (1.0-1.6 mm 

vs. 0.75-0.91 mm). The pharynx was swollen 

(131-158 |xm long by 127-140 jxm wide vs. 95- 

106 (Jim long by 78-106 |xm wide), and the 

esophagus was contracted in length but swollen 

in width (285-415 (xm long by 77-203 |xm wide 

vs. 547-640 (xm long by 80-105 |xm wide). In 

addition, the ceca were contracted slightly, 

reaching to the posterior level of the testes rather 

than beyond the posterior level of the testes, and 

the eggs were swollen or collapsed and therefore 

larger in whole mounts (109—117 |xm long by 

81-95 urn wide vs. 103-119 |xm long by 45-56 

(xm wide). The long side of the oval cirrus sac 

was oriented laterally rather than vertically, with 

the pore on the sinistral end rather than at the 

anterior end, and the testes were larger (411-560 

|xm long by 258-339 |xm wide compared with 

250-340 |xm long by 195-234 |xm wide). If the

methods of fixation had not been known, these 

differences would be great enough to suspect 

different species. 

Cyathocotylidae Poche, 1926 

Szidatia taiwanensis 

(Fischthal and Kuntz, 1975) comb. n. 

(Figs. 3-6) 

Redescription 

Based on holotype and 9 specimens from Vietnam: 

Szidatiinae Dubois, 1938. Body bipartite; 

forebody pyriform, 0.85 mm (0.81-1.15 mm) 

long, not accounting for slight curl; hindbody 

conical; entire body 1.20 mm (1.03-1.57 mm) 

long, 0.57 mm (0.47-0.93 mm) wide at maximum 

width just posterior to midforebody. Ratio 

of forebody to total body length 1:1.39 (1:1.27- 

1.36). Tegument with spines over entire forebody, 

with few or no spines on hindbody, with 

numerous minute papillae near posterior end. 

Oral sucker with subterminal mouth, (90-119) 

long, 119 (99-144) wide. Prepharynx very 

short. Pharynx oval, (60-75) long, (60-80) 

wide. Esophagus 94 (67-149) long, 31 (23-40) 

wide. Ceca bifurcating at about anterior Vs of 

forebody, extending to level of posterior margin 

of ovary. Acetabulum 82 (62-95) long, 98 (85- 

1 12) wide, situated between anterior % and anterior 

l/2 of forebody, always smaller than oral 

sucker. Tribocytic organ oval, protruding slightly 

from ventral surface, 327 (283-447) long, -250 

(246-423) wide, with medial-longitudinal slit 

with basal layer of dark-staining cells. 

Testes 2, oblique; anterior testis spanning both 

forebody and hindbody, -80 (149-213) long, 

124 (114-169) wide; posterior testis in hindbody 

91 (144-199) long, 142 (129-229) wide. Cirrus 

sac in posterior end of body, club-shaped, 410 

(332-362) long, 65 (60-73) wide, extending anteriorly 

to middle of anterior testis, containing 

internal seminal vesicle, pars prostatica with 

small prostatic cells, and short unspined ejaculatory 

duct (not true cirrus); internal seminal 

vesicle 241 (206-339) long, 45 (57-110) wide 

at thickest portion; pars prostatica 1 1 1 (43-198) 

long, —30 (23-25) wide, sinuous in some specimens, 

surrounded externally by free gland cells; 

ejaculatory duct 45 (24-74) long, 20 (8-14) 

wide, thin walled, eversible, surrounded by 

gland cells. 

Ovary nearly spherical, 85 (65-129) long, 75 

(70—99) wide. Vitellarium comprised of 2 pairs 


of lateral fields of follicles underlying tribocytic 

organ, with the 2 on each side overlying each 

other, ventral to ovary and testes; fields not confluent 

anteriorly; right field totaling —12-14 

(16-20) follicles; left field totaling -14 (16-23) 

follicles; follicles spherical to ovate, ranging 37— 

43 (50-99) long, >50 (55-124) wide. Vitelline 

reservoir lying posterior to ovary and between 

testes, 145 (119-149) long, 80 (78-144) wide, 

(159-167) thick (thickness measured from 2 laterally 

mounted specimens). Uterus reaching anteriorly 

to near level of anterior extent of vitellarium; 

distal portion an indistinct metraterm; 

metraterm slightly longer than cirrus sac (coiled 

in holotype and some other specimens), demarcated 

by transverse muscular band, surrounded 

by lining of free gland cells, emptying into genital 

atrium separate from ejaculatory duct; genital 

atrium an open funnel in distal portion of 

body, surrounded by gland cells. Eggs 5 (4-12) 

in number, 150 (140-169) long, 77 (80-99) 

wide, with indistinct operculum. 

Excretory vesicle V-shaped; arms united both 

anteriorly and medially; anterior junction at level 

of and dorsal to midesophagus; medial junction 

at level of anterior extent of tribocytic organ, 

ventral to ceca, leading to small bladder 

associated with base of acetabulum; excretory 

pore subterminal, opening ventral to and separate 

from constricted genital atrium. 


HOSTS: Enhydris chinensis (Gray, 1842), 

rear-fanged water snake or Chinese water snake 

(Colubridae), and Enhydris plumbea (Boie, 

1827), rear-fanged water snake, rice paddy 

snake, or plumbeous water snake (Colubridae). 

LOCALITIES: Ha Noi, Nam Ha, Thai Binh, 

Ha Nam, Nam Dinh, Hai Duong, and Hai Phong 

provinces in Vietnam. 

INFECTION SITE: Gallbladder (holotype USNPC 

No. 73148 from Taiwan in E. chinensis from 

intestine). 

PREVALENCE AND INTENSITY OF INFECTION: Sixteen 

of 43 specimens from intestine of E. chinensis 

(37%) each hosted 1-9 individual worms; 

24 of 51 specimens of E. plumbea (47%) each 

hosted 1-9 worms. 

SPECIMENS DEPOSITED: Voucher specimens 

USNPC Nos. 90039 and 90040 (E. chinensis), 

HWML No. 15390 (2 slides, E. chinensis). 



Remarks 

Fischthal and Kuntz (1975) described Mesostephanoid.es 

taiwanensis from the small intestine 

of E. chinensis from Taipei Prefecture in 

Taiwan on the basis of a single specimen. We 

examined that specimen, USNPC No. 73148, 

and, even though it was not from the gallbladder, 

we considered our specimens from Vietnam 

conspecific with it. Data for the holotype fit 

those for our specimens. We, however, interpret 

differently the terminal genitalia described and 

illustrated by Fischthal and Kuntz (1975) in their 

Figure 15. Perhaps they misinterpreted the nonillustrated 

muscular looping of the metraterm as 

spines. In any event, the cirrus sac of the holotype 

contained a club-shaped seminal vesicle 

measuring about 260 u,m long, a straight pars 

prostatica 113 u,m long, and a short ejaculatory 

duct 45 fjim long. The metraterm coiled once 

prior to descending to the genital atrium. We observed 

spines in none of these features. Because 

of the lack of a spined cirrus and lack of anterior 

confluence of the bands of vitelline follicles, we 

consider that the species belongs to Szidatia Dubois, 

1938, as Szidatia taiwanensis (Fischthal and 

Kuntz, 1975) comb. n. 

Dubois (1951) differentiated the monotypic 

genus Mesostephanoides Dubois, 1951, from 

other cyathocotylid genera on the basis of Mesostephanoides 

burmanicus (Chatterji, 1940), 

having a forebody to total body length ratio that 

we estimate as 1:1.14-1.18, a spined cirrus 300 

(Jim long, and vitelline follicles confluent anteriorly. 

We consider the first 2 features of questionable 

generic significance, and an examination 

of M. burmanicus and related species may 

show that Mesostephanoides is a synonym of 

Gogatea Lutz, 1935, a genus with species having 

anteriorly confluent vitelline follicles. 

Szidatia taiwanensis resembles its 2 congeners 

in that the vitelline follicles are in separate 

lateral fields, as opposed to being in a single 

confluent field underlying the tribocytic organ as 

reported for species of Gogatea, the only other 

genus in Szidatiinae. Szidatia taiwanensis most 

closely resembles Szidatia nemethi Dollfus, 

1953, from the water snake TV. maura (as N. viperina) 

from Charrat, Morocco, but differs from 

this species by having a much smaller tribocytic 

organ (283—447 jjim long by 246-423 |xm wide 

vs. 885 (Jim long by 765 u,m wide), larger eggs 

(140-169 jxm long by 80-99 (Jim wide vs. 105 


(xm long by 55 (Jim wide), and a club-shaped 

rather than a coiled seminal vesicle. Szidatia taiwanensis 

differs from S. joyeuxi, the type and 

remaining species in the genus, by having a 

longer esophagus (67-149 |JLm compared with 

40 |xm), ceca that reach only to the level of the 

anterior testis rather than beyond the posterior 

testis, an acetabulum that lies between the middle 

and anterior % of the forebody rather than 

exactly in the middle, and oblique rather than 

tandem testes; the anterior testis is almost entirely 

within the forebody rather than in the 

hindbody, the seminal vesicle is club-shaped 

rather than coiled, and the eggs are larger (140- 

169 (xm long by 80-99 (Jim wide vs. 100 (Jim 

long by 70 (xm wide). The minute papillae on 

the tegument at the posterior end, too small to 

illustrate to scale, probably serve a sensory 

function during mating or egg deposition. The 

intestine, the infection site of the specimen reported 

by Fischthal and Kuntz (1975), may have 

resulted from postmortem migration from the 

gallbladder. A few specimens of other species 

normally found in the gallbladder occasionally 

occur in the intestine normally. We do not think 

the difference in sites based on 1 specimen is 

significant. 

As with the echinostomate specimens of S. 

vietnamensis, the initial specimens of S. taiwanensis 

had been placed in fresh water prior to 

fixation in unheated formalin (see Fig. 6). Unlike 

those of S. vietnamensis, these specimens 

were not fixed under any pressure; nevertheless, 

their measurements and features varied dramatically 

from those obtained from specimens fixed 

with heat and used for the above description. 

Hindbody length was generally shorter (146- 

506 (Jim vs. 326—762 (Jim), and both the anterior 

and posterior testes occurred entirely in the forebody 

of specimens exposed to water, whereas 

the posterior testis was always in the hindbody 

of heat-killed specimens. The cirrus sac was 

generally straighter in osmotically stressed specimens 

and more bent or curled in the region of 

the pars prostatica in heat-killed specimens. In 

addition, the tribocytic organ was greatly enlarged 

(492-650 (Jim long by 520-685 |xm wide 

vs. 283-447 fxm long by 245-423 ujn wide) in 

the stressed specimens. The most important difference 

due to fixation techniques pertained to 

the configuration of the vitellarium. In the 

stressed specimens, vitelline follicles appeared 

confluent anteriorly (horseshoe-shaped distribu-

tion) but were in independent lateral bands in 

heat-killed specimens. Because the presence or 

absence of an anterior vitelline confluence differentiates 

Gotatea from Szidatia, and the appearance 

of confluence can be influenced in 

cold-killed or freshwater-soaked specimens, the 

methods of fixation clearly have a bearing on 

taxonomic interpretations. 

Discussion 

Classification of Singhiatrema 

The position of Singhiatrema within Echinostomatiformes 

La Rue, 1957, continues to be debated 

by taxonomists. Ommatobrephinae Poche, 

1926, was created for Ommatobrephus and 

Singhiatrema, and Parorchiinae Lai, 1936, was 

included in Ommatobrephidae on the basis of 

the generic level character of the collar spines 

(see Simha and Chattopadhyaya, 1967). Singhiatrematinae 

Simha, 1962, was later created to 

house Singhiatrema, presumably because the author 

considered the presence of collar spines to 

be of taxonomic importance at the subfamiliar 

rather than generic level. These changes resulted 

in Ommatobrephidae containing Ommatobrephinae, 

Parorchiinae, and Singhiatrematinae, each 

containing a single genus. Members of Ommatobrephinae 

and Singhiatrematinae differ only in 

the presence or absence of a single row of collar 

spines. We consider the presence or absence of 

collar spines to represent a generic feature only 

and, therefore, consider Singhiatrematinae a junior 

subjective synonym of Ommatobrephinae. 

Yamaguti (1971) considered Parorchiinae a subfamily 

in Philophthalmidae Travassos, 1918. We 

agree with Yamaguti in returning Parorchiinae 

to Philophthalmidae because members of that 

subfamily do not possess characters consistent 

with Ommatobrephidae. Most notably, parorchiines 

have tegumental spines, a conspicuous prepharynx, 

an external seminal vesicle, a spined 

cirrus, opposite testes that do not diverge anteriorly, 

and life cycles that involve birds and estuarine 

molluscs. Although no knowledge of the 

larval stages of species in Singhiatrema or Ommatobrephus 

is available, all known adults of 

species in these genera infect freshwater or terrestrial 

reptilian hosts, indicating a freshwater 

rather than an estuarine life cycle. 

Classification of Szidatia 

Contrary opinions regarding the status of key 

generic features of Gogatea and Szidatia have 


formed the basis for debates over whether the 2 

cyathocotylid subfamilies Szidatiinae and Gogatinae 

Mehra, 1947, should be synonymized 

(see Dubois, 1938, 1951, 1953; Mehra, 1947; 

Sudarikov, 1962; Yamaguti, 1971). Both subfamilies 

contain only a single genus, but the 

above discussion of Mesostephanoides should 

be noted. 

Lutz (1935) erected Gogatea and created the 

combination Gogatea serpentum (Gogate, 1932) 

for Prohemistomum serpentum Gogate, 1932, a 

parasite in the intestine of the colubrid snake X. 

piscator (as Natrix piscator) from the Union of 

Myanmar (as Burma). Lutz (1935) included Gogatea 

in his new subfamily Prohemistominae 

Lutz, 1935. Szidat (1936) added the new combination 

Gogatea joyeuxi (Hughes, 1929) for the 

cyathocotylid previously considered Prohemistomum 

joyeuxi (Hughes, 1929) from the colubrid 

water snake Natrix natrix scutata Pallas, 

1771 (as Tropidontus natrix persa), by Joyeux 

and Baer (1934). That species developed in the 

snake when fed diplostomula consistent with diplostomula 

described by Hughes (1929) from 

Tunis, Morocco (Joyeux and Baer, 1934). Gogatea 

indicum Mehra, 1947, was subsequently 

described from X. piscator in India. Dubois 

(1938) observed that the vitellarium of G. joyeuxi 

consisted of 2 distinct rows of vitelline follicles, 

1 on either side of the tribocytic organ. 

The vitellarium of other species in Gogatea consists 

of a confluent arch of follicles, lying dorsal 

to the tribocytic organ. Dubois (1938) erected 

Szidatia Dubois, 1938, on the basis of the differing 

configuration of the vitellaria and created 

the new combination S. joyeuxi for G. joyeuxi. 

Dubois (1938) also created the subfamily Szidatinae, 

emended as Szidatiinae by Yamaguti 

(1958), to house both Szidatia and Gogatea. 

Mehra (1947) did not consider that the difference 

in vitelline configuration warranted generic 

distinction and rejected Szidatia and therefore 

Szidatiinae. Instead, he created Gogatinae to 

contain all species of Gogatea, with G. serpentum 

as the type species. Dollfus (1953), Sudarikov 

(1962), and Yamaguti (1971) all accepted 

the generic distinctions established by Dubois 

(1938) and considered Gogatinae a junior synonym 

for Szidatiinae, retaining G. serpentum 

and S. joyeuxi as the type species for their respective 

genera. We concur with these authors 

and think that the different configuration of the 



vitellarium can be used as an informative generic 

level taxonomic distinction. 

Fixation techniques 

A wide variety of techniques has been used 

by parasitologists to fix specimens. Most of 

those techniques yield satisfactory and consistent 

results; however, those that produce results 

inconsistent with other procedures should be 

avoided. We advocate collecting digeneans by 

initially placing them live in physiological saline 

solution (7.7—8.5 g NaCl per liter of distilled water). 

They should never be exposed to distilled 

or tap water while alive. Unless specimens are 

to be used for ultrastructural or other specific 

puiposes, they should be killed rapidly with 

heat. They can be killed by a flame under a slide 

with worms in a small amount of saline or by 

pouring a relatively large volume of hot or boiling 

saline or tap water over specimens immersed 

in little or no saline. The specimens then should 

be transferred, without touching them with forceps, 

into 5—10% buffered formalin solution 

soon after being killed. Killing with hot formalin 

solution or other fixatives is acceptable but produces 

harsh fumes. Killing with heat produces 

consistently fixed specimens, ideal for making 

comparative measurements. Fixation with pressure 

may be useful for examining certain organ 

systems such as the female reproductive complex 

or the terminal genitalia, but specimens 

fixed under pressure should be used with care 

for taxonomic purposes because the entire specimen 

or specific structures may be distorted. 

Comparison of heat-killed specimens with 

specimens bathed in fresh water and then coldkilled 

revealed differences. Specimens of S. vietnamensis 

that were osmotically stressed prior to 

fixation were a different shape, they were wider, 

the pharynx and esophagus were distorted, the 

ceca were slightly shorter, the testes were larger, 

and the cirrus sac was oriented differently. Any 

one of these conditions might be used to misidentify 

a species, incorrectly describe specimens, 

or provide misleading information. 

Stressed specimens of 5. taiwanensis exhibited 

some distortions at the specific level, but more 

importantly, the confluent vitelline follicles represent 

a generic distinction for Gogatea. Lack of 

knowledge about the methods used on specimens 

for descriptions of some species of Gogatea 

and Szidatia shows the need to reevaluate 

those species with heat-killed specimens. Until 


such material is available, we prefer to treat the 

genera separately. 


We thank Cathy Schloss and Kristine Wilkie 

of the Gulf Coast Research Laboratory for helping 

with some of the literature. We also thank 

Eric Hoberg and Pat Pilitt from the United States 

National Parasite Collection, Beltsville, Maryland, 

for the loan of a specimen and Dr. Richard 

Heckmann of Brigham Young University for his 

interest in the project. Partial support for this 

study was provided by International Paper. 


Chakrabarti, K. K. 1967. On a new echinostome 

(Singhiatrema lali n. sp.) from the intestine of a 

turtle. Science and Culture 33:407-408. 

Chattopadhyaya, D. R. 1967. On a new trematode of 

the genus Singhiatrema Simha, 1954, from the 

cloaca of the Indian cobra in Hyderabad. Indian 


Dollfus, R. P. 1953. Miscellanea helminthologica 

Maroccana VII. Les Szidatia de Matrix viperina 

(Latreille, 1802) [Trematoda Digenea]. Archives 

de 1'Institut Pasteur du Maroc 4:505-512. 

Dubois, G. 1938. Monographic des Strigeida (Trematoda). 

Memoires de la Societe Neuchateloise des 

Sciences Naturelles 6:1—535. 

. 1951. Nouvelle cle de determination des groupes 

systematiques et des genres de Strigeida 

Poche (Trematoda). Revue Suisse de Zoologie 

58(39):639-691. 

-. 1953. Liste Systcmatique des Strigeida (Trematoda) 

de 1'Inde. Pages 77-88 in J. Dayal and K. 

Singh, eds. Thapar, G. S., Commemoration Volume. 

University of Lucknow, Lucknow, India. 

Dwivedi, M. P. 1968. On a new species of the genus 

Singhiatrema Simha, 1954 (Trematoda: Echinostomidae). 

Indian Journal of Helminthology 19: 

141-144. 

Fischthal, J. H., and R. E. Kuntz. 1975. Some trernatodes 

of amphibians and reptiles from Taiwan. 

Proceedings of the Helminthological Society of 

Washington 42:1-13. 

Hughes, R. C. 1929. Studies on the trematode family 

Strigeidae (Holostomidae) No. XIV. Two new species 

of diplostomula. Occasional Papers of the 

Museum of Zoology, University of Michigan 202: 

1-29 + 1 pi. 

Joyeux, C., and J. G. Baer. 1934. Sur un trematode 

de couleuvre. Revue Suisse de Zoologie 41:203- 

215. 

Langeron, M. 1924. Recherches sur les cercaires des 

piscines de Gafsa et enquete sur la bilharziose 

Tunisienne. Archives de FInstitut Pasteur de Tunis 

13:19-67. 

Lutz, A. 1935. Observances e consideracoes sobre 

Cyathocotylineas e Prohemistomineas. Memorias 

do Institute Oswaldo Cruz 30:157-182. 

Mehra, H. R. 1947. Studies on the family Cyathoco-

tylidae Poche: Part 2. A contribution to our 

knowledge of the subfamily Prohemistominae 

Lutz, 1935, with a discussion on the classification 

of the family. Proceedings of the National Academy 

of Sciences, India 17:1-52. 

Simha, S. S. 1954. On a new trematode genus Singhiatrema 

singhia n. g., n. s. from rat snake Ptyas 

(Zamenis) miicosus from Hyderabad-Deccan. Proceedings 

of the Indian Science Congress (1954) 

Part IV 41:32. 

. 1958. Studies on the trematode parasite of 

reptiles found in Hyderabad state. Zeitschrift fur 

Parasitenkunde 18:161-218. 

, and D. R. Chattopadhyaya. 1967. A discussion 

on the systematic position of the subfamily 

Singhiatreminae Simha, 1962. Indian Journal of 

Helminthology 18(seminar supplement):54-58. 


Symposium Announcement 

Sudarikov, V. E. 1962. Order Strigeidida (La Rue, 

1926) Sudarikov, 1959. Trematodes of Animals 

and Man. Principles of Trematodology, Academy 

of Sciences of the U.S.S.R. Helminthological Laboratory, 

Moscow 19:267-469. (In Russian.) 

Szidat, L. 1936. Parasiten aus Seeschwalben 1. Uber 

neue Cyathocotyliden aus dem Darm von Sterna 

hirundo L. und Sterna paradisea. Zeitschrift fur 

Parasitenkunde 8:285-317. 

Yamaguti, S. 1958. Systema Helminthum. Volume I. 

The Digenetic Trematodes of Vertebrates, Part I. 

Interscience Publishers, New York, New York. 

979 pp. 

. 1971. Synopsis of Digenetic Trematodes of 

Vertebrates. 2 volumes. Keigaku Publishing Company, 

Tokyo, Japan. 1074 pp. + 349 pis. 

PARASITOLOGY IN SCIENCE AND SOCIETY 

Sponsored by The Helminthological Society of Washington 

Saturday, October 27, 2001, 1:00-5:00 pm 

S. Dillon Ripley Center of the South Quadrangle, Room 3111 

National Museum of Natural History of the Smithsonian Institution 

Washington, D.C. 

The 676th meeting of the Helminthological Society of Washington, to be held on October 27, 2001 at 

the Smithsonian Institution, will be a special event. The Helminthological Society of Washington, in 

cooperation with The American Society of Parasitologists, will conduct a symposium entitled Parasitology 

in Science and Society. The purpose of the symposium is to assess the state of parasitology as a discipline, 

clearly define the role of parasitology in science, and explore ways that various parasitological and related 

societies and governmental agencies can work together to strengthen our discipline. Dr. Richard O'Grady, 

Executive Director of the American Institute of Biological Sciences, will speak on the power and importance 

of collaborative efforts in science. Dr. Eric P. Hoberg of the United States Department of Agriculture's 

Beltsville Agricultural Research Center, will discuss the seminal importance of phylogenetic studies 

and taxonomic inventories and collections to contemporary parasitology. Additionally, leaders of various 

parasitological and related societies and governmental agencies are being asked to clearly articulate the 

missions and visions of their organizations as they relate to parasitology. Because each society and agency 

is a unique entity, we all have special strengths to offer to our broader discipline. Further, participants are 

being asked to identify contemporary societal issues which the discipline of parasitology can address, and 

indicate specific ways that their organization is, and can aid in, addressing these issues. We invite you to 

make every effort to attend this important meeting of the Helminthological Society of Washington. 

INVITED SOCIETIES, AGENCIES, AND ORGANIZATIONS 

American Association of Veterinary Parasitologists 

American Heartworm Society 

American Institute of Biological Sciences 

American Society of Parasitologists 

American Society of Tropical Medicine and Hygiene 

Canadian Zoological Society 

Centers for Disease Control and Prevention 

Entomological Society of America 

Helminthological Society of Washington 

National Institutes of Health 

National Science Foundation 

Society of Nematologists 

Society of Protozoologists 

United States Department of Agriculture 

United States Department of the Interior 

Wildlife Disease Association 




Rhabdias ambystomae sp. n. (Nematoda: Rhabdiasidae) from the 

North American Spotted Salamander Ambystoma inaculatum 

(Amphibia: Ambystomatidae) 

YURIY KUZMIN,' VASYL V. TKACH,1-2 AND SCOTT D. SNYDER3'4 

1 Department of Parasitology, Institute of Zoology, Ukrainian National Academy of Sciences, 15 Bogdan 

Khmelnitsky Street, Kiev-30, MSP, 01601, Ukraine, 

2 Institute of Parasitology, Polish Academy of Sciences, Twarda Street 51/55, 00-818, Warsaw, Poland, and 

3 Department of Biology and Microbiology, University of Wisconsin Oshkosh, Oshkosh, Wisconsin 54901, 

U.S.A. (e-mail: snyder@uwosh.edu) 

ABSTRACT: Rhabdias ambystomae sp. n. is described on the basis of specimens found in the lungs and body 

cavity of the spotted salamander (Ambystoma maculatum) from northwestern Wisconsin, U.S.A. The new species 

differs from Rhabdias bermani in tail shape, arrangement of circumoral lips, and position of vulva, from Rhabdias 

tokyoensis in the morphology and size of the buccal capsule and the shape of the esophagus, and from 

Rhabdias americanus in the absence of pseudolabia at the cephalic extremity and the shape of the tail. Rhabdias 

ambystomae sp. n. is the first species of the genus described from salamanders in North America. 

KEY WORDS: Nematoda, Rhabdiasidae, Rhabdias ambystomae sp. n., salamanders, Ambystoma maculatum, 

Wisconsin, U.S.A. 

Nematodes of the genus Rhabdias Stiles and 

Hassall, 1905, are globally distributed lung parasites 

of amphibians and reptiles. Among amphibian 

hosts, the vast majority of Rhabdias species 

have been reported from anurans (frogs and toads), 

whereas only 2 species of the genus have been 

described from caudatans (salamanders): Rhabdias 

bermani Rausch, Rausch, and Atrashkevich, 1984, 

from the Siberian newt Salamandrella keyserlingii 

Dybowski, 1870, in the eastern Palearctic (Rausch 

et al., 1984) and Rhabdias tokyoensis Wilkie, 

1930, from Cynops spp. in Japan (Wilkie, 1930). 

In North America, Rhabdias spp. previously have 

been found in the lungs and body cavities of several 

species of salamanders (Lehmann, 1954; Dyer 

and Peck, 1975; Price and St. John, 1980; Coggins 

and Sajdak, 1982; Muzzall and Schinderle, 1992; 

Bolek and Coggins, 1998; Goldberg et al., 1998). 

These nematodes were identified as either Rhabdias 

sp., Rhabdias ranae Walton, 1929, or Rhabdias 

joaquinensis Ingles, 1935, the latter 2 species 

normally restricted to anuran amphibians. 

In the course of investigations of the helminth 

fauna of Wisconsin amphibians, infections by a 

species of Rhabdias were detected in the lungs and 

body cavities of 2 specimens of the spotted salamander 

Ambystoma maculatum (Shaw, 1802). 

Morphological examination revealed these worms 


228 


to represent a new species of the genus Rhabdias. 

This species is described herein as Rhabdias ambystomae 

sp. n. 


Amphibians were collected from a roadside wetland 

near Pigeon Lake, Bayfield County, Wisconsin, U.S.A. A 

total of 26 gravid and 110 subadult nematodes were 

found in 2 of 4 A. maculatum. Nematodes were fixed in 

hot formalin and postfixed in 70% ethanol. Prior to light 

microscopic examination, worms were cleared in glycerol 

by gradual evaporation from a 5% solution of glycerol in 

70% ethanol. Nematodes to be examined with scanning 

electron microscopy (SEM) were postfixed in ethanol, dehydrated 

in a graded series of ethanol and acetone, and 

critical point dried in a Desk II Critical Point Dryer® 

(Denton Vacuum, Inc., Moorestown, New Jersey, U.S.A.) 

with CO2 as the transition fluid. The specimens were 

mounted on stubs, coated with gold, and examined with 

a Hitachi 2460N® scanning electron microscope (Hitachi 

USA, Mountain View, California, U.S.A.) at an accelerating 

voltage of 10-15 kV 

Five specimens of R. bermani from S. keyserlingii collected 

in Magadanskaya Region, Russia, 10 specimens of 

R. tokyoensis from the brown newt Cynops ensicauda 

(Hallowell, 1860) collected on Okinawa Island, Japan, 20 

specimens of R. ranae from the northern leopard frog 

Rana pipiens (Schreber, 1782) collected in Wisconsin, 

U.S.A., and 18 specimens of Rhabdias americanus Baker, 

1978, from the American toad Bufo americanus Hoibrook, 

1836, collected in Wisconsin, U.S.A. were examined 

by light microscopy and measured after being 

cleared as above. All measurements are given in micrometers 

unless otherwise stated. Measurements are given 

for the holotype followed by minimum and maximum 

measurements of paratypes in parentheses.

KUZMIN ET AL.—RHABDIAS AMBYSTOMAE SP. N. 229 

Table 1. Measurements taken from gravid Rhabdias ambystomae sp. n. (type series; n = 18) (measurements 

in micrometers unless otherwise noted). 

Character 

Body length (mm) 

Body width 

Buccal capsule depth 

Buccal capsule width 

Width of esophagus, anterior end 

Width of esophagus, muscular region 

Minimum width of esophagus, glandular region 

Esophageal bulb width 

Distance, anterior end of esophagus to nerve ring 

Distance, anterior end of esophagus to nerve ring (as % of esophagus length) 


Esophagus length (as % of body length) 

Distance from anterior end to vulva (mm) 

Distance from anterior end to vulva (as % of body length) 

Tail length 

Tail length (as % of body length) 

Description 

Results 

Rhabdias ambystomae sp. n. 

(Figs. 1-15) 

Because both gravid and subadult nematodes 

were found in the same host specimens, each of 

these stages is described separately. 

GRAVID SPECIMENS (Table 1): Body elongated 

12.6 (6.8-13.0) mm long, 350 (210-430) wide. 

Anterior end rounded, posterior end tapered. Body 

cuticle swollen, especially on anterior and posterior 

thirds of body. Irregularly arranged transverse 

folds formed by cuticular surface. Round oral 

opening surrounded by 6 small lips, each bearing 

1 elongated conical inner papilla and 2 minute outer 

papillae. Inner papillae directed toward oral 

opening. Flat cuticular ring separating each lip 

from edge of oral opening. Buccal capsule cuplike 

in lateral view, round in apical view. Buccal 

capsule depth 15 (12-15), width 17 (15-17). 

Esophagus club-shaped, 560 (450-590) in length, 

with short anterior muscular portion and long posterior 

glandular portion. Nerve ring at level of border 

between muscular and glandular regions of 

esophagus, 160 (130-180) from esophagus anterior 

end. Large optically dense hypodermal cells 

prominent along glandular region of esophagus. 

Excretory glands indistinct, excretory duct short. 

Two large anterior coelomocytes situated between 

posterior end of esophagus and loop of anterior 

genital limb. Intestine thick, filled with brown or 

Holotype 

12.6 

350 

15 

17 

45 

50 

57 

90 

160 

28.6 

560 

4.5 

7 

56 

210 

1.7 

Paratypes 

(mean [min.— max.]) 

10.5 (6.8-13.0) 

342 (210-430) 

14 (12-15) 

17 (15-17) 

43 (40-45) 

54 (45-67) 

59 (42-67) 

85 (65-100) 

159 (130-180) 

28.9 (22.8-34.6) 

546 (450-590) 

5.4 (4.1-7.4) 

5.8 (3.8-7.4) 

55 (47.4-58.2) 

236 (190-300) 

2.3 (1.6-3.1) 

black contents. Intestinal walls thinner in posterior 

than in anterior region of body. Muscular sphincter 

between intestine and rectum present. Rectum 

lined with thick cuticle. Tail wide, conical, 210 

(190-300) in length. Vulva usually postequatorial 

with indistinct lips. Genital system amphidelphic. 

Ovaries straight or slightly twisted, lying along intestine. 

Proximal regions of both ovaries overlap 

level of vulva. Both limbs of genital system bend 

backward at level of oviducts. Anterior oviduct occasionally 

forms 2 loops as it bends. Seminal receptacles 

short, thick walled. Uteri wide, thin 

walled, filled with numerous eggs. Egg size 112- 

130 X 55-65. 

SUBADULT SPECIMENS (Table 2): Body length 

4.15 (3.3-4.8) mm, width 111 (100-120). Anterior 

end rounded, posterior end tapered. Body cuticle 

thin and smooth, slightly swollen at anterior and 

posterior ends. Head structures similar to those in 

gravid worms. Lateral lips situated farther from 

oral opening than submedian lips. Buccal capsule 

and esophagus shapes similar to those in adults. 

Buccal capsule 12 (10-12) deep, 17 (15-17) wide. 

Esophagus 432 (400-460) long. Two elongated 

narrow excretory glands stretch from posterior 

edge of nerve ring to anterior end of intestine. Pair 

of coelomocytes situated subventrally, close to anterior 

limb of genital system. Intestine thick, reddish. 

Rectum sclerotized. Tail elongated, 171 

(150-190) in length. Bulbous projection of body 

wall slightly posteriad to anal opening. Vulva postequatorial. 

Vulva lips indistinct. Genital system 



Figures 1-4. Rhabdias ainbystomae sp. n., adult. 1. Anterior end. 2. Head end, lateral view. 3. Cephalic 

extremity, apical view. 4. Posterior end. 1, 2, 4, holotype; 3, paratype. 


KUZMIN ET AL.—RHABD1AS AMBYSTOMAE SP. N. 231 

Table 2. Measurements of Rhabdias ambystomae sp. n. subadult specimens (n = 15) (measurement in 

micrometers unless otherwise noted). 

Character 

Body length (mm) 

Body width 

Buccal capsule depth 

Buccal capsule width 

Width of esophagus, anterior end 

Width of esophagus, muscular region 

Minimum width of esophagus, glandular region 

Esophageal bulb width 

Distance, anterior end of esophagus to nerve ring 

Distance, anterior end of esophagus to nerve ring (as % of esophagus length) 


Esophagus length (as % of body length) 

Distance from anterior end to vulva (mm) 

Distance from anterior end to vulva (as % of body length) 

Tail length 

Tail length (as % of body length) 

completely developed, but eggs absent. Proximal 

regions of gonads overlap level of vulva. Each gonad 

forms a single loop as it bends. Uteri narrow, 

lacking eggs. 


TYPE HOST: Spotted salamander Arnbystoma 

maculatum (Shaw, 1802). 

TYPE LOCALITY: Roadside wetland near Pigeon 

Lake, Bayfield County, Wisconsin, U.S.A.; 

46°20'84"N, 91°20'58"W. 

SITES OF INFECTION: Lungs, body cavity. 

TYPE SPECIMENS: The type series consists of 

the gravid specimens only. Holotype: U.S. National 

Parasite Collection, Beltsville, Maryland, 

U.S.A., USNPC 90869. Paratypes: USNPC 

90870 (9 specimens); Department of Parasitology, 

Institute of Zoology, Kiev, Ukraine, Vial N 

847 (8 specimens). 

ETYMOLOGY: The new species is named in 

reference to the generic name of its type host. 

PREVALENCE AND INTENSITY: Two of 4 specimens 

of spotted salamander; 20-116 specimens 

of R. ambystomae (5-21 adult nematodes in 

lungs and 15-95 subadults in body cavity). 

Remarks and Discussion 

Rhabdias ambystomae sp. n. is most similar 

morphologically to R. bermani and R. tokyoensis, 

the only other species of Rhabdias described 

from salamanders. The 3 species are similar in 

body size and shape and in egg size (Table 3). 

Rhabdias ambystomae sp. n. differs from R. ber- 

Mean 

4.2 

11 1 

12 

17 

37 

39 

42 

57 

149 

34.4 

432 

10.5 

2.4 

57.4 

171 

4.1 

Minimum 

3.3 

100 

10 

15 

35 

35 

37 

50 

110 

26.8 

400 

8.8 

1.9 

54.4 

150 

3.5 

Maximum 

4.9 

120 

12 

17 

40 

42 

47 

60 

170 

37.8 

460 

12.0 

2.8 

61.8 

190 

5.0 

mani in the absence of a lancet-like cuticular 

swelling of the posterior extremity characteristic 

of the latter species. In R. bermani, the circumoral 

lips are arranged in 2 lateral groups; in each 

group, the lateral lip is situated closer to the oral 

opening than are the submedian lips (Rausch et 

al., 1984). In contrast, the lateral lips of R. ambystomae 

sp. n. are situated farther from the oral 

opening than are the submedian lips, a character 

more prominent in subadult worms than in 

adults (Figs. 3, 7). Additionally, the vulva of R. 

ambystomae sp. n. is usually postequatorial, 

whereas it is equatorial in R. bermani (Rausch 

et al., 1984). The most apparent differences between 

R. ambystomae sp. n. and R. tokyoensis 

are the markedly smaller buccal capsule and narrower 

esophagus of R. ambystomae (Table 3). 

Rhabdias ambystomae sp. n. can be distinguished 

readily from 2 Rhabdias species common 

in North American amphibians, R. atnericanus 

and R. ranae, by the absence of lateral 

pseudolabia on the cephalic end. The presence 

and shape of pseudolabia in R. americanus and 

R. ranae were well documented by Baker (1978) 

and confirmed by examination of specimens collected 

as part of the present study. In addition, 

R. americanus has a more elongated tail (cf. 

Baker, 1978, fig. 3) compared with that of R. 

ambystomae sp. n. 

Parasitic nematodes of the genus Rhabdias 

have been reported in several species of salamanders 

from the midwestern U.S.A. Price and 

St. John (1980) reported finding an undeter- 



Figures 5-9. Rhabdias ambystomae sp. n., subadult. 5. Anterior end. 6. Head end, lateral view. 7. 

Cephalic extremity, apical view. 8. Posterior end. 9. Anterior loop of genital system and coelomocytes. 

mined species of Rhabdias in the smallmouth ( = subadult) nematodes were found in the body 

salamander Ambystoma texanum (Matthes, cavity. Bolek and Coggins (1998) found unde- 

1855) from Illinois. Adult specimens were found termined subadult Rhabdias in the body cavity 

in the lungs of the host, whereas "larval" of the lungless red-backed salamander Pletho- 



Figures 10-15. External morphology of Rhabdias ambystomae sp. n. 10. Cephalic extremity of adult. 

11. Cephalic extremity of subadult; note amphid marked with an arrow. 12. Vulva of adult. 13. Vulva of 

subadult; note obliteration of the female genital opening at this stage. 14. Anus of adult. 15. Anus of 

subadult. Scale bars = 



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rfon cinereus (Green, 1818) from collecting sites 

within a few kilometers of where R. ambystomae 

sp. n. was collected as part of the present study 

(J. R. Coggins, University of Wisconsin, Milwaukee, 

personal communication). Oddly 

enough, no Rhabdias were found in A. maculaturn 

collected by Bolek and Coggins (1998) 

from the same area. This information suggests 

that R. ambystomae sp. n. may also parasitize P. 

cinereus in northern Wisconsin but cannot reach 

maturity in this lungless amphibian. Further examinations 

will be necessary for confirmation. 

Other authors identified lung nematodes collected 

from salamanders in Wisconsin and Michigan 

as R. ranae (Coggins and Sajdak, 1982; 

Muzzall and Schinderle, 1992). This species has 

been recorded in a number of North American 

anurans, most of which belong to the family 

Ranidae (Baker, 1978; Yoder and Coggins, 

1996; Bursey and DeWolf, 1998). In our opinion, 

the determination of the material from salamanders 

as R. ranae is questionable because of 

the high level of host specificity demonstrated 

by most representatives of Rhabdias; members 

of this genus have never been found to parasitize 

hosts from more than a single order (Rausch et 

al., 1984). Similarly, the report by Goldberg et 

al. (1998) of a frog parasite, Rhabdias joaquinensis 

Ingles, 1935, from salamanders in California 

may also represent a misidentification and 

is in need of confirmation. 

Rhabdias ambystomae sp. n. is the first species 

of this genus described from North American 

salamanders. The rich salamander fauna of 

North America and the strict specificity of rhabdiasids 

to their hosts indicate that further studies 

of material from the field or museum collections 

may reveal the presence of more species of 

Rhabdias unique to the salamanders of the New 

World. 


We thank Dr. Gennadiy Atrashkevich, who 

kindly provided several specimens of 5. keyserlingii 

for our investigation, and Dr. Hideo Hasegawa 

for the loan of R. tokyoensis. We are 

grateful to Dr. Robert Wise for assistance with 

SEM. Collection of amphibians in Wisconsin 

was conducted under a permit provided by the 

Wisconsin Department of Natural Resources. 

This research was supported by a grant from the 

Vander Putten International Fund of the University 

of Wisconsin Oshkosh.


Baker, M. R. 1978. Morphology and taxonomy of 

Rhabdias spp. (Nematoda: Rhabdiasidae) from 

reptiles and amphibians of southern Ontario. Canadian 


Bolek, M. G., and J. R. Coggins. 1998. Helminth 

parasites of the spotted salamander Ambystoma 

maculatum and red-backed salamander Plethodon 

c. cinereus from northwestern Wisconsin. Journal 


65:98-102. 

Bursey, C. R., and W. R. DeWolf. 1998. Helminths 

of the frogs, Rana catesbeiana, Rana clamitans, 

and Rana palustris, from Coshocton County, 

Ohio. Ohio Journal of Science 98:28-29. 

Coggins, J. R., and R. A. Sajdak. 1982. A survey of 




Dyer, W. G., and S. B. Peck. 1975. Gastrointestinal 

parasites of the cave salamander, Eurycea lucifuga 

Rafinesque, from the south-eastern United States. 


Goldberg, S. R., C. R. Bursey, and H. Cheam. 1998. 

Composition and structure of helminth communities 

of the salamanders, Aneides lugubrls, Batrachoseps 

nigriventris, Ensatina eschscholtzii 

(Plethodontidae), and Taricha torosa (Salamandridae) 

from California. Journal of Parasitology 

84:248-251. 


Lehmann, D. L. 1954. Some helminths of west coast 

urodels. Journal of Parasitology 40:231. 

Muzzall, P. M., and D. B. Schinderle. 1992. Helminths 

of the salamanders Ambystoma t. tigrinum 

and Ambystoma laterale (Caudata, Ambystomatidae) 

from southern Michigan. Proceedings of the 


205. 

Price, R. L., and T. St. John. 1980. Helminth parasites 

of the small-mouth salamander, Ambystoma 

texamim Matthes, 1855, from William County, Illinois. 

Proceedings of the Helminthological Society 


Rausch, R. L., V. R. Rausch, and G. I. Atrashkevich. 

1984. Rhabdias bermani sp. n. (Nematoda, 

Rhabdiasidae) from the Siberian salamander (Hynobius 

keyseiiingi) from the north-east of Asia. 

Zoologicheskiy Zhurnal 63:1297-1303. (In Russian.) 

Wilkie, J. S. 1930. Some parasitic nematodes from 

Japanese Amphibia. Annals and Magazine of Natural 

History 6:606-614. 

Yamaguti, S. 1935. Studies on the helminth fauna of 

Japan. Part 10. Amphibian nematodes. Japanese 




crucifer Wied, and the wood frog, Rana 



63:211-214. 




Supplemental Diagnosis of Myxobolus gibbosus (Myxozoa), with a 

Taxonomic Review of Myxobolids from Lepomis gibbosus 

(Centrarchidae) in North America 

DAVID K. CONE 

Department of Biology, Saint Mary's University, Halifax, Nova Scotia, Canada B3H 3C3 

(e-mail: david.cone@stmarys.ca) 

ABSTRACT: Myxobolus gibbosus Herrick, 1941 (Myxosporea) is reported from the connective tissue of gills of 

Lepomis gibbosus (Centrarchidae) in Algonquin Park, Ontario, Canada. The new material (formalin-preserved) 

is used to supplement the original taxonomic diagnosis of nearly 60 yr ago. Spores are round to oval in valvular 

view, 11-14 jjim long and 10-11 jxm wide, with a distinctly blunt capsular region. The polar capsules are 

relatively large for the size of the spore, measuring 6-7 jxm long and 3.5-4.0 (Jim wide, and aligned almost 

parallel to each other. There are 8-12 loose filament coils lying up to 45° to the long axis of the capsule. The 

taxonomy of species of Myxobolus described or reported from L. gibbosus in North America is examined, and 

the following are considered to be valid taxa: Myxobolus dechtiari Cone and Anderson, 1977; M. gibbosus; 

Myxobolus magnasphcrus Cone and Anderson, 1977; Myxobolus osburni Herrick, 1936; Myxobolus paralintoni 

Li and Desser, 1985; and Myxobolus uvidiferus Cone and Anderson, 1977. Comparative photographs of spores 

accompany differential diagnoses of the 6 species. Myxobolus gibbosus Li and Desser, 1985, and Myxobolus Hi 

Desser, 1993, are junior synonyms of M. uvidiferus. Myxobolus lepomicus Li and Desser, 1985, is considered a 

species inquirendae, and the reports of Myxobolus cyprinicola Reuss, 1906, and Myxobolus poecilichthidis 

Fantham, Porter, and Richardson, 1939, from L. gibbosus are considered misidentifications. 

KEY WORDS: Myxobolus gibbosus, Myxosporea, redescription, differential diagnoses, pumpkinseed sunfish, 

Lepomis gibbosus, Centrarchidae, Algonquin Park, Canada. 

Myxobolus gibbosus Herrick, 1941 (Myxozoa) 

was described from connective tissue of the 

gill arch of pumpkinseed sunfish (Lepomis gibbosus 

(Linnaeus, 1758)) from the island region 

of western Lake Erie (Herrick, 1941). In subsequent 

surveys of myxosporean parasites of 

pumpkinseed (Cone and Anderson, 1977a, b; Li 

and Desser, 1985; Hayden and Rogers, 1997), 

the parasite was not encountered. However, during 

a new survey of myxosporeans of fish in 

Algonquin Park, a single pseudocyst of M. gibbosus 

was discovered. This rare find enabled the 

author to assess information provided in the 

original species description and to critically 

compare the parasite with other species of the 

genus reported from pumpkinseed. The present 

study describes the new material and reviews the 

taxonomy of myxobolids from pumpkinseed in 

North America. 


Nine pumpkinseed (6-8.9 cm in total length) were 

collected in baited trapnets set 20 June 1994 and 21 

June 1995 in the shallows of Lake Sasajewan 

(45°35'N; 78°30'W), Algonquin Park, Ontario, Canada. 

The fish were pithed and necropsied. All body organs 

and tissues were examined microscopically for 

myxosporean pseudocysts, and, when found, they were 

236 

fixed in 10% buffered formalin. Fixed pseudocysts 

were punctured and the spore contents stabilized in 

temporary mounts prepared with 1% agar (Lom, 

1969). Spores were photographed with interference 

contrast optics. Enlarged photographic prints of individual 

spores were used to determine spore dimensions. 

Descriptive terminology follows Lom and Dykova 

(1992). Measurements are presented in micrometers. 

The sample of M. gibbosus was compared with 

other species of Myxobolus in the author's collection, 

namely Myxobolus dechtiari Cone and Anderson, 

1977; Myxobolus magnaspherus Cone and Anderson, 

1977; Myxobolus osburni Herrick, 1936; Myxobolus 

paralintoni Li and Desser, 1985; and Myxobolus uvitliferus 

Cone and Anderson, 1977. Syntype slides of M. 

gibbosus (NMCICP 1984-0359), Myxobolus lepomicus 

(NMCICP 1984-0362), and M. paralintoni (NMCICP 

1984-0364) housed in the parasite collection of the Canadian 

Museum of Nature were also examined. A photo-voucher 

(negative film) is deposited in the United 

States National Parasite Collection (USNPC), Beltsville, 

Maryland, U.S.A. 

Results 

Myxobolus gibbosus Herrick, 1941 

(Figs. 1 and 2) 

Supplementary diagnosis 


Pseudocyst egg-shaped, gray-white and minute 

(250 long), embedded in connective tissue 

surrounding base of gill arch. Spores round to

Figure 1. Spores of Myxobolus gibbosus. Scale bar = 10 [Jim. 

oval in valvular view, with blunt capsular edge. 

Spores 11.8 ± 0.9 (11-14, « = 10) long and 

10.6 ± 0.5 (10-1 1) wide. Width-to-length ratio 

1:1.09 ± 0.08 (1.05-1.3). Polar capsules oval, 

6.8 ± 0.3 (6-7) long and 4.0 ± 0.3 (3.5-4) wide, 

aligned almost parallel to each other. Polar filaments 

in 8-12 loose coils, lying up to 45° to long 

axis of capsule. Capsulogenic nuclei prominent, 

triangular. Shallow intercapsular appendix evident 

in some spores. Sutural ridge thin and 

smooth. 


HOST: Pumpkinseed sunfish (Lepomis gibbosus) 

(Centrarchidae); total length 6.2 cm, 1 + 

yr old. 

LOCALITY/COLLECTION DATE: Lake Sasajewan, 

Algonquin Park, Ontario, Canada (45°35'N; 

78°30'W), 20 June 1994. 

CONE—DIAGNOSIS OF MYXOBOLUS 237 

SITE OF INFECTION: Connective tissue of gill 

arch. 

PREVALENCE AND INTENSITY OF INFECTION: One 

of 9 fish infected with 1 pseudocyst. 

SPECIMENS DEPOSITED: Photo-voucher USNPC 

No. 091157.00. 

Remarks 

Myxobolus gibbosus has not been reported in 

surveys of myxozoans in L. gibbosus from Algonquin 

Park (Cone and Anderson 1977a, b; Li 

and Desser, 1985). It was probably not overlooked, 

for the parasite has several distinct diagnostic 

features. It forms small but obvious 

pseudocysts in the connective tissue of the gill 

arch and produces round spores with a blunt 

capsular end. The polar capsules are relatively 

large, the length being about half the length of 

the spore, and they are arranged almost parallel 



Table 1. Comparison of pertinent taxonomic information about Myxobolus gibbosus reported in the original 

species description and that observed in the present study. 

Host 

Locality 

Tissue site 

Pseudocyst 

Spore length 

Spore width 

Spore thickness 

Polar capsule length 

Polar capsule width 

Polar filament coils 

Herrick (1941)* 

Lepomis gibbosus 

Lake Erie 

Connective tissue of gill 

Round, 0.75 mm 

10.6-12.3 

9.8-12.3 

6.5-8.2 

5.7-7.4 

3.3-4.1 

8-12 

* Based on fresh material in a hanging drop preparation. 

t Based on formalin-fixed material in asar wet mounts. 

to each other. It appears then that M. gibbosus 

is simply rare in this region. The dimensions of 

the preserved spores found in the present study 

are similar to those described by Herrick (1941) 

from fresh material (Table 1). It should be noted 

that dimensions of fixed spores are often smaller 

than those of fresh spores because shrinkage can 

take place during fixation. This means that fresh 

spores of M. gibbosus in Algonquin Park may 

be slightly larger than those described originally 

by Herrick (1941). 

Spores of other species of Myxobolus (M. dechtiari, 

M. magnaspherus, M. osburni, M. paralintoni, 

and M. uvuliferus) from L. gibbosus are 

presented for comparative purposes (Figs. 3—7). 

Each species has a distinct spore shape and specific 

tissue site in which it develops and is readily 

identified by these indicators. Myxobolus 

paralintoni (Fig. 4) has oval spores in frontal 

view and develops in the bulbus arteriosus of the 

heart (Hayden and Rogers, 1997; Cone and 

Overstreet, 1998). Myxobolus dechtiari (Fig. 5) 

has spores that are broadly pyriform in frontal 

view and develops in gill tissue (Cone and Anderson, 

1977a). Myxobolus uvuliferus has slightly 

compressed spores in frontal view usually 

with the width greater than length, often has polar 

capsules dissimilar in the length, and develops 

in the connective tissue capsule surrounding 

the metacercaria of Uvulifer ambloplites 

(Hughes, 1927) Dubois, 1938 (Cone and Anderson, 

1977a). Myxobolus osburni has round 

spores in frontal view and develops in the exocrine 

tissue of the pancreas (Cone and Anderson, 

1977a). Myxobolus magnaspherus has round 

spores in frontal view that are huge, often 20 


Present studyt 

Lepomis gibbosus 

Lake Sasajewan 

Connective tissue of gill 

Round, 0.25 mm 

11-14 

10-11 

— 

6-7 

3.5-4 

8-11 

|xm in diameter, and develops in connective tissue 

of the body, including the peritoneum (Cone 

and Anderson, 1977a). 

Taxonomic Key to the Species of Myxobolus 

Infecting Pumpkinseed Sunfish 

la. Spore length more than 16 jjim 

M. magnaspherus (Fig. 8) 

Ib. Spore length less than 16 |xm 2 

2a. Polar capsules aligned more or less parallel - 

M. gibbosus (Fig. 9) 

2b. Polar capsules converged anteriorly 3 

3a. Spore circular in frontal view M. osburni (Fig. 10) 

3b. Spore not circular in frontal view 4 

4a. Spore width greater than spore length 

M. uvuliferus (Fig. 11) 

4b. Spore width less than spore length 5 

5a. Spore oval in frontal view M. paralintoni (Fig. 12) 

5b. Spore broadly pyriform in frontal view 

M. dechtiari (Fig. 13) 

Discussion 

Ten species of Myxobolus Butschli, 1882 

(Myxosporea) have been reported from L. gibbosus 

in North America (Herrick, 1936, 1941; 

Cone and Anderson, 1977a, b; Ingram and 

Mitchell, 1982; Li and Desser, 1985; Desser, 

1993; Cone and Overstreet, 1998). The author 

has necropsied L. gibbosus from Algonquin Park 

and from Lake Erie and has to date encountered 

6 of the 10 species, namely M. dechtiari, M. 

gibbosus, M. magnaspherus, M. osburni, M. 

paralintoni, and M. uvuliferus. 

The reports of Myxobolus cyprinicola Reuss, 

1906, and Myxobolus poecilichthidis Fantham, 

Porter, and Richardson, 1939, from the brain and 

heart and from the gills, respectively, of L. gib-


Figures 2-7. Photographs of spores in frontal view of species of Myxobolus known to parasitize Lepomis 

gibbosus in North America. 2. Myxobolus gibbosus (formalin-preserved). 3. Myxobolus paralintoni (formalin-preserved). 

4. Myxobolus dechtiari (formalin-preserved). 5. Myxobolus uvuliferus (formalin-preserved). 

6. Myxobolus osburni (formalin-preserved). 7. Myxobolus magnaspherus (fresh spore). Scale bar 

= 10 jjim and applies to all figures. 

bosus in Algonquin Park (Li and Desser, 1985) 

are considered misidentifications. Both of these 

species have similarities in tissue site and spore 

morphology to M. dechtiari and M. paralintoni 

and could have easily been confused with them. 

Li and Desser (1985) were apparently unaware 

of the study by Cone and Anderson (1977a) in 

nearby Ryan Lake. 



8 

12 

Figures 8-13. Drawings of spores in frontal view of species of Myxobolus known to parasitize Lepomis 

gibbosus in North America. 8. Myxobolus gibbosus. 9. Myxobolus paralintoni. 10. Myxobolus dechtiari. 11. 

Myxobolus uvuliferus. 12. Myxobolus osburni. 13. Myxobolus magnaspherus. Scale bar = 5 (Jim and applies 

to all figures. 

The type material of M. lepomicus Li and 

Desser, 1985, described from a variety of organs 

of L. gibbosus, has deteriorated, and spores are 

not to be found on the slide. The species description 

includes a schematic drawing of the 

spore. Until additional samples are obtained the 

species is considered a species inquirendae. 

Myxobolus gibbosus Li and Desser, 1985, is a 

homonym of M. gibbosus Herrick, 1941. Desser 

(1993) proposed Myxobolus Hi as a nomen novum 

to replace M. gibbosus Li and Desser, 1985. 

However, Landsberg and Lom (1991) considered 

M. gibbosus Li and Desser, 1985, to be a 

junior synonym of M. uvuliferus, and thus both 

M. gibbosus and M. Hi become junior synonyms 

of M. uvuliferus. The report by Hoffman (1998) 

that M. gibbosus Li and Desser, 1985, is a junior 

synonym of M. osburni cannot be supported on 

the basis of spore shape. 

The 6 confirmed species of Myxobolus mentioned 

above are known to parasitize L. gibbosus 

or related centrarchid fishes in North America. 

Myxobolus magnaspherus and M. paralintoni 

have been found in redear sunfish (Lepomis microlophus 

(Giinther, 1859)) in Mississippi, 

U.S.A. (D. K. Cone, Saint Mary's University, 


and R. M. Overstreet, Gulf Coast Research Laboratory, 

unpublished data) and redbreast sunfish 

(Lepomis auritus (Linnaeus, 1758)) in Maryland, 

U.S.A. (Hayden and Rogers, 1997), respectively. 

Myxobolus osburni has been reported 

(Herrick, 1936; Otto and Jahn, 1943) from bluegill 

sunfish (Lepomis macrochirus Rafinesque, 

1819), smallmouth bass (Micropterus dolomieu 

Lacepede, 1802), and black crappie (Pomoxis nigromaculatus 

(Lesueur, 1829)). The genus clearly 

has undergone a diverse radiation in these 

hosts, and it is of ecological interest that all 6 

species are found in L. gibbosus in Algonquin 

Park and that all occupy distinct and very specific 

tissue sites in this host species. 


The research was funded by a Natural Sciences 

and Engineering Research Council of 

Canada (NSERC) Research Grant awarded to 

the author. Thanks are extended to the staff of 

the Harkness Research Laboratory for their help 

and hospitality and to Sherwin Desser, University 

of Toronto, for providing constructive comments 

on a draft of the paper.


Cone, D. K., and R. C. Anderson. 1977u. Myxosporidan 

parasites of pumpkinseed (Lepomis gibbosus 

L.) from Ontario. Journal of Parasitology 

63:657-666. 

, and . 1977b. Parasites of pumpkinseed 

(Lepomis gibbosus L.) from Ryan Lake, Algonquin 

Park, Ontario. Canadian Journal of Zoology 

55:1410-1423. 

-, and R. M. Overstreet. 1998. Species of Myxobolus 

(Myxozoa) from the bulbus arteriosus of 

centrarchid fishes in North America, with a description 

of two new species. Journal of Parasitology 

84:371-374. 

Desser, S. S. 1993. Myxobolus Hi nom. nov.: a replacement 

for M. gibbosus Li and Desser, 1985 (Myxosporea: 

Myxozoa), preoccupied. Canadian Journal 

of Zoology 71:1461. 

Hayden, K. J., and W. A. Rogers. 1997. Redescription 

of Myxobolus paralintoni (Myxosporea: Myxobolidae), 

with notes regarding new host and locality. 


Herrick, J. A. 1936. Two new species of Myxobolus 

from fishes of Lake Erie. Transactions of the 

American Microscopical Society 55:194-198. 

. 1941. Some myxosporidian parasites of Lake 

Erie fishes. Transactions of the American Microscopical 

Society 66:163-170. 


Hoffman, G. L. 1998. Parasites of North American 

Freshwater Fishes, 2nd ed. Comstock Publishing 

Associates, Cornell University Press, Ithaca, New 

York, U.S.A., and London, U.K. 539 pp. 

Ingram, K. M., and L. G. Mitchell. 1982. Pancreatic 

infections of Myxobolus osburni Herrick (Myxozoa: 

Myxosporea) in the pumpkinseed, Lepomis 

gibbosus (Linnaeus), in Iowa. Journal of Wildlife 

Diseases 18:75-80. 

Landsberg, J. H., and J. Lorn. 1991. Taxonomy of 

the genera of the Myxobolus/Myxosoma group 

(Myxobolidae: Myxosporea), current listing of 

species and revision of synonyms. Systematic Parasitology 

18:165-186. 

Li, L., and S. S. Desser. 1985. The protozoan parasites 

of fish from two lakes in Algonquin Park, 

Ontario. Canadian Journal of Zoology 63:1846- 

1858. 

Loin, J. 1969. On a new taxonomic character in 

Myxosporidia, as demonstrated in descriptions of 

two new species. Folia Parasitologica 16:97-103. 

, and I. Dykova. 1992. Protozoan Parasites of 

Fishes. Developments in Aquaculture and Fisheries 

Science 26. Elsevier, Amsterdam, London, 

New York, Tokyo. 315 pp. 

Otto, G. R., and T. L. Jahn. 1943. Internal myxosporidian 

infections of some fishes of the Okobojii 

region. Proceedings of the Iowa Academy of Science 

50:323-335. 

2001-2002 MEETING SCHEDULE OF THE 

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27 October 2001 

28 October 2001 

28 November 2001 

January 2002 

March 2002 

May 2002 

Symposium—Parasitology in Science. 1:00 PM. Smithsonian Institution, 

Quad Building, (Room 3111), Washington, DC 

(Contact Persons: Bill Moser, 202-357-2473 or Dennis Richardson, 203- 

582-8607). 

Presidential Summit of Parasitology Societies. 8:30 AM. 

Smithsonian Institution, Quad Building, (Room 3112), Washington, DC 

(Contact Persons: Bill Moser, 202-357-2473 or Dennis Richardson, 203- 

582-8607). 

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(Contact Person: Bill Moser, 202-357-2473). 

Date, time, and place to be announced. 






Cuticular Changes in Fergusobiid Nematodes Associated with 

Parasitism of Fergusoninid Flies 

ROBIN M. GiBLiN-DAVis,1-5 KERRIE A. DAViES,2 DONNA S. WiLLiAMS,3 AND 

TED D. CENTER4 

1 Ft. Lauderdale Research and Education Center, University of Florida, 3205 College Avenue, Davie, Florida 

33314-7799, U.S.A. (e-mail: giblin@ufl.edu), 

2 Department of Applied and Molecular Ecology, Adelaide University, Glen Osmond, 5064 South Australia 

(e-mail: kerrie.davies@adelaide.edu.au), 

3 Microbiology and Cell Science, University of Florida, P.O. Box 1 10700, Gainesville, Florida 3261 1-0700, 

U.S.A. (e-mail: dswill@ufl.edu), and 

4 United States Department of Agriculture-Agricultural Research Service, Invasive Plant Research Laboratory, 

3205 College Avenue, Davie, Florida 33314-7799, U.S.A. (e-mail: tcenter@ars.usda.gov). 

ABSTRACT: In the stylet-bearing nematode Fergusobia sp. (Tylenchida: Neotylenchidae), we hypothesize an 

additional separation (apolysis) and loss (ecdysis) of the adult cuticle, without the formation of a new cuticle, 

during the transition from the preparasitic to parasitic female. This pattern is in direct contrast to the characteristic 

4-molt pattern accepted for most nematodes. Transmission electron microscope comparisons of the cuticle of an 

adult parthenogenetic female, male, and preparasitic female from the plant-parasitic phase of the fergusobiid life 

cycle revealed a relatively simple cuticle with an epicuticle, amorphous cortical/median zone, and a striated 

basal zone that is underlain by a relatively thin epidermis and striated somatic muscles. In contrast, the parasitic 

female from the adult fly was without its stylet and cuticle, the epidermis was enlarged, the outer edges of the 

epidermis were modified into microvilli, and the somatic muscles and esophagus were degenerate. The apparent 

hypertrophy and development of epidermal microvilli greatly expand the surface area of the parasitic female 

and presumably increase the nematode's ability to absorb nutrients directly through the epidermis from the host's 

hemolymph without cuticular interference. 

KEY WORDS: Fergusobia, parasitism, Fergusonina, cuticle, epidermis, TEM, molting, nematode, fly, Myrtaceae, 

Australia. 

In the only known mutualistic association between 

nematodes and insects (Maggenti, 1982), 

nematodes of the genus Fergusobia Currie, 

1937, together with flies of the genus Fergusonina 

Currie, 1937, induce galls in young meristematic 

tissues of myrtaceous hosts in Australasia 

(Giblin-Davis et al., 2001). The nematode is apparently 

responsible for gall induction (Currie, 

1937; R. M. Giblin-Davis, unpublished data), 

and the fly for dispersal and sustenance of the 

nematode. The female fly deposits its eggs and 

juvenile nematode parasites in plant tissue (Currie, 

1937). As these nematodes feed, a gall begins 

to form, and the nematodes develop into 

parthenogenetic females that lay eggs giving rise 

to amphimictic male and female nematodes. Inseminated 

preparasitic females are infective and 

invade mature female third-instar fly larvae. Inside 

the fly, the nematodes develop into parasitic 

females that deposit eggs in the fly's hemolymph. 

Juvenile nematodes that hatch from these 


242 


eggs move to the oviducts of the adult fly and, 

together with the fly's eggs, are deposited into 

appropriate plant tissue to begin the next generation. 

During dissections of mature third-instar fly 

larvae from a variety of myrtaceous hosts 

(swamp bloodwood Corymbia ptychocarpa (F. 

Mueller, 1859), South Australian blue gum Eucalyptus 

leucoxylon F. Mueller, 1855, and broadleaved 

paperbark Melaleuca quinquenervia (Cavanilles, 

1797) S. T. Blake, 1958), we observed 

apparent separation (apolysis) and loss (ecdysis) 

of the adult cuticle during transition from the 

preparasitic to parasitic female without the formation 

of a new cuticle (K. A. Davies, unpublished 

data). This assumes that the first molt occurs 

in the egg in Fergusobia, as with other Tylenchida, 

and that 3 molts occur after emergence 

from the egg through to the preparasitic female. 

This pattern is surprising because nematodes 

characteristically undergo 4 molts in their development 

from the juvenile to the adult stage 

(Bird and Bird, 1991). We report on the ultra-

G1BLIN-DAVIS ET AL.—CUTICULAR CHANGES IN FERGUSOBIID NEMATODES 243 

structure of the cuticle of adults at different 

phases of the life cycle of Fergusobia. 


Multilocular flower bud galls of undescribed species 

of Fergusobia and Fergusonina were collected on 9 

August 1999 from C. ptychocarpa at the Sherwood 

Arboretum in Sherwood, Queensland, Australia 

(27°32.06'S; 152°58.39'E). Galls were dissected. Adult 

parthenogenetic female and amphimictic male and preparasitic 

infective female nematodes present in the 

plant tissue were placed separately into Trump's fixative 

for transmission electron microscopy or in formalin-aceto-alcohol 

fixative (Southey, 1970). Mature 

fly larvae (third-instar) and adults were dissected from 

the galls in phosphate-buffered saline (pH 7.2). Parasitic 

female nematodes were removed from the hemocoel 

and placed into Trump's fixative. Specimens 

were postfixed in 2% formaldehyde (prepared from 

paraformaldehyde), 2% glutaraldehyde in 0.1 M cacodylate 

buffer at pH 7.2 for 18 hr at 4°C. After repeated 

rinsing in buffer, specimens were postfixed in 

2% OsO4 in 0.1 M cacodylate buffer at pH 7.2 for 3 

days at 4°C. Nematodes were rinsed in water, fixed 

with 1 % aqueous uranyl acetate, dehydrated through 

100% ethanol into 100% acetone, and infiltrated with 

Spurr's epoxy resin. Blocks were sectioned on an 

RMC® ultramicrotome. Sections were poststained with 

5% aqueous uranyl acetate and lead citrate before 

viewing on a Zeiss EM 10® transmission electron microscope 

at 80 kV. 

Results 

Examination of the cuticle of an adult parthenogenetic 

female and a male nematode revealed 

a relatively simple cuticle with a striated basal 

zone, an amorphous cortical/median zone, and a 

distinct epicuticle (Figs. 1, 2). It is underlain by 

relatively thin epidermis that covers the striated 

somatic muscles. 

Comparisons of the preparasitic female nematode 

from the plant gall and the parasitic female 

nematode from the adult fly show dramatic differences 

(Figs. 3-7). The preparasitic female has 

cuticle, epidermis, and muscles similar to those 

described for the male and parthenogenetic female 

from the plant host (Figs. 3, 4). However, 

the cuticle appears thinner (200-250 nm vs. 

450-550 nm for the parthenogenetic female and 

630—680 nm for the male). The parasitic form 

of the nematode from the adult fly has no cuticle. 

The epidermis is greatly enlarged, and the 

outer edge of the epidermis appears to be modified 

into microvilli (Figs. 5—7). The somatic 

muscles appear degenerated (Fig. 6). 

During the transition from the preparasitic to 

the parasitic female nematode in the larval fly, 

the stylet is lost and the esophagus and intestine 

appear to degenerate. In a parasitic female from 

a fly larva, the remnant of the adult epicuticle 

was present (Fig. 5), but it was not present in 

the parasitic female from an adult fly (Figs. 6, 

7). The apparent hypertrophy and development 

of epidermal microvilli greatly expand the surface 

area of the parasitic female and presumably 

increase the nematode's ability to absorb nutrients 

directly through its epidermis from the 

host's hemolymph without cuticular interference. 

Interestingly, the cuticle represents a form 

of protection against insect host defense mechanisms. 

However, these mechanisms may be 

modified or lacking in the female larva, pupa, 

and adult fly in this mutualistic association. 

Whether there is a strong defense system in male 

flies to prevent parasitism by Fergusobia or the 

nematodes fail to penetrate the male fly larvae 

is not known. 

Discussion 

Riding (1970) reported that microvilli were 

present on the outside of the parasitic female 

stage of Howardula husseyi Richardson, Hesling, 

and Riding, 1977 (=Bradynema sp.) (Allantonematidae), 

a tylenchid parasite of the 

phorid fly, Megaselia halterata Wood, 1910. A 

cuticle was not observed in this stage of the 

nematode, suggesting that the microvilli were of 

epidermal origin and that there could have been 

an additional apolysis and ecdysis without cuticular 

replacement, as appears to occur in Fergusobia. 

The epidermis in this nematode was hypertrophied. 

In addition, the stylet and esophagus 

are not present in this form of H. husseyi 

(Poinar, 1979). Subbotin et al. (1994) reported 

that entomoparasitic females of Wachekitylenchus 

bovieni (Wachek, 1955) Slobodyanyuk, 

1986 (Parasitylenchidae), and Bradynema rigidum 

(von Siebold, 1836) zur Strassen, 1892 (Allantonematidae), 

had similar body wall morphology 

to H. husseyi. Entomoparasitic females 

of the tylenchid Skarbilovinema laumondi Chizhov 

and Zakharenkova, 1991 (lotonchiidae), 

exhibited a body wall composed of a "spongy" 

layer of the epidermis formed by interwoven 

and fused microvilli without a cuticle (Subbotin 

et al., 1993). 

In contrast, the epicuticle is apparently retained 

by the entomoparasitic amphimictic female 

of Paraiotonchium nicholasi Slobodyanyuk, 

1975 (=Heterotylenchus sp.) (lotonchiidae) 

(Nicholas, 1972). Ultrastructural differences 



629 nm 

;./::•" •':6,'-, -,:•••::•.,;-; 

mkdM \• 

Figures 1, 2. Longitudinal sections of the cuticle of Fergusobia sp. ex Corymbia ptychocarpa from galled 

flower buds. 1. Adult parthenogenetic female. 2. Adult male. BS = basal striations; C/M = cortical/median 

zone; E = epicuticle. 

were observed between the amphimictic female, 

parthenogenetic female, and amphimictic-phase 

juvenile of P. nicholasi from the body cavity of 

the Australian bush fly Musca vetustissima 


Walker, 1857 (Nicholas, 1972). Fourth-stage juvenile 

(J4) nematodes are deposited into cow 

dung where they mate, "metamorphose" into 

the infective form, penetrate the fly larva, and

- 

GIBLIN-DAVIS ET AL.—CUTICULAR CHANGES IN FERGUSOBIID NEMATODES 245 

.. 

'•••':«•' 

- 

244 nm 

•'?"... - 

• . 

Figures 3, 4. Longitudinal sections of the cuticle of a preparasitic adult female of Fergusobia sp. ex 

Corymbia ptychocarpa from galled flower buds. 3. Close-up showing epidermal folds. 4. Different section. 

BS = basal striations; C/M = cortical/median zone; E = epicuticle; Ep = epidermis; M = muscle. 



CR Mv 

V/"7f:.:v r^25"/ '••'•:%6&^!£^^±-^ 

- > V - : : . : ;•- '

GIBLIN-DAVIS ET AL.—CUTICULAR CHANGES IN FERGUSOBIID NEMATODES 247 

harenkova and Chizhov, 1991, and Allantonema 

rnirabile Leuckart, 1884 (Allantonematidae), 

were similar to those of P. nicholasi, being composed 

of a hypertrophied epidermis with microvilli 

that was covered by a cuticle-like layer 

(Subbotin et al., 1994). 

Insect hemolymph characteristically has high 

levels of amino acids, trehalose, other nonamino 

organic acids, and salts (Chapman, 1972), making 

it a nutrient-rich environment for parasites 

that can overcome innate host defense mechanisms. 

Insect-parasitic tylenchid nematodes have 

adapted to the challenges of obtaining nutrition 

from a living insect host in a variety of ways, 

including acquisition per os (through the 

mouth), through a modified or absent cuticle, or 

through prolapsis and modification of the uterus 

as in the Sphaerulariinae (Sphaerulariidae). Tylenchids 

from the Neotylenchidae, Allantonematidae, 

lotonchiidae, and Parasitotylenchidae 

have insect-parasitic forms that are obese and 

have degenerate esophagi, intestines that are degenerate 

or modifed as storage organs, and the 

stylet often sunken into the body or even lacking 

(Siddiqi, 2000), suggesting that they employ 

some form of transcuticular or transepidermal 

uptake. 

Deladenus (Neotylenchidae), Paraiotonchium 

(lotonchiidae), Howardula (Allantonematidae), 

Skarbilovinerna (lotonchiidae), and Fergusobia 

(Neotylenchidae) may represent contemporary 

examples of an evolutionary trend from per os 

to transepidermal nutrient acquisition in insectparasitic 

Tylenchida. Of course, this is a highly 

speculative exercise until more information 

about the transition between preparasitic and 

parasitic females is known and some independent 

phylogenetic data are available. The evolutionary 

trend is hypothesized to be: 1) Per os 

acquisition via a stylet, esophagus, and gut. This 

strategy takes advantage of the existing stylet for 

feeding on fungi, plants, or other invertebrates. 

It is a less energy- and time-efficient method of 

nutrient acquisition for a hemocoelic parasite because 

obtaining food through the stylet requires 

expending energy to maintain and operate its 

esophagus and intestine. 2) Per os acquisition 

with thinning and partial apolysis of the cuticle 

and coincident epidermal folding to increase surface 

area for supplemental transcuticular uptake 

of nutrients (possibly Deladenus spp.). 3) Early 

per os acquisition followed by apolysis, partial 

absorption of the cuticle without the creation of 

a new cuticle, and folding of the epidermis such 

that uptake is transcuticular and somatic muscles, 

esophagus, and gut degenerate (e.g., Paraiotonchium). 

4) Early per os acquisition followed 

by full apolysis and ecdysis without the 

creation of a new cuticle. There is hypertrophy 

and folding of the epidermis, and nutrient uptake 

is transepidermal, somatic muscles and esophagus 

degenerate, and the gut degenerates or is 

transformed into a storage organ (e.g., Howardula, 

Skarbilovinema, and Fergusobia). The epidermal 

hypertrophy and folding are superficially 

similar to the formation of plicae (epidermal 

folds) during the development of a new cuticle 

(Bird and Bird, 1991) but are more extensive 

and apparently are not accompanied by the formation 

of a new cuticle. 


We thank Drs. Bill Howard and Thomas 

Weissling for review of the manuscript and Matthew 

Purcell, Jeff Makinson, and Dr. John 

Goolsby for making the senior author's visit to 

the Australian Biological Control laboratory in 

Indooroopilly, Queensland, Australia, such a 

productive and enjoyable experience. This project 

was funded in part by USDA-ARS Specific 

Cooperative Agreement No. 58-6629-9-004 

from the USDA Invasive Plant Research Laboratory 

in Davie, Florida, U.S.A. This is Florida 

Agricultural Experiment Station Journal Series 

No. R-07870. 


Bird, A. F., and J. Bird. 1991. The Structure of Nematodes, 

2nd ed. Academic Press, Inc., New York, 

U.S.A. 316 pp. 

Chapman, R. F. 1972. The Insects: Structure and 

Function. American Elsevier Publishing Co., Inc., 

New York, U.S.A. 819 pp. 

Currie, G. A. 1937. Galls on Eucalyptus trees: A new 

type of association between flies and nematodes. 

Proceedings of the Linnean Society of New South 

Wales 62:147-174. 

Giblin-Davis, R. M., K. A. Davies, G. S. Taylor, and 

W. K. Thomas. 2001. Entomophilic nematode 

models for studying biodiversity and cospeciation. 

Pages 00-00 in Z. X. Chen, S. Y. Chen, and D. 

W. Dickson, eds. Nematology, Advances and Perspectives. 

Tsinghua University Press/Springer- 

Verlag, New York, U.S.A. (In press.) 

Maggenti, A. R. 1982. General Nematology. Springer- 

Verlag, New York. 372 pp. 

Nicholas, W. L. 1972. The fine structure of the cuticle 

of Heterotylenchus. Nematologica 18:138-140. 

Poinar, G. O., Jr. 1979. Nematodes for Biological 



Control of Insects. CRC Press, Inc., Boca Raton, Subbotin, S. A., V. N. Chizhov, and N. N. Zakhar- 

Florida, U.S.A. 277 pp. 

Riding, I. L. 1970. Microvilli on the outside of a nematode. 

Nature 226:179-180. 

Siddiqi, M. R. 2000. Tylenchida: Parasites of Plants 

and Insects, 2nd ed. CABI Publishing, St. Albans, 

U.K. 848 pp. 

Southey, J. F., ed. 1970. Laboratory methods for work 

with plant and soil nematodes, 5th ed. Technical 

Bulletin 2 of the Ministry of Agriculture, Fisheries 

and Food. Her Majesty's Stationery Office, London, 

U.K. 148 pp. 

enkova. 1993. Ultrastructure of the body wall of 

parasitic and infective females of Skarbilovinema 

laumondi (Tylenchida: lotonchiidae). Fundamental 

and Applied Nematology 16:1-4. 

, , and . 1994. Ultrastructure of 

the integument of parasitic females in entomoparasitic 

tylenchids. 1. Two species of the genus 

Wachekitylenchus, Allantonema mirabile and Bradynema 

rigidum. Russian Journal of Nematology 

2:105-112. 

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Tegumentary Ultrastructure (SEM) of Preadult and Adult 

Lobatostoma jungwirthi Kritscher, 1974 (Trematoda: Aspidogastrea) 

ANALIA PAOLA' AND MARIA CRISTINA DAMBORENEA2 

Departamento Cientffico Zoologia Invertebrados, Facultad de Ciencias Naturales y Museo, 

Paseo del Bosque s/n° -1900- La Plata, CONICET.PIP 4728/96 Argentina 

(e-mail: 'apaola@museo.fcnym.unlp.edu.ar; 2cdambor@museo.fcnym.unlp.edu.ar) 

ABSTRACT: Larval Lobatostoma jungwirthi Kritscher, 1974 (Trematoda: Aspidogastrea) parasitize the digestive 

gland of Heleobia parchappii (d'Orbigny, 1835) (Mollusca: Hydrobiidae) and, as adults, the posterior intestine 

of the chameleon cichlid Cichlasotna facetum (Jenyns, 1842) (Pisces, Cichlidae). Currently, L. jungwirthi is the 

only aspidogastrid reported from freshwater fishes in Argentina. Tegumentary structures of preadults and adults 

of L. jungwirthi were observed under scanning electron microscopy. In the preadult, 2 types of sensory receptors 

were observed: monociliate papillae of intermediate length on the walls and crests of the ventral adhesive disc 

as well as on the disc periphery and oral lobules, and nonciliate dome-shaped papillae on the crests of the 

ventral adhesive disc, neck, and oral lobules. In adults, other types of sensory receptors could be observed: in 

the posterior dorsal region, monociliate papillae with longer cilia than those found in the preadult, and a multiciliate 

structure in the dorsal region at the posterior third of the body. This is the first record of a surface 

multiciliate receptor in aspidogastreans. The pores of marginal glands were found only between the anterior 

alveoli. 

KEY WORDS: Aspidogastrea, Lobatostoma jungwirthi, tegument, sensory papillae, SEM, Argentina. 

Lobatostoma jungwirthi Kritscher, 1974 

(Trematoda: Aspidogastrea), is the only species 

of the genus that parasitizes freshwater fishes. It 

was first found in 1974, in the stripefin eartheater 

Gymnogeophagus rhabdotus (Hensel, 

1870), in the Sinus River, Brazil (Kritscher, 

1974). Lunaschi (1984) found it in the posterior 

intestine of the chameleon cichlid Cichlasoma 

facetum (Jenyns, 1842) (Pisces: Cichlidae) at 2 

localities in Buenos Aires Province. Later, Zylber 

and Ostrowski de Nunez (1999) described 

the larval stages of L. jungwirthi from the gonad 

of Heleobia castellanosae (Gaillard, 1974) (Gastropoda: 

Hydrobiidae) collected in an artificial 

pond in Buenos Aires City. 

To date, the morphology of the larval (Zylber 

and Ostrowski de Nunez, 1999) and adult 

(Kritscher, 1974; Lunaschi, 1984) stages of this 

species is known only at the light microscopy 

level. Several investigators have described the 

tegumentary ultrastructure of adult aspidogastrids, 

such as Aspidogaster conchicola Baer, 

1826 (Halton and Lyness, 1971), and Cotylogaster 

occidentalis Nickerson, 1902 (Ip and 

Desser, 1984). The variability of tegumentary 

sensory structures of the cotylocidia of C. occidentalis 

(Fredericksen, 1978), the development 

and growth of the ventral adhesive disc of C. 


249 

occidentalis and A. conchicola (Fredericksen, 

1980), and the sensory receptors of the larval 

stage of Lobatostoma manteri Rohde, 1973 

(Rohde and Watson, 1989a, b, 1992), and Multicotyle 

purvisi Dawes, 1941 (Rohde and Watson, 

1990b, c, d), were also studied. The aim of 

the present paper is to describe the tegumentary 

ultrastructure of juvenile and adult specimens of 

L. jungwirthi under scanning electron microscopy 

(SEM). 


Parasites removed from the posterior intestine of C. 

facetum were identified as L. jungwirthi on the basis 

of the descriptions of Kritscher (1974) and Lunaschi 

(1984). Juvenile stages were found in the digestive 

gland of Heleobia parchappii (d'Orbigny, 1835) (Mollusca: 

Hydrobiidae), and adult specimens were obtained 

from the posterior intestine of C. facetum. Both 

host species were naturally parasitized by this aspidogastrid 

in Saladita Pond, Avellaneda District, Buenos 

Aires. 

The specimens were fixed in 10% formalin and 

washed in distilled water. They were dehydrated by 2 

changes in 35, 50, 70, and 90% acetone for 15 min 

each and 3 changes in 100% acetone. The material was 

critical point dried, then mounted on stubs and coated 

for SEM observation (JEOL 100). 

The immature stage was named the postacetabular 

juvenile, following the nomenclature used by Fredericksen 

(1980). Two stages could be distinguished according 

to the development of the ventral adhesive 

disc: a recently formed postacetabular juvenile, with 

little differentiation of alveoli and the buccal opening 



without oral lobules; and a preadult characterized by 

the presence of oral lobules and a distinct differentiation 

between alveoli, with the external morphology 

similar to that of the adult. 

Results 

Postacetabular juvenile 

In the recently formed postacetabular juvenile 

(Fig. 1), the oral disc characteristic of the adult 

phase is not observed. The mouth is a simple 

opening, without lobules (Fig. 2). 

The preadult shows a rough dorsal cone. Central 

and marginal alveoli, completely differentiated 

and varying in number, are observed on the 

ventral adhesive disc (Fig. 3). The posterior alveoli 

are less developed. Monociliate sensory 

papillae are found in the internal wall of the 

marginal alveoli (Fig. 4). The pores of the marginal 

bodies can be observed on the external 

border between the marginal alveoli. They are 

more developed in the anterior region of the 

ventral disc (Figs. 4, 5). The oral disc has 3 ventral 

and 2 dorsal lobules as in the adult, though 

they are not completely developed (Fig. 6). 

Monociliate sensory papillae and dome-shaped 

papillae are observed on the posterior surface of 

the oral disc. In this region, there is no regular 

distribution pattern of sensory structures (Fig. 

7). 

In the dorsal region of the neck, immediately 

behind the oral disc, there are pores and domeshaped 

papillae (Fig. 8). The monociliate papillae 

each have a cilium emerging from a bulbous 

surface. 

Adult 

The ventral adhesive disc has 16 marginal 

pairs and 32 central alveoli. The limit between 

both groups of central alveoli cannot be clearly 

observed (Fig. 9). The anterior region of the 

ventral adhesive disc shows a neat differentiation 

among the alveoli, with many sensory structures 

(Fig. 10). Completely differentiated pores 

of the marginal bodies are found between the 

marginal alveoli (Fig. 11). Monociliate papillae 

are located on the internal wall of each alveolus, 

arranged in 2 concentric circles. Two rows of 

dome-shaped papillae occur on the external edge 

of the marginal alveoli (Figs. 11-13). Monociliate 

and dome-shaped papillae (Fig. 14) are 

present on both sides of the transverse dividing 

line between the central alveoli. The excretory 

pore can be seen in the dorsal cone (Fig. 15). 


Two kinds of monociliate receptors (Fig. 16) and 

a single multiciliate structure (Fig. 17) were observed 

in the posterior dorsal region. 

Discussion 

The external morphology of the preadult of L. 

jungwirthi is very similar to that of the adult 

from C. facetum. Both juveniles and adults show 

a single excretory pore that ends at the channel 

formed by the union of the lateral ducts, as described 

by Lunaschi (1984). In agreement with 

the observations of Kritscher (1974) and Lunaschi 

(1984), the adult stage does not show a pore 

of Laurer's canal. 

Four types of sensory receptors were observed 

by SEM: 

MONOCILIATE PAPILLAE WITH A SHORT CILIUM: 

This type of receptor was irregularly distributed 

on the dorsal tegument, in the posterior surface 

of the oral lobules, and in the neck of juvenile 

L. jungwirthi. A denned pattern of distribution 

was observed only on the edges of the alveoli 

of the ventral adhesive disc. Rohde and Watson 

(1992) described this structure as a receptor 

formed by a cilium of intermediate length, being 

the most common type on the surface of L. manteri. 

Halton and Lyness (1971) described this 

type of papilla as the most frequent receptor on 

the body surface of A. conchicola. This receptor 

is more abundant in the oral lobules and in the 

central, marginal, and peripheral regions of the 

ventral adhesive disc of L. jungwirthi. This distribution 

agrees with that observed by Halton 

and Lyness (1971) in A. conchicola. The type of 

monociliate papilla found in L. jungwirthi may 

also correspond to that described by Fredericksen 

in the juvenile acetabulum of C. occidentalis 

and the simple uniciliate sensory structures observed 

in the cotylocidium larva of the same 

species (Fredericksen, 1978). Likewise, they are 

similar to type I sensilla found in adult C. occidentalis 

(Ip and Desser, 1984). Monociliate receptors 

were observed in Lobatostoma sp. (Rohde, 

1972), and the type I receptor has been observed 

in the tegument of posterior suckerlets of 

larval Multicotyle purvisi (Rohde and Watson, 

1990c). Monociliate tegumental receptors were 

also found in the buccal complex of Polylabroides 

australis (Murray, 1931) (Monogenea, Microcotylidae) 

(Rohde and Watson, 1995b) and in 

Udonella sp. (Platyhelminthes) (Rohde et al., 

1989). 

MONOCILIATE PAPILLAE WITH A LONG CILIUM:

PAOLA AND DAMBORENEA—TEGUMENT OF LOBATOSTOMA JUNGWIRTHI 251 

Figures 1-5. Immature stages of Lobatostoma jungwirthi. 1. Postacetabular juvenile showing a simple 

buccal cavity (without lobules) and the ventral adhesive disc with poorly differentiated alveoli. 2. Frontal 

view of the buccal cavity of the postacetabular juvenile. 3. Alveoli of the ventral adhesive disc, oral lobules, 

and a rough posterior cone (dorsal) (black arrow) in latero-ventral view of the preadult. 4. Marginal 

alveoli of the ventral adhesive disc of the preadult, with sensory receptors of the monociliate type (white 

arrow), dome-shaped receptor (arrowhead), and the pores of the marginal glands not yet formed between 

the posterior alveoli (black arrow). 5. Pore of a marginal gland (^marginal body) of the preadult. Scales: 

1, 2 = 10 |xm; 3 = 50 fim; 4 = 25 |xm; 5 = 10 |xm. 

These were found only on the surface of the 

dorsal cone of the adult. This type of receptor 

may correspond to the receptor with a long cilium 

in the larva of L. manteri (Rohde and Watson, 

1992). It is also similar to the receptor type 

A described by Ip et al. (1982) in adult C. occidentalis. 

NONCILIATE DOME-SHAPED RECEPTORS: These 

were found on the posterior surface of the oral 

lobules and in the neck of the juvenile. They 



Figures 6-10. Preadult and adult of Lobatostoma jungwirthi. 6. Frontal view of the preadult showing 

the 5 oral lobules. 7. Posterior surface of the oral lobules showing monociliate papillae (arrow) and domeshaped 

papillae arranged without defined pattern. 8. Neck region (ventral) with pores (white arrow) (in 

some cases with secretion) and dome-shaped papillae. 9. General ventral view of the adult. A clear differentiation 

of the longitudinal septum of the ventral adhesive disc cannot be observed. 10. Anterior end 

of the ventral adhesive disc showing dome-shaped and monociliate papillae (arrow) on the edge of the 

walls. Scales: 6 = 50 u,m; 7, 8 = 10 u,m; 9 = 25 u,m; 10 = 100 u,m. 

also were distributed on the borders between the 

alveoli of the adhesive disc of both juvenile and 

adult worms. Nonciliate sensory receptors were 

found by Rohde and Watson (1990b) in the external 

ventral tegument of the ventral suckerlets 

in M. purvisi. This structure may correspond to 

the nonciliate disc-shaped receptors or to the 


nonciliate type with a long root described by 

Rohde and Watson (1992) in the larva of L. 

manteri. Nonciliate tegumental receptors were 

also found in the juvenile of Astramphilina elongata 

Johnston, 1931 (Monogenea) (Rohde and 

Watson, 1990a). 

A SINGLE MULTICILIATE RECEPTOR: This Was

PAOLA AND DAMBORENEA—TEGUMENT OF LOBATOSTOMA JUNGWIRTHI 253 

Figures 11-14. Ventral adhesive disc of adult Lobatostoma jungwirthi. 11. General view of the marginal 

alveoli on the marginal region at midbody level of the ventral adhesive disc. 12. Walls of the marginal 

alveoli showing 2 circles of dome-shaped papillae (arrow) and marginal gland pore. 13. Detail of the 

internal wall with almost 1 complete circle of monociliate papillae (arrows). 14. Ventral view: transverse 

septum with monociliate (black arrow) and nonciliate papillae (white arrow). Scales: 11 = 50 u,m; 12 = 

25 u,m; 13, 14 = 10 jrni. 

observed in the posterior third of the dorsal region 

of the adult of L. jungwirthi. Paired multiciliate 

receptor complexes with 10 short cilia 

were described by Rohde and Watson (1990d) 

to be located dorsally to the mouth cavity of 

larval M. purvisi, but our record is the first of a 

surface multiciliate receptor in aspidogastreans. 

However, multiciliate receptors are found in other 

groups of parasitic Platyhelminthes, i.e., in the 

taste organ of the buccal complex of Pricea tnultae 

Chauhan, 1945 (Monogenea, Gastrocotylidae). 

Rohde and Watson (1996) found these receptors 

concentrated in small pits. Additionally, 

multiciliate pit-receptors were observed in the 

buccal complex of Polylabroides australis (Rohde 

and Watson, 1995b) and in specimens of an 

undescribed species of Proseriata (Monocelididae: 

Minonidae) (Rohde and Watson, 1995a). 

Pores of the marginal bodies were observed 

on the external border between the marginal alveoli 

of juveniles and adults of L. jungwirthi. 

Kritscher (1974) found similar structures in the 

adult of the species. These pores were described 

as part of a glandular system in the adhesive disc 

of aspidogastrids (Rohde, 1994). 

Unlike the preadult, the youngest juvenile 

does not have oral lobules. A few less-developed 

alveoli were observed on the ventral adhesive 

disc. Two sensory receptors were found in the 

preadult of L. jungwirthi'. a ciliate receptor with 

a cilium of intermediate length, and a nonciliate 

dome-shaped receptor. These correspond to 2 of 



Figures 15-17. Tegument of posterior dorsal region 

of Lobatostoma jungwirthi. 15. Excretory pore 

on the dorsal cone. 16. Type of monociliate papilla 

in the posterior cone region. 17. Multiciliate sensory 

receptor in the posterior dorsal region. Scales: 15, 

17 = 5 (Jim; 16 = 10 fjim. 

the 4 types of receptors described by Rohde and 

Watson (1992) in the larva of L. manteri. In addition, 

a third kind of receptor was observed that 

corresponds to the marginal body complex in the 

adult. However, the complex formed by the marginal 

bodies was observed as described for the 

adult, though the pores of these glands are not 

yet developed in the posterior region of the ventral 

adhesive disc in the preadult. 

The most common receptors in the preadult 


and in the adult were the intermediate length 

monociliate and the nonciliate dome-shaped papillae. 

The monociliate structures may correspond 

to some of the types I to V of monociliate 

dome-shaped papillae described for adult L. 

manteri (Rohde and Watson, 1989a). The nonciliate 

dome-shaped papillae may be homologous 

to those of type VI A and B found in the 

adult of L. manteri. Both types of receptors were 

abundant over the entire body, arranged in circles 

on the ventral adhesive disc. Longer monociliate 

papillae and multiciliate receptors were 

found exclusively in the adult. Probably more 

than one type of nonciliate and monociliate papillae 

may exist. It would be interesting to study 

these papillae further under transmission electron 

microscopy. 


Fredericksen, D. 1978. The fine structure and phylogenetic 

position of the cotylocidium larva of Cotylogaster 

occidentalis Nickerson, 1902 (Trematoda, 

Aspidogastrea). Journal of Parasitology 64: 

961-966. 

. 1980. Development of Cotylogaster occidentalis 

Nickerson, 1902 (Trematoda, Aspidogastridae) 

with observations on the growth of the ventral 

adhesive disc in Aspidogaster conchicola 

Baer, 1827. Journal of Parasitology 66:973-984. 

Halton, D. W., and R. A. W. Lyness. 1971. Ultrastructure 

of the tegument and associated structures 

of Aspidogaster conchicola (Trematoda: Aspidogastrea). 


Ip, H., and S. Desser. 1984. Transmission electron 

microscopy of the tegumentary sense organs of 

Cotylogaster occidentalis (Trematoda: Aspidogastrea). 


, , and I. Weller. 1982. Cotylogaster 

occidentalis (Trematoda, Aspidogastrea): scanning 

electron microscopic observations of sense organs 

and associated surface structures. Transactions of 

the American Microscopical Society 101:253- 

261. 

Kritscher, V. E. 1974. Lobatostoma jungwirthi nov. 

spec. (Aspidocotylea, Aspidogastridae) aus Geophagus 

brachiurus Cope 1894 (Pise., Cichlidae). 

Annalen des Naturhistorischen Museums in Wien 

78:381-384. 

Lunaschi, L. 1984. Helmintos parasites de peces de 

agua dulce de Argentina. II. Presencia de Lobatostoma 

jungwirthi Kritscher, 1974 (Trematoda: 

Aspidogastrea) en Cichlasoma facetum (Jenyns). 

Neotropica 20:187-193. 

Rohde, K. 1972. Sinnesrezeptoren von Lobatostoma 

n. sp. (Trematoda, Aspidogastrea). Die Naturwissenschaften 

59:168-169. 

. 1994. The Aspidogastrea, especially Multicotyle 

purvisi Dawes, 1941. Advances in Parasitology 

10:78-151. 

, and N. Watson. 1989a. Sense receptors in

PAOLA AND DAMBORENEA—TEGUMENT OF LOBATOSTOMA JUNGWIRTH1 255 

Lobatostoma manteri (Trematoda, Aspidogastrea). 

International Journal for Parasitology 19:847-858. 

, and . 1989b. Ultrastructure of the marginal 

glands of Lobatostoma manteri (Trematoda, 

Aspidogastrea). Zoologischer Anzeiger 223:301- 

310. 

, and . 1990a. Ultrastructural studies of 

juvenile Austramphilina elongata: transmission 

electron microscopy of sensory receptors. Parasitology 

Research 76:336-342. 

, and . 1990b. Non-ciliate sensory receptors 

of larval Multicotyle purvisi (Trematoda, 

Aspidogastrea). Parasitology Research 76:585- 

590. 

, and . 1990c. Uniciliate sensory receptors 

of larval Multicotyle purvisi (Trematoda, Aspidogastrea). 

Parasitology Research 76:591-596. 

, and . 1990d. Paired multiciliate receptor 

complexes in larval Multicotyle purvisi (Trematoda, 

Aspidogastrea). Parasitology Research 76: 

597-601. 

, and . 1992. Sense receptors of larval 

Lobatostoma manteri (Trematoda, Aspidogastrea). 


New Books Available 

-, and -. 1995a. Sensory receptors and 

epidermal structures of a meiofaunal turbellarian 

(Proseriata: Monocelididae: Minoninae). Australian 


, and . 1995b. Ultrastructure of the buccal 

complex of Polylabroides australis (Monogenea, 

Polyopisthocotylea, Microcotylidae). International 


, and . 1996. Ultrastructure of the buccal 

complex of Pricea multae (Monogenea, Polyopisthocotylea, 

Gastrocotylidae). Folia Parasitologica43:117-132. 

-, and F. Roubal. 1989. Ultrastructure 

of flame bulbs, sense receptors, tegument and 

sperm of Udonella (Platyhehninthes) and the phylogenetic 

position of the genus. Zoologischer Anzeiger 

222:143-157. 

Zylber, M. I., and M. Ostrowski de Nunez. 1999. 

Some aspects of the development of Lobatostoma 

jungwirthi Kritscher, 1974 (Aspidogastrea) in 

snails and cichlids fishes from Buenos Aires, Argentina. 

Memorias do Instituto Oswaldo Cruz 94: 

31-35. 

The Flagellates: Unity, Diversity and Evolution. Barry S. C. Leadbeater and J. C. Green, Editors. 2000. 

The Systematics Association Special Volume Series 59, Taylor and Francis Limited, 29 West 35th Street, 

New York, NY 10001-2299. i-xi, 401 pp. ISBN 0-7484-0914-9. 7" X 9%", hard cover. Cost: US$130.00 

or Canadian $195.00, per copy plus shipping and handling. Abstract: 35 Authors have contributed to the 

17 chapters of this examination of the blood parasite group. "[The] book sets out to examine flagellates 

from a multidisciplinary standpoint. Of primary concern are the unifying structures, mechanisms and 

processes involved in flagellate biology. [It begins] with a review of the complex history of flagellate 

studies from the first use of microscopes ... to the present. [This is] followed by a series of chapters on 

common aspects of flagellates, . . . [including] a discussion of the problems inherent in being a flagellate, 

and reviews of the structure and function of the flagellum itself, the cytoskeleton, surface structures and 

sensory mechanisms. The diversity of flagellates is recognized in the next series of chapters, which include 

reviews of trophic strategies of both free-living and parasitic groups, and contributions on ecology, biogeography 

and population genetics. The final chapters . . . are concerned with the occurrence and loss of 

organelles, and other aspects of flagellate evolution and phylogeny." 

Interrelationships of the Platyhelminths. D. T. J. Littlewood and R. A. Bray, Editors. 2001. The Systematics 

Association Special Volume Series 60, Taylor and Francis Limited, 29 West 35th Street, New 

York, NY 10001-2299. i-xii, 356 pp. ISBN 0-7484-0903-3. 8W X 11", hard cover. Cost: US$125.00 or 

Canadian $188.00, per copy plus shipping and handling. Abstract: This book's 27 chapters have been 

prepared by no fewer than 50 different expert contributors. It "has been split into four sections, rather 

dissimilar in length, that highlight the underlying goals [of bringing together workers from divergent areas 

to produce a work unified by modern approaches to phylogenetic analysis]. The first section takes a broader 

perspective on the status of the Platyhelminthes, its monophyly, placement in relation to other Metazoa 

and the nature of the basal taxa. The second section deals with the interrelationships of major free-living 

taxa, and the third on symbiotic and parasitic taxa. The final section encompasses contributions that view 

phylogeny and phylogenetic inference from the point of view of particular characters or techniques." 




Research Note 

Infectivity and Comparative Pathology of Echinostoma caproni, 

Echinostoma revolutum, and Echinostoma trivolvis (Trematoda) in the 

Domestic Chick 

SHANNON K. MULLIGAN,' JANE E. HUFFMAN,1-3 AND BERNARD FRIED2 

1 Department of Biological Sciences, Fish and Wildlife Microbiology Laboratory, East Stroudsburg University, 

East Stroudsburg, Pennsylvania 18301, U.S.A. (e-mail: jhuffman@po-box.esu.edu) and 

2 Department of Biology, Lafayette College, Easton, Pennsylvania 18042, U.S.A. 

ABSTRACT: We examined the clinical and pathological 

effects of 3 species of 37-collar-spined Echinostoma in 

domestic chicks. Three groups of 6 chicks each were 

infected with 50 metacercariae of either Echinostoma 

caproni, Echinostoma revolutum, or Echinostoma trivolvis. 

A group of 6 chicks was not infected and 

served as the uninfected controls. The chicks were 

necropsied on day 14 postinfection (PI). Infectivity and 

worm recovery rates for E. caproni were 100% and 

24%, respectively; for E. revolutum, they were 67% 

and 9%, respectively; and for E. trivolvis, they were 

83% and 15%, respectively. Echinostoma caproni was 

located in the middle third of the small intestine, 

whereas E. revolutum and E. trivolvis were located in 

the lower third, showing that niche selection of the 

different echinostomes varied. The echinostomes became 

ovigerous on days 10, 12, and 14 PI for E. caproni, 

E. trivolvis, and E. revolutum, respectively. Goblet 

cell proliferation in the host intestinal mucosa occurred 

in all infections. 

KEY WORDS: Echinostoma caproni, Echinostoma 

trivolvis, Echinostoma revolutum, Trematoda, domestic 

chicks, echinostomiasis, pathology, clinical effects, 

goblet cell, infectivity. 

Because echinostomiasis has produced significant 

mortality in ducks raised for commercial 

production in Europe and Asia (Kishore and 

Sinha, 1982), studies on experimental avian 

models to define the clinical and pathological 

features of the echinostomes are needed. Except 

for the experimental studies by Kim and Fried 

(1989) on gross and histopathological effects of 

Echinostoma caproni Richard, 1964, in an experimental 

avian model, such studies are lacking. 

In North America, avian hosts in the wild are 

often infected with Echinostoma trivolvis (Cort, 

1914) and Echinostoma revolutum (Froelich, 

1802) and species of Echinoparyphium (43- and 


256 


45-collar-spined echinostomes). Interestingly, 

the habitat of species of Echinoparyphium in the 

gut of birds is more anteriad than that of either 

E. trivolvis or E. revolutum. Echinostoma caproni 

also tends to localize more anteriad in the 

avian gut than either E. trivolvis or E. revolutum, 

and may serve as a useful model for Echinoparyphium 

infections. Therefore, information 

obtained from single infections of the 3 echinostome 

species examined in this study may be 

useful to wildlife studies of birds naturally infected 

with 3 or more species of echinostomes. 

The objectives of this study were to determine 

the following parameters in E. caproni', E. revolutum-, 

and E. trivolvis-infected birds: packed 

cell volume, hemoglobin concentration, and the 

relative splenic and hepatic weights of infected 

and noninfected domestic chicks. Parasite recovery 

and location were recorded from infected 

animals. We also examined tissues grossly and 

microscopically for evidence of pathological 

changes. Metacercarial cysts of E. caproni and 

E. trivolvis were obtained from the kidneys and 

pericardial sacs of laboratory-infected Biornphalaria 

glabrata (Say, 1816) snails (Huffman 

and Fried, 1990). Metacercarial cysts of E. revolutum 

were obtained from experimentally infected 

Lymnaea elodes (Say, 1821) snails (Sorenson 

et al., 1997). Twenty-four-d-old unfed 

domestic chicks were obtained from Reich Poultry 

Farm (Marietta, Pennsylvania, U.S.A.). All 

chicks were infected on day 1 prior to feeding. 

All animals were provided food (Country Egg 

Producer®, Agway Inc., Syracuse, New York, 

U.S.A.) and water ad libitum throughout the 

study. Group A (N = 6) was not infected and 

served as controls for the study. Chicks in 

Groups B-D each received 50 metacercarial 

cysts per os of either E. caproni (Group B, N = 

6), E. trivolvis (Group C, N = 6), or E. revolu-

RESEARCH NOTES 257 

Table 1. Mean worm recovery, mean percentage of recovery, percentage infected, and range of parasites 

recovered from chicks infected with Echinostoma caproni (Group B, TV = 6), Echinostoma trivolvis (Group 

C, N = 6), and Echinostoma revolutum (Group D, N = 6). Uninfected controls are represented by Group 

A (TV = 6). 

Group 

A 

B 

C 

D 

No. cysts 

administered 

0 

50 

50 

50 

Mean no. worms 

recovered (range) 

0(0) 

12 (8-20) 

8 (0-21) 

5 (0-16) 

turn (Group D, /V = 6). Fecal samples were collected 

from each infected animal and checked 

for echinostome eggs daily, starting on day 8 

postinfection (PI). Approximately 1 g of feces 

was emulsified in distilled water and examined 

by light microscopy. Animals were weighed every 

2 d to monitor weight gain in the chicks. 

Blood samples were collected from the jugular 

veins of all chicks on day 14 PI into tubes 

containing 0.13 M sodium citrate, refrigerated, 

and processed within 24 hr. Packed cell volume 

and hemoglobin concentration were measured 

and recorded. 

All chicks (Groups A-D) were necropsied on 

day 14 PI. The small intestines, ceca, and cloacas 

were opened, and the location and number 

of echinostomes in the infected chicks were recorded. 

Hepatic, splenic, and intestinal tissue 

samples from all groups were fixed in 10% neutral 

buffered formalin for 48 hr and then dehydrated 

in a series of graded alcohols, cleared in 

xylene, embedded in paraffin, and sectioned at 

6 (xm. At necropsy, the relative spleen and liver 

weights were determined. 

Infected and control tissues were stained in 

hematoxylin and eosin to evaluate histopathological 

effects. The occurrence of immunological 

cells from the infected and control tissues 

was also evaluated. 

Echinostome eggs were first seen in the fecal 

samples of all chicks from Group B on day 10, 

of 5 chicks from Group C on day 12, and of 4 

chicks from Group D on day 14 PI. All chicks 

exposed to E. caproni became infected. The 

number of worms recovered from their intestines 

ranged from 8 to 20, with a 24% recovery. The 

number of parasites recovered from Group C 

ranged from 0 to 21, a 15% recovery. Four of 6 

chicks (67%) exposed to E. revolutum cysts 

were infected, and the numbers of worms recov- 

Mean percentage 

of worms recovered 

0 

24 

15 

9 

Day of first 

appearance 

of eggs in feces 

0 

10 

12 

14 

Percentage of 

chicks infected 

0 

100 

83 

67 

ered ranged from 0 to 16, averaging 9%. These 

data are summarized in Table 1. 

The locations of the recovered worms from 

the infected chicks varied according to species. 

Echinostoma caproni was located in the midthird 

of the intestine, between the pylorus and 

the cloaca. Both E. trivolvis and E. revolutum 

were found toward the end of the intestine near 

the cloaca, but typically, E. revolutum was found 

more posteriad than E. trivolvis. No differences 

were noted between the number of parasites recovered 

per chick and the location of the worms. 

Echinostoma caproni tended to be in groups, but 

single worms were also found. Echinostoma revolutum 

and E. trivolvis were found singly and 

in groups. 

There was no significant weight loss (P > 

0.05) in chicks infected with any species of echinostome 

versus the uninfected (control) group. 

No differences were seen in liver or spleen 

weights of the infected chicks versus the control 

group, nor were there differences in liver or 

spleen weights between the infected groups. 

There were no notable differences in either the 

measured packed cell volume or hemoglobin 

concentration of the infected chicks compared 

with the uninfected chicks. 

Histologically, the liver and spleen tissues of 

Groups B-D showed no sign of immunological 

response or damage from the parasites. Damage 

to the intestinal villi of chicks infected with E. 

caproni was observed at the site of parasite attachment, 

villi were atrophic, and the circular 

musculature was hypertrophied with collagenlike 

fibers present. There was a proliferation of 

goblet cells. Hemorrhage occurred at the site of 

attachment. In chicks infected with E. trivolvis, 

damage to villi also occurred, with lymphocytic 

infiltration and goblet cell proliferation but with 

no hemorrhage noted. The response to E. revo- 



lutuin was less severe than with the other 2 parasites, 

but damage to the intestinal villi was observed. 

The presence of eggs in the host's feces is 

used for diagnosis of echinostomiasis. The time 

of deposition of eggs in the feces will vary 

among species (Huffman and Fried, 1990). In 

this study, E. caproni eggs were first noted in 

the chick's feces on day 10 PI, followed by E. 

trivolvis eggs on day 12 PI, and eggs of E. revolutum 

were found on day 14 PI. 

Infectivity in the chicks varied between the 

different Echinostoma species in this study. Factors 

such as age, size of cyst inoculum, pretreatment 

of metacercarial cysts, and host-gut emptying 

time influence the infectivity of E. trivolvis 

in experimentally infected chicks (Fried et al., 

1997). Huffman and Fried (1990) reported E. 

trivolvis to have infectivity varying between 50 

and 69%. Fried (1984) reported 100% infectivity 

when preselected cysts were used. In this study, 

the E. trivolvis metacercariae administered to the 

chicks averaged 83% infectivity. Experimental 

infection with E. caproni cysts in this study resulted 

in 100% infectivity, agreeing with a previous 

study on this species conducted by Fried 

et al. (1988), which reported 97% infectivity. 

Echinostoma revolutum infectivity in this study 

(67%) is congruent with the report of Humphries 

et al. (1997) on the species in experimentally 

infected chicks (64%). 

Huffman and Fried (1990) summarized findings 

of average worm recoveries for E. trivolvis 

in experimentally infected chicks. These results 

varied between 6 and 21%. In this study, a mean 

of 15% of the flukes were recovered from chicks 

infected with E. trivolvis. Echinostoma caproni 

infections resulted in 24% worm recovery, concurring 

with the report by Fried et al. (1988) of 

28% worm recovery. An average of 9% of the 

administered cysts were recovered as adult 

worms in the E. revolutum infections. This differed 

from the 32% worm recovery of E. revolutum 

reported by Humphries et al. (1997) and 

the 21 % worm recovery reported for the same 

species of Echinostoma in domestic chicks 

(Fried et al., 1997). 

Echinostoma trivolvis distribution along the 

intestine of the domestic chick varied in numerous 

past studies (Huffman and Fried, 1990). On 

day 14 PI of our study, E. trivolvis was found 

mainly in the lower intestine near the cloaca. 

Echinostoma revolutum also was seen in the 


posterior aspect of the intestine, congruent with 

results by Humphries et al. (1997) and Fried et 

al. (1997). Echinostoma caproni was found 

more anteriad than the other echinostomes in 

this study, mainly clustered in the midthird of 

the intestine. 

Weight gain, spleen and liver weights, packed 

cell volume, and hemoglobin concentrations of 

the infected chicks compared with the controls 

were not affected by the presence of any of the 

echinostomes. As noted in previous studies, liver 

or spleen tissue damage did not occur. Huffman, 

Iglesias, and Fried (1986) noted increased pathology 

in golden hamsters infected with echinostomes 

and increased pathology when greater 

numbers of parasites infected the host. Infectivity 

in the present study was less compared with 

other studies. In the study by Fried and Wilson 

(1981), high worm burdens caused a decrease in 

chick weight. Changes in blood parameters and 

tissue damage (Huffman, Michos, and Fried, 

1986) have been noted in rodent hosts infected 

with echinostomes. 

In conclusion, there are differences in the 

host—parasite relationships for each of the echinostomes 

used in this study. An understanding 

of these differences will contribute to better understanding 

the biosystematics of the 37-collarspined 

echinostome group. Some of these differences 

may help elucidate species distinctions 

when echinostomes are recovered from naturally 

infected hosts in both single and multiple infections. 


Fried, B. 1984. Infectivity, growth, and development 

of Echinostoma revolutum (Trematoda) in the domestic 

chick. Journal of Helminthology 58:241- 

244. 

, R. A. Donovick, and S. Emili. 1988. Infectivity, 

growth, and development of Echinostoma 

liei (Trematoda) in the domestic chick. International 


, T. J. Mueller, and B. A. Frazer. 1997. Observations 

of Echinostoma revolutum and Echinostoma 

trivolvis in single and concurrent infections 

in domestic chicks. International Journal for 


-, and B. D. Wilson. 1981. Decrease in body 

weight of domestic chicks infected with Echinostoma 

revolutum (Trematoda) or Zygocotyle lunata 

(Trematoda). Proceedings of the Helminthological 


Huffman, J. E., and B. Fried. 1990. Echinostoma and 

echinostomiasis. Advances in Parasitology 29: 

215-269.

, D. Iglesias, and B. Fried. 1986. Echinostoma 

revolutum pathology of intestinal infections of the 

golden hamster. International Journal for Parasitology 

18:873-874. 

-, C. Michos, and B. Fried. 1986. Clinical and 

pathological effects of Echinostoma trivolvis (Digenea: 

Echinostomatidae) in the golden hamster, 

Mesocricetus auratus. Parasitology 93:505=515. 

Humphries, J. E., A. Reddy, and B. Fried. 1997. 

Infectivity and growth of Echinostoma revolutum 

(Froelich, 1802) in the domestic chick. International 




Research Note 


Kim, J., and B. Fried. 1989. Pathological effects of 

Echinostoma caproni (Trematoda) in the domestic 

chick. Journal of Helminthology 63:227-230. 

Kishore, N., and D. P. Sinha. 1982. Observations of 

Echinostoma revolutum infection in the rectum of 

domestic ducks (Anas platyrhynchos domesticiis). 

Agricultural Science Digest 2:57-60. 

Sorenson, R. E., I. Kanev, B. Fried, and D. Minchella. 

1997. The occurrence and identification of 

Echinostoma revolutum from North American 

Lymnaea elodes snails. Journal of Parasitology 83: 

169-170. 

Excystation and Distribution of Metacercariae of Echinostoma 

caproni in ICR Mice 

BERNARD FRIED,' ADITYA REDDY, AND CAROLINE D. BALFOUR 

Department of Biology, Lafayette College, Easton, Pennsylvania 18042, U.S.A. (e-mail: friedb@lafayette.edu) 

ABSTRACT: In vivo excystation and distribution of 

newly excysted metacercariae of Echinostoma caproni 

Richard, 1964, were studied in 16 ICR mice, each fed 

400 metacercarial cysts and necropsied at various intervals 

from 1 to 24 hr postinfection (p.i.). Excysted 

metacercariae were recovered from the stomach and 

intestine (duodenum and jejunum) at 1 hr p.i. In vivo 

excystation in this echinostome occurred in the stomach 

and the anterior part of the small intestine. Encysted 

metacercariae were recovered from the stomach, 

small intestine, and cecum-large intestine at 1 and 

2 hr p.i. Recovery of encysted metacercariae was rare 

at 3 hr and nil at 4 and 24 hr. At 3, 4, and 24 hr, the 

encysted metacercariae had either excysted or were 

voided. Excysted metacercariae were widely scattered 

throughout the small intestine at all times, with about 

75% located in segments 1, 2, and 3 (duodenal-jejunum 

zone) of the small intestine at 3, 4, and 24 hr p.i. 

KEY WORDS: trematodes, in vivo excystation, Echinostoma 

caproni, metacercariae, ICR mice. 

Although information is available (Fried and 

Emili, 1988; Fried, 1994; Ursone and Fried, 

1995) on chemical excystation of metacercarial 

cysts of Echinostoma caproni Richard, 1964, 

there are no studies on in vivo excystation of 

this echinostome in mice. Metacercariae of most 

intestinal digeneans excyst in the vertebrate 


small intestine, but details on in vivo excystation 

and the microhabitat where excystation occurs 

are poorly understood in the Digenea. Simonsen 

et al. (1989) stated that E. caproni metacercarial 

cysts excysted in the duodenum of the mouse 

and the newly excysted metacercariae migrated 

to the posterior third of the small intestine. However, 

Simonsen et al. did not do experimental 

studies on in vivo excystation of the metacercarial 

cyst. 

The purpose of this research was to examine 

in vivo excystation of E. caproni metacercariae 

at various times up to 24 hr postinfection (p.i.) 

and to determine the distribution of newly excysted 

metacercariae in ICR mice. The ICR 

mouse is widely used as a laboratory host for 

this echinostome (see review in Fried and Huffman, 

1996). The only previous study that has 

reported distribution of preovigerous worms of 

E. caproni is that of Manger and Fried (1993), 

who showed that, by 2 days p.i., more than 90% 

of the juvenile worms were located in segment 

3 posterior to the pylorus (equivalent to the jejunum) 

in the ICR mouse. 

Metacercarial cysts of E. caproni were removed 

from the pericardial cavity and kidney of 

experimentally infected Biomphalaria glabrata 

Say, 1818, snails and fed (400 cysts/mouse) via 

stomach tube to 16 6-8-wk old, outbred, female 



Table 1. Percentage of encysted (EN) and excysted (EX) metacercariae (M) of Echinostoma caproni from 

16 mice, each fed 400 cysts. 

Time 

postinfection 

Group* (hr) 

A 

B 

C 

D 

E 

1 

2 

3 

4 

24 

M 

EN 

EX 

EN 

EX 

EN 

EX 

EN 

EX 

EN 

EX 

Stomach 

0.9 

1.0 

0.3 

0.8 

0 

0.5 

0 

0.2 

0 

0 

1 

1.5 

1.9 

0.5 

2.4 

0 

2.6 

0 

2.3 

0 

1.6 

Segments of the small intestine 

2 

1.3 

1.6 

0.5 

1.8 

0.1 

2.9 

0 

2.6 

0 

2.7 

3 

0.7 

2.5 

0.4 

1.3 

0 

3.1 

0 

2.9 

0 

3.3 

* Groups A and B each with 2 mice; Groups C, D, and E each with 4 mice. 

ICR mice (Hosier and Fried, 1991). Preliminary 

studies on 2 mice each fed 100 metacercarial 

cysts and necropsied at 1 and 2 hr p.i. showed 

metacercarial recoveries (combined data of excysted 

and encysted metacercariae) of about 

10% at necropsy. The preliminary work showed 

the inherent difficulties in recovering these small 

organisms (excysted metacercariae measuring 

about 250 (xm in length and encysted metacercariae 

about 150 (Jim in diameter) from the intestinal 

tract. Moreover, the color, size, and motility 

of the villi made it difficult to distinguish 

them from excysted metacercariae. Empty cysts 

were also seen at necropsy but were not counted. 

On the basis of our experiences with the preliminary 

study, we increased the cyst inoculum to 

400 per host in the study reported herein. 

Groups of 2 mice each were necropsied at 1 

and 2 hr p.i. (Groups A and B, respectively; Table 

1), and groups of 4 mice each were necropsied 

at 3, 4, and 24 hr p.i. (Groups C, D, and E, 

respectively; Table 1). The numbers of encysted 

and excysted metacercariae in the stomach, in 5 

intestinal segments of equal length (approximately 

10 cm each), beginning at the pylorus 

and ending at the ileocecal valve, and in the 

combined cecum—large intestine were counted. 

Empty cysts were seen but not counted in hosts 

necropsied at 1 and 2 hr p.i. The numbers were 

converted to percentages and the information is 

presented in Table 1. In Group A, the greatest 

percentage of encysted metacercariae was in 

segment 1 of the small intestine, and excysted 

metacercariae were recovered as far posteriad as 

segment 4 of the small intestine. Most of the 

4 

0.4 

0.5 

0.5 

1.7 

0.1 

2.0 

0 

1.7 

0 

2.2 


5 

0.4 

0 

0.2 

0.6 

0.1 

0.3 

0 

0.4 

0 

0.5 

Cecumlarge 

intestine 

0.2 

0 

0.1 

0 

0.1 

0 

0 

0 

0 

0 

Total 

5.4 

7.5 

2.5 

8.6 

0.4 

11.4 

0 

10.1 

0 

10.3 

excysted metacercariae recovered at 1 hr p.i. 

were located in segment 3. Some excysted metacercariae 

were in the stomach at 1 hr and were 

alive and active. These organisms either had excysted 

in the stomach or, possibly, could have 

excysted in the small intestine and migrated anteriad 

to the stomach. In Group A, the finding 

of most excysted metacercariae in segments 1, 

2, and 3 of the small intestine (duodenum-jejunum 

region) provides support for claims that in 

vivo excystation takes place in the anterior part 

of the small intestine. This finding supports the 

statement of Simonsen et al. (1989) referenced 

above. 

With time, the ratio of encysted to excysted 

metacercariae declined (see last column in Table 

1), and by 4 hr p.i., encysted metacercariae were 

not found. These findings suggest that by 4 hr 

p.i. most of the encysted metacercariae had excysted 

or were voided. The idea of metacercariae 

being voided by 4 hr p.i. is consistent with 

the fact that the usual transit time for ingested 

food in the mouse digestive tract is 4 hr (Barrachina 

et al., 1997). Fecal examinations to determine 

the possible presence of excysted or encysted 

metacercariae in the stool were not made. 

About 75% of the excysted metacercariae in 

Groups C, D, and E were located in segments 1, 

2, and 3. Hence, newly excysted juveniles, up to 

at least 24 hr p.i., are more dispersed in the gut 

than are older worms. Manger and Fried (1993) 

showed that by day 2 p.i. more than 90% juvenile 

E. caproni were localized in segment 3 (the 

jejunum), and by day 4 and beyond, worms 

tended to migrate even more posteriad, with

most being found in segment 4 (jejunum-ileum 

zone). 

In conclusion, this study provides information 

on in vivo excystation of E. caproni metacercariae 

during the first day after infection in ICR 

mice. The in vivo studies show that most metacercariae 

excyst in the duodenum and migrate to 

the jejunum to become juveniles. The distribution 

of excysted metacercariae is quite variable 

during the first 24 hr of infection. The low recovery 

rates of organisms (only about 10% 

when mice received 100 cysts, and from 10.1% 

to 12.9% when mice received 400 cysts) attest 

to the fact that these organisms are difficult to 

detect in the first 24 hr after excystation. Perhaps 

some of these missing larvae are in sites other 

than the intestinal lumen, e.g., ducts or crypts 

associated with the intestine or other unknown 

locations. Manger and Fried (1993) did not report 

recovery percentages of juvenile worms of 

E. caproni from ICR mice at 2 d p.i. They did 

report a wide range of worm recoveries (18 to 

95%) from 2 to 8 d p.i. The fact that their recoveries 

were higher than the 10—13% in this 

study would suggest that some of the newly excysted 

metacercariae had been overlooked or 

had reemerged from extra-luminal sites. 

Support for this work was provided in part by 

funds from the Kreider Professorship to Dr. Ber- 



Research Note 


nard Fried. We thank Ms. Vivienne R. Felix for 

typing the manuscript. 


Barrachina, M. D., V. Martinez, J. Y. Wei, and Y. 

Tache. 1997. Leptin-induced decrease in food intake 

is not associated with changes in gastric emptying 

in lean mice. American Journal of Physiology 

272:1007-1011. 

Fried, B. 1994. Metacercarial excystment of trematodes. 

Advances in Parasitology 33:91-144. 

, and S. Emili. 1988. Excystation in vitro of 

Echinostoma liei and E. revolutum (Trematoda) 

metacercariae. Journal of Parasitology 74:98-102. 

, and J. E. Huffman. 1996. The biology of the 

intestinal trematode Echinostoma caproni. Advances 

in Parasitology 38:311-368. 

Hosier, D. W., and B. Fried. 1991. Infectivity, 

growth, and distribution of Echinostoma caproni 

(Trematoda) in the ICR mouse. Journal of Parasitology 

77:640-642. 

Manger, P. M., Jr., and B. Fried. 1993. Infectivity, 

growth and distribution of preovigerous adults of 

Echinostoma caproni in ICR mice. Journal of Helminthology 

67:158-160. 

Simonsen, P. E., E. Bindseil, and M. K0ie. 1989. 

Echinostoma caproni in mice: studies on the attachment 

site of an intestinal trematode. International 


Ursone, R. L., and B. Fried. 1995. Light microscopic 

observations of Echinostoma caproni metacercariae 

during in vitro excystation. Journal of Helminthology 

69:253-257. 

Helminths Collected from Rattus spp. in Bac Ninh Province, Vietnam 

XUAN-DA PHAM,1'5 Cffl-LlEM TRAN,2 and HIDEO HASEGAWA3'4 

1 Department of Infectious Disease Control, Oita Medical University, Hasama, Oita 879-5593, Japan 

(e-mail: DA@oita-med.ac.jp), 

2 Ministry of Health, 138A Giang Vo, Ba Dinh, Hanoi, Vietnam, and 

3 Department of Biology, Oita Medical University, Hasama, Oita 879-5593, Japan 

(e-mail: hasegawa@oita-med.ac.jp) 

ABSTRACT: Helminthological examination was made 

on 35 rats (12 Rattus tanezumi, 14 Rattus argentiventer, 

and 9 Rattus losea) captured in 3 different habitats, 

i.e., residential, paddy field, and hilly areas, all in Bac 

Ninh Province, northern Vietnam. One trematode (Notocotylus 

sp.), 2 cestodes (Raillietina celebensis, Hymenolepis 

diminuta), 6 nematodes (Strongyloides ratti, 

Strongyloides venezuelensis, Nippostrongylus brasi- 

liensis, Orientostrongylus cf. tenorai, Syphacia muris, 

Gongylonema neoplasticum), and 1 acanthocephalan 

(Moniliformis monilifortnis) were collected. The species 

composition and prevalence of these helminths 

differed among the habitats, apparently because of biological 

characters of the parasites and environmental 

conditions of the localities. 

KEY WORDS: helminths, rat, Rattus tanezumi, Rattus 



argentiventer, Rattus losea, Trematoda, Notocotylus 

sp., Cestoda, Raillietina celebensis, Hymenolepis diminuta, 

Nematoda, Strongyloides ratti, Strongyloides venezuelensis, 

Nippostrongyhis brasiliensis, Orientostrongylus 

cf. tenorai, Syphacia muris, Gongylonema 

neoplasticum, Acanthoccphala, Moniliformis moniliformis, 

prevalence, ecology, Vietnam. 

There have been only limited reports on the 

parasites of rats from Vietnam (see Segal et al., 

1968). Most previous surveys were carried out 

before 1970, and no data are available to assess 

the parasitological condition of rats at the present 

time. In 1999, we had an opportunity to 

examine helminths collected from rats trapped 

near Hanoi, northern Vietnam. Ten helminth 

species, including some of taxonomic and ecological 

interest, were found as recorded herein. 

Rats were collected with live traps in 3 different 

habitats, i.e., residential areas, paddy 

fields, and low hilly areas, all in Bac Ninh Province, 

Vietnam, in December 1999. They were 

anesthetized with ether and killed. Their viscera 

were fixed in 10% formalin solution and transported 

to the Oita Medical University, Oita, Japan. 

Their heads were also fixed in 10% formalin 

for species identification on the basis of 

skull morphology. On examination, the lung, 

heart, and liver were minced in water with fine 

forceps under a stereomicroscope to detect helminths 

parasitic in these organs. Then, the alimentary 

canal was cut open and washed on a 

stainless steel sieve with aperture size of 0.1 

mm. The residues left on the sieve were transferred 

to a Petri dish and observed under a stereomicroscope 

to recover helminths. The stomach 

wall was observed under a stereomicroscope 

with transillumination to find nematodes dwelling 

in the wall. 

Helminths collected were cleared in a glycerol-alcohol 

solution by evaporating alcohol 

and mounted on glass slides with a 50% glycerol 

solution. Some trematodes were stained 

with alum carmine or Heidenhain's iron hematoxylin, 

dehydrated in an alcohol series 

with ascending concentration, cleared in xylene 

and creosote, and mounted with Canada 

balsam. Voucher parasite specimens and host 


5 Present address: Department of Parasitology, Hanoi 

Medical College, 1st Ton That Tung, Dong Da, 

Hanoi, Vietnam (e-mail: thuda2001@yahoo.com). 


skulls are deposited in the National Science 

Museum, Tokyo (NSMT), Japan, with the accession 

numbers NSMT-P1 5073-5076, 

NSMT-As 2944-2953, and NSMT-M 31601- 

31606. 

The 35 rats examined included 12 black rats 

Rattus tanezumi Temminck, 1844 ( = so-called 

Asian-type Rattus rattus Linnaeus, 1758; cf. 

Musser and Carleton (1993)), 14 ricefield rats 

Rattus argentiventer (Robinson and Kloss, 

1916), and 9 lesser ricefield rats Rattus losea 

(Swinhoe, 1871). Helminths were not detected 

from the lung and liver, although Angiostrongylus 

cantonensis (Chen, 1935) and Calodium 

hepaticum (Bancroft, 1893) (syn. Capillaria 

hepatica (Bancroft, 1893)) have been previously 

recorded from those organs of rats in 

Vietnam (cf. Segal et al., 1968). Meanwhile, 

10 helminth species, comprising 1 trematode, 

2 cestodes, 6 nematodes, and 1 acanthocephalan, 

were collected from the alimentary canal 

(Table 1). Most of these helminths are common 

rat parasites, being widely distributed in 

the surrounding countries (Myers and Kuntz, 

1964, 1969; Ow-Yang, 1971; Singh and 

Cheong, 1971; Wiroreno, 1978; Sinniah, 1979; 

Ow-Yang and Durette-Desset, 1983; Hasegawa, 

1990; Hasegawa et al., 1992, 1994; Hasegawa 

and Syafruddin, 1995). Orientostrongylus 

sp. was found only in 2 rats from the 

paddy fields. Because no males were found, 

species identification is withheld, although it 

is strongly suggested to be Orientostrongylus 

tenorai Durette-Desset, 1970, a common rat 

parasite widely distributed from Afghanistan 

to Taiwan (Durette-Desset, 1970; Ow-Yang 

and Durette-Desset, 1983; Ohbayashi and Kamiya, 

1980; Hasegawa, 1990; Hasegawa et al., 

1994; Hasegawa and Syafruddin, 1995). 

Among the parasites recovered, 2 cestodes, 

Hymenolepis diminuta (Rudolphi, 1819) and 

Raillietina celebensis (Janicki, 1902); 1 nematode, 

Gongylonema neoplasticum (Fibiger et 

Ditlevsen, 1914); and 1 acanthocephalan, Moniliformis 

monilifonnis (Bremser, 1811) have 

been recorded previously from Vietnamese 

rats (Segal et al., 1968). Notocotylus sp. is of 

special interest because trematodes of this genus 

have been reported only rarely from rats 

of the subfamily Murinae, although some 

members have often been recorded from voles 

of the subfamily Arvicolinae (cf. Yamaguti,


Table 1. Helminthic infections among rats collected from 3 different habitats in Bac Ninh Province, 

Vietnam. 

Rattus species: 

No. rats examined: 

Head and trunk length (cm) range: 

(Mean): 

Trematoda 

Notocotylus sp. 

Cestoda 

Raillietina celebensis 

Hymenolepis diminuta 

Nematoda 

Strongyloides spp.f 

Nippostrongylus brasiliensis 

Orientostrongylus cf. tenorai 

Syphacia murls 

Gongylonema neoplasticum 

Acanthocephala 

Moniliformis monilifonnis 

Habitat: Residential 

R. tanezumi 

10 

14-19 

(16.7) 

— 

2 (20) 

— 

— 

1 (10) 

— 

6 (60) 

4 (40) 

1 (10) 

* No. rats infected (prevalence in parentheses). 

t Strongyloides ratti and/or S. venezuelensis. 

1971). The morphological characteristics of 

Notocotylus sp. are summarized below. 

DESCRIPTION: Notocotylus sp. Trematoda: 

Notocotylidae. Body foliate, attenuated anteriorly, 

1.39-2.64 mm long by 0.57-0.91 mm 

wide; 3 longitudinal rows of prominent ventral 

glands present on ventral surface, each row 

composed of 12 to 14 glands; oral sucker terminal; 

esophagus short; ceca diverticulated, 

terminating just posterior to ovary; testes 

lobed, located lateral to terminal portion of 

ceca; genital pore immediately posterior to 

oral sucker; ovary intertesticular, lobed; vitellaria 

in lateral fields, extending from middle 

of body to anterior margins of testes; metraterm 

and cirrus sac almost equal in length; egg 

ellipsoidal, 22-24 by 11-12 |xm, with polar 

filaments, 2 to 3 times longer than egg length. 

HOSTS: Rattus argentiventer (Robinson et 

Kloss, 1916) and Rattus losea (Swinhoe, 1871). 

SITE IN HOSTS: Intestine. 

LOCALITY: Paddy field and low hilly area in 

Bac Ninh Province (21°7'N; 105°59'E), Vietnam. 

SPECIMENS DEPOSITED: National Science Museum, 

Tokyo, NSMT-P1 5073, 5074. 

REMARKS: In the neighboring areas of Vietnam, 

only 2 Notocotylus species have been 

recorded from mammals: Notocotylus mamii 

Hsu, 1954, of which adults were experimen- 

R. argentiventer 

1 

12-20 

(15.4) 

1 (14%)* 

— 

1 (14) 

2 (29) 

6 (86) 

1 (14) 

4 (57) 

— 

— 

Paddy field Low hilly area 

R. losea 

1 

10-18 

(14.2) 

2 (29) 

— 

— 

3 (43) 

3 (43) 

1 (14) 

3 (43) 

— 

R. tanezumi 

2 

17-17.5 

(17.3) 

— 

— 

— 

— 

2 (100) 

— 

2 (100) 

— 

R. argentiventer 

1 

16-21 

(17.5) 

3 (43) 

— 

— 

1 (14) 

7 (100) 

— 

2 (29) 

— 

• 

R. losea 

2 

13-14 

(13.5) 

1 (50) 

— 

— 

— 

2 (100) 

— 

2 (100) 

— 

tally raised in rabbits, and Notocotylus ratti 

Yie, Qiu, Weng, Li, and Li, 1956, the only 

representative exclusively known from Rattus, 

both from southern China (Hsu, 1954; Yie et 

al., 1956). Notocotylus mamii resembles the 

present form in the position of the genital pore 

but is readily distinguished in that the ceca are 

simple and the eggs have extremely elongated 

filaments, often more than 10 times longer 

than the egg length (Hsu, 1954, 1957). Although 

N. ratti resembles the present species 

in having diverticulated ceca, it is clearly distinguished 

by possessing a genital pore located 

posterior to the cecal bifurcation and only 5 to 

6 ventral glands in the median row (Yie et al., 

1956). Presumably, the present worms represent 

a new species. However, proposal of a 

new taxon is withheld because the present 

worms were contracted by unsuitable fixation, 

obscuring some of the key structures. 

The species composition and prevalence of 

the rat helminths differed greatly among the 

habitats surveyed. Only Nippostrongylus brasiliensis 

(Travassos, 1914) and Syphacia muris 

(Yamaguti, 1935) were common to all 3 habitats. 

Raillietina celebensis, G. neoplasticum, 

and M. moniliformis were collected only from 

the rats captured around houses. Strongyloidid 

and trematode infections were not found in the 

rats captured around houses. Notocotylus sp. 


—


was recovered from the rats collected in the 

paddy and hilly areas but not from those in the 

residential area. 

The differences in species composition and 

prevalence among the habitats could be explained 

by the biological characters of each 

helminth and by the environmental conditions. 

Because members of Notocotylus require a 

freshwater snail as the intermediate host (cf. 

Yamaguti, 1975), their presence in the paddy 

fields is reasonable. The hilly area surveyed in 

the present study contained many small ponds 

that might provide habitats for the snails. Also 

not unexpected is that G. neoplasticum and M. 

moniliformis were found only in the rats captured 

around the houses, because their intermediate 

hosts, usually cockroaches, are abundant 

in the residential areas. Nippostrongylus 

brasiliensis and Strongyloides spp. require 

moist soil for their embryonic and larval development 

and transmission. Their low prevalence 

or absence in the residential areas 

seems to be due to the dried soil around the 

houses in that season. Syphacia muris, the 

only helminth with relatively stable prevalence 

among the habitats, has a quite simple 

life cycle passed nearly entirely within the 

body of the host, and hence, its prevalence is 

less affected by the external environment. 

We are deeply indebted to Dr. G. G. Musser, 

American Museum of Natural History, for his 

kindness in identifying the rat species and to 

Dr. S. Kamegai, Meguro Parasitological Museum, 

for his kind consultation on Notocotylus 

sp. 


Durette-Desset, M. C. 1970. Caracteres primitifs de 

certains Nematodes Heligmosomes parasites de 

Murides et de Cricetides orientaux. Definition 



Research Note 


Nematodes of the Tribe Cyathostominea (Strongylidae) Collected 

from Horses in Scotland 

J. RALPH LlCHTENFELS,1'5 AlOBHINN MCDONNELL,2 SANDY LOVE,3 AND 

JACQUELINE B. MATTHEWS4 

1 Biosystematics Unit, Parasite Biology, Epidemiology, and Systematics Laboratory, The Henry A. Wallace 

Beltsville Agricultural Research Center, Agricultural Research Service, U.S. Department of Agriculture, 

Beltsville, Maryland 20705-2350, U.S.A. (rlichten@anri.barc.usda.gov), 

2 Department of Veterinary Parasitology, University of Glasgow Veterinary School, Bearsden Road, 

Glasgow G61 1QH, United Kingdom, 

3 Weipers Centre for Equine Welfare, University of Glasgow Veterinary School, Bearsden Road, 

Glasgow G61 1QH, United Kingdom (S.Love@vet.gla.ac.uk), and 

4 Division of Equine Studies, Department of Veterinary Clinical Science and Animal Husbandry, Faculty of 

Veterinary Science, University of Liverpool, Leahurst CH64 7TE, United Kingdom (J.B.Matthews@liv.ac.uk) 

ABSTRACT: Nematodes of the tribe Cyathostominea 

are important parasites of horses. They live in large 

numbers in the large intestine and include over 50 species 

worldwide. This report describes an enumeration 

study of species found in a small population of horses 

in western Scotland. As found previously in a wide 

range of geographic regions, the 7 most abundant species 

of Cyathostominea, of the 18 recorded in this 

study, accounted for over 94% of the total population. 

One major exception to the results of previous studies 

was the presence of the most common species in this 

population, Cylicocyclus ashworthi. This species has 

not been recorded in the U.K. since its original description 

in 1924 and is morphologically very similar 

to another member of the same genus, Cylicocyclus 

nassatus, from which it has not been distinguished in 

previous studies in this geographical region. A rare 

species, Tridentoinfundibulum gobi, was found in low 

numbers in 3 of 4 horses. 

KEY WORDS: Nematoda, Cyathostominea, species 

survey, prevalence, intensity, horses, morphological 

identification, Scotland. 

Nematodes of the tribe Cyathostominea are 

the most common helminth parasites of the 

horse and are ubiquitous in all breeds. Members 

of the tribe Cyathostominea (Strongylidae) have 

been commonly referred to as small strongyles, 

cyathostomins (for the tribe), or cyathostomes 

(for the genus Cyathostomum Molin, 1861) 

(Hartwich, 1986). However, in order to avoid 

possible confusion with members of the nema- 


tode genus Cyathostoma Blanchard, 1849 (Syngamidae), 

which are sometimes referred to as 

cyathostomes, we will use the common name 

cyathostomins to refer to the 51 species included 

in the tribe Cyathostominea as listed by Lichtenfels 

et al. (1998). Infections with these nematodes 

are complex: 51 species of cyathostomins 

have been recorded in horses, donkeys, and zebras 

worldwide (Lichtenfels et al., 1998), but 10 

of these species have been reported only from 

zebras or donkeys, and a few other species are 

known to have very limited distributions. However, 

most horses carry a burden of 5 to 10 common 

species, including many thousands (sometimes 

more than 100,000) of lumen-dwelling 

adult nematodes and as many larval stages in the 

walls of the large intestine (Reinemeyer et al., 

1984; Bucknell et al., 1995). Clinically, cyathostomins 

are associated with various syndromes, 

the most dramatic of which is larval cyathostominosis, 

a fatal enteritis that occurs secondary to 

synchronized reactivation of arrested larvae 

from the intestinal mucosa (Giles et al., 1985; 

van Loon et al., 1995). The major obstacles to 

understanding, and therefore controlling, these 

parasites are their complexity, our inability to 

identify eggs in the feces, and the difficulty in 

identifying larvae on pasture. Until recently, the 

parasitic stages of cyathostomins could be identified 

only by adult worm morphology. However, 

recent studies have examined the molecular relationship 

of these species with a view to developing 

molecular probes for use in identifica- 



tion of both preparasitic and parasitic stages. For 

such studies, it is critical that the cyathostomins 

be identified and classified as consistently as 

possible. Modern identification manuals exist 

(Lichtenfels, 1975; Hartwich, 1986; Dvojnos 

and Kharchenko, 1994), but problems in identifying 

several species persist (Lichtenfels et al., 

1997). In 1997, workers convened an international 

workshop to clarify the systematics of the 

Cyathostominea (Sun City, Republic of South 

Africa), and an agreement was reached on a consensus 

recommendation for the taxonomy of 51 

species as detailed in Lichtenfels et al. (1998). 

Despite the importance of these parasites, information 

is still lacking on species prevalence 

in Britain, especially since the development of 

widespread anthelmintic resistance. The last detailed 

study of species prevalence and infection 

intensity in the U.K. was in 1976 (Ogbourne, 

1976). The present report describes the species 

of cyathostomins present in the large intestine of 

a population of ponies from western Scotland. 

The nematodes were collected to provide DNA 

sequence information for the development of diagnostic 

tools and for phylogenetic analysis of 

the nematodes. 

Adult parasites were collected from intestinal 

contents of 4 Welsh-Shetland cross ponies aged 

from 9 to 15 mo originating from a local horse 

population. The history of anthelmintic treatment 

of the ponies is unknown. These animals 

were euthanized at the University of Glasgow 

Veterinary School for reasons other than parasite 

infestation. Intestinal contents were coarse-filtered 

with household plastic sieves. After sieving, 

the contents were passed through a Baermann 

apparatus with milk filters and then 

through SS-jjim wire-mesh sieves. Individual 

adult parasites were washed in sterile phosphatebuffered 

saline (137 mM NaCl, 8.1 mM 

Na2HPO4, 2.7 mM KC1, 1.47 mM KH2PO4, pH 

7.2). Where possible, a total of 200 parasites 

were collected from the ventral and dorsal colon; 

however, the cecum often contained fewer 

than 50 adult parasites. With the aid of a dissection 

microscope, the heads were excised with a 

scalpel because the bodies were subsequently 

used for DNA extraction. The heads were stored 

in 200 (JL! of 5% buffered formalin, then mounted 

on glass slides in a few drops of phenolalcohol 

(80% melted phenol crystals and 20% 

absolute ethanol) to which glycerine had been 

added at about 5% of the volume, and studied 


with an Olympus BX50® differential interference 

contrast microscope. The parasites were 

identified according to the key of Lichtenfels 

(1975), supplemented by more recent descriptions 

of certain species (Lichtenfels and Klei, 

1988; Kharchenko et al., 1997; Lichtenfels et al., 

1997, 1999). The taxonomy used in this report 

follows the checklist of genera and species recommended 

by the 1997 international workshop 

(Lichtenfels et al., 1998). Representative heads 

of 14 species of cyathostomins and Craterostomum 

acuticaudatum and photomicrographs of 2 

species of cyathostomins have been deposited in 

the U.S. National Parasite Collection, U.S. Department 

of Agriculture, Beltsville, Maryland 

20705-2350 as accession numbers 90698- 

90714. Two species of cyathostomins, Cylicocyclus 

elongatus and Cylicocyclus radiatus, and 

Gyalocephalus capitatus could not be documented 

by either method. 

Eighteen cyathostomin species, representing 5 

genera, were identified. Table 1 shows the total 

numbers of parasite species per animal identified 

morphologically and the relative numbers of 

each species collected from each animal. Eight 

species occurred in all 4 ponies. One rare species, 

Tridentoinfundibulum gobi, was found in 

•Scotland for the first time. It had been reported 

previously only in Asia and North America 

(Lichtenfels et al., 1998). In addition, individuals 

of the genera Craterostomum and Gyalocephalus 

were isolated but in smaller numbers 

than most of the cyathostomin species. The 7 

most abundant cyathostomin species were, in 

descending order, Cylicocyclus ashworthi, Cyathostomum 

catinatum, Cylicostephanus longibursatus, 

Cyclostephanus minutus, Cylicocyclus 

nassatus, Cylicocyclus insigne, and Cylicostephanus 

goldi. These species comprised over 

94% of the total cyathostomin burden. These results 

are similar to recent enumeration studies 

performed in several geographically distinct regions, 

for example in the U.S.A., Europe, and 

Australia (Reinemeyer et al., 1984; Mfitilodze 

and Hutchinson, 1985; Bucknell et al., 1995; 

Gawor, 1995). In addition, in terms of local studies 

performed previously in the U.K., the species 

identified here were very similar to those reported 

by Mathieson in Scotland (1964); Ogbourne 

in southwest England (1976), and Love 

and Duncan in Scotland (1992). Ogbourne 

(1976) performed the most extensive study and 

identified 21 species in 86 horses of various ages


Table 1. Numbers of specimens of nematodes, by species, collected from 4 ponies from western Scotland. 

Parasite species 

Cylicocyclus ashworthi (Le Roux, 1924) Mclntosh, 1933 

Cylicocycliis nassatus (Looss, 1900) Chaves, 1930 

Cylicocyclus insigne (Boulenger, 1917) Chaves, 1930 

Cylicocyclus ultrajectinus (Ihle, 1920) Ershov, 1939 

Cylicocyclus leptostomum (Kotlan, 1920) Chaves, 1930 

Cylicocyclus radiatus (Looss, 1900) Chaves, 1930 

Cylicocyclus elongatus (Looss, 1 900) Chaves 1 930 

Cyathostomum car ina turn Looss, 1900 

Cyathostomum pateratum (Yorke and Macfie, 1919) K'ung, 1964 

Coronocyclus coronatus (Looss, 1900) Hartwich, 1986 

Coronocyclus labiatus (Looss, 1900) Hartwich, 1986 

Cylicostephanus calicatus (Looss, 1900) Ihle, 1922 

Cylicostephanus longibitrsatiis (Yorke and Macfie, 1918) Cram, 1924 

Cylicostephanus minutiis (Yorke and Macfie, 1918) Cram, 1924 

Cylicostephanus goldi (Boulenger, 1917) Lichtenfels, 1975 

Cylicostephanus bidentatus (Ihle, 1925) Lichtenfels, 1975 

Cylidodontophoms bicoronatus (Looss, 1900) Ihle, 1922 

Tridentoinfundibulum gobi Tshoijo, in Popova, 1958 

Craterostomum acuticaudatum (Kotlan, 1919) Ihle, 1920 

Gyalocephalus capitatus Looss, 1900 

and breeds. In the latter study, 80% of these 

horses had C. longibursatus, C. goldi, C. calicatus, 

C. catinatum, C. coronatus, and C. nassatus. 

The most notable exception between the 

current study and all of these previous studies is 

the presence of C. ashworthi, the most prevalent 

species identified in our population. Cylicocyclus 

ashworthi was last reported in the U.K. as 

a new species (Le Roux, 1924) and has not been 

reported there since. Of note is that, in a comparable 

study performed several years earlier on 

worm populations derived from the same pastures 

as those used here, Love and Duncan 

(1992) identified 6 species, and C. nassatus was 

one of the most numerous. Cylicocyclus ashworthi 

and C. nassatus are morphologically very 

similar, and it is highly likely that these and other 

workers misidentified C. ashworthi as C. nassatus 

prior to the recent redescriptions of these 

species (Lichtenfels et al., 1997). Cylicocyclus 

nassatus is characterized by a cuticular shelf on 

the inner surface of the buccal capsule, a dorsal 

gutter that is as long as 50% of the buccal capsule 

depth, and 20 elements in the external leaf 

crown. Cylicocyclus ashworthi can be distinguished 

from C. nassatus by the absence of the 

shelf from the inner surface of the buccal capsule, 

by its much shorter dorsal gutter, and by 

25-29 external leaf crown elements that differ 

in shape from those of C. nassatus (Lichtenfels 

1 

94 

10 

24 

5 

0 

0 

0 

20 

9 

1 

0 

1 

24 

32 

26 

2 

0 

1 

4 

2 

2 

77 

51 

6 

1 

1 

0 

0 

51 

0 

2 

0 

2 

1 1 

55 

13 

4 

2 

1 

3 

0 

Pony 

3 

81 

47 

43 

1 

3 

1 

1 

80 

6 

0 

0 

3 

85 

6 

19 

9 

0 

3 

0 

0 

4 

83 

15 

14 

0 

0 

0 

0 

73 

0 

0 

1 

0 

23 

49 

4 

1 

0 

0 

1 

0 

et al., 1997). The ability to clearly observe the 

cuticular shelf in the buccal capsule is dependent 

on the clearing agent used, and this may have 

contributed to the difficulty in identifying this 

unique feature in previous studies. 

In addition to the historical difficulty in separating 

C. nassatus and C. ashworthi, C. ashworthi 

has also been misidentified as C. triramosus, 

which has also been confused with C. 

nassatus prior to its recent redescription (Kharchenko 

et al., 1997). We now know that C. triramosus 

is exclusively a parasite of zebras. It is 

imperative that C. nassatus and C. ashworthi be 

correctly differentiated because they are 2 of the 

most common nematodes found in the ventral 

colon of horses, and if DNA probes are to be 

developed on the basis of morphological delineation, 

then consistent identification is a prerequisite. 

Interestingly, Hung et al. (1997) performed 

sequencing of the first (ITS-1) and second 

(ITS-2) internal transcribed spacers of 5.8S 

ribosomal DNA of these species and found that 

C. nassatus and C. ashworthi, differentiated by 

head morphology, were sufficiently different at 

the DNA level to assign them to separate species. 

These results are similar to work performed 

on the intergenic spacer region of the nuclear 

DNA, where over 50% DNA sequence difference 

was found between these 2 species (Kaye 

et al., 1998), with low intraspecific variation 


268 COMPARATIVE PARASITOLOGY, 68(2), JULY 2001 

(0.3% for C. ashworthi and 1.9% for C. nassatus). 

Subsequently, oligoprobes designed from 

these IGS sequences have been used successfully 

to distinguish DNA of individual C. nassatus 

and C. ashworthi (Hodgkinson et al., 2001). Furthermore, 

pairwise evolutionary distances, calculated 

under maximum likelihood with a GTR 

model estimated from data from the mitochondrial 

large ribosomal RNA subunit and ITS-2 

DNA, showed a 9.5% difference between the 2 

species (McDonnell et al., 2000). 

A consistent feature of all of the species incidence 

studies, including the present work, is 

that a small number of species, usually 5 to 10, 

constitute more than 80% of the infective load. 

Furthermore, the proportions of species have remained 

remarkably stable over many years, despite 

the widespread use of anthelmintics and 

the development of resistance. The rarer species 

are found at less consistent levels, but this is 

expected because the small populations may 

have gone undetected in cases where small subsamples 

of the worm populations have been examined 

for practical reasons. Consequently, 

much of the data on the least commonly discovered 

species are certain to underestimate true 

prevalence. Chapman et al. (1999) reported that 

9 to 15 species were found in a single animal 

when 200 specimens were identified, but the 

number increased to 20 to 29 when all nematodes 

in an entire 5% aliquot were identified. 

In the current study, Strongylus species were 

not found. This is probably indicative of the efficacy 

of anthelmintics and their strategic use in 

parasite control programs. In older studies, 100% 

prevalence of Strongylus species was reported 

(Le Roux, 1924; Foster and Ortiz, 1937). More 

recently, Bucknell et al. (1995) reported a prevalence 

of 38% Strongylus species in a study in 

which the deworming history of the horses was 

not known. Here, it was observed that, whereas 

relatively few species occurred exclusively in one 

or other parts of the intestines, most followed distinct 

site distributions (not shown), strongly biased 

in favor of a particular region, and (with the 

exception of C. ashworthi) these distributions 

were as described by Ogbourne (1976). Here, as 

in Ogbourne's study, the majority of C. pateraturn, 

C. insigne, and C. longibursatus individuals 

were found in the dorsal colon, whereas most of 

the C. nassatus, C. ultrajectinus, C. catinatum, 

and C. goldi adults were found in the ventral colon. 

Some caution must be taken in the interpre- 


tation of these data because some of the species 

were found only in very low numbers. Also consistent 

with the findings of Ogbourne (1976) was 

that the cecum was the most sparsely populated 

region of the large intestine. 

This work was supported by a project grant 

funded by the Home of Rest for Horses near 

Lacey Green, Princes Risborough, Buckinghamshire, 

U.K. We thank James McGoldrick 

and colleagues at the Department of Veterinary 

Parasitology, University of Glasgow, for assistance 

in preparing the nematode heads. We 

thank Patrick Shone of Olympus for providing 

a microscope. 


Bucknell, D. G., R. B. Gasser, and I. Beveridge. 

1995. The prevalence and epidemiology of gastrointestinal 

parasites of horses in Victoria, Australia. 


Chapman, M. R., D. D. French, and T. R. Klei. 

1999. Intestinal helminths of ponies; a comparison 

of species prevalent in Louisiana pre- and postivermectin. 

P. 74 in Proceedings of the American 

Association of Veterinary Parasitologists 44th Annual 

Meeting, New Orleans, Louisiana, July 10- 

13, 1999. (Abstract.) 

Dvojnos, G. M., and V. A. Kharchenko. 1994. Strongylidae 

in domestic and wild horses. Publishing 

House Naukova Dumka, Kiev, Ukraine. 234 pp. 

[In Russian.] 

Foster, A. O., and P. Ortiz O. 1937. A further report 

on the parasites of a selected group of equines in 

Panama. Journal of Parasitology 23:360-364. 

Gawor, J. J. 1995. The prevalence and abundance of 

internal parasites in working horses autopsied in 

Poland. Veterinary Parasitology 58:99-108. 

Giles, C. J., K. A. Urquhart, and J. A. Longstaffe. 

1985. Larval cyathostomiasis (immature trichonema-induced 

enteropathy): a report of 15 clinical 

cases. Equine Veterinary Journal 17:196-201. 

Hartwich, G. 1986. On the Strongylus tetracanthus 

problem and the systematics of the Cyathostominae 

(Nematoda, Strongyloidea). Mitteilungen aus 

dem Zoologischen Museum in Berlin 62:61-102. 

Hodgkinson, J. E., S. Love, J. R. Lichtenfels, S. Palfreman, 

Y. H. Ramsey, and J. B. Matthews. 

2001. Evaluation of the specificity of five oligoprobes 

for identification of Cyathostomin species 

from horses. International Journal for Parasitology 

31:197-204. 

Hung, G. C., N. B. Chilton, I. Beveridge, A. Mc- 

Donnell, J. R. Lichtenfels, and R. B. Gasser. 

1997. Molecular delineation of Cylicocyclus nassatus 

and C. ashworthi (Nematoda: Strongylidae). 


Kaye, J. N., S. Love, J. R. Lichtenfels, and J. B. 

McKeand. 1998. Comparative sequence analysis 

of the intergenic spacer region of cyathostome 

species. International Journal for Parasitology 28: 

831-836.

Kharchenko, V. A., G. M. Dvojnos, and J. R. Lichtenfels. 

1997. A redescription of Cylicocyclus triramosus 

(Nematoda: Strongyloidea): a parasite of 

the zebra, Equus burchelli antiquorum. Journal of 


Le Roux, P. L. 1924. Helminths collected from 

equines in Edinburgh and in London. Journal of 

Helminthology 2:111-134. 

Lichtenfels, J. R. 1975. Helminths of domestic equids. 

Illustrated keys to the genera and species with emphasis 

on North American forms. Proceedings of 

the Helminthological Society of Washington 

42(special issue): 1-92. 

, V. A. Kharchenko, R. C. Krecek, and L. 

M. Gibbons. 1998. An annotated checklist by genus 

and species of 93 species level names for 51 

recognized species of small strongyles (Nematoda: 

Strongyloidea: Cyathostominae) of horses, asses 

and zebras of the world. Veterinary Parasitology 

79:65-79. 

-, C. Sommer, and M. Ito. 1997. Key 

characters for the microscopical identification of 

Cylicocyclus nassatus and Cylicocyclus ashworthi 

(Nematoda: Cyathostominea) of the horse, Equus 

caballus. Journal of the Helminthological Society 


, and T. R. Klei. 1988. Cylicostephanus torbertae 

sp. n. (Nematoda: Strongyloidea) from Equus 

caballus with a discussion of the genera Cylicostephanus, 

Petrovinema and Skrjabinodentus. 

Proceedings of the Helminthological Society of 

Washington 55:165-170. 

, P. A. Pilitt, V. A. Kharchenko, and G. M. 

Dvojnos. 1999. Differentiation of Coronocychis 



Research Note 


sagittatus and Coronocychis coronatus (Nematoda: 

Cyathostominea) of horses. Journal of the Helminthological 


Love, S., and J. L. Duncan. 1992. Development of 

cyathostome infection of helminth naive foals. 

Equine Veterinary Journal Supplement 13:93-98. 

Mathieson, A. O. 1964. A study into the distribution 

of, and host tissue responses associated with, 

some internal parasites of the horse. Thesis, University 

of Edinburgh, Edinburgh, U.K. 160 pp. 

McDonnell, A., S. Love, A. Tait, J. R. Lichtenfels, 

and J. B. Matthews. 2000. Phylogenetic analysis 

of partial mitochondrial cytochrome oxidase C 

subunit I and large ribosomal RNA sequences and 

nuclear internal transcribed spacer I sequences 

from species of Cyathostominae and Strongylinae 

(Nematoda, Order Strongylida), parasites of the 

horse. Parasitology 121:649-659. 

Mfitilodze, M. W., and G. W. Hutchinson. 1985. 

Prevalence and abundance of equine strongyles 

(Nematoda: Strongyloidea) in tropical Australia. 


Ogbourne, C. P. 1976. The prevalence, relative abundance 

and site distribution of nematodes in the 

subfamily Cyathostominae in horses killed in Britain. 


Reinemeyer, C. R., S. A. Smith, A. A. Gabel, and 

R. P. Herd. 1984. The prevalence and intensity 

of internal parasites in horses in the USA. Veterinary 


van Loon, G., P. Deprez, E. Muylle, and B. Sustronck. 

1995. Larval cyathostomiasis as a cause 

of death in two regularly dewormed horses. Journal 

of Veterinary Medicine, Series A 42:301-306. 

Helminth Parasites of the Green Frog (Rana clamitans} from 

Southeastern Wisconsin, U.S.A. 

H. RANDALL YODER,' JAMES R. CoooiNS,2 AND J. CORBETT REINBOLD 

Department of Biological Sciences, University of Wisconsin-Milwaukee, P.O. Box 413, Milwaukee, 

Wisconsin 53201, U.S.A. (e-mail: 2coggins@uwm.edu) 

ABSTRACT: Between 13 August and 3 September 1999, 

26 green frogs Rana clamitans Rafinesque, 1820, were 

collected from 2 ponds at the University of Wisconsin- 

Milwaukee Field Station in Ozaukee County, Wisconsin, 

U.S.A. Hosts were euthanized and organs were examined 

for helminth parasites. All host individuals were infected 


of Biology, Lamar University, P.O. Box 10037, 

Beaumont, Texas 77710, U.S.A. (e-mail: 

y oderhr @ hal .lamar.edu) 

with 1 or more helminth parasites. A total of 11 helminth 

species infected ./?. clamitans at this location: 9 platyhelminths 

(7 trematodes, 2 cestodes) and 2 nematodes. Mean 

abundance of infection was 65.5 ± 79.7 worms per host 

(range = 1-330). This is the first report of Clinostomum 

sp. from green frogs in Wisconsin. 

KEY WORDS: Amphibia, aquatic, Cestoda, Clinostomum, 

ephemeral pond, Haematoloechus varioplexus, 

green frog, helminth, Nematoda, parasites, Rana 

clamitans, survey, temporary pond, Trematoda, Wisconsin, 

U.S.A. 



The green frog Rana clamitans Rafmesque, 

1820, occurs from Newfoundland, where the 

population was introduced (Conant and Collins, 

1991), to western Ontario, Canada, in the northern 

extent of its range and from North Carolina 

to eastern Oklahoma, U.S.A. in the south (Vogt, 

1981). Although reports of green frog parasites 

are numerous, only 3 studies have been conducted 

in Wisconsin, U.S.A. (Williams and Taft, 

1980; Coggins and Sajdak, 1982; Bolek, 1998). 

A total of 26 green frogs were collected by hand 

between 13 August and 3 September 1999 from 2 

temporary ponds at the University of Wisconsin- 

Milwaukee Field Station, Ozaukee County, Wisconsin 

(43°23'N; 88°2'W). Frogs were transported 

to the laboratory and euthanized in MS-222 (ethyl 

m-aminobenzoate sulfonic acid). Body surface, 

mouth, eustachian tubes, celom, lungs, stomach, 

small intestine, colon, urinary bladder, liver, kidneys, 

and leg musculature in individual containers 

were examined with a dissecting microscope for 

the presence of helminth parasites. Nematodes 

were preserved in 70% ethanol and mounted in 

glycerin for identification. Larval and adult platyhelminths 

were fixed in alcohol-formalin-acetic 

acid, stained with acetic carmine, and mounted in 

Canada balsam. Voucher specimens were deposited 

at the H. W. Manter Helminth Collection, University 

of Nebraska, Lincoln, Nebraska (Table 1). 

Use of ecological terms follows the suggestions of 

Bush et al. (1997). 

All host individuals were infected with 1 or 

more helminths (prevalence = 100%). The component 

community of green frogs consisted of 11 

helminth species: 7 trematodes, 2 cestodes, and 2 

nematodes (Table 1). Overall mean abundance of 

helminths was 65.5 ± 79.7 worms per frog (range 

= 1—330). Haematoloechus varioplexus occurred 

with highest mean abundance, mean intensity, and 

prevalence of infection (Table 1). Nematodes occurred 

in low numbers and in few hosts (Table 1). 

Adult green frogs breed in a variety of permanent 

bodies of water (May-July in Wisconsin) and 

inhabit the periphery of these aquatic habitats 

throughout the summer (Vogt, 1981). During this 

time, adult frogs feed upon a variety of animals, 

including several species of insects with aquatic 

life histories (Jenssen and Klimstra, 1966). Whereas 

green frogs are known to migrate prior to hibernation, 

they are thought to seek out aquatic 

habitats that are well oxygenated and do not freeze 

entirely in winter (Lamoureux and Madison, 

1999). The ponds sampled in the present study are 


ephemeral. Even in years when some water remains 

over winter, these ponds freeze solid. The 

green frogs that we collected seem to have moved 

into these ponds as a place to feed prior to hibernating 

in other areas. 

The species composition and numbers of helminths 

in green frog infracommunities at this location 

were similar to those reported previously 

(Rankin, 1945; Bouchard, 1951; Najarian, 1955; 

Campbell, 1968; Williams and Taft, 1980; Coggins 

and Sajdak, 1982; Muzzall, 1991; McAlpine, 

1997; Bolek, 1998; McAlpine and Burt, 1998). 

The aquatic habitat and diet of green frogs correspond 

with helminth communities consisting mostly 

of platyhelminths with indirect life cycles and 

relatively few direct life cycle nematodes. In the 

present study, H. varioplexus occurred with the 

highest values of prevalence, mean intensity, and 

mean abundance. These values are also high compared 

with those reported in previous studies. 

Muzzall (1991) reported 57% of 120 green frogs 

infected with H. parviplexus, synonymous with H. 

varioplexus (Kennedy, 1981), with a mean intensity 

of 29. Najarian (1955) reported 48% of 40 

green frogs infected with H. parviplexus and 42% 

prevalence for H. breviplexus but did not provide 

values for intensity or abundance of infection. Bolek 

(1998) reported a prevalence of 44% for H. 

varioplexus from 75 green frogs with a mean intensity 

of 5.3. Others have reported prevalence values 

of 25% or less for Haematoloechus spp. from 

R. clamitans (Rankin, 1945; Bouchard, 1951; 

Campbell, 1968; Williams and Taft, 1980; Mc- 

Alpine and Burt, 1998). Haematoloechus varioplexus 

has been reported previously from wood 

frogs (Rana sylvatica Le Conte, 1825) and spring 

peepers (Pseudacris crucifer Wied, 1839) from the 

same ponds sampled in the current study (Yoder 

and Coggins, 1996). It is therefore likely that infected 

intermediate hosts are present in these 

ponds. Additionally, large numbers of immature H. 

varioplexus were recovered from green frogs, indicating 

that hosts are being infected while feeding 

at these locations. Odonates serve as second intermediate 

hosts for species of Haematoloechus. 

Muzzall (1991) reported that the absence of fish 

predators may have increased the number of adult 

odonates emerging from Turkey Marsh, Michigan, 

U.S.A., resulting in richer helminth communities 

than those occurring in habitats where both frogs 

and fish occur. The absence of fish from these 

ephemeral ponds may have had a similar result in 

terms of high values of parasitism by H. vario-

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'.frici/i- Thomas, 1939 (HWML 


Stafford, 1900 (HWML 1 

Gorgoderina bilobata 

Rankin, 1937 (HWML 

Megalodiscus temperatus 

Stafford, 1905 (HWI 

Clinostomum sp. 

Leidy, 1856 (HWML 15382) 

Metacercariat 

sstoda 

Proteocephalus sp. Weinland, 1858 (HWML 1 

Mesocestoides sp. 

Vaillant, 1863 (HWML 153 

ematoda 

Oswaldocruzia / npiens Walton, 1929 (HWML 

Cosmocercoides sp. Wilkie, 1930 (HWML 15: 


plexus. This is the first report of Clinostomum sp. 

from Wisconsin green frogs. 

We thank the U.W.M. Field Station Committee 

and staff for their support of this project. 


Bolek, M. G. 1998. A seasonal comparative study of 

helminth parasites in nine Wisconsin amphibians. 

M.S. Thesis, University of Wisconsin-Milwaukee, 

Milwaukee, Wisconsin, U.S.A. 134 pp. 

Bouchard, J. L. 1951. The Platyhelminthes parasitizing 

some northern Maine Amphibia. Transactions 

of the American Microscopical Society 70: 

245-250. 





Campbell, R. A. 1968. A comparative study of the 

parasites of certain Salientia from Pocahontas 

State Park, Virginia. Virginia Journal of Science 

19:13-20. 

Coggins, J. R., and R. A. Sajdak. 1982. Survey of 




Conant, R., and J. T. Collins. 1991. A Field Guide 

to Reptiles and Amphibians: Eastern and Central 

North America. Houghton Mifflin Company, Boston, 

Massachusetts, U.S.A. 450 pp. 

Jenssen, T. A., and W. D. Klimstra. 1966. Food habits 

of the green frog, Rana clamitans, in southern 

Illinois. American Midland Naturalist 76:169- 

182. 

Kennedy, M. J. 1981. A revision of the genus Haematoloechus 

Looss, 1899 (Trematoda: Haemato- 



Research Note 

loechidae) from Canada and the United States. 


Lamoureux, V. S., and D. M. Madison. 1999. Overwintering 

habitats of radio-implanted green frogs, 

Rana clamitans. Journal of Herpetology 33:430- 

435. 

McAlpine, D. F. 1997. Helminth communities in bullfrogs 

(Rana catesbeiand), green frogs (Rana 

clamitans), and leopard frogs (Rana pipiens) from 

New Brunswick, Canada. Canadian Journal of Zoology 

75:1883-1890. 

, and M. D. B. Burt. 1998. Helminths of bullfrogs, 

Rana catesbeiana, green frogs, R. clamitans, 

and leopard frogs, R. pipiens in New Brunswick. 

Canadian Field-Naturalist 112:50-68. 

Muzzall, P. M. 1991. Helminth infracommunities of 

the frogs Rana catesbeiana and Rana clamitans 

from Turkey Marsh, Michigan. Journal of Parasitology 

77:366-371. 

Najarian, H. H. 1955. Trematodes parasitic in the Salientia 

in the vicinity of Ann Arbor, Michigan. 

American Midland Naturalist 55:195-197. 

Rankin, J. L. 1945. An ecological study of the helminth 

parasites of amphibians and reptiles of 

western Massachusetts and vicinity. Journal of 


Vogt, R. C. 1981. Natural History of Amphibians and 

Reptiles of Wisconsin. Milwaukee Public Museum 

and Friends of the Museum, Milwaukee, Wisconsin, 

U.S.A. 205 pp. 

Williams, D. D., and S. J. Taft. 1980. Helminths of 

anurans from NW Wisconsin. Proceedings of the 

Helminthological Society of Washington 47:278. 



c. crucifer Weid, and the wood frog, Rana 



63:211-214. 

Gastrointestinal Helminths of Spinner Dolphins Stenella longirostris 

(Gray, 1828) (Cetacea: Delphinidae) Stranded in La Paz Bay, 

Baja California Sur, Mexico 

ROGELIO AcuiLAR-AouiLAR,1 ROSA GRISELDA MORENO-NAVARRETE, GUILLERMO SALGADO- 

MALDONADO, and BERNARDO VILLA-RAMIREZ 

Universidad Nacional Autonoma de Mexico, Institute de Biologia, Apartado Postal 70-153, Mexico, D. F, 

CP 04510, Mexico 

ABSTRACT: Thirty-one spinner dolphins Stenella longirostris 

stranded in La Paz Bay, Baja California Sur, 

Mexico, were examined for endoparasitic helminths. 

The following species were identified: Zalophotrema 


pacificum and Hadwenius tursionis (Digenea); Strobilocephalus 

triangularis, Trigonocotyle sp., and Tetraphyllidea 

gen. sp. larva (Cestoda); immature Bolbosoma 

hamiltoni (Acanthocephala); and Anisakis typica

(Nematoda). Except for H. tursionis, all the identified 

helminths are reported for the first time in Mexico. 

Stenella longirostris represents a new host for H. tursionis 

and A. typica. 

KEY WORDS: Cetacea, spinner dolphin, Stenella 

longirostris, parasites, Digenea, Cestoda, Nematoda, 

Acanthocephala, Gulf of California, Mexico. 

Although cetaceans, including dolphins, are 

common in marine waters of Mexico, their parasite 

fauna is poorly known. To date, only 2 reports 

on the helminth parasites of cetaceans in 

Mexico have been published. Lamothe-Argumedo 

(1987) identified the trematode Hadwenius 

tursionis (Marchi, 1873) in the intestine of 

the vaquita Phocoena sinus Norris and Mc- 

Farland, 1958 (Phocoenidae), from the northern 

Gulf of California, and Morales-Vela and Olivera-G6mez 

(1993) reported the trematode Nasitrema 

globicephala Neiland, Rice, and Holden, 

1970, and the nematodes Stenurus globicephalae 

Baylis and Daubney, 1925, Stenurus minor 

(Kuhn, 1829), and Crassicauda sp. in the pilot 

whale Globicephala macrorhynchus Gray, 1846 

(Delphinidae), from Cozumel Island, Quintana 

Roo (Caribbean Sea). The present report provides 

data on helminth occurrence in spinner 

dolphins Stenella longirostris (Gray, 1828) from 

the state of Baja California Sur, Mexico. 

In August 1993, 31 spinner dolphins were 

stranded in La Paz Bay (24°07'-24°21'N; 

110°17'-110°40'W), 20 km SW of the city of La 

Paz, Baja California Sur. The stranded dolphins 

consisted of 17 males (total length 130-188 cm, 

weight 19—57 kg, ages 1-18 yr) and 14 females 

(161-186 cm, 33-45 kg, 5.5-15 yr). The animals 

died of unknown causes during the strand, 

and they were kept deep frozen ( —22°C) until 

examination. During necropsy, the digestive 

tract of each animal was separated from its other 

viscera and examined for parasites. Trematodes 

and cestodes were fixed with Bouin's fluid and 

preserved in 70% ethanol, and acanthocephalans 

and nematodes were fixed and preserved in 70% 

ethanol. All helminths identified during the examination 

have been deposited in the Coleccion 

Nacional de Helmintos (National Helminth Collection) 

(CNHE) of the Universidad Nacional 

Autonoma de Mexico. 

Seven helminth species were recovered from 

1 Corresponding author (e-mail: 

raguilar@mail.ibiologia.unam.mx). 


the 31 dolphins. These include 2 trematodes: 

Zalophotrema pacificum Dailey and Perrin, 1973 

(bile ducts, prevalence 19%, mean intensity 6 

worms per parasitized host, range 1—16, CNHE 

No. 4018) and Hadwenius tursionis (Marchi, 

1873) (intestine, 6%, 1, 1-1, CNHE No. 4017); 

3 cestodes: Strobilocephalus triangularis (Diesing, 

1850) (rectum, 6%, 2, 2-2, CNHE No. 

4019), Trigonocotyle sp. (intestine, 90%, 5, 1- 

27, CNHE No. 4021; the poor condition of specimens 

preclude identification of species), and 

larval stages of Tetraphyllidea (intestine, 16%, 

31, 5-69, CNHE No. 4020); the nematode Anisakis 

typica (Diesing, 1860) (stomach, 77%, 18, 

1-98, CNHE No. 4023); and the immature acanthocephalan 

Bolbosoma hamiltoni Baylis, 1929 

(posterior intestine, 51%, 4, 1-9, CNHE No. 

4022). 

The helminth parasites of 5. longirostris have 

been reported by Delyamure (1955), Dailey and 

Brownell (1972), and Dailey and Perrin (1973). 

The previously recorded helminth fauna for this 

dolphin species includes the following: the trematodes 

Oschmarinella laevicaecum (Yamaguti, 

1942), Campula rochebruni (Poirier, 1886), Delphinicola 

tenuis Yamaguti, 1933, Lecithodesmus 

nipponicus Yamaguti, 1942, and Z. pacificum; 

the cestodes Diphyllobothrium fuhrmanni Hsu, 

1935, S. triangularis, Tetrabothrium forsteri 

(Krefft, 1871), Phyllobothrium delphini (Bosc, 

1802), Phyllobothrium sp., Monorygma grymaldii 

(Moniez, 1881), and Monorygma sp.; the 

nematodes Anisakis simplex (Rudolphi, 1809), 

Halocercus delphini Baylis and Daubney, 1925, 

and Mastigonema stenellae Dailey and Perrin, 

1973; and the acanthocephalans Bolbosoma vasculosum 

(Rudolphi, 1819), Bolbosoma balaenae 

(Gmelin, 1790), and Corynosoma sp. In the 

present study, the previously recorded Z. pacificum 

and S. triangularis are identified, and new 

host records are reported for H. tursionis, Trigonocotyle 

sp., tetraphyllidean B. hamiltoni, and 

A. typica. All but 1 of the identified species (H. 

tursionis) are recorded for the first time in Mexico. 

We thank Dr. Luis Fleischer and Hector 

Perez-Cortes, Centro Regional de Investigacion 

Pesquera, La Paz, Baja California Sur, for permission 

to examine dolphins; Dr. Tomas Scholz 

for confirmation of cestodes; Dr. Brent Nickol 

for review of the manuscript; Dr. Krzysztof 

Zdzitowiecki for providing helpful comments 

about the acanthocephalan; and Alejandro San- 



chez-Rios, Alejandra Nieto, and Francisco Anguiano 

for technical assistance. Collection of cetaceans 

was permitted by the Secretaria de Pesca, 

Mexico (authorization number 2275). 


Dailey, M. D., and R. L. Brownell. 1972. A checklist 

of marine mammal parasites. Pages 528-589 in S. 

H. Ridgeway, ed. Mammals of the Sea: Biology 

and Medicine. Charles C. Thomas, Springfield, Illinois, 

U.S.A. 

, and W. F. Perrin. 1973. Helminth parasites 

of porpoises of the genus Stenella in the eastern 

tropical Pacific, with description of two new species: 

Mastigonema stenellae gen. et sp. n. (Nematoda: 

Spiruroidea) and Zalophotrerna pacificurn 

n. sp. (Trematoda: Digenea). Fishery Bulletin 71: 

455-471. 



Research Note 

Delyamure, S. L. 1955. The Helminth Fauna of Marine 

Mammals. Ecology and Phylogeny. Izdatel'stov 

Akademii Nauk SSSR. Translated 1968, 

Israel Program for Scientific Translations, Jerusalem, 

Israel. 522 pp. 

Lamothe-Argumedo, R. 1987. Trematodos de mamfferos 

III. Hallazgo de Synthesium tursionis (Marchi, 

1873) Stunkard y Alvey, 1930 en Phocoena 

sinus (Phocoenidae) en el Golfo de California, 

Mexico. Anales del Instituto de Biologia, Universidad 


Zoologia 58:11-20. 

Morales-Vela, D., and L. D. Olivera-Gomez. 1993. 

Varamiento de calderones Globicephala macrorhynchus 

(Cetacea: Delphinidae) en la Isla de 

Cozumel, Quintana Roo, Mexico. Anales del Instituto 

de Biologia, Universidad Nacional Autonoma 

de Mexico, Serie Zoologia 64:177-180. 

The Lung Nematodes (Metastrongyloidea) of the Virginia Opossum 

Didelphis virginiana in Southern California, U.S.A. 

VICTORIA E. MATEY,M BORIS I. KUPERMAN,' JOHN M. KiNSELLA,2 G. F. LLOYD,' AND 

S. J. LANE3 

1 Department of Biology and Center for Inland Waters, San Diego State University, San Diego, California 92182, 

U.S.A. (e-mail: kuperman@sunstroke.sdsu.edu), 

2 Department of Pathobiology, College of Veterinary Medicine, University of Florida, Gainesville, Florida 32611, 

U.S.A. (e-mail: wormdwb@aol.com), and 

3 Project Wildlife, P.O. Box 80696, San Diego, California 92138, U.S.A (e-mail: sjlane7@home.com) 

ABSTRACT: The lungworm Heterostrongylus heterostrongylus 

(Nematoda: Metastrongyloidea) is reported 

for the first time from the Virginia opossum Didelphis 

virginiana in North America. Seventeen of 31 opossums 

(55%) examined from San Diego County, California, 

U.S.A., were infected with H. heterostrongylus, 

with intensities ranging from 8 to 128 worms per host 

(mean 41). Another species of metastrongyloid nematode, 

Didelphostrongylus hayesi, was found in 74% of 

the lungs examined, with intensity ranging from 2 to 

1,328 worms per host (mean 312). 

KEY WORDS: lungworm, Heterostrongylus heterostrongylus, 

Nematoda, opossum, Didelphis virginiana, 

Didelphostrongylus hayesi, California, U.S.A. 

The Virginia opossum Didelphis virginiana 

Ken; 1792, is the only marsupial inhabiting 



North America, occurring in tropical, subtropical, 

and temperate habitats from southern Canada 

to Costa Rica (Gardner, 1973). California, 

U.S.A., was outside the original range of D. virginiana 

until its accidental introduction into Los 

Angeles County and the San Jose area from various 

eastern states between 1890 and 1910. By 

1958, D. virginiana was distributed widely in all 

the areas of California below 1,500 m altitude 

(Hunsaker, 1977). 

Until recently, the metastrongyloid lungworms 

of D. virginiana have been studied only 

in the midwestern and eastern U.S.A. Alden 

(1995) reviewed helminth records from the Virginia 

opossum and listed records of Didelphostrongylus 

hayesi Prestwood, 1976, from North 

Carolina, Georgia, Louisiana, and Tennessee, 

U.S.A. Subsequently, Baker et al. (1995) record-


Figure 1. Cephalic end of the lung nematode Heterostrongylus heterostrongylus from the Virginia opossum 

Didelphis virginiana. Male, frontal view. SEM. Am = amphid; Cl = collarette; Cp = cephalic papilla; 

L = lip; M = mouth opening; R = ring surrounding mouth. Scale bar = 20 u,m. 

ed D. hayesi from the Virginia opossum from 

Sacramento County in northern California. The 

objective of our study was to determine the identity 

and prevalence of lung parasites in feral Virginia 

opossums from southern California. 

Thirty-one Virginia opossums from San Diego 

County, California, killed by cars or euthanized 

after trauma, were examined for lung parasites 

from March 1999 to June 2000. All samples 

were obtained from a local nonprofit organization, 

Project Wildlife, Opossum Team, 

members of which also carried out the necropsy 

of the animals. All specimens were categorized, 

on the basis of weight, into juveniles (0.14-0.90 

kg) or adults (1.2-3.4 kg). The lungs with attached 

trachea of 7 juveniles and 24 adults were 

examined grossly and under the dissecting microscope. 

The trachea, bronchi, and bronchioles 

were split, and the lung parenchyma was teased 

apart gently. Worms recovered were fixed in 5% 

formalin or alcohol-formalin-acetic acid (AFA). 

For light microscopy, worms were examined as 

temporary whole mounts in glycerine after 

clearing in glycerine-alcohol with a Diastar® 

microscope equipped with a Photostar® camera 

system and were measured in micrometers. For 

scanning electron microscopy (SEM), the specimens 

were postfixed in 1% osmium tetroxide, 

followed by dehydration in an ethanol series, 

critical point dried with liquid CO2, sputter coated 

with gold-palladium, and examined with a 

Hitachi S-2700® scanning electron microscope. 

Voucher specimens of nematodes were deposited 

in the H. W. Manter Laboratory of Parasitology, 

University of Nebraska State Museum, Lincoln, 

Nebraska, U.S.A. (accession numbers 

15617-15619). 

In all, 7,381 lungworms were found in adult 

and juvenile animals examined. The parasites 

were identified as metastrongyloid nematodes. 

Of these, 91.1% were identified as D. hayesi and 

8.9% were identified as Heterostrongylus heterostrongylus 

Travassos, 1925. This is the first 

record of H. heterostrongylus from D. virginiana 

and the first record of this nematode in 

North America. Previous records of H. heterostrongylus 

were from another species of opossum, 

Didelphis marsupialis Linnaeus, 1758, 



from Colombia and Brazil, South America (Travassos, 

1925; Vaz and Pereira, 1934; Anderson 

et al., 1980). 

Morphologic and morphometric features of H. 

heterostrongylus in the opossums from southern 

California resembled those of specimens from 

South America described by Anderson et al. 

(1980). In D. virginiana, the male worms were 

slightly smaller, and the female worms were 

larger than in D. marsupialis. The mean length 

and width of H. heterostrongylus males from D. 

virginiana were correspondingly 6.5 mm (5.0— 

7.2) and 280 |xm (245-320). For females, the 

mean length was 9.7 mm (8.6-13.4) and the 

mean width was 380 jxm (350-475). SEM study 

showed some features of the cephalic structures 

that were not noted by Anderson et al. (1980). 

The 6 lips are completely fused, and each of 

them, in addition to 2 cephalic papillae, bears an 

amphid opening on the surface by an amphidial 

canal (Fig. 1). The shape of the mouth opening 

varies from triangular to circular (Fig. 1). A delicate 

ring surrounding the mouth opening and a 

collarette formed by a dilated cuticle are considered 

as permanent structures (Fig. 1). A new 

character of the female caudal extremity found 

in our study is a pair of small caudal papillae 

near the tip of the short blunt tail. The morphology 

of the male bursa, with its large lobe 

formed by the dorsal ray and short (93-100 |xm), 

complex and slightly arcuate spicules, is the 

same as previously reported. 

In the infected opossums, H. heterostrongylus 

were found lying freely in the bronchi. In 1 case, 

worms were recovered from the trachea. 

Seventeen of 31 opossums (55%) were infected 

with H. heterostrongylus, and intensities 

of infection ranged from 8 to 128 (mean 41). 

Infections were found in 58% of adult animals, 

with 12 to 128 worms per host (mean 41), and 

43% of juveniles, with intensities of 8 to 80 

worms per host (mean 44). The other species of 

lung nematodes, D. hayesi, was found under the 

pleura and was piercing lung tissue in 23 of 31 

opossums (74%). Intensities of infection ranged 

from 2 to 1,328 worms per host (mean 312). All 

animals infected by H. heterostrongylus were 

also infected by D. hayesi. 

Baker et al. (1995) reported on the prevalence 

and treatment of D. hayesi infections in opos- 


sums collected in Yolo, Solano, and Sacramento 

counties in northern California. Infections were 

found in 23 of 33 opossums (70%). Because 

most of their infections were diagnosed by the 

presence of metastrongylid larvae in the feces 

and only 2 infections were confirmed by necropsies, 

it is possible that mixed infections of H. 

heterostrongylus and D. hayesi were also present 

in that area of California. Examination of 

additional host samples is desirable both in California 

and the eastern U.S.A. to determine the 

local range of H. heterostrongylus. 

We are deeply indebted to the Opossum Team 

of Project Wildlife, especially its leader, Blair 

Lee, and coleader, Beverly Rosas, and Meryl 

Faulkner, Mary Platter-Reiger, and numerous 

volunteers for collecting and providing specimens 

of the Virginia opossum for our studies. 

We are very grateful to D.V.M. Alfonso Guajardo, 

San Diego County Veterinarian Office, and 

Julie Irwin, student at San Diego State University 

(SDSU), for the necropsy of the opossums. 

Thanks are also due to SDSU student Susan 

Henderson for her technical assistance. We also 

thank 2 anonymous reviewers whose suggestions 

improved this paper. 


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 


Anderson, R. C., M. D. Little, and U. R. Strelive. 

1980. The unique lungworms (Nematoda: Metastrongyloidea) 

of the opossum (Didelphis marsupialis 

Linnaeus). Systematic Parasitology 2:1-8. 

Baker, D. G., L. F. Cook, E. M. Johnson, and N. 

Lamberski. 1995. Prevalence, acquisition, and 

treatment of Didelphostrongylus hayesi (Nematoda: 

Metastrongyloidea) infection in opossums (Didelphis 

virginiana). Journal of Zoo and Wildlife 

Medicine 26:403-408. 

Gardner, A. L. 1973. The systematics of the genus 

Didelphis (Marsupialia: Didelphidae) in North and 

Middle America. Special Publications of the Museum, 

Texas Tech University 4:3-45. 

Hunsaker, D. 1977. Ecology of New World marsupials. 

Pages 95-156 in D. Hunsaker II, ed. The 

Biology of Marsupials. Academic Press, New 

York-San Francisco-London. 

Travassos, L. 1925. Un nouveau type de Metastrongylidae. 

Comptes Rendus des Seances de la Societe 

de Biologic, Paris 93:1259-1262. 

Vaz, Z., and C. Pereira. 1934. Some new Brazilian 

nematodes. Journal of the Washington Academy 

of Sciences 24:54.



Research Note 


New Host and Distribution Records of Cosmocephalus obvelatus 

(Creplin, 1825) (Nematoda: Acuariidae), with Morphometric 

Comparisons 

JULIA I. DiAZ,1 GRACIELA T. NAVONE, AND FLORENCIA CREMONTE 

Centre de Estudios Parasitologicos y de Vectores (CONICET-UNLP), Calle 2 NQ 584, 1900 La Plata, 

Argentina (e-mail: cepave@netverk.com.ar) 

ABSTRACT: This is the first record of Cosmocephalus 

obvelatus (Creplin, 1825) Seurat, 1919 (Nematoda: 

Acuariidae), from Argentina (Valdes Peninsula, province 

of Chubut) and from the Magellanic penguin 

Spheniscus magellanicus (Aves: Spheniscidae). The 

prevalence of this parasite was 31.3% and the mean 

intensity was 5.4. Despite the wide geographic distribution 

and the great variety of hosts parasitized by C. 

obvelatus (14 families belonging to 8 orders), there 

were no significant differences in morphological characteristics 

and measurements from previous records. 

Both the wide distribution and the morphometrical stability 

of C. obvelatus may be explained by its ecology 

and mode of transmission. 

KEY WORDS: Cosmocephalus obvelatus, Acuariidae, 

Nematoda, Spheniscus magellanicus, Spheniscidae, 

marine birds, Argentina. 

Cosmocephalus obvelatus (Creplin, 1825) 

Seurat, 1919, an acuariid nematode with a wide 

distribution, has been previously reported in Europe, 

Asia, Africa, New Zealand, and North 

America (Wong and Anderson, 1982). In South 

America, there is only 1 record of C. obvelatus, 

described as Cosmocephalus tanakai by Rodrigues 

de Olivera and Vicente (1963) from the 

black-backed gull Larus dominicanus Lichtenstein, 

1823, in Brazil. Later, C. tanakai was synonymized 

with C. obvelatus by Anderson and 

Wong (1981). This parasite has a wide range of 

hosts, having been previously recorded in members 

of Lariidae, Pelecanidae, Rynchopidae, 

Sternidae, Anatidae, Podicipedidae, Phalacrocoracidae, 

Gaviidae, Ardeidae, Stercoraridae, 

Haematopodidae, Treschiornitidae, and Accipitridae 

(Barus and Majudmar, 1975; Borgsteede 

and Jansen, 1980; Anderson and Wong, 1981; 

1 Corresponding author (e-mail: jid027@yahoo. 

com.ar). 

Tuggle and Schmeling, 1982). Among members 

of the Spheniscidae, C. obvelatus has been cited 

only from the rockhopper penguin Eudyptes 

crestatus (Miller, 1784) caught in Chile and 

transferred to the Japanese Zoological Garden 

(Azuma et al., 1988). 

This note reports the first record of C. obvelatus 

in the Magellanic penguin Spheniscus magellanicus 

(Forster, 1781) (Aves: Spheniscidae). 

It is also the first time that C. obvelatus has been 

found in Argentina. Measurements of the specimens 

in this study are compared with those given 

by previous authors. Morphological details 

seen in the scanning electron microscope (SEM) 

and dates of prevalence and mean intensity are 

provided. 

At irregular intervals from 1996 to 2000, 16 

specimens of S. magellanicus, all of which had 

recently died of unknown but presumably natural 

causes, were collected along the coasts of the 

Valdes Peninsula (42°04'-42°53'S, 63°38'- 

64°30'W), province of Chubut, Argentina. After 

dissection, the digestive tract was fixed in 10% 

formalin. Acuariid nematodes were removed 

from the esophagus and stored in 70% ethanol. 

The specimens were cleared in lactophenol and 

studied under the light microscope. Some specimens 

were dried by the critical point method, 

examined by SEM (Jeol/SET 100®), and photographed. 

Voucher specimens were deposited in 

the Helminthological Collection of the Museo de 

La Plata (CHMLP), La Plata, Argentina (Accession 

no. 4811). 

The measurements of our specimens and 

those given by previous authors are listed in Table 

1. Morphological details are shown in Figures 

1-6. The prevalence was 31.25% and the 

mean intensity was 5.4. The esophagus was the 

only site of infection. We observed several de- 


Table 1. Comparative measurements of Cosmocephalus obvelatus from different hosts and localities. 

Rodrigues de 

Olivera and Anderson and Wong 

Vicente (1963) (1981) 

Cram (1927) Khalil (1931) Rao (1951) 

La 

Larus sp. 

Host* Several Pelecanus sp. Larus sp. 

Ne 

Larus delawarensis 

Canada 

Brazil 

Locality Europe Egypt Canada 

4 

10 

27 

— 

39 

45 

— 

— 

— 

3.3 

En 

44 

10 

19.4 (15.8-22.3) 

393 (320-500) 

615 (570-730) 

684 (640-770) 

685 (610-790) 

813 (705-940) 

1.3 (1.2-1.5) 

4.7 (4.1-5.1) 

6.0 (5.2-6.6) 

End of lateral alae 

8.4 (7.4-10.4) 

1 

12.5 

277 

363 

— 

— 

— 

0.77 

3.43 

4.2 

Female 

n — 1 immature — 

Total length (mm) 9.7-20 5.7 37122 

Maximum width (|xm) 300-380 — 200-400 

Buccal capsule ((Am) — — — 

Nerve ring (n-m) — — — 

Deirids (u.m) 490 — — 

Excretory pore (|xm) — — — 

Muscular esophagus (mm) — — — 

Glandular esophagus (mm) — — — 

Total esophagus (jxm) — 680 — 

Postdeirids — — — 

Vulva (from anterior end) (mm) 5.5 Midbody Midbody 

6.2 

— 

— 

39 

21 

— 

Long 

Short 

43 (40-45) 

25 

301 (220-380) 

— 

— 

36 

19 

175 

Vagina vera (pun) — — — 

Vagina uterina (|xm) — — — 

Egg length (p,m) 36 — 35-37 

Egg width (fi,m) 20 — 17-18 

Tail ((i,m) 230 180 — 

10 

10 

27 

— 

— 

46 

55 

— 

— 

5.2 

60 

10 

12.4 (9.9-14.3) 

279 (200-350) 

418 (380-510) 

474 (420-530) 

450 (350-540) 

583 (500-680) 

1.1 (1.0-1.3) 

4.0 (3.6-4.3) 

5.1 (4.6-5.4) 

End of lateral alae 

2 

9.5-12 

250-280 

369 

462 

399 

532-630 

0.98-1.32 

2.85-4.61 

3.83-5.93 

— 

Male 

n — 2 — 

Total length (mm) 5.7-12.2 7.6 37085 

Maximum width (jjum) 240-255 — 150-270 

Buccal capsule (\j.m) — — — 

Nerve ring ((JUTI) — — — 

Deirids (p,m) 430 — — 

Excretory pore (u,m) — — — 

Muscular esophagus (mm) — — — 

Glandular esophagus (mm) — — — 

Total esophagus (|j,m) — 930 — 

Postdeirids — — — 


COMP>

c 

Tj ;^ fn 

in 

ON 

o 

m 

ON 

"3 

s 

ON 

gcd 

u 

r? >ri ? 

— i r- r- 

7 i T 

CN r~ oc 

C- S C 

CN NO NC 

NO CN •r-i 

in


Figures 1-6. Cosmocephalus obvelatus from Spheniscus magellanicus. 1. Apical view. 2. Anterior extremity 

showing lateral alae (arrow), dorsal view. 3. Anterior extremity showing cephalic papillae (arrow), 

inflexions of chordons and deirid (arrow), lateral view. 4. Detail of deirid. 5. Detail of postdeirid located 

near end of lateral alae (arrow). 6. Posterior extremity of male with left spicule protruded and showing 

papillae arrangement with proximal pair of precloacal papillae lying out of line of distribution (arrow), 

latero-ventral view. Scale bars: 1 = 50 ixm, 2 = 500 (mm, 3 = 100 u.m, 4 = 10 (xm, 5 = 20 (Jim, and 6 = 

200 fjim. 


ticeps (Rudolphi, 1819), which also is cosmopolitan 

but has a narrower range of hosts, Etchegoin 

et al. (2000) found differences in measurements 

among specimens from different localities. 

The shape of the cordons, the morphology and 

size of the cervical papillae, and the location in 

the definitive host (habitats) seem to be of fundamental 

importance when establishing relationships 

in the acuariid group (Barus and Majudmar, 

1975). The cordons of Cosmocephalus have 

a complex structure and are relatively wide. 

Cosmocephalus obvelatus is always located in 

the esophagus of the host. The genera Synhimantus 

and Cosmocephalus are closely related; 

they have similar cordons and cervical papillae, 

but members of the former genus live under the 

cuticle of the gizzard. Etchegoin et al. (2000) 

reported morphometric differences that they 

considered as intraspecific variations in specimens 

from different hosts and localities. However, 

we observed that C. obvelatus varies little, 

even in different hosts and localities. This morphometrical 

stability may indicate that Cosmocephalus 

is better adapted to different hosts and 

diverse localities because all hosts have similar 

environmental and feeding habits (eating fish). 

The intermediate hosts of C. obvelatus are amphipods, 

and it uses fish as paratenic hosts (Anderson, 

1992). These characteristics may play an 

important role in the cosmopolitan distribution 

of C. obvelatus. Moreover, the distributions of 

many species of fish-eating birds overlap in their 

breeding and/or wintering grounds. 

The intensity of infection recorded here is 

similar to the intensities found by Keppner 

(1973) from the California gull Larus californicus 

Lawrence, 1854 (Lariidae) (prevalence [P] 

= 23.5% and mean intensity [I] = 4.25), by 

Courtney and Forrester (1974) from the brown 

pelican Pelecanus occidentalis Linnaeus, 1776 

(Pelecanidae), in North America (P = 40% and 

I = 4), and by Lafuente et al. (1999) from Audouin's 

gull Larus audouinii Payraudeau, 1826 

(Lariidae), in the Mediterranean Sea (P = 

82.76% and I = 5.08). This intensity is higher 

than that given by Threlfall (1968) from the 

black-backed gull Larus marinus Linnaeus, 

1758 (Lariidae), in Newfoundland (P = 9.39% 

and I = 1). 

Boero and Led (1970) described a new species, 

Cosmocephalus argentinensis, from 1 female 

specimen found in a Magellanic penguin 


in the Zoological Garden in La Plata, Argentina. 

We consider this acuariid as a species inquirendae 

because the description is very poor and no 

type materials (which were never deposited in a 

museum collection) are available for our examination. 

We gratefully acknowledge the staff of the 

Servicio de Microscopia Electronica de Barrido, 

Museo de La Plata, for their technical assistance 

and Lucy Shirlaw for revision of the English. 

This study was funded by the Consejo Nacional 

de Investigaciones Cientfficas y Tecnicas (CON- 

ICET) and by the Comision de Investigaciones 

Cientfficas de la Provincia de Buenos Aires 

(CIC). 


Anderson, R. C. 1992. Nematode Parasites of Vertebrates. 

Their Development and Transmission. 

CAB International, Wallingford, Oxon, U.K. 578 

pp. 

, and P. L. Wong. 1981. Redescription of Cosmocephalus 

obvelatus (Creplin, 1825) (Nematoda: 

Acuarioidea) from Larus delawarensis Ord (Laridae). 

Canadian Journal of Zoology 59:1897- 

1902. 

Azuma, H., M. Okamoto, M. Ohbayashi, Y. Nishine, 

and T. Mukai. 1988. Cosmocephalus obvelatus 

(Creplin 1825) (Nematoda: Acuariidae) collected 

from esophagus of rockhopper penguin, Eudyptes 

crestatus. Japanese Journal of Veterinary 

Research 36:73-77. 

Barus, V., and G. Majudmar. 1975. Scanning microscopic 

studies on the cordon structures of Acuariid 

genera (Nematoda: Acuariidae). Folia Parasitologica 

22:125-131. 

Boero, J. J., and J. E. Led. 1970. El parasitismo de 

La Fauna Autoctona. VI. Los parasites de la avifauna 

argentina I. Actas de las 5a Jornadas de Veterinaria, 

Facultad de Ciencias Veterinarias, Universidad 

Nacional de La Plata: 65-71. 

Borgsteede, F. H. M., and J. Jansen. 1980. Spirurata 

in wild birds in The Netherlands. Tropical and 

Geographic Medicine 32:91-92. 

Bowie, J. Y. 1981. Redescription of Cosmocephalus 

tanakai Rodriguez and Vicente (Nematoda-Acuariidae) 

a parasite of the southern black-backed 

gull in New Zealand. New Zealand Journal of Zoology 

8:249-253. 

Courtney, C. H., and D. J. Forrester. 1974. Helminth 

parasites of the brown pelican in Florida 

and Louisiana. Proceedings of the Helminthological 


Cram, E. B. 1927. Bird parasites of the suborders 

Strongylata, Ascaridata and Spirurata. Bulletin of 

the United States National Museum 140:1-465. 

Etchegoin, J. A., F. Cremonte, and G. T. Navone. 

2000. Synhimantus (Synhimantus) laticeps (Rudolphi, 

1819) Railliet, Henry et Sisoff, 1912 

(Nematoda, Acuariidae) parasitic in Tyto alba 



(Gmelin) (Aves, Tytonidae) in Argentina. Acta 

Parasitologica 45:99-106. 

Keppner, E. J. 1973. Some parasites of California 

gull, Lams californicus Lawrence, in Wyoming. 

Transactions of the American Microscopical Society 

92:288-291. 

Khalil, M. B. 1931. On two new species of nematodes 

from Pelecanus onocrotalus. Annals of Tropical 

Medicine and Parasitology 25:455-460. 

Lafuente, M., V. Roca, and E. Carbonell. 1999. Cestodos 

y nematodos de la gaviota de Audouin, Lams 

audouinii Payraudeau, 1826 (Aves: Laridae) 

en las Islas Chafarinas (Mediterraneo sudoccidental). 

Boletin de la Real Sociedad Espanola de Historia 

Natural (Sec. Biol.) 95:13-20. 

Rao, N. 1951. Cosmocephalus firlottei n. sp. (family 

JOHN S. MACKIEWICZ 

Elected to Life Membership in the Helnlinthological 

Society of Washington November 15, 2000 


Acuariidae) from the sea gull Larus argentatus. 


Rodrigues de Olivera, H., and J. J. Vicente. 1963. 

Nova especie do genero Cosmocephalus Molin, 

1858 (Nematoda, Spiruroidea). Revista Brasilera 

de Biologia 23:389-392. 

Threlfall, W. 1968. The helminth parasites of three 

species of gulls in Newfoundland. Canadian Journal 

of Zoology 46:827-830. 

Tuggle, B. N., and S. K. Schmeling. 1982. Parasites 

of the bald eagle (Haliaetus leucocephalus) of 

North America. Journal of Wildlife Diseases 18: 

501-506. 

Wong, P. L., and R. C. Anderson. 1982. The transmission 

and development of Cosmocephalus obvelatus 

(Nematoda: Acuarioidea) of gulls (Laridae). 


GRAHAM C. KEARN 

Elected to Life Membership in the Helminthological 

Society of Washington November 15, 2000



Anniversary Award 

The Helminthological Society of Washington 

NANCY D. PACHECO 

Harley G. Sheffield, left, presents the 2000 Anniversary Award to Nancy D. Pacheco 

As Chairman of the Anniversary Award Committee of the Helminthological Society of Washington, 

it is my duty to present the 2000 Anniversary Award to Nancy Pacheco. Not only is it a 

duty, it is an honor and a pleasure to be able to present this award to such an outstanding member 

of the Society. 

The Award is authorized by the Society's Constitution and is to be given to a member for one 

or more achievements of the following nature: an outstanding contribution to the science of parasitology 

or related sciences that brings honor and credit to the Society, an exceptional paper read 

at a meeting of the Society or published in the Society's journal, outstanding service to the Society, 

or another achievement or contribution of distinction that warrants the highest recognition by the 

Society. The Awards Committee determined that Nancy qualifies in all of the above categories. 

Nancy was born and educated in Kansas. During her junior year of high school, she was fortunate 

to be an exchange student under the American Field Service Program and lived 6 months in New 

Zealand. She subsequently received the Bachelor of Science Degree from Washburn University in 

Topeka. Following graduation, she came to the National Institutes of Health and received a position 

as a biologist in the National Heart Institute. Working with physicians and postdocs, Nancy was 

engaged in studies on cardiac muscle physiology. Maybe there was a lot of twitching in that job 

because for some reason, she saw the light and soon turned to parasitology. In 1968, and for the 

283 



next 8 years, she worked as a biologist in the Animal Parasitology Institute (API) of the U.S. 

Department of Agriculture doing research on poultry and bovine parasites. Shortly after her arrival 

at API, I received a call from her supervisor, Dr. Vetterling, saying that he had hired a new person 

for his electron microscopy lab, and I should come over to meet her. That began our long scientific 

and social association. In 1976, Nancy moved to the Naval Medical Research Institute as a research 

microbiologist. There, she studied parasite immunology in relation to the development of malaria 

vaccines and later worked on cytokine regulation in wound repair. In 1994, she somehow slipped 

out of parasitology, without seeking advice of the Helminthological Society of Washington, and 

worked in the Wound Repair Program at NMRI until her retirement in 1997. 

Nancy's resume lists numerous publications. They illustrate her research contributions in the 

ultrastructure of intracellular parasites, techniques for isolation of large numbers of malaria parasites, 

which is a prerequisite to vaccine development, development of an oral vaccine against Campylobacter 

infection, development of a method to study local inflammatory action, and the study of the 

effects of cytokines in preventing translocation of bacteria in hemorrhagic shock. 

A predominant factor in the Committee's decision was Nancy's excellent service to the Society, 

of which, undoubtably, most of you are aware. She has a record that is hard to beat. To the best 

of my knowledge, and with a little help from her curriculum vitae, Nancy has served in every 

office and has been on every committee of the Society, with the exception of Editor and the Editorial 

Committee. After holding the position of vice president in 1980, she moved up to president the 

next year. She must have done something right because she was re-elected president in 1991, a feat 

that, with one exception, has been unmatched in recent Society history. In 1999, she was elected 

to the position of corresponding secretary-treasurer, which she currently holds. 

In spite of the considerable time that she has donated to the Society, Nancy has offered her 

expertise in various roles in other societies such as the American Society of Parasitologists and the 

American Society of Tropical Medicine and Hygiene. Outside of the scientific area, she has been 

active in many church-related events with her husband Jim. There is one other activity that might 

be noted. As mentioned before, Nancy spent a number of years working with poultry coccidia. You 

all know what people in that field do—they search through chicken droppings and count the coccidial 

oocysts that they find. Well, Nancy must have developed a high degree of excellence in 

counting because she has steadily moved upward through the ranks in the H&R Block organization 

and is now a senior tax preparer. 

Nancy, I am very pleased to present to you, on the behalf of the Anniversary Awards Committee, 

the Helminthological Society of Washington's Anniversary Award for 2000. 



MINUTES 

Harley G. Sheffield, Ph.D. 

November 15, 2000 

Six Hundred Seventy-First through the Six Hundred Seventy-Fifth 

Meeting of the Helminthological Society of Washington 

671st Meeting: George Washington University, meeting. Dr. Eckerlin introduced the speakers. 

Washington, DC, 12 October 2000. President Robert Gwadz gave an overview of the National 

Dennis Richardson conducted the business Institutes of Health (NIH)-funded malaria remeeting. 

Ralph Eckerlin welcomed members search in Mali. This NIH program funds elecand 

their guests and introduced the President, lives for students with interests in either basic 

who briefly summarized the Executive Council sciences or clinical aspects of malaria. Albert 


Nieto reviewed recent advances in the use of 

synthetic peptides for cystic hydatid disease serology. 

John Hawdon provided an account of his 

research on potential hookworm vaccine candidates. 

Finally, Dr. Eckerlin reviewed his studies 

of fleas from flying squirrels in Virginia. New 

and renewal members included Russell C. Van 

Horn (U.S.A.), Eric Panitz (U.S.A.), and Riccardo 

Fiorillo (U.S.A.). 

672nd Meeting: 94th Aero Squadron, College 

Park, Maryland, 15 November 2000. The anniversary 

dinner meeting and program were presided 

over by President Dennis Richardson. The 

slate of officers for 2001 were elected and installed 

by the membership in attendance: Dennis 

J. Richardson, president; William E. Moser, vice 

president; W. Patrick Carney and Nancy D. Pacheco 

continue as reporting secretary and corresponding 

secretary-treasurer, respectively. 

Nancy Pacheco was presented the Anniversary 

Award by Harley Sheffield. John S. Mackiewicz 

and Graham C. Kearn were elected to Life 

Membership (accepted for him by Gene Hayunga) 

and Honorary Membership (accepted for 

him by Sherman Hendrix), respectively. 

673rd Meeting: Nematology Laboratory, Agricultural 

Research Service, USDA, Beltsville, 

Maryland, 17 January 2001. President Dennis 

Richardson presided over the business meeting, 

and Vice President Lynn Carta presided over the 

scientific session. Peter Maser summarized his 

paper "Comparative Biochemistry of Parasitic 

and Free-Living Nematodes." William Wergin 

gave an overview of "Low Temperature Scanning 

Microscopy and its Application in Agriculture." 

Dr. Carta finished the session with "Integrating 

Nematode Morphology and Molecules 

in Plant-Parasitic and Free-Living Nematodes." 

Two new members were announced: Wellington 

MINUTES 285 

A. Oyibo (Nigeria) and Analfa Cristina Paola 

(Argentina). 

674th Meeting: Walter Reed Army Institute of 

Research/Naval Medical Research Center, Silver 

Spring, Maryland, 12 March 2001. President 

Dennis Richardson presided over the business 

meeting and Eileen Franke-Villasante presided 

over the scientific session. Ed Rowton reviewed 

the "Status of the Recent Outbreak of Canine 

Leishmaniasis in the United States." His paper 

was followed by David Fryauff's report on "A 

New Twist on an Old Drug: Recent DOD Studies 

of Primaquine for Malaria Prophylaxis." 

Kent Kester summarized "Advances in Pre- 

Erythrocytic Malaria Vaccine Development," 

and Dr. Ling presented the final paper on "The 

Labor Involved in Malaria Field Studies in Indonesia." 

675th Meeting: Biology Department, Gettysburg 

College, Gettysburg, Pennsylvania, 5 May 

2001. President Richardson presided over the 

business meeting. Robin Overstreet, vice president 

of the American Society of Parasitologists 

(ASP), was welcomed and introduced to the 

members and guests by the president. Dr. Overstreet 

advised members that ASP has resources 

to provide travel grants for students who present 

papers at ASP meetings and to support speakers 

who are invited to present papers at meetings of 

affiliated societies. Sherman Hendrix presided 

over the scientific session. The first paper, presented 

by Dr. Overstreet, covered "Shrimp Parasites 

and Diseases," followed by Eric Hoberg's 

paper on the "Ancient Mariner—A History of 

Seabirds and Tapeworms on the Deep Blue 

Sea." Ann Barse summarized "New Host and 

Geographic Records of Monogenea of Billfishes." 

Sherman Hendrix presented the final paper 

on "Some Aspects of Biology of Bothitrema 

bothi (Platyhelminthes: Monogenea)." 



68(2), 2001, p. 286 

Abaigar, T., 134 

Abrams, A., 177 

Agarwal, N., 87 

Aguilar-Aguilar, R., 204, 272 

Aguirre-Macedo, M. L., 42, 76, 

190 

Al-Sady, R. S. S., 108 

Amin, O. M., 103, 108 

Arjona-Torres, G., 42, 190 

Baez-Vale, R., 196 

Balfour, C. D., 126, 259 

Bassat, S. E, 108 

Ben Hassine, O. K., 91 

Boeger, W. A., 66 

Bolek, M. G., 164 

Brailovsky, D., 196 

Brooks, D. R., 177 

Burge, A. N., 52 

Bursey, C. R., 21, 138 

Cabanas-Carranza, G., 196, 204 

Caira, J. N., 52 

Cano, M., 134 

Carreno, R. A., 177 

Caspeta-Mandujano, J. M., 196, 204 

Center, T. D., 242 

Coggins, J. R., 164, 269 

Cone, D. K., 236 

Cremonte, E, 277 

Criscione, C. D., 149, 156 

Curran, S. S., 219 

Curtis, M. A., 112 

Damborenea, M. C., 249 

Dang, T. T., 219 

Davies, K. A., 242 

Demarais, S., 116 

Diaz, J. I., 277 

Dumailo, S., 42, 190 

Durden, L. A., 177 

Eckerlin, R. P., 143 

Espeso, G., 134 



AUTHOR INDEX FOR VOLUME 68 

Euzet, L., 91 

Fedynich, A. M., 116 

Font, W. E, 149, 156 

Forrester, D. J., 130, 173 

Foster, G. W., 130, 173 

Fried, B., 126, 256, 259 

Garcia-Prieto, L., 9 

Garijo, M. M., 134 

Giblin-Davis, R. M., 242 

Goldberg, S. R., 21, 138 

Gonzalez-Sob's, D., 42, 190 

Hasegawa, H., 261 

Hoberg, E. P., 177 

Hogue, C. C., 36 

Huffman, J. E., 256 

Jimenez-Ruiz, E A., 9 

Joy, J. E., 185 

Kharchenko, V. A., 97 

Kinsella, J. M., 130, 274 

Krecek, R. C., 97 

Kritsky, D. C., 66, 87 

Kuperman, B. I., 274 

Kuzmin, Y., 228 

Lane, S. J., 274 

Lichtenfels, J. R., 97, 265 

Lloyd, G. E, 274 

Love, S., 265 

Matey, V. E., 274 

Matthews, J. B., 265 

Mayen-Pena, E., 196 

McDonnell, A., 265 

Mendoza-Garfias, B., 9 

Mhaisen, E T., 108 

Milek, J. A., 122 

Monasmith, T., 116 

Moreno-Navarrete, R. G., 204, 272 

Mullican, S. K., 256 

Navone, G. T., 277 

Neifar, L., 91 

Nguyen, T. L., 219 

Ortiz, J. M., 134 

Overstreet, R. M., 219 

Paola, A., 249 

Parmelee, J. R., 21 

Peng, J. S., 36 

Perez-Ponce de Leon, G., 1, 9 

Pham, X.-D., 261 

Posel, P., 42, 190 

Reddy, A., 259 

Reinbold, J. C., 269 

Robaldo, R. B., 66 

Rossi, M. L., 126 

Ruiz de Ybanez, M. R., 134 

Salgado-Maldonado, G., 196, 204, 

272 

Sanchez-Nava, P., 204 

Scholz, T., 42, 76, 190 

Seville, R. S., 122 

Siu-Estrada, E., 42, 190 

Snyder, S. D., 228 

Soto-Galera, E., 196, 204 

Thul, J. E., 173 

Tkach, V. V., 228 

Tran, C.-L., 261 

Tucker, R. B., 185 

Vidal-Martinez, V. M., 42, 76, 190 

Villa-Ramirez, B., 272 

Williams, D. S., 242 

Wright, M. E., 112 

Yadav, V. S., 87 

Yoder, H. R., 269 

KEYWORD AND SUBJECT INDEX FOR VOLUME 68 

Acanthobothrium dollyae sp. n., 52 

Acanthobothrium maryanskii sp. 

n., 52 

Acanthobothrium royi sp. n., 52 

Acanthocephala, 9, 36, 103, 108, 

130, 190, 196, 204, 261, 272 

286 


Acanthocephalan cystacanth, 21 

Acuariidae gen. sp., 21, 196 

Adenomera andreae, 21

Adenomera hylaedactyla, 21 

Algansea tincella, 204 

Algonquin Park, Canada, 236 

Allocreadiidae gen. sp., 190 

Allocreadium mexicanum, 204 

Ancyrocephalus etropli, 87 

Ambystoma maculatum, 228 

American toad, 228 

Amphibia, 1, 21, 149, 164, 185, 

228, 269 

Amphilophus alfari, 42, 76, 190 

Anas platyrhynchus, 1 

Anisakis sp., 36 

Anisakis typica, 272 

Anolis carolinensis, 149 

Anura, 1, 21, 149, 164, 185 

Apharyngostrigea sp., 42 

Apicomplexa, 122 

Aplectana hylambatis, 21 

Aponurus sp., 36 

Archocentrm nigrofasciatus, 42, 

76 

Arctic chair, 112 

Arctic lakes, 112 

Argentina, 249, 277 

Ascocotyle (Ascocotyle) tenuicollis, 

42 

Ascocotyle (Phagicola) diminuta, 

42 

Ascocotyle (Phagicola) mollienisicola, 

42 

Ascocotyle (Phagicola) nana, 42 

Ashworthius sp., 177 

Aspidogastrea, 249 

Astyanax fasciatus, 1, 42, 190, 

196 

Athene cunicularia, 130 

Atherinella crystallina, 204 

Australia, 242 

Autonomous Region of the South 

Atlantic, Nicaragua, 42 

Aves, 1, 130, 173, 277 

Awaous tajasica, 204 

Aythya collaris, 173 

Azteca chub, 204 

Baja California Sur, Mexico, 272 

Balsas River, Mexico, 196 

Balsas catfish, 196 

Balsas shiner, 196 

Balsas splitfin, 196 

Barn owl, 130 

Barred owl, 130 

Basin white-lipped frog, 21 

Batracholandros magnavulvaris, 

185 

Batracholandros spectatus, 21 

Beireis' treefrog, 21 

Biodiversity, 1, 177 

Biogeography, 149 

Biomphalaria glabrata, 126 

Black-bellied garter snake, 9 

Blackbelt cichlid, 42, 76, 190 

Blackfin goodea, 196, 204 

Blackfin silverside, 204 

Blood parasites, 173 

Bluegill, 204 

Bolbosoma hamiltoni, 272 

Bolivian bleating frog, 21 

Bolivian white-lipped frog, 21 

Boophilus microplus, 177 

Bothriocephalus acheilognathi, 9, 

196, 204 

Brachylairna mcintoshi, 130 

Brachylecithum rarum, 130 

Brazil, 66 

Brevimulticaecum sp., 21, 42 

Brilliant-thighed poison frog, 21 

Broad-leaved paperbark, 242 

Brown egg frog, 21 

Brown newt, 228 

Bubo virginianus, 130 

Buenos Aires Province, Argentina, 

249 

Bufo americanus, 228 

Bufo glaberrimus, 21 

Bufo marinus, 1, 21 

Bufo typhonius, 21 

Burchell's zebra, 97 

Burrowing owl, 130 

Butterfly cichlid, 42, 76, 190 

California, U.S.A., 36, 274 

Canada, 112, 236 

Cane toad, 1, 21 

Capillaria cyprinodonticola, 196 

Capillaria dispar, 130 

Capillaria falconis, 130 

Capillaria sp., 130 

Capillaria tenuissima, 130 

Capillariidae gen. sp., 204 

Capillariinae gen. sp., 9 

Casmerodius albus, 1 

Catenotaenia sp., 116 

Centrocestus formosanus, 204 

Centropomidae, 66 

Centropomus undecimalis, 66 

Centrorhynchiis kuntzi, 130 

Centrorhynchus spinosus, 130 

Centrocestus formosanus, 196 

Cepedietta michiganensis, 185 

Cestoda, 21, 42, 52, 112, 116, 

130, 138, 149, 156, 164, 196, 

261, 269, 272 

Cestoidea, 9, 36 


INDEX 287 

Chandleronema longigutturata, 

130 

Channel catfish, 196 

Chelonia mydas, 1 

Checklist, 204 

Chihuahuan Desert, Mexico, 116 

Chirostoma humboldtianiim, 204 

Chirostoma jordani, 204 

Chirostoma labarcae, 204 

Chirostoma riojai, 206 

Choanotaenia speotytonis, 130 

Chubut, Argentina, 277 

Cichlasoma beani, 204 

Cichlasoma geddesi, 1 

Cichlasoma faceturn, 249 

Cichlasoma istlanum, 196 

Cichlasoma maculicauda, 42, 76, 

190 

Cichlasoma managuense, 42, 76, 

190 

Cichlasoma nigrofasciatum, 196 

Cichlasoma synspillum, 1 

Cichlasoma urophthalmus, 1 

Ciliata, 185 

Cladocystis trifolium, 42 

Clethrionomys gapperi, 122 

Clinical effects, 256 

Clinostoinum complanatum, 42, 

196, 204 

Clinostoinum sp., 42, 269 

Coccidia, 134 

Coccidiosis, 134 

Colostethus marchesianus, 21 

Colubridae, 219 

Common carp, 204 

Common oval frog, 21 

Common snook, 66 

Community ecology, 116 

Component communities, 1 16 

Contracaecum sp., 9, 42, 196, 204 

Convict cichlid, 42, 76, 190, 196 

Copepods, 112 

Coronocyclus coronatus, 265 

Coronocyclus labiatus, 265 

Coryrnbia ptychocarpa, 242 

Coiynosoma sp., 36 

Cosmocephalus argentinensis, 277 

Cosmocephalus obvelatus, 277 

Cosmocerca brasiliense, 21 

Cosmocerca parva, 21 

Cosmocerca podicipinus, 21 

Cosmocercella phyllomedusae, 21 

Cosmocercoides sp., 164, 269 

Costa Rica, 177 

Cosymbotus platyurus, 138 

Cow Knob salamander, 185 

Crassicutus cichlasomae, 190


Craterostomum acuticaudatum, 

265 

Crepidostomum sp., 190 

Crowding effects, 156 

Crump's treefrog, 21 

Cryptogonimidae gen. sp., 42, 204 

Ctenophryne geayi, 21 

Cundinamarca toad, 21 

Cuticle, 242 

Cuvier's gazelle, 134 

Cyathocotylidae, 219 

Cyathostominea, 265 

Cyathostomurn catinatum, 265 

Cyathostomum montgomeryi, 97 

Cyathostomurn pateratum, 265 

Cylicocyclus ashworthi, 265 

Cylicocyclus elongatus, 265 

Cylicocyclus insigne, 265 

Cylicocyclus leptostomum, 265 

Cylicocyclus nassatus, 265 

Cylicocyclus radiatus, 265 

Cylicocyclus ultrajectinus, 265 

Cylicostephanus bidentatiis, 265 

Cylicostephanus calicatus, 265 

Cylicostephanus longibursatus, 

265 

Cylicostephanus minutus, 265 

Cylicostephanus goldi, 265 

Cylidodontophorus bicoronatus, 

265 

Cylindrotaenia americana, 21 

Cynops ensicauda, 228 

Cyprinus carpio, 204 

Cyst storage, 126 

Cysticercus, 116, 177 

Dactylogyridae, 76, 87 

Darkedged splitfin, 204 

Demerara Falls treefrog, 21 

Dermatemys mawii, 1 

Development, 149 

Diaptomus (Aglaodiaptornus) leptopus, 

112 

Diaptomus (Leptodiaptornus) minutus, 

112 

Didelphis virginiana, 1, 274 

Didelphostrongylus hayesi, 274 

Digenea, 1, 9, 36, 42, 190, 196, 

204 

Diphyllobothrium dendriticum, 

112 

Diplectanidae, 66 

Diplobatis ommata, 52 

Diplostomum sp., 204 

Diplostomum (A.) compactutn, 42, 

196 

Diplostomum (Tylodelphys) sp., 9 

Dipodomys merriami, 116 

Dispharynx affinis, 130 

Dispharynx nasuta, 130 

Domestic chicks, 256 

Dorcas gazelle, 134 

Dracunculus ophidensis, 9 

Dull rocket frog, 21 

Eastern screech-owl, 130 

Echinostoma caproni, 126, 256, 

259 

Echinostoma revolutum, 256 

Echinostoma trivolvis, 256 

Echinostomiasis, 256 

Ecology, 261 

Edalorhina perezi, 21 

Eimeria clethrionomyis, 122 

Eimeria elegatis, 134 

Eimeria gazella, 134 

Eimeria pallida, 134 

Eirunepe snouted treefrog, 21 

Elachistocleis avails, 21 

Elasmobranchii, 52, 91 

Electron microscopy, 52, 228, 

242, 249, 274, 277 

Eleutherodactylus cruralis, 21 

Eleutherodactylus fenestratus, 21 

Eleutherodactylus imitatrix, 21 

Eleutherodactylus peruvianus, 21 

Eleutherodactylus toftae, 21 

Ency station, 126 

Enhydris chinensis, 219 

Enhydris plumbea, 219 

Epidermis, 242 

Epipedobates femoralis, 21 

Epipedobates pictus, 21 

Equus burchelli antiquorum, 97 

Zssctt: americanus, 103 

Estado de Mexico, Mexico, 9, 

196, 204 

Etroplus suratensis, 87 

Eucalyptus leucoxylon, 242 

Eustrongylides sp., 9, 42, 196, 

204 

Excisa excisiformis, 130 

Excretion pattern, 134 

Excystation, 126, 259 

Experimental infection, 112 

Falcaustra affinis, 9 

Falcaustra mexicana, 9 

Falcaustra wardi, 9 

Falcaustra sp., 42 

Fence lizards, 149 

Fergusobia sp., 242 

Fergusonina sp., 242 

Filaria sp., 116 

Fixation artifacts, 156, 219 

Flat-bodied house gecko, 138 


Florida, U.S.A., 130, 149, 173 

Fourth International Congress of 

Nematology, 172 

Fourth International Symposium 

on Monogenea, 142 

Freshwater mullet, 108 

Gazella cuvieri, 134 

Gazella dama mhorr, 134 

Gazella dorcas neglecta, 134 

Gazelle, 134 

Gehyra mutilata, 138 

Gekkonidae, 138, 149 

Genyonemus lineatus, 36 

Georgia, U.S.A., 103 

Girardinichthys multiradiatus, 204 

Glossocercus auritus, 196 

Glypthelmins parva, 21 

Glypthelmins quieta, 164, 269 

Gnathostoma sp., 204 

Goblet cell, 256 

Gold-striped frog, 21 

Golden hamster, 112 

Gongylonema dipodomysis, 116 

Gongylonema neoplasticum, 261 

Gongylonema pulchrum, 177 

Goodea atripinnis, 196, 204 

Gorgoderina bilobata, 269 

Great horned owl, 130 

Green anole, 149 

Green chromide, 87 

Green frog, 164, 269 

Guanajuato, Mexico, 204 

Guerrero, Mexico, 196 

Gulf of California, 52 

Gunther's banded treefrog, 21 

Guppy, 196 

Gussevia herotilapiae sp. n., 76 

Gyalocephalus capitatus, 265 

Gymnura altavela, 91 

Gyrodactylus elegans, 204 

Gyrodactylus sp., 196, 204 

Hadwenius tursionis, 212 

Haemaphysalis juxtakochi, 177 

Haematoloechus varioplexus, 164, 

269 

Haemoproteus nettionis, 173 

Halipegus eccentricus, 164, 269 

Hamptophryne boliviano, 21 

Hedruris siredonis, 9 

Heleobia parchappii, 249 

Helminth larvae, 42 

Helminthological Society of 

Washington: 

Anniversary Award, 283 

Dues Increase, 163 

Editors' Acknowledgments, 155

Honorary Membership, 282 

Life Membership, 282 

Meeting Schedule, 121 

Mission and Vision Statements, 

147 

Symposium Announcement, 

227 

Helminths, 9, 36, 116, 130, 149, 

156, 177, 185, 190, 196, 204, 

219, 228, 242, 249, 256, 259, 

261, 265, 269, 272, 274, 277 

Hematozoa, 173 

Hemidactylus frenatus, 138 

Hemidactylus garnotii, 138, 149 

Hemidactylus turcicus, 149, 156 

Hemiphyllodactylus typus, 138 

Henle's snouted treefrog, 21 

Herotilapia multispinosa, 42, 76, 

190 

Heterandria bimaculatus, 196 

Heteromoxyuris deserti, 116 

Heteronchocotyle gymnurae sp. 

n., 91 

Heterophyidae gen. sp., 42 

Heterostrongylus heterostrongylus, 

274 

Hexabothriidae, 91 

Horses, 265 

Host specificity, 149 

House gecko, 138 

Hybopis boucardi, 196 

Hyla boans, 21 

Hyla brevifrons, 21 

Hyla calcarata, 21 

Hyla fasciata, 21 

Hyla granosa, 21 

Hyla koechlini, 21 

Hyla leali, 21 

Hyla leucophyllata, 21 

Hyla marmorata, 21 

Hyla parviceps, 21 

Hyla rhodopepla, 21 

Hyla schubarti, 21 

Hymenolepis diminuta, 261 

Hysterothylacium sp., 196 

Ictalunis balsanus, 196 

Ictalurus punctatus, 196 

Ilyodon whitei, 196 

Imitator robber frog, 21 

India, 87 

Indo-Pacific geckos, 138, 149 

Infectivity, 126, 256 

Insecta, 242 

Iraq, 108 

Ixodes affinis, 177 

Jaguar cichlid, 42, 76, 190 

Jaguar leaf frog, 21 

Jalisco, Mexico, 9, 196, 204 

Japan, 228 

Jeweled splitfin, 204 

Juvenile nematodes, 36 

Key, 236 

Kinosternon hirtipes, 9 

Koechlin's treefrog, 21 

Lacistorhynchus dollfusi, 21 

La Paz robber frog, 21 

Leal's treefrog, 21 

Lecithochirium sp., 21 

Lemdana wernaati, 130 

Lepidochelys olivacea, 1 

Lcpidodactylus aurcolincatus, 138 

Lepornis gibbosus, 236 

Lepomis macrochirus, 206 

Leptodactylus bolivianus, 21 

Leptodactylus leptodactyloides, 21 

Leptodactylus mystaceus, 21 

Leptodactylus pentadactylus, 21 

Leptodactylus petersii, 21 

Leptodactylus rhodonotus, 21 

Lerma chub, 204 

Lerma livebearer, 196, 204 

Lerma-Santiago river basin, Mexico, 

204 

Leucocytozoon simondi, 173 

Life cycle, 112 

Ligula intestinalis, 204 

Lithodytes lineatiis, 21 

Liza abu, 108 

Lobatostoma jungwirthi, 249 

Long-whiskered catfish, 190 

Los Angeles Harbor, U.S.A., 36 

Louisiana, U.S.A., 149, 156 

Lowland tropical bullfrog, 21 

Madre de Dios treefrog, 21 

Magellanic penguin, 277 

Magnivitellinum simplex, 190, 196 

Mammalia, 1, 97, 112, 116, 122, 

126, 134, 177, 259, 261, 265, 

272, 274 

Manaus slender-legged treefrog, 

21 

Manteriella sp., 36 

Marbled treefrog, 21 

Margotrema bravoae, 204 

Maritrema sp., 130 

Mastophorus dipodomis, 116 

Mediterranean geckos, 149, 156 

Mediterranean Sea, 91 

Megalodiscus temperatus, 269 

Melaleuca quinquenervia, 242 

Merriam's kangaroo rat, 116 


INDEX 289 

Mesa Central, Mexico, 9 

Mesa silverside, 204 

Mesocestoides sp., 164, 269 

Mesocricetus auratus, 112 

Metacercariae, 42, 126, 164, 259, 

269 

Mexican garter snake, 9 

Mexican molly, 196, 204 

Mexican mud turtle, 9 

Mexican tetra, 196 

Mexico, 1, 9, 52, 196, 204, 272 

Michoacan, Mexico, 9, 196, 204 

Microfilaria, 173 

Microphallus sp., 130 

Mississippi, U.S.A., 149 

Mohor gazelle, 134 

Mollusca, 126, 249 

Molly, 190 

Molting, 242 

Moniliformis moniliformis, 261 

Monogenea, 9, 76, 91, 196, 204 

Monogenoidea, 66, 87 

Morelos, Mexico, 196 

Morphology, 97, 156, 265, 277 

Mugilidae, 108 

Mus musculus, 126, 259 

Myrtaceae, 242 

Myxobolus cyprinicola, 236 

Myxobolus dechtiari, 236 

Myxobolus gibbosus, 236 

Myxobolus Hi, 236 

Myxobolus lipomicus, 236 

Myxobolus magnaspherus, 236 

Myxobolus osburni, 236 

Myxobolus paralintoni, 236 

Myxobolus poecilichthidis, 236 

Myxobolus uvuliferus, 236 

Myxozoa, 236 

Namibia, 97 

Napo's tropical bullfrog, 21 

Natalus mexicanus, 1 

Nayarit, Mexico, 204 

Nematoda, 9, 21, 42, 97, 116, 

130, 138, 164, 185, 190, 196, 

204, 228, 242, 261, 265, 269, 

272, 274, 277 

Neodiplostomum americanum, 130 

Neodiplostomum reflexum, 130 

Neoechinorhynchidae, 103, 108 

Neoechinorhynchus didelphis sp. 

n., 103 

Neoechinorhynchus golvani, 190, 

196, 204 

Neoechinorhynchus iraqensis sp. 

n., 108 

New books, 102, 107, 176, 255 

New combination, 87, 219


New genus, 87 

New geographical record, 9, 21, 

42, 52, 66, 76, 87, 122, 130, 

134, 138, 173, 177, 185, 190, 

196, 204, 236, 261, 265, 269, 

272, 274, 277 

New host record, 9, 21, 36, 42, 

52, 66, 76, 87, 112, 122, 130, 

134, 138, 164, 177, 185, 204, 

261, 272, 274, 277 

New Mexico, U.S.A., 116 

New species, 52, 76, 91, 103, 

108, 219, 228 

New synonymy, 236 

Nicaragua, 42, 76, 190 

Nippostrongylus brasiliensis, 261 

Northern leopard frog, 228 

Notocotylus sp., 261 

Notropis sallei, 204 

Nycticorax nycticorax, 1 

Oaxaca, Mexico, 196 

Obituary: 

W. Henry Leigh, 143 

Obituary Notice: 

Julius Feldmesser, 41 

Ocellated electric ray, 52 

Ochetosoma brevicaecum, 9 

Ochoterenella vellardi, 21 

Odocoileus virginianus, 177 

Oligogonotylus manteri, 42, 190 

Ommatobrephidae, 219 

Ondoorstepoort Helminthological 

Collection, 195 

Ontario, Canada, 236 

Oochoristica javaensis, 138, 149, 

156 

Oocyst, 122, 134 

Ophidascaris sp., 21 

Orientostrongylus cf. tenorai, 261 

Orinoco lime treefrog, 21 

Osteocephalus taurinus, 21 

Oswaldocruzia lopesi, 21 

Oswaldocruzia pipiens, 164, 269 

Otus asio, 130 

Pachitea robber frog, 21 

Parapharyngodon maplestoni, 138 

Parelaphostrongylus tennis, 177 

Parvitaenia cochlearii, 196 

Parvitaenia macropeos, 196 

Pastel cichlid, 42, 76, 190 

Pathology, 256 

Perciformes, 66, 87 

Perez's snouted frog, 21 

Pernambuco, Brazil, 66 

Peru, 21 

Peru robber frog, 21 

Peru white-lipped frog, 21 

Petenia splendida, 1 

Peter's frog, 21 

Phalacrocorax olivaceus, 1 

Philander opossum, 1 

Philippine Islands, 138 

Phrynohyas coriacea, 21 

Phrynohyas venulosa, 21 

Phyllomedusa atelopoides, 21 

Phyllomedusa pcdliata, 21 

Phyllomedusa tomopterna, 21 

Phyllomedusa vaillanti, 21 

Physaloptera sp., 21 

Physalopteroides venancioi, 21 

Pipa pipa, 21 

Pisces, 1, 36, 42, 66, 76, 87, 103, 

108, 112, 190, 196, 204, 236, 

249 

Plagiorchis sp., 130 

Platyhelminthes, 1 

Plethodon punctatus, 185 

Plethodon wehrlei, 185 

Pneumatophilus variabilis, 9 

Poecilia reticulata, 196 

Poecilia sphenops, 196, 204 

Poecilia veil/era, 42, 190 

Poeciliopsis gracilis, 196 

Poeciliopsis infans, 196, 204 

Poeciliopsis sp., 204 

Polymorphic brevis, 9, 204 

Polystornoidella oblonga, 9 

Porrocaecum depressmn, 130 

Porrocaecum sp., 21 

Porthole livebearer, 196 

Posthodiplostomurn minimum, 9, 

42, 196, 204 

Postorchigenes ovatus, 138 

Prescribed fire, 116 

Procamallanus (Spirocamallanus) 

neocaballeroi, 190 

Procamallanus (Spirocamallanus) 

rebecae, 190 

Prosthenhystera obesa, 190 

Prosthogonirnus ovatus, 130 

Proteocephalidea gen. sp., 42, 204 

Proteocephalus sp., 269 

Proteocephalus variabilis, 9 

Proterodiplostomidae gen. sp., 42 

Proterodiplostomum sp., 204 

Pseudis paradoxa, 21 

Pseudocapillaria tomentosa, 204 

Pterygodermatites dipodomis, 116 

Puebla, Mexico, 196 

Puerto Rico, 66 

Pumpkinseed sunfish, 236 

Quebec, Canada, 112 

Queretero, Mexico, 204 


Queensland, Australia, 242 

Raillietina celebensis, 261 

Raillietnema gubernaculatum, 21 

Rana clamitans, 269 

Rana clamitans melanota, 164 

Rana montezumae, 1 

Rana pipiens, 228 

Rana sphenocephala, 149 

Rana vaillanti, 1 

Ransom Trust Report, 248 

Rattus argiventer, 261 

Rattus losea, 261 

Rattus tanezumi, 261 

Rear-fanged water snake, 219 

Redescription, 236 

Redfin pickerel, 103 

Redside cichlid, 196 

Red-skirted treefrog, 21 

Red-snouted treefrog, 21 

Reptilia, 1, 9, 138, 149, 156, 219 

Reserva Cuzco Amazonico, 21 

Rhabdias ambystomae sp. n., 228 

Rhabdias americanus, 228 

Rhabdias bermani, 228 

Rhabdias fuscovenosa, 9 

Rhabdias ranae, 228 

Rhabdias tokyoensis, 228 

Rhabdiasidae, 228 

Rhabdochona canadensis, 196 

Rhabdochona kidderi kidderi, 190, 

196 

Rhabdochona lichtenfelsi, 196 

Rhabdochoma mexicana, 196 

Rhabdosynochus hargisi sp. n., 

66 

Rhabdosynochus hudsoni sp. n., 

66 

Rhabdosynochus rhabdosynochus, 

66 

Rhamdia nicaraguensis, 42, 190 

Ring-necked duck, 173 

Rio Mamore robber frog, 21 

River goby, 204 

Russia, 228 

Rusty treefrog, 21 

Saccocoelioides sogandaresi, 190, 

196 

Saccocoelioides spp., 190 

Salamanders, 185, 228 

Salamandrella kaiserlingii, 228 

Salmonidae, 112 

Salvelinus alpinus, 112 

Sarayacu treefrog, 21 

Scarthyla ostinodactyla, 21 

Sceloporus u. iindulatus, 149 

Schrankiana inconspicata, 21

Schrankiana larvata, 21 

Schrankiana schrankai, 21 

Schrankianella brasili, 21 

Schubart's Rondonia treefrog, 21 

Sciadicleithrum bicuensc sp. n., 

76 

Sciadicleithrum maculicaudae sp. 

n., 76 

Sciadicleithrum meekii, 76 

Sciadicleithrum mexicanum, 76 

Sciadicleithrum nicaraguense sp. 

n., 76 

Sciadicleithrum sp., 204 

Scinax garbei, 21 

Scinax icterica, 21 

Scinax pedromedinai, 21 

Scinax ruba, 21 

Sclerocleidoid.es gen. n., 87 

Sclerocleidoides etropli comb, n., 

87 

Scotland, 265 

Seasonal study, 164 

Sensory papillae, 249 

Serpinema trispinosum, 9, 42 

Sharpnose silverside, 204 

Shortfin silverside, 204 

Siberian newt, 228 

Sinaloan cichlid, 204 

Singhiatrema vietnamensis sp. n., 

219 

Skrjabinelazia machidai, 138 

South Africa, 97 

South American bullfrog, 21 

South American common toad, 21 

South Australian blue gum, 242 

Southern leopard frog, 149 

Southern red-backed vole, 122 

Spain, 134 

Spauligodon hemidactylus, 138 

Sphaenorhynchus lacteus, 21 

Spheniscus magellanicus, 211 

Spinitectus osorioi, 9 

Spinner dolphin, 272 

Spiny butterfly ray, 91 

Spirocamallanus sp., 36 

Spiroxys contorta, 9 

Spiroxys sp., 42, 196, 204 

Spiroxys susanae, 9 

Spirurids, 130 

Splendidofilaria fallisensis, 173 

Spot-legged poison frog, 21 

Spottail chub, 204 

Spottail killifish, 196 

Spotted salamander, 228 

Stenella longirostris, 272 

Stephana stomum californicum, 36 

Stephanostomum tristephanum, 36 

Stomylotrema vicarium, 130 

Strigea elegans, 130 

Strix varia, 130 

Strobilocephalus triangularis, 272 

Strongylidae, 97 

Strongyloides ratti, 261 

Strongyloides sp., 130 

Strongyloides venezuelensis, 261 

.Stump-toed gecko, 138 

Stunkardiella minima, 42 

Subulura forcipata, 130 

Subulura reclinata, 130 

Surinam golden-eyed treefrog, 21 

Surinam toad, 21 

Survey, 9, 21, 42, 76, 130, 138, 

164, 173, 177, 185, 190, 196, 

261, 265, 269 

vSwamp bloodwood, 242 

Swimming frog, 21 

Synhimantus laticeps, 130 

Syphacia muris, 261 

Szidatia taiwanensis comb, n., 

219 

Tadarida brasiliensis, 1 

Taxonomy, 9, 52, 66, 76, 87, 91, 

97, 103, 108, 149, 156, 204, 

219, 228, 236 

Tegument, 249 

Telorchis corti, 9 

Tetra, 42, 190 

Tetrameres microspinosa, 130 

Tetrameres strigiphila, 130 

Tetraphyllidea, 36, 52, 272 

Thailand, 138 

Thamnophis eques, 9 

Thamnophis melanogaster, 9 

Ticks, 177 

Tiger-striped leaf frog, 21 


INDEX 291 

Toady leaf frog, 21 

Toluca silverside, 204 

Tree gecko, 138 

Trematoda, 21, 126, 130, 138, 

164, 249, 256, 259, 261, 269, 

272 

Tribolium castaneum, 149 

Tridentoinfundibulum gobi, 265 

Trigonocotyle sp., 272 

Troschel's treefrog, 21 

Tunisia, 91 

Tyto alba, 130 

Ultrastructure, 242, 249 

Urocledoides cf. costaricensis, 

196 

Uvulifersp., 42, 196 

U.S.A., 36, 103, 116, 122, 132, 

149, 156, 164, 173, 185, 228, 

269, 274 

Valipora minuta, 196 

Variation, 156 

Veined treefrog, 21 

Vietnam, 219, 261 

Virginia opossum, 274 

Waltonella sp., 164 

Wehrle's salamander, 185 

West Virginia, U.S.A., 185 

White croaker, 36 

White-lined leaf frog, 21 

White-tailed deer, 177 

Wisconsin, U.S.A., 164, 228, 269 

Worm recovery, 126 

Wyoming, U.S.A., 122 

Xenotoca variatus, 204 

Yellow-lined smooth-scaled gecko, 

138 

Yellow-snouted treefrog, 21 

Yucatan molly, 42 

Yurirea alta, 204 

Zalophotrema pacificum, 212 

Zebra, 97 

Zoogonoides sp., 36

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VOLUME 68 JULY 2001 NUMBER 2 

CONTENTS 

(Continued from Front Cover) 

CONE, D. K. Supplemental Diagnosis ofMyxobolus gibbosus (Myxozoa), with a Taxonomic Review of 

Myxobolids from Lepomis gibbosus (Centrarchidae) in North America . 236 

GIBLIN-DAVIS, R. M., K. A. DAVIES, D. S. WILLIAMS, AND T. D. CENTER. Cuticular Changes in Fergusobiid 

Nematodes Associated with Parasitism of Fergusoninid Flies 242 

PAOLA, A., AND M. C. OAMBORENEA. Tegumentary infrastructure (SEM) of Preadult and Adult Lobatostoma 

jungwiithi Kritscher, 1974 (Trematoda: Aspidogastrea) _ 249 

RESEARCH NOTES 

MULLIGAN, S. K., J. E. HUFFMAN, AND B. FRIED. Infectivity and Comparative Pathology of Echinostoma 

caproni, Echinostoma revolution, and Echinostoma trivolvis (Trematoda) in the Domestic Chick 256 

FRIED, B., A. REDDY, AND C. D. BALFOUR. Excystation and Distribution of Metacercariae of Echinostoma 

caproni in ICR Mice ... 259 

PHAM, X.-D., C.-L. TRAN, AND H. HASEGAWA. Helminths Collected from Rattus spp. in Bac Ninh 

Province, Vietnam . 261 

LICHTENFELS, J. R., A. MCDONNELL, S. LOVE, AND J. B. MATTHEWS. Nematodes of the Tribe Cyathostominea 

(Strongylidae) Collected from Horses in Scotland 265 

YODER, H. R., J. R. COGGINS, AND J. C. RjEiNBOLD. Helminth Parasites of the Green Frog (Rana clamitans) 

from Southeastern Wisconsin, U.S.A 269 

AGUILAR-AGUILAR, R., R. G. MORENO-NAVARRETE, G. SALGADO-MALDONADO, AND B. VILLA-RAM!REZ. 

Gastrointestinal Helminths of Spinner Dolphins Stenella longirostris (Gray, 1828) (Cetacea: Delphinidae) 

Stranded in La Paz Bay, Baja California Sur, Mexico . 272 

MATEY, V E., B. I. KUPERMAN, J. M. KINSELLA, G. F. LLOYD, AND S. J. LANE. The Lung Nematodes 

(Metastrongyloidea) of the Virginia Opossum Didelphis virginiana in Southern California, U.S.A. 274 

DIAZ, J. I., G. T. NAVONE, AND F. CREMONTE. New Host and Distribution Records of Cosmocephahis 

obvelatus (Creplin, 1925) (Nematoda: Acuariidae), with Morphometric Comparisons 277 

ANNOUNCEMENTS 

EDITORS ' ACKNOWLEDGMENTS 15 5 

NOTICE OF DUES INCREASE 163 

FOURTH INTERNATIONAL CONGRESS OF NEMATOLOGY . . 172 

NEW 'BOOKS AVAILABLE 176,255 

ONDERSTEPOORT HELMINTHOLOGICAL COLLECTION . 195 

SYMPOSIUM: PARASITOLOGY IN SCIENCE AND SOCIETY , 227 

HELMINTHOLOGICAL SOCIETY OF WASHINGTON MEETING SCHEDULE 241 

REPORT ON THE BRAYTON H. RANSOM MEMORIAL TRUST FUND , 248 

LIFE MEMBERSHIP: JOHN S. MACKIEWICZ . ...... 282 

HONORARY MEMBERSHIP: GRAHAM C. KEARNE 282 

PRESENTATION OF 2000 ANNIVERSARY AWARD 283 

MINUTES OF THE MEETINGS OF THE HELMINTHOLOGICAL SOCIETY OF WASHINGTON 284 

AUTHOR INDEX FOR VOLUME 68 286 

KEY WORD AND SUBJECT INDEX FOR VOLUME 68 286 

MEMBERSHIP APPLICATION OF THE HELMINTHOLOGICAL SOCIETY OF WASHINGTON 292 

Date of publication, 31 July 2001 

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PRINTED BY ALLEN PRESS, INC., LAWRENCE, KANSAS 66044, U.S.A.

Comparative Parasitology 68(2) 2001 - Peru State College

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