Comparative Parasitology 67(2) 2000 - Peru State College

July 2000 Number 2 

Comparative Parasitology 

Formerly the 

Journal of the Helminthological Society of Washington 

CONTENTS 

KRITSKY, D. C., F. A. JIMENEZ-RUIZ, AND O. SEY. Diplectanids (Monogenoidea: Dactylogyridea) 

from the Gills of Marine Fishes of the Persian Gulf off Kuwait 145 

DAILEY, M. D., AND S. R. GOLDBERG. Langeronia burseyi sp. n. (Trematoda: Lecithodendriidae) 

from the California Treefrog, Hyla cadaverina (Anura: Hylidae), with 

Revision of the Genus Langeronia Caballero and Bravo-Hollis, 1949 165 

BOUAMER, S., AND S. MoRAND. Oxyuroids of Palearctic Testudinidae: New Definition 

'of the Genus Thaparia Ortlepp, 1933 (Nematoda: Pharyngodonidae), Redescription 

of Thaparia thapari thapari, and Descriptions of Two New Species 169 

MUZZALL, P. M. Parasites of Farm-Raised Trout in Michigan, U.S.A. . 181 

BULLARD, S. A., G. W. BENZ, R. M. OVERSTREET, E. H. WILLIAMS, JR., AND J. HEMDAL. 

Six New Host Records and an Updated List of Wild Hosts for Neobenedenia melleni 

(MacCallum) (Monogenea: Capsalidae) . _ 

DUCLOS, L. M., AND D. J. RICHARDSON. Hymenolepis nana in Pet Store Rodents 

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

Helminth Parasites from the Eastern American Toad, Bufo americanus americanus, 

from Southeastern Wisconsin, U.S.A 202 

MACHADO, P. M., S. C. DE ALMEIDA, G. C. PAVANELLI, AND R. M. TAKEMOTO. Ecological 

Aspects of Endohelminths Parasitizing Cichla monoculus Spix, 1831 (Perciformes: 

Cichlidae) in the Parana River near Porto Rico, State of Parana, Brazil 210 

LYONS, E. T, T. R. SPRAKER, K. D. OLSON, S. C. TOLLIVER, AND H. D. BAIR. Prevalence 

of Hookworms (Uncinaria lucasi Stiles) in Northern Fur Seal (Callorhinus ursinus 

Linnaeus) Pups on St. Paul Island, Alaska, U.S.A.: 1986-1999 218 

HASEGAWA, H., T. Doi, A. FUJISAKI, AND A. MIYATA. Life History of Spiroxys hanzaki 

Hasegawa, Miyata, et Doi, 1998 (Nematoda: Gnathostomatidae) 224 

BOCZON, K., AND B. WARGIN. Inducible Nitric Oxide Synthase in the Muscles of Trichinella 

sp.-Infected Mice Treated with Glucocorticoid Methylprednisolone 230 

FUJINO, T., T. SHINOHARA, K. FUKUDA, H. ICHIKAWA, T. NAKANO, AND B. FRIED. The 

Expulsion of Echinostoma trivolvis: Worm Kinetics and Intestinal Cytopathology 

in Jirds, Meriones unguiculatus „ •. 236 

DARAS, M. R., S. SISBARRO, AND B. FRIED. Effects of a High-Carbohydrate Diet on 

Growth of Echinostoma caproni in ICR Mice _ . 241 

KOGA, M., H. AKAHANE, R. LAMOTHE-ARGUMEDO, D. OSORIO-SARABIA, L. GARC{A-PRIETO, 

J. M. MARTINEZ-CRUZ, S. P. DlAZ-CAMACHO, AND K. NODA. Surface Ultrastructure 

of Larval Gnathostoma cf. binucleatum from Mexico ... ..... 244 

(Continued on Outside Back Cover) 

Copyright © 2011, The Helminthological Society of Washington

THE HELMINTHOLOGICAL SOCIETY OF WASHINGTON 

THE SOCIETY meets approximately five times per year for the presentation and discussion of papers 

in any and all branches of parasitology or related sciences. All interested persons are invited to attend. 

Persons interested in membership in the Helminthological Society of Washington may obtain application 

blanks in recent issues of COMPARATIVE PARASITOLOGY. A year's subscription to 

COMPARATIVE PARASITOLOGY is included in the annual dues of $25.00 for domestic membership 

and $28.00 for foreign membership. Institutional subscriptions are $50.00 per year. Applications for 

membership, accompanied by payments, may be sent to the Corresponding Secretary-Treasurer, Nancy 

D. Pacheco, 9708 DePaul Drive, Bethesda, MD 208(17, U:S,A. 

The HelmSoc internet home page is located at http://www.gettysburg.edu/~shendrix/helmsoc.html 

OFFICERS OF THE SOCIETY FOR 2000 

President: DENNIS J. RICHARDSON 

Vice President: LYNN K. CARTA 

Corresponding Secretary-Treasurer: NANCY D. PACHECO 

Recording Secretary: W. PATRICK CARNEY 

Archivist/Librarian: PATRICIA A. PILITT 

Custodian of Back Issues: J. RALPH LICHTENFELS 

Representative to the American Society of Parasitologists: ERIC P. HOBERG 

Executive Committee Members-at-Large: RALPH P. ECKERLIN, 2000 

WILLIAM E. MOSER, 2000 

ALLEN L.RICHARDS, 2001 

BENJAMIN M. ROSENTHAL, 2001 

Immediate Past President: ERIC P. HOBERG 

COMPARATIVE PARASITOLOGY 

COMPARATIVE PARASITOLOGY is published semiannually at Lawrence, Kansas by the 

Helminthological Society of Washington. Papers need not be presented at a meeting to be published in 

the journal. Publication of COMPARATIVE PARASITOLOGY is supported in part by the Brayton H. 

Ransom Memorial Trust Fund. 

MANUSCRIPTS should be sent to the EDITORS, Drs. Willis A. Reid, Jr., and Janet W. Reid, 6210 

Hollins Drive, Bethesda, MD 20817-2349, email: jwrassoc@erols.com. Manuscripts must be typewritten, 

double spaced, and in finished form. Consult recent issues of the journal for format and style. The 

original and three copies are required. Photocopies of drawings may be submitted for review purposes 

but glossy prints of halftones are required; originals will be requested after acceptance of the manuscript. 

Papers are accepted with the understanding that they will be published only in the journal. 

REPRINTS may be ordered from the PRINTER at the same time the corrected proof is returned to 

the EDITORS. 

AUTHORS' CONTRIBUTIONS to publication costs (currently S50/page for members, $100/page 

for non-members) will be billed by Allen Press and are payable to the Society. 

BACK ISSUES and VOLUMES of COMPARATIVE PARASITOLOGY are available, Inquiries concerning 

back volumes and current subscriptions should be directed to the business office. 

BUSINESS OFFICE. The Society's business office is at Lawrence, Kansas. All inquiries concerning 

subscriptions or back issues and all payments for dues, subscriptions, and back issues should be 

addressed to: Helminthological Society of Washington, % Allen Press, Inc., 1041 New Hampshire St., 

Lawrence, Kansas 66044, U.S.A. 

2000 

ROY C.-ANDERSON 

RALPH P. ECKERLIN 

ROBIN N. HUETTEL 

FUAD M. NAHHAS 

DANNY B. PENCE 

JOSEPH F. URBAN 

EDITORIAL BOARD 

WILLIS A. REID, JR. & JANET W. REID, Editors 

2001 

WALTER A BOEGER 

WILLIAM F. FONT 

DONALD FORRESTER 

J. RALPH LICHTENFELS 

JOHN S. MACKIEWICZ 

BRENT NICKOL 

© The Helminthological Society of Washington 2000 

ISSN 1049-233X 

2002 

DANIEL R. BROOKS 

HIDEO HASEGAWA 

SHERMAN S. HENDRIX 

JAMES E. JOY 

DAVID MARCOGLIESE 

DANTE S. ZARLENGA 

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


Comp. Parasitol. 

67(2), 2000 pp. 145-164 

Diplectanids (Monogenoidea: Dactylogyridea) from the Gills of 

Marine Fishes of the Persian Gulf off Kuwait 

DELANE C. KRITSKY,M F. AGUSTIN JiMENEZ-Ruiz,2 AND OTTO SEY3 

1 Department of Health and Nutrition Sciences, College of Health Professions, Box 8090, Idaho State 

University, Pocatello, Idaho 83209, U.S.A. (e-mail: kritdela@isu.edu), 

2 Laboratorio de Helmintologfa, Institute de Biologia, National Autonomous University of Mexico (UNAM), 

Apartado Postal 70-153, Ciudad de Mexico D.F., Mexico, and 

3 Department of Zoology, University of Kuwait, P.O. Box 5969, Safat 13060, Kuwait; current address: H-7633 

Pecs, Ercbanyasz u. 10, Hungary 

ABSTRACT: Seventeen species of Diplectanidae were collected from the gills of 17 species of marine fishes from 

the Persian Gulf off Kuwait. Lepidotrema kuwaitcnsis sp. n. from Terapon puta (Teraponidae), Lamellodiscus 

furcillatus sp. n. from Diplodus noct (Sparidae), and Protolamellodiscus senilobatus sp. n. from Argyrops spinifer 

and A. filamentosus (Sparidae) are described. Diplectanum cazauxi from Sphyraena jello and S. obtusata (Sphyraenidae) 

(new host and geographic records), D. sillagonum from Sillago siharna (Sillaginidae) (new geographic 

record), Pseudolarnellodiscus sphyraenae from Sphyraena chrysotaenia (Sphyraenidae) (new host and geographic 

records), and Calydiscoides flexuosus from Nemipterus peronii and N. bipunctatus (Nemipteridae) (new host 

and geographic records) are redescribed. An incidental geographic record for C. flexuosus on N. japonicus from 

the western coast of India is included. Ten diplectanid species from 8 hosts were unidentified for lack of sufficient 

specimens. Diplectanum longipenis (synonym: Squamodiscus longipenis) is transferred to Lepidotrema. Squamodiscus 

is removed from synonymy with Diplectanum and becomes a junior subjective synonym of Lepidotrema. 

Calydiscoides indianus (synonyms: Lamellospina Indiana and C. indicus) is a junior subjective synonym 

of C. flexuosus. 

KEY WORDS: Monogenoidea, monogenean, Diplectanidae, Calydiscoides flexuosus, Diplectanum cazauxi, Diplectanum 

sillagonum, Diplectanum sp., Lamellodiscus furcillatus sp. n., Lamellodiscus sp., Lepidotrema kuwaitensis 

sp. n., Lepidotrema longipenis comb, n., Protolamellodiscus senilobatus sp. n., Pseudolamellodiscus 

sphyraenae, Pseiidorhabdosynochus sp., Acanthopagrus berda, Acanthopagrus bifasciatus, Acanthopagrus latus, 

Argyrops filamentosus, Argyrops spinifer, Diplodus noct, Epinephelus areolatus, Epinephelus tauvina, Hemiramphus 

marginatus, Nemipterus bipunctatus, Nemipterus peronii, Otolithes argenteus, Sillago sihama, Sphyraena 

chrysotaenia, Sphyraena jello, Sphyraena obtusata, Terapon puta, Persian Gulf, Kuwait. 

A survey of the helminth parasites infesting described by Kritsky et al. (1986). Measurements, all 

marine fishes off the Kuwaiti coast by O.S. was in njicrometers, were made with a filar micrometer ac- 

_. . *nn~ ?n i- cording to procedures of Mizelle and Klucka (1953); 

conducted between October 1992 and December average measurements are followed by ranges and 

1996. Species of Diplectanidae (Monogenoidea) number (n) of specimens measured in parentheses; unwere 

found on the gills of 17 marine fishes rep- stained flattened specimens mounted in Gray and 

resenting the Hemiramphidae, Nemipteridae, Wess' medium were used to obtain measurements of 

Sciaenidae, Serranidae, Sillaginidae, Sparidae, the hap'oral sclerites and copulatory complex; other 

„ . . , , m • j rr,, . 

Sphyraemdae, and Teraponidae. The present pa- 

^ J f . . 

per includes descriptions and taxonomic considmeasurements 

were obtained from unflattened specit 

. , . „ ., t . , . v, . 

mens stained in Gomon s tnchrome and mounted in 

Canada balsam; the dimension of the pyriform ovary 

erations of 3 new and 5 previously described is the greatest width. Numbering of hook pairs follows 

species. the scheme proposed by Mizelle (1936; see Mizelle 

and Price, 1963). Type specimens of new species and 

Materials and Methods voucher specimens of previously described species 

, . , . i i , /- . i . «• 

Hosts were obtained from the local fish market, Ku- 

, , ,. . ,. , . . . . ' . 

wait, and examined directly tor helminth parasites. Diplectanids 

were removed from the gills of respective 

were deposited in the United States National Parasite 

_, „ . /T TOXT~,x r» i* -n A* , ^ ^ , 

Collection (USNPC), Beltsville, Maryland, and the 

, , - , ,, • r- i. TT • • AT i , n 

helminth collection of the University of Nebraska State 

hosts, fixed, and stored as described by Sey and Nah- Museum (HWML), Lincoln, Nebraska, U.S.A., as mhas 

(1997); vials containing the helminths were then dlcated m the respective species accounts. For cornshipped 

to Idaho State University. Methods of staining, Paratlve Purposes, the following specimens were exmounting, 

and illustration of diplectanids were those ammed: 3 voucher specimens of Lepidotrema tenue 

Johnston and Tiegs, 1922 (USNPC 63156); 4 voucher 

specimens of Lepidotrema bidyana Murray, 1931 

4 Corresponding author. (USNPC 63157); 5 voucher specimens of Lepidotrema 

145 


146 COMPARATIVE PARASITOLOGY, 67(2), JULY 2000 

angusta (Johnston and Tiegs, 1922) (USNPC 63158); 

holotype, 22 paratypes of Pseudolamellodiscus sphyraenae 

Yamaguti, 1953 (Meguro Parasitological Museum, 

Tokyo, Japan [MPM] 22556); holotype, numerous 

paratypes of Lamellodiscus convolutus Yamaguti, 

1953 (MPM 22558); holotype, 27 paratypes of Lamellodiscus 

flexuosus Yamaguti, 1953 (MPM 22557); 

holotype, 11 paratypes of Squamodisciis longipenis 

Yamaguti, 1934 (labeled as S. longiphallus) (MPM 

22564); and 37 voucher specimens of Calydiscoides 

flexuosus Yamaguti, 1953 (USNPC 89024). Host 

names and synonyms follow those provided by the 

FAO Fish Base at http://www.fao.org/waicent/faoinfo/ 

fisher y/fi shbase/fishbase .htm. 

Results 

A total of 17 species of Diplectanidae was 

found on 17 species of marine fishes collected 

off the Kuwaiti coast. Specimens of only 7 of 

the 17 diplectanid species were sufficient for 

identification and description. Ten unidentified 

diplectanids and their hosts are listed in Table 1. 

Class Monogenoidea Bychowsky, 1937 

Order Dactylogyridea Bychowsky, 1937 

Diplectanidae Monticelli, 1903 

Diplectanum cazauxi Oliver and Paperna, 

1984 

(Figs. 1-8) 

REDESCRIPTION (measurements of specimens 

from Sphyraena obtusata Cuvier, 1929, follow 

those from Sphyraena jello Cuvier, 1829 in 

brackets): Diplectaninae. Body 964 (729- 

1,080; n = 4) [824 (608-1,070; n = 4)] long, 

fusiform; greatest width 170 (123-242; n = 4) 

[156 (97-229; n = 4)] usually in posterior trunk 

at level of testis. Tegument smooth. Cephalic 

margin tapered; 2 terminal, 2 bilateral cephalic 

lobes poorly developed; head organs numerous; 

cephalic glands numerous in cephalic area, 2 bilateral 

groups posterolateral to pharynx. Eyes 4; 

members of posterior pair slightly larger, farther 

apart than anterior members; 1 anterior eye occasionally 

absent; granules small, ovate; accessory 

granules absent to numerous in cephalic region. 

Mouth subterminal, ventral to pharynx; 

pharynx 52 (39-68; n = 4) [42 (32-48; n = 4)] 

wide, ovate to subrectangular in dorsoventral 

view; esophagus short or nonexistent; intestinal 

ceca blind. Peduncle short to elongate. Haptor 

81-82 (n = 2) [70 (69-72; n = 3)] long, 127 

(113-140; n = 2) [130 (120-137; n = 3)] wide, 

bilaterally lobed; squamodiscs similar, each 49 

(36-60; n = 6) [50 (46-61; n = 7)] long, 77 

Copyright © 2011, The Helminthological Society of Washington 

(61-88; n = 6) [72 (64-86; n = 7)] wide, with 

17-19 concentric rows of dumbbell-shaped rodlets, 

each with anterior lightly sclerotized blunt 

spinelet. Ventral anchor 30 (29-32; n = 11) [31 

(29-32; n = 6)] long, with elongate deep root, 

knob-like superficial root, straight shaft, moderately 

long point extending slightly past level 

of tip of superficial root; anchor base 9 (8-10; 

n = 3) [7-8 (n = 1)] wide. Dorsal anchor 23 

(22-24; n = 10) [23 (22-25; n = 8)] long, with 

subtriangular base, slightly curved shaft, point 

extending past level of tip of superficial anchor 

root; anchor base 7-8 (n = 6) wide. Ventral bar 

72 (58-85; n = 10) [66 (62-76; n = 6)] long, 

subrectangular, with tapered ends, ventral 

groove; paired dorsal bar 42 (36-46; n — 10) 

[40 (37-44; n = 8)] long, spatulate medially. 

Hooks similar; each 10 (9-11; n = 19) [10 (9- 

11; n = 11)] long, with protruding thumb with 

slightly depressed tip, delicate point, shank; 

hook pair 1 lying medial to anchors on short 

haptoral peduncles, pairs 2—4, 6 submarginal on 

lateral haptoral lobes, pair 5 associated with distal 

shaft of ventral anchor, pair 7 on dorsal surface 

of lateral haptoral lobe; filamentous booklet 

(FH) loop shank length. Male copulatory organ 

41 (39-44; n = 4) [36 (31-40; n = 3)] long, 

weakly sclerotized, C shaped, with slightly enlarged 

base, nipple-like termination. Accessory 

piece absent. Testis 261 (185-300; n = 4) [207 

(129-292; n = 4)] long, 92 (70-130; n = 4) [82 

(65—105; n = 4)] wide, pyriform; course of vas 

deferens not observed; seminal vesicle a simple 

dilation of vas deferens, lying along body midline 

dorsal to vagina; 2 small prostatic reservoirs 

immediately anterior to male copulatory organ, 

saccate. Ovary 42 (31-56; n = 3) [40-41 (n = 

1)] wide, elongate pyriform, looping right intestinal 

cecum, lying transversely anterior to testis; 

oviduct elongate; ootype ventral, a small dilated 

portion of female duct; uterus delicate, extending 

along seminal vesicle; seminal receptacle not 

observed; vagina nonsclerotized, aperture sinistroventral 

near level of male copulatory organ; 

vitellaria throughout trunk, except absent in regions 

of major reproductive organs. 

HOSTS AND LOCALITY: Pickhandle barracuda, 

Sphyraena jello Cuvier, 1829 (Sphyraenidae): 

Persian Gulf off Kuwait (15 October 1993). Obtuse 

barracuda, Sphyraena obtusata Cuvier, 

1829 (Sphyraenidae): Persian Gulf off Kuwait (9 

July 1993). 

PREVIOUS RECORDS: Yellowtail barracuda,

Table 1. Unidentified diplectanids infesting marine fishes off Kuwait. 

Host 

Acanthopagrus berda 

(Forsskal, 1775) 

(Sparidae) 

Acanthopagrus bifaxciatus 


(Sparidae) 

Acanthopagrus latus 

(Houttnyn, 1782) 

(Sparidae) 

Diplodus noct 

(Valenciennes, 1830) 

(Sparidae) 

Epinephelus arcolatus 


(Serranidae) 

Epinephelus tauvina 


(Serranidae) 

Hemiramphus marginatus 


(Hemiramphidae) 

Otolithcs argenteus 

(Cuvier, 1830) 

(Sciaenidae) 

Date of collection 

30 November 1996 

10 May 1995 

10 October 1995 

28 March 1995 

23 March 1996 


29 July 1993 

16 June 1993 


15 June 1993 

10 March 1994 

8 May 1995 


5 April 1996 


Sphyraena flavicauda Riippell, 1838 (Sphyraenidae): 

Gulf of Aqaba (Golfe D'Aquaba [sic]), 

Gulf of Suez (Egypt), Indian Ocean off Malindi 

(Kenya) (all Oliver and Paperna, 1984). 

SPECIMENS STUDIED: 12 voucher specimens 

from S. jello, USNPC 89010, HWML 15023; 8 

voucher specimens from S. obtusata, USNPC 

89009. 

REMARKS: Diplectanum cazauxi is known 

only from species of barracuda (Sphyraenidae). 

Our report of this species on 5. jello and S. obtusata 

from the Persian Gulf represents new host 

and geographic records. The known geographic 

distribution of D. cazauxi currently includes the 

western Indian Ocean and adjacent regions including 

the northern gulfs of the Red Sea and 

the Persian Gulf. 

The original description of D. cazauxi is 

based on morphometrics of the squamodisc and 

sclerotized haptoral and copulatory structures. 

Although Oliver and Paperna (1984) mentioned 

that the ovary loops the right intestinal cecum, 

a symplesiomorphic feature for all members of 

the Diplectanidae, other details of the internal 

anatomy were not considered. Our redescription 

KRITSKY ET AL.—DIPLECTANIDS FROM KUWAIT 147 

Parasite 

Lamellodiscus sp. 1 






Pseudorhabdosynochus sp. 1 

Pseudorhabdosynochus sp. 2 


Diplectanum sp. 1 

Diplectanurn sp. 2 

USNPC no. 

89011 

89012 

89013 

89014 

89015 

89016 

89017 

89018 

89019 

89030 

89029 

89031 

89034 

89033 

89035 

89032 

adds information on soft-tissue features of the 

reproductive, digestive, and nervous systems. 

The morphometrics of the haptoral sclerites 

and squamodisc in our specimens are in general 

agreement with those reported by Oliver and 

Paperna (1984) in the original description of D. 

cazauxi. Mounting media (Gray and Wess' medium, 

Malmberg's medium, and Hoyer's medium) 

commonly used to visualize the sclerites of 

monogenoideans apply pressure on the specimen. 

In D. cazauxi, this pressure results in significant 

distortion of the lightly sclerotized male 

copulatory organ. The copulatory organs of D. 

cazauxi shown in Figure 11 of Oliver and Paperna 

(1984) are clearly distorted, as were our 

specimens mounted in Gray and Wess' medium. 

Such artifacts are minimized when specimens 

are mounted in Canada balsam, which does not 

result in significant coverslip pressure on the 

specimen (compare Fig. 4 with Fig. 11 of Oliver 

and Paperna, 1984). 

The copulatory complex, dorsal anchor, haptoral 

bars, and squamodisc of Diplectanum cazauxi 

closely resemble those of Laterocaecum 

pearsoni Young, 1969, suggesting that these 



Figures 1-8. Diplectanum cazauxi Oliver and Paperna, 1984. 1. Whole mount (composite, ventral; 

dorsal squamodisc not shown). 2. Ventral anchor. 3. Dorsal anchor. 4. Copulatory complex. 5. Dorsal bar. 

6. Hook. 7. Ventral bar. 8. Ventral view of haptor showing ventral squamodisc and positions of hook 

pairs. All figures are drawn to the 25-jjim scale, except Figures 1 and 8 (200-jjim and 50-u.m scales, 

respectively). 

species likely share a common evolutionary history. 

Laterocaecum was proposed by Young 

(1969) for a diplectanid collected from the obtuse 

barracuda, S. obtusata, from Moreton Bay, 

Queensland, Australia. Young (1969) differentiated 

the genus from other diplectanid genera 


by species possessing lateral diverticula of the 

intestinal ceca (lateral diverticula absent in all 

other species of Diplectanidae) and 12 (6 pairs) 

hooks in the adult. If D. cazauxi actually shares 

a phylogenetic history with L. pearsoni as suggested 

by their similar morphology and host

preferences, separation of Laterocaecum from 

Diplectanum may not be justified, and the 2 

unique characters presented by L. pearsoni may 

represent secondarily derived features within Diplectanum. 

We do not formally propose synonymy 

of the 2 genera at this time, however, because 

hypotheses on phylogenetic relationships 

within the Diplectanidae are lacking and Diplectanum 

may represent a paraphyletic group (see 

"Discussion"). Diplectanum cazauxi differs 

from L. pearsoni by having a knob-like superficial 

root on the ventral anchor (root elongate 

in L. pearsoni) and by possessing 7 pairs of 

hooks in the adult (6 pairs in L. pearsoni). 

Diplectanum sillagonum Tripathi, 1957 

(Figs. 9-15) 

REDESCRIPTION (Tripathi's [1957] original 

measurements and counts are in brackets following 

respective parameters of specimens from the 

Persian Gulf): Diplectaninae. Body 755 (694- 

815; n = 4) [623-1,058] long, fusiform, somewhat 

flattened dorsoventrally; greatest width 131 

(110-153; n = 4) [114-144] usually in anterior 

trunk near level of copulatory organ. Tegument 

smooth. Cephalic margin tapered; 2 terminal, 2 

bilateral cephalic lobes poorly developed; subspherical 

ventral pouch lying anterior to pharynx, 

opening to exterior via simple midventral 

pore. Head organs numerous; distributed in 3 

poorly defined groups; anterior posterior groups 

associated with respective cephalic lobes. Cephalic 

glands lateral to pharynx, extending posteriorly 

past level of esophageal bifurcation. 

Eyes 4; members of posterior pair larger, closer 

together than anterior members; granules small, 

ovate; accessory granules numerous, distributed 

throughout cephalic, anterior trunk regions. 

Mouth subterminal, ventral to pharynx; pharynx 

47 (40-53; n = 4) [41-49] wide, subspherical; 

esophagus short or absent; intestinal ceca blind. 

Peduncle short, broad. Haptor 124 (113-137; 

n = 4) [57] long, 159 (150-170; n = 4) [133- 

152] wide, bilaterally lobed; squamodiscs similar, 

each 73 (62-83; n = 12) [57-76] in diameter, 

subcircular, with 13—15 [11—15] concentric 

rows of dumbbell-shaped rodlets, each with anterior 

lightly sclerotized blunt spinelet. Ventral 

anchor 44 (38-50; n = 14) [49-53] long, with 

elongate roots (deep root longest), straight shaft, 

recurved point extending slightly past level of 

tip of superficial anchor root; anchor base 14 


(11-16; n = 8) wide. Dorsal anchor 40 (38-44; 

n = 13) [41-49] long, with subtriangular base, 

slightly curved shaft, recurved point extending 

past level of tip of superficial anchor root; anchor 

base 12 (10-14; n = 7) wide. Ventral bar 

74 (67-86; n = 10) [60-72] long, with tapered 

ends, ventral groove; median anterior constriction. 

Paired dorsal bar 69 (63-75; n = 11) [57- 

64] long, medial end expanded, bilobed. Hooks 

similar; each 12 (11—13; n = 29) long, with protruding 

thumb with slightly depressed tip, delicate 

point, slender shank; hook pair 1 at level of 

tips of ventral bar, medial to anchors; pairs 2—4, 

6, 7 submarginal in lateral haptoral lobes; pair 5 

associated with distal ventral anchor shaft; FH 

loop shank length. Male copulatory organ 34 

(30-39; n = 6) [41-45] long, a sigmoid tube 

originating from ring-like sclerotized base, with 

fine recurved tip. Accessory piece variable, 

comprising 2 articulated subunits, 1 subunit with 

bilobed proximal end articulating to other subunit. 

Testis 70 (69-71; n = 2) [38-53 X 76- 

152] in diameter, subspherical; course of vas deferens 

not observed; seminal vesicle a simple 

elongate dilation of vas deferens, lying along 

body midline dorsal to seminal receptacle; prostatic 

reservoir saccate, posterior to male copulatory 

organ, frequently containing granules 

only at anterior end. Ovary 57 (42-71; n = 2) 

[38 X 57] wide, elongate pyriform, looping right 

intestinal cecum, lying transversely anterior to 

testis; oviduct elongate; ootype, uterus not observed; 

seminal receptacle ovate, originating 

from short tubular vagina; vagina with small 

bead-like sclerotization having cupped proximal 

end; vaginal aperture sinistral; vitellaria throughout 

trunk, except absent in regions of major reproductive 

organs. 

HOSTS AND LOCALITY: Silver sillago, Sillago 

sihama (Forsskal, 1775) (Sillaginidae): Persian 

Gulf off Kuwait (31 December 1993, 18 April 

1996). 

PREVIOUS RECORDS: Sillago sihama: Chandipore, 

Chilka Lake, Puri, all Bay of Bengal, 

India (Tripathi, 1957). Sillago sihama: Burdekin 

River, Duyfken Point, Point Samson, and Darwin, 

Australia; Phuket and Bang Saen, Thailand; 

Gendering and Kula Lumpur, Malaysia; Bali, Indonesia; 

Aberdeen market and Sai Kung, Hong 

Kong; Ring Ring, Kapa Kapa, and Sinapa, Paupua 

New Guinea; and Madras, India (all Hayward, 

1996). Slender sillago, Sillago attenuata 

McKay, 1985: Ras Lanura, Saudi Arabia (Hay- 



9 

Figures 9-15. Diplectanum sillagonum Tripathi, 1957. 9. Whole mount (composite, body ventral, haptor 

dorsal), showing position of hook pairs. 10. Hook. 11. Copulatory complex. 12. Ventral bar. 13. Dorsal 

bar. 14. Ventral anchor. 15. Dorsal anchor. All figures are drawn to the 25-u.m scale, except Figure 9 

(200-|xm scale). 

10 


15

ward, 1996). Vincent's sillago, Sillago vincenti 

McKay, 1980: Kavanad, Kerala, India (Hayward, 

1996). 

SPECIMENS STUDIED: 14 voucher specimens, 

USNPC 89007, 89008, HWML 15022. 

REMARKS: Diplectanum sillagonum was described 

by Tripathi (1957) from the gills of 

S. sihama from western coastal localities on the 

Bay of Bengal, India. His description of this species 

is of marginal value for species determination. 

Nonetheless, the original drawings of the 

copulatory complex, anchors, bars, and whole 

mount, while diagrammatic, strongly suggest 

conspecificity with our collection from the Persian 

Gulf. Persian Gulf specimens were obtained 

from the same host species as that of the type 

series, and respective measurements of specimens 

from the Persian Gulf and India are comparable. 

However, the types of D. sillagonum 

were not available for confirmation. General 

morphology of the sclerotized haptoral structures 

and copulatory complex generally corresponds 

to figures of this species offered by Hayward 

(1996). However, Hay ward (1996) did not 

mention the presence of the midventral pouch 

located anterior to the pharynx in his redescription 

of the species. 

Lepidotrema kuwaitensis sp. n. 

(Figs. 16-23) 

DESCRIPTION: Diplectaninae. Body 504 

(452—603; n — 8) long, fusiform; greatest width 

105 (90-121; n = 9) near body midlength. Tegument 

smooth. Cephalic margin tapered; 2 terminal, 

2 bilateral cephalic lobes poorly developed; 

3 bilateral pairs of head organs with anterior, 

posterior pairs associated with respective 

cephalic lobes; cephalic glands not observed. 

Eyes 4, equidistant; members of posterior pair 

larger than anterior members; anterior eyes frequently 

absent, 1 or both posterior eyes occasionally 

dissociated; granules small, ovate, numerous 

accessory granules at eye level. Mouth 

subterminal, ventral to pharynx; pharynx 23 

(19-26; n = 9) wide, ovate to subspherical; 

esophagus short to absent; intestinal ceca blind. 

Peduncle short to elongate. Haptor 83 (65-100; 

n = 9) long, 139 (124-151; n = 9) wide, bilaterally 

lobed. Squamodiscs similar, each 37 (26- 

43; n = 4) long, 40 (27-48; n = 6) wide, subcircular, 

with 8-10 concentric rows of dumbbellshaped 

rodlets becoming progressively more 


delicate in posterior rows; 2-4 rows (layers) of 

elongate delicate spinelets wrap around posterior 

margin of both squamodiscs, spinelets frequently 

absent. Ventral anchor 42 (39-45; n = 25) 

long, with elongate roots (deep root longest), 

evenly curved shaft with terminal indentation at 

articulation with recurved point; point extending 

slightly past level of tip of superficial anchor 

root; anchor base 9 (7-11; n = 13) wide. Dorsal 

anchor 37 (32-40; n = 23) long, with narrow 

base, long deep root, curved shaft, point extending 

past level of tip of superficial root of anchor 

base; anchor base 7 (6-9; n = 11) wide. Ventral 

bar 59 (52-66; n = 21) long, with tapered ends, 

ventral groove; paired dorsal bar 55 (47—60; n 

= 23) long, spatulate, with posteromedial spine. 

Hook 10 (9-12; n = 37) long, with protruding 

thumb with slightly depressed end, delicate 

point, shank dilated slightly in some specimens. 

Hook pair 1 lying medial to haptoral lobes, posterior 

to bars; pairs 2-4, 7 in lateral haptoral 

lobes; pair 5 associated with shaft of ventral anchor; 

pair 6 at level of or just anterior to dorsal 

bar. FH loop shank length. Male copulatory organ 

68 (60-74; n =11) long, a sigmoid tube 

with wall of varying thickness along length, 

acute tip. Accessory piece absent. Testis subspherical, 

54 (42-65; n = 9) in diameter; course 

of vas deferens not observed; seminal vesicle a 

simple dilation of vas deferens, lying along body 

midline dorsal to ootype; prostatic reservoirs 3, 

saccate; anterior prostatic vesicles bilateral to 

male copulatory organ, with prostatic ducts 

fused prior to entering base of male copulatory 

organ via common duct; posterior reservoir caudal 

to male copulatory organ, apparently emptying 

independently into base of male copulatory 

organ. Ovary 23 (19—25; n = 3) wide, pyriform, 

anterodorsal to testis, looping right intestinal 

cecum; oviduct elongate; ootype ventral, a 

small dilated portion of female duct; uterus delicate, 

extending anteriorly to left of prostatic reservoirs; 

seminal receptacle not observed; vaginal 

aperture sinistroventral, with circular muscular 

rim; vagina funnel-shaped, narrowing to 

short tube; vagina with proximally thickened 

walls; vitellaria throughout trunk, except absent 

in regions of major reproductive organs. Egg 

83-84 (n = 1) long, 56-57 (n = 1) wide, ovate, 

with short proximal filament. 

TYPE HOST: Small-scaled terapon, Terapon 

puta (Cuvier, 1829) (Terapontidae). 



Figures 16—23. Lepidotrema kuwaitensis sp. n. 16. Whole mount (composite, ventral; dorsal squamodisc 

not shown), showing positions of hook pairs. 17. Copulatory complex. 18. Hooks. 19. Enlargement of worm 

at level of reproductive organs (composite, ventral). 20. Dorsal bar. 21. Ventral bar. 22. Ventral anchor. 

23. Dorsal anchor. All figures are drawn to the 25-u.m scale, except Figures 16 and 19 (100-u.m and 50ujm 

scales, respectively). 

TYPE LOCALITY: Persian Gulf off Kuwait (9 

July 1993, 15 October 1993, 26 March 1996). 

INFECTION SITE: Gills. 

DEPOSITED SPECIMENS: Holotype, USNPC 

89020; 25 paratypes, USNPC 89021, 89022, 

89023, HWML 15025. 

ETYMOLOGY: This species is named for the 

country of Kuwait. 

REMARKS: The primary distinguishing feature 

of Lepidotrema Johnston and Tiegs, 1922, is the 

presence of groups of elongate spinelets forming 

fan-like structures on the posterior portions of the 


squamodiscs (Oliver, 1987). The genus currently 

includes 6 species, all from freshwater teraponids 

in Australia: Lepidotrema therapon Johnston and 

Tiegs, 1922; L. angusta; L. bidyana; Lepidotrema 

fidiginosum Johnston and Tiegs, 1922; Lepidotrema 

simplex Johnson and Tiegs, 1922; and L. tentie. 

Our finding of L. kuwaitensis on T. puta (Teraponidae) 

in the Persian Gulf is the first report of 

a member of Lepidotrema from a marine host. 

Existing descriptions of the 6 freshwater species 

are of marginal value for comparison with L. kuwaitensis, 

and most species require redescription

(see Johnston and Tiegs, 1922; Murray, 1931; 

Young, 1969). 

In L. kuwaitensis, the posterior spinelets are 

delicate (or frequently absent, an apparent artifact 

resulting from deterioration of the specimen before 

fixation) and resemble those of L. angusta 

as depicted by Young (1969). These species are 

easily separated by the comparative morphology 

of the copulatory complexes (sigmoid in L. kuwaitensis; 

coiled with about 1 ring in L. angusta). 

Examination of the types of Diplectanum longipenis 

(Yamaguti, 1934) Yamaguti, 1963 

(=Squamodiscus longipenis Yamaguti, 1934), 

confirmed that L. kuwaitensis shares many features 

(general morphology and arrangement of 

the sclerotized haptoral and copulatory sclerites 

and internal reproductive organs) with this species 

and may be more closely aligned to it than 

to those from fresh water. While staining procedures 

used by Yamaguti (1934) did not allow 

us to see spinelets near the posterior margin of 

the squamodisc in D. longipenis, similarities in 

the morphology of sclerotized structures and the 

general organization of the reproductive organs 

suggest that the 2 species are congeneric. Thus, 

we propose the transfer of D. longipenis to Lepidotrema 

as L. longipenis (Yamaguti, 1934) 

comb. n. Squamodiscus Yamaguti, 1934, is removed 

from synonymy with Diplectanum and 

becomes a junior subjective synonym of Lepidotrema. 

Lepidotrema kuwaitensis differs from 

L. longipenis by having delicate anchors (base 

of dorsal anchor in L. kuwaitensis narrow; broad 

in D. longipenis) and by the number of rodlet 

rows in the squamodisc (8—10 rows in L. kuwaitensis; 

18-21 in D. longipenis). 

Pseudolamellodiscus sphyraenae Yamaguti, 

1953 

(Figs. 24-34) 

REDESCRIPTION: Diplectaninae. Body 1196 

(1020-1354; n = 17) long, flattened dorsoventrally; 

greatest width 244 (196-289; n = 16) 

usually in anterior trunk. Trunk with anterior 

dextroventral sclerite, posterior dextroventral 

sclerite, sinistroventral spined pit. Anterior dextroventral 

sclerite 58 (48-72; n = 26) long, with 

lobulate base, rod-shaped distal end protruding 

from small ventral pore, spined; number of 

spines variable. Posterior dextroventral sclerite 

37 (32-45; n = 27) long, spatulate, with incised 

distal margin; sinistroventral pit blind, with 4—6 


spines, opening ventrally via small aperture 

through tegument; tips of spines usually protruding 

through pore. Tegument smooth. Cephalic 

margin tapered; cephalic lobes poorly developed; 

head organs numerous along anterolateral 

margins of cephalic area; cephalic glands 

posterolateral to pharynx. Eyes 4; members of 

posterior pair larger, slightly farther apart than 

anterior members; 1 member of each pair occasionally 

absent; granules small, irregular; accessory 

granules uncommon in cephalic region. 

Mouth subterminal, ventral to anterior portion of 

pharynx; pharynx 61 (53-70; n = 19) wide, 

elongate, ovate; esophagus short to nonexistent; 

intestinal ceca blind. Peduncle broad. Haptor 

337 (260-421; n = 18) wide, 114 (93-148; 

n = 18) long, bilaterally lobed; squamodiscs 

similar, each 61 (47-70; n = 17) long, 249 

(200-310; n = 17) wide, with approximately 45 

longitudinal parallel rows of dumbbell-shaped 

spines in anterior portion of squamodisc; posterior 

portion with numerous spine-like scales. 

Ventral anchor 41 (36-44; n = 25) long, with 

elongate deep root, knob-like superficial root, 

slightly curved shaft, recurved point extending 


base 11 (10—12; n = 3) wide. Dorsal anchor 

33 (31-36; n = 31) long, with short deep 

root, triangular superficial root perpendicular to 

anchor base, curved shaft, point extending past 

level of tip of superficial root of anchor base; 

anchor base 9 (8-11; n = 6) wide. Ventral bar 

268 (218-328; n = 22) long, narrowed medially, 

ends tapered, recurved anteriorly; ventral groove 

present. Paired dorsal bar 69 (59-87; n = 25) 

long, club-shaped. Hooks similar; each 10-11 (n 

= 26) long, with protruding depressed thumb, 

delicate point, shank. Hook pair 1 submarginal, 

lying posterior to bars near base of haptoral 

lobes; pairs 2-7 located on lateral haptoral 

lobes; FH loop shank length. Male copulatory 

organ 33 (31-35; n = 9) long, with large base, 

bent shaft, acute bent tip, subbasal pointed projection. 

Accessory piece absent. Common genital 

pore absent; male genital pore lying ventrally 

to left of body midline slightly posterior to male 

copulatory organ; uterine pore ventral, slightly 

posterior to level of male genital pore, somewhat 

dextral to body midline. Testis 165 (144-184; n 

= 15) long, 81 (59-98; n = 16) wide, ovate; 

course of vas deferens not observed; 2 seminal 

vesicles simple dilations of vas deferens; proximal 

vesicle elongate, fusiform, lying along mid- 



5 1 34 

Figures 24-34. Pseudolamellodiscus sphyraenae Yamaguti, 1953. 24. Whole mount (composite, ventral; 

dorsal squamodisc not shown), showing positions of hook pairs. 25. Anterior dextroventral sclerite. 26. 

Copulatory complex. 27. Posterior dextroventral sclerite. 28. Posterodorsal accessory sclerite (lateral view). 

29. Sinistroventral spinous cavity (lateral view). 30. Hook. 31. Dorsal bar. 32. Dorsal anchor. 33. Ventral 

anchor. 34. Ventral bar. All figures are drawn to the 25-u.m scale, except Figures 24 and 34 (200-jim and 

50-fjim scales, respectively). 

line of body posterior to male copulatory organ; 

distal vesicle anterior to male copulatory organ, 

short, pyriform; prostatic reservoir saccate, anterior 

to male copulatory organ. Ovary 81 (62- 

107; n = 16) wide, forming lobed cap on anterior 

margin of testis, with proximal sinistral loop 

before extending around right intestinal cecum; 

oviduct narrowing to small tube before joining 


slightly expanded ootype; uterus delicate, extending 

to right of body midline; vaginal aperture 

sinistroventral; vagina tubular, frequently 

containing apparent spermatophore, joining 

small seminal receptacle lying to left of ootype; 

vitellaria throughout trunk, except absent in regions 

of reproductive organs. 

HOST AND LOCALITY: Yellowstrip barracuda,

Sphyraena chiysotaenia Klunzinger, 1884 

(Sphyraenidae): Persian Gulf off Kuwait (16 October 

1996). 

PREVIOUS RECORDS: Sphyraena sp.: Macassar, 

Celebes (Yamaguti, 1953). Great barracuda, 

Sphyraena barracuda (Walbaum, 1972): Nosy 

Be, Madagascar (Rakotofiringa and Maillard, 

1979). 

SPECIMENS STUDIED: 34 voucher specimens, 

USNPC 89028, HWML 15020. 

REMARKS: While Yamaguti (1953) did not 

adequately describe the haptoral sclerites, male 

copulatory organ, and trunk sclerites of Pseudolamellodiscus 

sphyraenae, our examination of 

the holotype and paratypes of this species confirmed 

that our specimens were conspecific with 

P. sphyraenea. In the account of P. sphyraenea 

from Madagascar by Rakotofiringa and Maillard 

(1979), the morphology of the haptoral and 

trunk sclerites were also not presented, but their 

figure of the trunk region, which includes a 

small drawing of the male copulatory organ, 

clearly supports their identification. However, 

both Yamaguti (1953) and Rakotofiringa and 

Maillard (1979) confused the vagina with the 

uterus. This error is supported by some of our 

specimens that contained a spermatophore in the 

tube that these authors described as the "uterus" 

and a developing egg in the tube they labeled 

"vagina." 

The slide containing the types of P. sphyraenea 

includes the holotype, 23 paratypes, and 

several fragments of specimens. Included in the 

23 paratypes are 2 specimens clearly of an undescribed 

Pseudolamellodiscus species, characterized 

by having 1 large ventral trunk sclerite 

with a bifurcated, foliated proximal end. 

Lamellodiscus furcillatus sp. n. 

(Figs. 35-42) 

DESCRIPTION: Lamellodiscinae. Body 1,092 

(924-1,294; n = 4), long, fusiform; greatest 

width 207 (183-226; n = 4), usually in posterior 

trunk near level of testis. Tegument smooth. Cephalic 

margin tapered; 2 terminal, 2 bilateral cephalic 

lobes poorly developed; 3 bilateral pairs 

of head organs with anterior, posterior pairs associated 

with respective cephalic lobes; cephalic 

glands posterolateral to pharynx. Eyes 4; equidistant; 

members of posterior pair larger than 

anterior members; anterior eyes occasionally absent; 

granules ovate, variable in size; accessory 


granules common in cephalic region. Mouth 

subterminal, ventral to pharynx; pharynx 54 

(46-60, n = 4) wide, ovate to subspherical; bilateral 

pair of prepharyngeal (buccal) glands anterior 

to pharynx; esophagus short to nonexistent; 

intestinal ceca blind. Peduncle broad. Haptor 

163 (148-180; n = 4) wide, 111 (104-117; 

n = 4) long, bilaterally lobed; lobes short. Lamellodiscs 

similar; each 58 (52-62; n = 4) long, 

42 (40-46; n — 4) wide, ovate, with 10 lamellar 

rings; anterior (deep) lamella forming complete 

ring; intermediate lamellae superficially incomplete, 

medially indented; posterior (superficial) 

lamella indented, complete. Ventral anchor 58 

(54-61; n — 8) long, with elongate deep root, 

short depressed superficial root, evenly curved 

shaft, recurved point; point extending slightly 


base 11-12 (n = 2) wide. Dorsal anchor 48 

(45-52; n = 8) long, with elongate deep root, 

erect knob-like superficial root, evenly curved 

shaft, nonrecurved point; anchor base 14-15 

(n = 2) wide. Ventral bar 76 (70-82; n = 8) 

long, plate-like, with ends constricted subterminally, 

ventral groove. Paired dorsal bar 62 

(56-68; n = 8) long, morphologically complex, 

broad. Hooks similar; each 12 (11-13; n = 7) 

long, with protruding slightly depressed thumb, 

delicate point, shank; FH loop shank length. 

Hook pair 1 submedial at level of posterior margin 

of ventral bar; pairs 2-4 submarginal in lateral 

haptoral lobes; pair 5 associated with ventral 

anchor shafts; pairs 6, 7 dorsal at level of tip of 

deep root of ventral anchor. Male copulatory organ 

60 (58-65; n = 7) long, a sigmoid tube with 

acute recurved tip. Accessory piece 53 (49—58; 

n = 4) long, with subterminal elongate branch. 

Testis subspherical, 111 (110-113; n = 2) in diameter, 

course of vas deferens not observed; 

seminal vesicle a simple dilation of vas deferens, 

lying to left of body midline dorsal to seminal 

receptacle; prostatic reservoir saccate, lying anterior 

to copulatory complex. Ovary 65 (64—67; 

n = 2) wide, elongate pyriform, diagonal, looping 

right intestinal cecum, overlapping testis; 

oviduct elongate; ootype ventral, a small dilated 

portion of female duct; uterus delicate, seminal 

receptacle small. Vaginal aperture sinistral; vagina 

short, frequently containing apparent subspherical 

spermatophore. Vitellaria throughout 

trunk, except absent in regions of reproductive 

organs. 



Figures 35-42. Lamellodiscus furcillatus sp. n. 35. Whole mount (composite, dorsal; ventral lamellodisc 

not shown). 36. Dorsal bar. 37. Ventral bar. 38. Copulatory complex. 39. Ventral anchor. 40. 

Hook. 41. Dorsal anchor. 42. Dorsal view of haptor showing dorsal lamellodisc and positions of hook 

pairs. All figures are drawn to the 25-fjim scale, except Figures 35 and 42 (500-u,m and 50-fj.m scales, 


TYPE HOST: Red Sea seabream, Diplodus 

noct (Valenciennes, 1830) (Sparidae). 


October 1995, 23 March 1996). 


DEPOSITED SPECIMENS: Holotype, USNPC 

89036; 7 paratypes, USNPC 89037, HWML 

15024. 

ETYMOLOGY: The specific name is from Lat- 


in (furcillatus = a small fork) and refers to the 

accessory piece of the copulatory complex. 

REMARKS: Lamellodiscus furcillatus sp. n. 

resembles Lamellodiscus baeri Oliver, 1974, in 

the morphology of the paired dorsal bars and 

general morphology of the copulatory complex. 

Oliver's (1974) description of L. baeri from the 

common seabream, Pagrus pagrus (Linnaeus, 

1758), Sparidae, is brief and does not include

figures of the internal anatomy (whole mount), 

hooks, or lamellodisc. However, L. furcillatus is 

easily differentiated from L. baeri by the presence 

of a nonrecurved point of the dorsal anchor 

(point of dorsal anchor recurved in L. baeri). 

Calydiscoides flexuosus (Yamaguti, 1953) 

Young, 1969 

(Figs. 43-52) 

REDESCRIPTION (Table 2 for measurements): 

Lamellodiscinae. Body long, fusiform; greatest 

width usually at level of testis. Tegument 

smooth. Cephalic margin tapered; 2 terminal, 2 

bilateral cephalic lobes poorly developed; 3 bilateral 

groups of head organs with anterior, posterior 

groups associated with respective cephalic 

lobes; cephalic glands posterolateral to pharynx. 

Eyes 4; members of posterior pair larger, usually 

closer together than anterior members; 1 member 

of anterior pair occasionally absent; granules 

usually ovate, variable in size; accessory granules 

common in cephalic region. Mouth subterminal, 

ventral to pharynx; pharynx ovate to subspherical; 

esophagus short; intestinal ceca blind. 

Peduncle broad. Haptor bilaterally lobed; lamellodiscs 

similar, each with 10 "telescoping" lamellae, 

posterior lamellae incomplete forming 

posterior superficial opening. Ventral anchor 

with elongate roots (superficial root longest) 

usually overlying one another (Fig. 51), evenly 

curved shaft, point recurved, not reaching level 

of tip of superficial anchor root. Dorsal anchor 

with short deep root, triangular superficial root, 

curved shaft, point extending past level of tip of 

superficial root. Ventral bar with tapered ends 

directed anterolaterally, ventral groove. Paired 

dorsal bar with bilobed medial end. Hooks similar; 

each with protruding thumb with slightly 

depressed end, delicate point, shank; hook pair 

1 lying medial to ventral anchor at level of anterior 

margin of ventral bar, pairs 2 (anterior), 3 

lateral to ventral lamellodisc, pairs 4, 6 submarginal 

in haptoral lobe, pair 5 associated with 

ventral anchor shaft, pair 7 lateral to dorsal lamellodisc; 

FH loop nearly shank length. Male 

copulatory organ, accessory piece nonarticulated. 

Male copulatory organ C-shaped, variably 

sclerotized, with acute termination, 2 subterminal 

branches embedded in wall of genital atrium 

present or absent. Accessory piece variable, flattened. 

Testis ovate; course of vas deferens in relation 

to gut not observed; vas deferens tortuous 


(Fig. 46), with anterior loop, expanded to form 

seminal vesicle; prostatic reservoir not observed. 

Ovary elongate pyriform, looping right 

intestinal cecum, lying transversely to diagonally 

anterodorsal to testis; oviduct elongate; ootype 

an expanded portion of female duct, surrounded 

by numerous glands; uterus with thick 

wall, ventral to proximal portion of vas deferens, 

extending dorsal to anterior loop of vas deferens. 

Vaginal aperture sinistroventral; vagina variable, 

funnel-shaped, narrowing to short tortuous tube; 

seminal receptacle absent, or represented by 

small expansion of vaginal duct prior to emptying 

into female duct; vaginal funnel with sclerotized 

clasp-like wall. Vitellaria coextensive 

with gut, absent in regions of reproductive organs. 

SYNONYMS: Lamellodiscus flexuosus Yamaguti, 

1953; Lamellospina Indiana Karyakarte 

and Das, 1978; Calydiscoides indicus Venkatanarsaiah 

and Kulkarni, 1980. 

HOST AND LOCALITIES: Notchedfin threadfin 

bream, Nemipterus peronii (Valenciennes, 1830) 

(originally identified as N. tolu [Valenciennes, 

1830]) and Delagoa threadfin bream, Nemipterus 

bipunctatus (Valenciennes, 1830) (originally 

identified as N. delagoae Smith, 1941) (Nemipteridae): 

Persian Gulf off Kuwait (31 December 

1993, 10 January 1994, respectively). Japanese 

threadfin bream, Nemipterus japonicus 

(Bloch, 1791) (Nemipteridae): Port of Okha, 

Gujarat, India. 

PREVIOUS RECORDS: Ornate threadfin bream, 

Nemipterus hexodon (Quoy and Gaimard, 1824) 

(originally identified as Synagris taeniopterus 

(Valenciennes, 1830): Macassar, Celebes (Yamaguti, 

1953). Nemipterus japonicus: Ratnagiri, 

west coast, Maharashtra, India (Karyakarte and 

Das, 1978); Kakinada, Bay of Bengal, India 

(Venkatanarsaiah and Kulkarni, 1980). 

SPECIMENS STUDIED: 18 voucher specimens 

from N. peronii, USNPC 89025, HWML 

15019; 14 voucher specimens from N. bipunctatus, 

USNPC 89026; 37 voucher specimens 

from N. japonicus, USNPC 89024. 

REMARKS: Yamaguti (1953) described Lamellodiscus 

flexuosus from the gills of Synagris taeniopterus 

(=Nemipterus hexodon) collected at 

Macassar, Celebes. Young (1969) transferred this 

helminth to Calydiscoides Young, 1969, based on 

the presence of "telescoping lamellae" in the lamellodisc, 

and Oliver (1987) recognized Young's 

reassignment of L. flexuosus to Calydiscoides. In 



Figures 43-52. Calydiscoides flexuosus (Yamaguti, 1953) Young, 1969. 43. Whole mount (composite, 

ventral; dorsal lamellodisc not shown), showing positions of hook pairs. 44, 45. Copulatory complexes. 46. 

Enlargement of reproductive organs (composite, ventral). 47. Dorsal bar. 48. Ventral bar. 49. Hook. 50. 

Dorsal anchor. 51, 52. Ventral anchors. All figures are drawn to the 25-fj.m scale, except Figures 43 and 

46 (200-jJiin and 50-(xm scales, respectively). 

addition, Oliver (1987) transferred Lamellospina 

indiana Karyakarte and Das, 1978, to Calydiscoides 

as C. indianus, considered Lamellospina 

Karyakarte and Das, 1978, a junior synonym of 

Calydiscoides, and placed C. indicus Venkatanarsaiah 

and Kulkarni, 1980, in synonymy with C. 


52 

indianus. Present findings support Young's (1969) 

and Oliver's (1987) taxonomic proposals. Our examination 

of the holotype and paratypes of L. 

flexuosus and new voucher specimens of C. flexuosus 

collected from the Indian coast and Kuwait 

confirmed that L. flexuosus Yamaguti, 1953, and


Table 2. Comparative measurements (in micrometers) of Calydiscoides flexuosus (Yamaguti, 1953) 

Young, 1969, from 3 species of Nemipterus (Nemipteridae) from the Persian Gulf and Indian Ocean. 

Body 

Length 

Width 

Haptor 

Length 

Width 

Lamellodisc 

Length 

Width 

Pharynx 

Width 

Copulatory organ 

Length 

Accessory piece 

Length 

Dorsal anchor 

Length 

Base width 

Ventral anchor 

Length 

Base width 

Bar length 

Dorsal 

Ventral 

Hook 

Length 

Germarinum 

Width 

Testis 

Length 

Width 

N. peronii Kuwait 

771 (675-843; n = 8) 

142 (110-162; n= 11) 

94 (78-103; /; = 10) 

107 (97-114; /; = 10) 

49 (37-60; n = 10) 

41 (34-46; n = 10) 

42 (35-48; /; 

32 (29-37; n = 6) 

21 (17-26; n = 6) 

35 (32-40; n = 14) 

11 (10-12; n = 4) 

46 (43-48; n = 14) 

21(1 6-26; n = 12) 

45 (39-54; n = 14) 

54 (45-65; // = ID 

12 (11-13; n = 9) 

28 (24-34; n = 3) 

149 (123-177; 

63 (42-76; n 

= 11) 

n = 9) 

= 9) 

N. bipunctatus Kuwait 

776 (680-987; n = 9) 

134 (107-162: n = 9) 

102 (78-128; n = 9) 

104 (88-1 16; n = 7) 

49 (43-58; n = 7) 

37 (31-45; n - 6) 

38 (33-44; n = 8) 

36 (32-39; n = 7) 

19(1 5-24; n = 5) 

35 (30-39; n - 13) 

10 (9-1 1; n = 

46 (43-52; n = 14) 

22 (19-25; /; = 10) 

43 (37-51; n = 14) 

52 (47-58; n = ID 

11 (10-13; // = 6) 

29 (26-33; n = 3) 

149 (121-178; 

70 (51-89; n 

Calydiscoides indianus (Karyakarte and Das, 

1978) Oliver, 1987, and its synonyms Lamellospina 

Indiana Karyakarte and Das, 1978, and Calydiscoides 

indicus Venkatanarsaiah and Kulkarni, 

1980, are synonyms of C. flexuosus (Yamaguti, 

1953) Young, 1969. 

Specimens of Calydiscoides flexuosus from 

India, Kuwait, and the Celebes are morphologically 

indistinguishable. However, we did observe 

some differences in dimensions of the 

body and haptoral sclerites (Table 2). Specimens 

from the Celebes were somewhat smaller than 

those from India, while those from Kuwait were 

intermediate in size. These differences are not 

considered sufficient to separate the collections 

: 2) 

n = 9) 

= 9) 

N. japonicus India 

893 (781-1,044; n = 17) 

136 (100-187; n = 19) 

126 (113-142; n = 12) 

121 (91-149; n '= 

8) 

68 (54-76; n = 

42 (36-53; n = 

48 (23-56; n = 14) 

36 (29-48; n = 

11 (9-14; n = 

49 (42-56; n = 

21 (18-25; n = 

48 (38-66; n = 

56 (49-64; n = 

12-13 (/i = 

41 (37-45; n = 

13) 

16) 

27) 

10) 

30) 

13) 

18) 

10) 

13) 

: 4) 

183 (149-219; n = 9) 

79 (62-93; n = 9) 

N. hcxodon Celebes 

528 (411-796; 

74 (62-9 1 ; n 

88 (69-113; n = 17) 

93 (76-119; n = 16) 

43 (31-56; n = 15) 

31 (27-36; n = 20) 

23 (19-27; n = 18) 

27 (25-29; n = 19) 

9 (8-10; n = 

39 (35-43; n = 21) 

17 (15-20; n = 3) 

37 (34-42; n = 25) 

43 (39-48: n = 16) 

11 (10-12; n = 9) 

— 

— 

n = 16) 

= 20) 

: 2) 

into distinct species, and could result from effects 

of different environmental and host factors 

on the parasite. All previous descriptions of this 

species lack detail and clarity of the morphological 

features necessary to identify the species; 

our redescription provides details of the morphology 

of the sclerotized parts of the haptor 

and copulatory complex. 

Protolamellodiscus senilobatus sp. n. 

(Figs. 53-60) 

DESCRIPTION (measurements of specimens 

from A. filamentosus follow those from the type 

host in brackets): Lamellodiscinae. Body 

1,065 (720-1318; n = 8) [714 (673-755; n = 



Figures 53-60. Protolamellodiscus senilobatus sp. n. 53. Whole mount (composite, dorsal; ventral lamellodisc 

not shown). 54. Hook. 55. Copulatory complex. 56. Ventral anchor. 57. Ventral bar. 58. Dorsal 

bar. 59. Dorsal anchor. 60. Dorsal view of haptor showing dorsal lamellodisc and positions of hook pairs 

(ventral lamellodisc not shown). All figures are drawn to the 25-fJim scale, except Figures 53 and 60 (100fxm 

and 50-u.m scales, respectively). 


60

2)] long, slender, fusiform; greatest width 185 

(120-240; n = 9) [164 (148-179; n = 2)] at 

level of testis. Tegument smooth. Cephalic margin 

narrow; 2 terminal, 2 bilateral cephalic lobes 

poorly developed; 3 bilateral pairs of head organs 

with anterior, posterior pairs associated 

with respective cephalic lobes; cephalic glands 

posterolateral to pharynx. Eyes 4; members of 

posterior pair slightly larger, closer together than 

anterior members; anterior pair frequently absent; 

granules irregular, variable in size; accessory 

granules common in cephalic region. 

Mouth subterminal, ventral to pharynx; pharynx 

73 (61-89, n = 10) [60 (51-70; n = 2)] wide, 

ovate or somewhat truncated posteriorly; esophagus 

short to nonexistent; intestinal ceca blind. 

Peduncle narrow, elongate. Haptor 206 (169- 

235; n = 8) [155 (150-161; n = 2)] wide, 111 

(104-117; n = 8) [84 (79-89; n = 2)] long, with 

3 bilateral pairs of lobes containing respective 

hook pairs 2, 3, 4 near apices; anterior lobes 

about half the length of more posterior lobes; 

lamellodiscs similar, each 44 (37-53; n = 10) 

[37 (35-39; n = 2)] long, 32 (29-38; n = 10) 

[30 (29-31; n = 2)] wide, with 1 complete, 8 

incomplete lamellae lacking medial indentation; 

lamellae appear to telescope somewhat in dorsoventral 

view. Ventral anchor 45 (38-49; n = 

11) [42-43 (n = 1)] long, with elongaite roots 

(deep root longest), evenly curved shaft, point 

acutely recurved not reaching level of tip of superficial 

root; base 14 (9-16; n = 8) wide. Dorsal 

anchor 41 (37-44; n = 17) [35-36 (n = 1)] 

long, with elongate deep root, short thickened 

superficial root, straight shaft, point reaching 

past level of tip of superficial root; base 9 (8- 

10; n = 13) wide. Ventral bar 41 (34-47; n = 

17) [36 (34-38; n = 2)] long, plate-like, with 

short knob-like ends; dorsal bar 40 (35-46; n 

= 23) [36 (34-38; n = 3)] long, with medial 

bend, spinous projection at proximal end. 

Hooks similar; each 10 (9-11; n = 28) [9-10 

(n = 3)] long, with protruding slightly depressed 

thumb, delicate point, shank; hook pair 

1 lying near base of ventral anchor; pairs 2, 3, 

4 at apices of respective haptoral lobes; pair 5 

posterior to ends of ventral bar; pair 6 near 

point of dorsal anchor; pair 7 near base of dorsal 

anchor. FH loop nearly shank length. Copulatory 

complex comprising articulated male 

copulatory organ, accessory piece. Male copulatory 

organ 45 (38-53; n = 22) [42-43 (n = 

1)] long, a curved heavily sclerotized tube with 

KRITSKY ET AL.~DIPLECTANIDS FROM KUWAIT 161 

subterminal recurved spine, distal loop terminating 

broadly; base of male copulatory organ 

lacking sclerotized margin. Accessory piece 28 

(18-34; n = 16) [32-33 (n = 1)] long, comprising 

flattened proximal portion, bifurcating 

near midlength to terminally acute elongately 

striated branch, spatulate branch frequently 

folded upon itself distally. Testis 107 (101-113; 

n = 2) long, 52 (48-55; n = 2) wide, ovate; 

vas deferens looping left intestinal cecum; seminal 

vesicle fusiform, simple dilation of vas deferens, 

lying slightly to left of body midline; 

prostatic reservoir saccate, lying anterior to 

copulatory complex. Ovary 47 (44-56; n = 5) 

wide, pyriform, looping right intestinal cecum, 

lying transversely to diagonally anterior to testis; 

oviduct elongate; ootype, uterus not observed; 

vaginal aperture sinistrodorsal, submarginal; 

vagina short, nonsclerotized, with proximal 

chamber containing apparent spermatophore, 

opening into medial seminal receptacle; 

vitellaria dense throughout trunk, except absent 

in regions of reproductive organs. One egg (deformed 

during mounting) infrequently present 

in uterus, with short proximal filament. 

TYPE HOST: King soldierbream, Argyrops 

spinifer (Forsskal, 1775) (Sparidae). 


January 1994). 


OTHER RECORD: Soldierbream, Argyrops filamentosus 

(Valenciennes, 1830) (Sparidae): Persian 

Gulf off Kuwait (18 October 1995). 

SPECIMENS STUDIED: Holotype, USNPC 

89005; 28 paratypes from A. spinifer, USNPC 

89006, HWML 15021; 3 voucher specimens 

from A. filamentosus, USNPC 89027. 

ETYMOLOGY: The specific name is from Latin 

(sen/i — six + lobat/o = lobe) and refers to 

the 6 bilateral lobes of the haptor. 

REMARKS: Oliver (1987) recognized 3 species 

of Protolamellodiscus from hosts of 3 marine 

teleost families: Protolamellodiscus serranelli 

(Euzet and Oliver, 1965) Oliver, 1969, from 

the comber, Serranus cabrilla (Linnaeus, 1758), 

the brown comber, Serranus hepatus (Linnaeus, 

1758), and the painted comber, Serranus scriba 

(Linnaeus, 1758), Serranidae; Protolamellodiscus 

convolutus (Yamaguti, 1953) Oliver, 1987, 

from N. hexodon, Nemipteridae; and Protolamellodiscus 

raibauti Oliver and Radujkovic, 

1987, from the annular seabream, Diplodus annularis 

(Linnaeus, 1758), Sparidae. The fourth 



species, P. senilobatus sp. n., occurs on sparid 

hosts (Argyrops spp.)- The new species most 

closely resembles P. raibauti in the comparative 

morphology of the copulatory complex but differs 

from this species by possessing a subterminal 

spine arising from the male copulatory organ, 

3 bilateral pairs of haptoral lobes (lobes 

lacking in P. raibauti), a flattened subrectangular 

ventral bar (bar rod-shaped in P. raibauti), 

and each dorsal bar with a proximal spine (see 

Oliver and Radujkovic, 1987). Protolamellodiscus 

senilobatus differs from P. serranelli in the 

comparative morphology of the copulatory complex. 

While Yamaguti's (1953) description of 

P. convolutus lacks details of the sclerotized 

structures of the haptor and copulatory complex, 

P. senilobatus is distinguished from this species 

by possessing 3 bilateral pairs of haptoral lobes. 

Oliver and Radujkovic (1987) described the 

vagina of P. raibauti as opening sublaterally on 

the left side of the body. In P. senilobatus, the 

vaginal aperture is submarginal on the sinistrodorsal 

body surface, midway between the ovary 

and copulatory complex. In P. senilobatus, the 

vas deferens loops the left intestinal cecum, 

while Euzet and Oliver (1965) reported the vas 

deferens to be intercecal in P. serranelli. Oliver 

and Radujkovic (1987) did not observe the 

course of the vas deferens relative to the intestine 

in P. raibauti. Confirmation of these 2 characters 

as potential diagnostic features of Protolamellodiscus 

is required. 

Members of Protolamellodiscus Oliver, 1969, 

and Calydiscoides Young, 1969, are characterized, 

in part, by having a ventral and a dorsal 

lamellodisc, each with several concentric unpaired 

lamellae, with the most anterior lamella 

forming a complete circle. Calydiscoides is, in 

part, diagnosed by the presence of telescoping 

lamellae. Depending on the orientation of the 

lamellodisc when examined microscopically, 

specimens of P. senilobatus occasionally show 

that the deeper lamellae telescope, although not 

to the extent exhibited in described species of 

Calydiscoides. While outside the scope of the 

present study, it is possible that Protolamellodiscus 

and Calydiscoides are synonyms. Further 

study of all species in these genera combined 

with a phylogenetic analysis is necessary to clarify 

synonymy and/or validity of the genera. 

Discussion 

In his revision of the Diplectanidae, Oliver 

(1987) divided the family into 4 subfamilies 


based primarily on the morphology and presence/absence 

of the accessory adhesive organs 

of the haptor. He recognized the Diplectaninae 

Monticelli, 1903 ("squamodiscs" composed of 

concentric rows of sclerotized rodlets): Lamellodiscinae 

Oliver, 1969 ("lamellodiscs" composed 

of concentric lamellae); Rhabdosynochinae 

Oliver, 1987 (lateral "placodiscs" unarmed); 

and Murraytrematoidinae Oliver, 1982 

(accessory adhesive organs absent). 

Oliver (1987) removed the then monotypic 

Rhamnocercinae Monaco, Wood, and Mizelle, 

1954, from the Diplectanidae, elevated it to familial 

level, and placed it in the poorly supported 

superfamily Heterotesioidea Euzet and DOS- 

SOU, 1979 (see Kritsky and Boeger, 1989), apparently 

because some previous descriptions of 

species of Rhamnocercus stated that the intestinal 

ceca are "apparently" united posterior to the 

gonads (Hargis, 1955, in R. bairdiella Hargis, 

1955; subsequently by Luque and lannacone 

[1991] in R. oliveri Luque and lannacone, 1991). 

However, Monaco et al. (1954) and Seamster 

and Monaco (1956) did not mention the intestine 

in the respective descriptions of R. rhamnocercus 

Monaco, Wood, and Mizelle, 1954, and 

R. stichospinus Seamster and Monaco, 1956. 

Luque and lannacone (1991) stated that the intestinal 

ceca end blindly in Rhamnocercoides 

menticirrhi Luque and lannacone, 1991, and 

Rhamnocercus stelliferi Luque and lannacone, 

1991. It appears that errors have been made concerning 

the morphology of the gut in some species 

of Rhamnocercinae, and the value of this 

character in determining familial relationships is 

limited. Along with Diplectaninae, Lamellodiscinae, 

Rhabdosynochinae, and Murraytrematoidinae, 

we tentatively consider the Rhamnocercinae 

a member of the Diplectanidae, based on 

general haptoral and internal morphology. However, 

these subfamilies all lack evolutionary support 

(phylogenetic analyses are lacking), and 

some or all may be unnatural. 

With the exception of Diplectanum Diesing, 

1858, and Lamellodiscus Johnston and Tiegs, 

1922, all diplectanid genera are defined by derived 

autapomoiphic features, suggesting that 

Diplectanum and Lamellodiscus are unnatural 

(paraphyletic) and currently serve as "catchall" 

groups for species lacking obvious derived characters. 

Kritsky and Boeger (1989) and Kritsky 

and Kulo (1992) discussed the probability of the

creation of paraphyletic taxa when new taxa are 

based primarily on autapomorphic features. 

Oliver (1987) considered Diplectanwn to include 

species having a squamodisc composed of 

concentric U-shaped rows of rodlets. Other diplectanine 

genera were diagnosed with characters 

thought to be lacking in Diplectanwn, such 

as closed circular rows of rodlets (Cycloplectanum 

Oliver, 1968 [=Pseudorhabdosynochus Yamaguti, 

1958, see Kritsky and Beverley-Burton 

1986]), divergent rows of rodlets (Heteroplectanum 

Rakotofiringa, Oliver, and Lambert, 

1987), lateral intestinal diverticula (Latericaecum 

Young, 1969), a row of elongate spines posterior 

to the squamodisc (Lepidotrema Johnston 

and Tiegs, 1922), 1 "squamodisc" (Monoplectanum 

Young, 1969), modified anchors (Pseudodiplectanum 

Tripathi, 1957), and parallel rodlets 

(Pseudolamellodiscus Yamaguti, 1953). 

Similarly, Oliver (1987) defined Lamellodiscus 

by species having paired (apparently incomplete) 

lamellae forming the lamellodiscs. The remaining 

genera in the Lamellodiscinae include 

forms with the following features absent in species 

of Lamellodiscus'. Calydiscoides Young, 

1969, with species having unpaired telescoping 

lamellae; Furnestinia Euzet and Audoin, 1959, 

with species lacking 1 "lamellodisc"; Protolamellodiscus 

Oliver, 1969, with species having 

closed or "O-shaped" lamella in the lamellodisc; 

and Telegamatrix Ramalingam, 1955, with 

a reproductive appendix containing the copulatory 

complex and vagina. Some of these genera 

might not be valid, as suggested by the apparent 

close relationship of Diplectanum cazauxi with 

Laterocaecum pearsoni and Protolamellodiscus 

senilobatus with species of Calydiscoides (see 

remarks under D. cazauxi and P. senilobatus). 

That Diplectanum and Lamellodiscus are paraphyletic 

is supported by observations on specimens 

in the present study. Both genera include 

species with varying characters, which were not 

considered generic features by Oliver (1987), 

but that could be used to determine monophyletic 

groups in the 2 genera. Features such as 

presence/absence of an accessory piece, position 

of the vaginal aperture, and morphology of the 

copulatory complex, among others, may have 

value in determining monophyletic groupings in 

these genera. In Lepidotrema, which is characterized 

by species possessing a posterior shield 

of elongate spines as its autapomorphic character, 

at least 1 species, L. longipenis, apparently 


lacks these structures. Thus, even some of the 

unique characters defining some of these genera 

(posterior spinous shield in Lepidotrema; gut diverticula 

in Laterocaecum) may not be valid for 

defining monophyletic groups within the Diplectanidae. 

Acknowledgments 

The authors are grateful to Dr. J. R. Lichtenfels 

(USNPC) and Dr. J. Araki (MPM) for allowing 

access to type and voucher specimens in 

their care. Dr. N. Agarwal, Department of Zoology, 

University of Lucknow, Lucknow, India, 

contributed specimens of Calydiscoides flexuosus 

from the western Indian coast for use in the 

present study. 

Literature Cited 

Euzet, L., and G. Oliver. 1965. Lamellodiscus serranelli 

n. sp. (Monogenea) parasite de Teleosteens 

du genre Serranus. Annales de Parasitologie Humaine 

et Comparee 40:261-264. 

Hargis, W. J. 1955. Monogenetic trematodes of Gulf 

of Mexico fishes. Part III. The superfamily Gyrodactyloidea. 

Quarterly Journal of the Florida 

Academy of Sciences 18:33-47. 

Hay ward, C. J. 1996. Revision of diplectanid monogeneans 

(Monopisthocotylea, Diplectanidae) in 

sillaginid fishes, with a description of a new species 

of Monoplectanum. Zoologica Scripta 25: 

203-213. 

Johnston, T. H., and O. W. Tiegs. 1922. New gyrodactyloid 

trematodes from Australian fishes, together 

with a reclassification of the super-family 

Gyrodactyloidea. Proceedings of the Linnean Society 

of New South Wales 47:83-131. 

Karyakarte, P. P., and S. R. Das. 1978. A new 

monogenetic trematode, Lamellospina Indiana 

n. gen., n. sp. (Monopisthocotylea: Diplectanidae) 

from the marine fish, Nemipterus japonicus (Gunther) 

in India. Rivista di Parassitologia 39:19-22. 

Kritsky, D. C., and M. Beverley-Burton. 1986. The 

status of Pseudorhabdosynochm Yamaguti, 1958, 

and Cycloplectanum Oliver, 1968 (Monogenea: 

Diplectanidae). Proceedings of the Biological Society 

of Washington 99:17-20. 

, and W. A. Boeger. 1989. The phylogenetic 

status of the Ancyrocephalidae Bychowsky, 1937 

(Monogenea: Dactylogyroidea). Journal of Parasitology 

75:207-211. 

, and S.-D. Kulo. 1992. Schilbetrematoides 

pseudodactylogyrus gen. et sp. n. (Monogenoidea, 

Dactylogyridae, Ancyrocephalinae) from the gills 

of Schilbe intermedium (Siluriformes, Schilbeidae) 

in Togo, Africa. Journal of the Helminthological 

Society of Washington 59:195-200. 

, V. E. Thatcher, and W. A. Boeger. 1986. 

Neotropical Monogenea. 8. Revision of Urocleidoides 

(Dactylogyridae, Ancyrocephalinae). Pro- 



ceedings of the Helminthological Society of 

Washington 53:1-37. 

Luque, J. L., and J. lannacone. 1991. Rhamnocercidae 

(Monogenea: Dactylogyroidea) in sciaenid 

fishes from Peru, with description of Rhamnocercoides 

menticirrhi n. gen., n. sp. and two new species 

of Rhamnocercus. Revista de Biologia Tropical 

39:193-201. 

Mizelle, J. D. 1936. New species of trematodes from 

the gills of Illinois fishes. American Midland Naturalist 

17:785-806. 

, and A. R. Klucka. 1953. Studies on monogenetic 

trematodes. XIV. Dactylogyridae from 

Wisconsin fishes. American Midland Naturalist 

49:720-733. 

, and C. E. Price. 1963. Additional haptoral 

hooks in the genus Dactylogyrus. Journal of Parasitology 

49:1028-1029. 

Monaco, L. H., R. A. Wood, and J. D. Mizelle. 1954. 

Studies on monogenetic trematodes. XVI. Rhamnocercinae, 

a new subfamily of Dactylogyridae. 

American Midland Naturalist 52:129-132. 

Murray, F. V. 1931. Gill trematodes from some Australian 

fishes. Parasitology 23:492-506. 

Oliver, G. 1974. Nouveaux aspects du parasitisme des 

Diplectanidae Bychowsky, 1957 (Monogenea, 

Monopisthocotylea) chez les Teleosteens Perciformes 

des cotes de France. Comptes Rendus de 

1'Academic des Sciences (Paris, Serie D) 279: 

803-805. 

. 1987. Les Diplectanidae Bychowsky, 1957 

(Monogenea, Monopisthocotylea, Dactylogyridea) 

Systematique: biologie: ontogenie: ecologie: essai 

de phylogenese. Doctor of Science Thesis, 

1'Universite des Sciences et Techniques du Languedoc, 

Academic de Montpellier, France, 433 pp. 

, and I. Paperna. 1984. Diplectanidae Bychowsky, 

1957 (Monogenea, Monopisthocotylea), 

parasites de Perciformes de Mediterranee orientale, 

de la mer Rouge et de 1'ocean Indien. Bul- 

Obituary Notice 

MICHAEL J. PATRICK 

March 9, 1962-March 10, 2000 

Elected to Membership in 1989 


letin du Museum National d'Historic Naturelle, 

Paris, 4th series, 6 (section A, No. l):49-65. 

, and B. Radujkovic. 1987. Protolamellodiscus 

raibauti n. sp., une nouvelle espece de Diplectanidae 

Bychowsky, 1957 (Monogenea, Monopisthocotylea) 

parasite de Diplodus annularis 

(Linnaeus, 1758) (Sparidae). Annales de Parasitologie 

Humaine et Comparee 62:209-213. 

Rakotofiringa, S., and C. Maillard. 1979. Helminthofaune 

des Teleostei de Madagascar: revision du 

genre Pseudolamellodiscus Yamaguti, 1953 

(Monogenea). Annales de Parasitologie (Paris) 54: 

507-518. 

Seamster, A., and L. H. Monaco. 1956. A new species 

of Rhamnocercinae. American Midland Naturalist 

55:180-183. 

Sey, O., and F. M. Nahhas. 1997. Digenetic trematodes 

of marine fishes from the Kuwaiti coast of 

the Arabian Gulf: Family Monorchiidae Odhner, 

1911. Journal of the Helminthological Society of 


Tripathi, Y. R. 1957. Studies on the parasites of Indian 

fishes. II. Monogenea, family: Dactylogyridae. 

Indian Journal of Helminthology 7:5—24. 

Venkatanarsaiah, J., and T. Kulkarni. 1980. New 

monogenetic trematode of the genus Calydiscoides 

Young, 1969 from the gills of Neinipterus japonicus. 

Proceedings of the Indian Academy of 

Parasitology 1:20-22. 

Yamaguti, S. 1934. Studies on the helminth fauna of 

Japan. Part 2. Trematodes of fishes, 1. Japanese 

Journal of Zoology 5:249-541. 

. 1953. Parasitic worms mainly from Celebes. 

Part 2. Monogenetic trematodes of fishes. Acta 

Medicinae Okayama 8:203-256 (with 9 plates). 

Young, P. C. 1969. Some monogenoideans of the 

family Diplectanidae Bychowsky, 1957 from Australian 

teleost fishes. Journal of Helminthology 43: 

223-254.


67(2), 20(K) pp. 165-168 

Langeronia burseyi sp. n. (Trematoda: Lecithodendriidae) from the 

California Treefrog, Hyla cadaverina (Anura: Hylidae), with Revision 

of the Genus Langeronia Caballero and Bravo-Hollis, 1949 

MURRAY D. DAiLEY1-3 AND STEPHEN R. GOLDBERG2 

1 The Marine Mammal Center, Marin Headlands, Sausalito, California, U.S.A. 94965 

(e-mail: daileym@tmmc.org) and 

2 Department of Biology, Whittier College, Whittier, California, U.S.A. 90608 (e-mail: 

sgoldberg @ whittier.edu) 

ABSTRACT: Langeronia burseyi sp. n. (Trematoda: Lecithodendriidae), a new trematode from the small intestine 

of Hyla cadaverina Cope, 1866, is described and illustrated. One (0.03%) of 36 adult specimens of H. cadaverina 

collected from Orange County, California, U.S.A., harbored 83 specimens of L. burseyi sp. n. Langeronia bursevi 

sp. n. is distinguished from all other species in the genus by body size, location of the cirrus, length of the ceca, 

placement of the vitellaria, and the shape of the excretory bladder. This is the first report of a species of 

Langeronia from a member of the Hylidae. An emended diagnosis and key to the genus Langeronia are presented. 

KEY WORDS: Digenea, Lecithodendriidae, Langeronia burseyi, new species description, taxonomy, California 

treefrog, Hyla cadaverina, Orange County, California, U.S.A. 

The taxonomic statuses of the genera Langeronia 

Caballero and Bravo-Hollis, 1949, and 

Loxogenes Stafford, 1904, have been the subject 

of much controversy. Caballero and Bravo-Hollis 

(1949) erected the genus Langeronia for a 

new species, Langeronia macrocirra, from the 

northern leopard frog, Rana pipiens Schreber, 

1782, in Mexico. A second species, Langeronia 

provitellaria, was described by Sacks (1952) 

from the Florida leopard frog, Rana sphenocephala 

Cope, 1886, in Florida, U.S.A. Yamaguti 

(1958) considered Langeronia synonymous with 

the genus Loxogenes Stafford, 1905. Brenes et 

al. (1959) examined specimens recovered from 

the cane toad, Bufo marinus (Linnaeus, 1758) in 

Costa Rica and disagreed with Yamaguti, concluding 

that Langeronia was a valid genus. Ubelaker 

(1965) collected trematodes from B. marinus 

in Nicaragua and published a redescription 

of L. macrocirra, concluding that L. provitellaria 

should be considered a synonym of that species. 

He also supported Yamaguti and his 1958 

synonymy of the 2 genera. Christian (1970) 

studied specimens collected from the intestines 

of R. pipiens in Wisconsin, Ohio, and Vermont, 

U.S.A., which he identified as L. provitellaria, 

Loxogenes sp., and a new species, Langeronia 

parva, respectively. Christian (1970) disagreed 

with Yamaguti's (1958) opinion synonymizing 

3 Corresponding author. 

165 

Langeronia and Loxogenes and supported Brenes 

et al. (1959) in validating the generic status 

of Langeronia. Christian (1970) did not mention 

the article by Ubelaker (1965) and the synonomy 

of L. macrocirra and L. provitellaria. However, 

Christian (1970) did state that in his opinion, 

according to the description and measurements 

given by Brenes et al. (1959) for "L. macrocirra," 

they were actually redescribing L. 

provitellaria. This would tend to explain why 

Ubelaker (1965) synonymized the 2 species, 

comparing the overlap of measurements from 

his specimens with the measurements given by 

Brenes et al. (1959). Yamaguti (1958) cited Loxogenes 

s. str. and Langeronia as subgenera of 

the genus Loxogenes s. lat. Babero and Golling 

(1974) reported 3 species of L. provitellaria 

from 2 bullfrogs (Rana catesbiana Shaw, 1802) 

collected in Nye County, Nevada, U.S.A. 

Materials and Methods 

One of 36 California treefrogs, Hyla cadaverina 

Cope, 1866, examined (LACM No. 88937) from 

Orange County, California was infected with 83 trematodes 

in the large intestine. All H. cadaverina specimens 

had been collected between 1952 to 1967 and 

deposited in the herpetology collection of the Natural 

History Museum of Los Angeles County (LACM). 

They were originally preserved in 10% formalin and 

later stored in 70% ethanol. 

Worms were removed from the large intestine, 

rinsed in 70% ethanol, stained in Delafield's hematoxylin, 

dehydrated in ethanol, and mounted in Canada 



balsam. Subsequent examination of the trematode 

specimens indicated that they represented an undescribed 

species of the genus Langeronia. Drawings were 

made with the aid of a drawing tube. Measurements 

are in micrometers unless otherwise indicated. The 

range is followed by the mean in parentheses. Type 

specimens were deposited in the United States National 

Parasite Collection (USNPC), Beltsville, Maryland, 

U.S.A. 

Some of the cotype specimens of L. macrocirra 

(USNPC No. 37127), the type specimens of L. provitellaria 

(USNPC No. 47569), and the type and paratype 

specimens of L. parva (USNPC No. 70557, 

70558) were examined during this study. 

Description 

Results 

Langeronia burseyi sp. n. 

(Fig. 1) 

Based on 10 of 83 specimens: Lecithodendriidae 

(Liihe, 1901) Odhner, 1910; Pleurogenetinae 

Travassos, 1921. Body small, pyriform, 

0.60-0.75 mm (0.66) long, maximum width 

0.38-0.55 mm (0.49) at testicular level. Tegument 

thin, spinose. Oral sucker subterminal, 95— 

105 (102) long by 70-93 (81) wide. Prepharynx 

absent. Pharynx 60-68 (63) long by 38-45 (42) 

wide. Esophagus 23-28 (24) long by 12-15 (13) 

wide. Ceca bifurcate just anterior to midbody 

and extending posteriorly to anterior of testes. 

Acetabulum approximating size of oral sucker, 

75-105 (92) long by 78-98 (90) wide. Cirrus 

pouch 225-287 (261) long by 53-70 (59) wide, 

arching transversely over acetabulum, then 

twisting medioventrally and opening into shallow 

thin-walled atrium with genital pore. Testes 

smooth, opposite, transversely oval, in posterior 

third of body. Right testis 92-155 (119) long by 

98-163 (138) wide, left testis 110-125 (113) 

long by 125-188 (146) wide. Ovary round to 

oval, at acetabular level, anterior to right testis, 

56-90 (68) long by 56-100 (77) wide. Seminal 

receptical ovoid to spherical, 70 long by 45 

wide. Mehlis' gland directly postacetabular, 

Laurer's canal not observed. Vitellaria dorsal, 

follicular, extending from just posterior to pharynx 

to anterior half of ceca on either side of 

esophagus. Uterus with irregular transverse 

loops filling post-testicular space. Eggs smooth, 

elliptical, 23-28 (24) long by 12-15 (13) wide. 

Excretory bladder V-shaped, excretory pore terminal. 


Taxonomic summary 

TYPE HOST: California treefrog, Hyla cadaverina 

Cope, 1866, deposited in Natural History 

Museum of Los Angeles County as LACM 

88937. 

TYPE LOCALITY: Harding Canyon, Orange 

County, California, U.S.A. (33°42'N, 

117°38'W). 

COLLECTION DATE: 16 June 1965. 

SITE OF INFECTION: Large intestine. 

DEPOSITED SPECIMENS: Holotype and paratypes 

USNPC No. 89628 

ETYMOLOGY: This species is named for 

Charles R. Bursey, Pennsylvania State University, 

Shenango, Pennsylvania, U.S.A., in recognition 

of his many contributions to the parasitology 

of amphibians and reptiles. 

Langeronia Caballero and Bravo-Hollis, 

1949 

EMENDED DIAGNOSIS: Lecithodendriidae, 

Pleurogenetinae. Body spatulate to pyriform, 

spined. Oral sucker well-developed, terminal or 

subterminal. Prepharynx present or absent; pharynx 

well-developed. Esophagus present; ceca 

wide, extending to midbody. Acetabulum equatorial. 

Testes symmetrical, postacetabular, and 

intercecal. Cirrus pouch elongate, twisted, extending 

transversely intercecally in space between 

intestinal bifurcation and acetabulum. 

Genital pore preacetabular, ventral to or on internal 

border of left cecum. Ovary dextral to acetabulum, 

pretesticular. Uterine coils lateral, 

postequatorial; eggs smooth, operculate. Vitellaria 

follicular, in shoulder area, on either side 

of esophagus, not confluent. Excretory bladder 

Y- or V-shaped. Intestinal parasites of amphibians. 

TYPE SPECIES: Langeronia macrocirra Caballero 

and Bravo-Hollis, 1949, from R. pipiens 

in Mexico. 

OTHER SPECIES: In addition to L. macrocirra, 

the genus currently contains 2 other species, 

Langeronia provitellaria Sacks, 1952, and L. 

parva Christian, 1970. Langeronia burseyi sp. n. 

differs from all members of the genus in its 

small size (smallest in the genus), placement of 

cirrus at the acetabular level, cecal length, and 

V-shaped bladder. It most closely resembles L. 

provitellaria in the position of the vitellaria, 

with both beginning at the pharyngeal level. 

However, in L. burseyi the vitellaria end just

DAILEY AND GOLDBERG—LANGERONIA BURSEYI SP. N. FROM TREEFROGS 167 

Figure 1. Langeronia burseyi sp. n. from Hyla cadaverina. Entire worm, ventral view. 

posterior to the cecal bifurcation, while in L. (size, position of vitellaria and pharynx, length 

provitellaria, they extend to the anterior of the of ceca) distinguish L. macrocirra and L. procirrus. 

The new species resembles L. macrocirra vitellaria as separate species, 

and L. parva in that the ovary and testes are not 

deeply lobed as in L. provitellaria. The deeply Key to the Species of Langeronia 

lobed ovary and testes along with other features la. Body length more than 1.30 mm 2 



lb. Body length less than 1.30 mm 3 

2a. Ovary and testes deeply lobed .. L. provitellaria 

2b. Ovary and testes not deeply lobed 

L. macrocirra 

3a. Prepharynx present L. parva 

3b. Prepharynx absent L. burseyi 

Remarks and Discussion 

During this study, the specimens treated by 

Ubelaker (1965) and Christian (1970) were not 

available. However, after examination of all other 

known available specimens, we agree with 

Christian (1970) that Langeronia is a valid genus 

and that L. macrocirra and L. provitellaria 

are valid species. The differences between Loxogenes 

and Langeronia are as follows: In Loxogenes, 

the vitellaria are confluent, while in 

Langeronia they are not. In Loxogenes, the testes 

are on the same level as the acetabulum, and 

the ovary is always preacetabular; while in Langeronia, 

the testes are postacetabular, and the 

ovary is at the same level or just postacetabular. 

In Loxogenes, the cirrus pouch is extracecal and 

club-shaped, and the genital pore is extracecal, 

anterior and dorsal. In Langeronia, the cirrus 

pouch is elongate, not club-shaped, twisted intercecally, 

and preacetabular, while the genital 

pore is lateral and ventral to the left cecum. In 

Loxogenes, the intestinal ceca do not extend to 

the acetabulum, while in Langeronia, they always 

extend past the acetabulum. In Loxogenes, 

the uterine coils are arranged in an anterior-posterior 

configuration, while in Langeronia, the 

uterine coils are lateral loops confined to the 

posterior half of the body. 


The authors thank Robert L. Bezy, Natural 

History Museum of Los Angeles County, for 


ALAN F. BIRD 

February 11, 1928-December 13, 1999 

Elected to Honorary Membership in 1997 


permission to examine Hyla cadaverina; J. 

Ralph Lichtenfels, United States National Parasite 

Collection, for the loan of type material; and 

Lynn Hertel, University of New Mexico, for her 

help with illustrations. 


Babero, B. B., and K. Coiling. 1974. Some helminth 

parasites of Nevada bullfrogs, Rana catesbiana 

Shaw. Revista de Biologia Tropical, Universidad 

de Costa Rica 21:207-220. 

Brenes, R. R., G. Arroyo-Sancho, and E. Delgado- 

Flores. 1959. Helmintos de la Republica de Costa 

Rica XI. Sobre la validez del genero Langeronia 

Caballero y Bravo, 1949 (Trematoda: Lecithodendriidae) 

y hallazgo de Ochetosoma miladelarocai 

Caballero y Vogelsang, 1947. Revista de 

Biologfa Tropical, Universidad de Costa Rica 7: 

81-87. 

Caballero, E., and M. Bravo-Hollis. 1949. Description 

d'un nouveau genre de Pleurogeninae (Trematoda: 

Lecithodendriidae) de grenouilles du Mexique 

(1) Langeronia macrocirra n. g., n. sp. Annales 

de Parasitologie Humaine et Comparee 24: 

193-199. 

Christian, F. A. 1970. Langeronia pai~va sp. n. (Trematoda: 

Lecithodendriidae) with a revision of the 

genus Langeronia Caballero and Bravo-Hollis, 

1949. Journal of .Parasitology 56:321-324. 

Sacks, M. 1952. Langeronia provitellaria (Lecithodendriidae), 

a new species of trematode from Rana 

pipiens sphenocephala. Transactions of the American 

Microscopical Society 71:267—269. 

Ubelaker, J. E. 1965. The taxonomic status of Langeronia 

Caballero and Bravo-Hollis, 1949 with 

the synonymy of Loxogenes provitellaria Sacks, 

1952 with Loxogenes macrocirra Caballero and 

Bravo-Hollis, 1949. Transactions of the Kansas 

Academy of Science 68:187-190. 

Yamaguti, S. 1958. Systema Helminthum. Vol. 1. The 

Digenetic Trematodes of Vertebrates, Parts I and 

II. Interscience Publishers, Inc., New York.



Oxyuroids of Palearctic Testudinidae: New Definition of the Genus 

Thaparia Ortlepp, 1933 (Nematoda: Pharyngodonidae), Redescription 

of Thaparia thapari ihapari, and Descriptions of Two New Species 

SALAH BouAMER1-3 AND SERGE MoRAND2 

1 Groupe Pluridisciplinaire des Sciences, Universite de Perpignan, Av. de Villeneuve, 66860 Perpignan, France 

(e-mail: bouamer@univ-perp.fr) and 

2 Centre de Biologie et d'Ecologie Tropicale et Mediteraneenne, Laboratoire de Biologic Animale (UMR 5555 

CNRS), Universite de Perpignan, Av. de Villeneuve, 66860 Perpignan, France (e-mail: morand@univ-perp.fr) 

ABSTRACT: The generic diagnosis of Thaparia Ortlepp, 1933, is emended based on the study and redescription 

of Thaparia thapari thapari (Dubinina, 1949) from the cecum of Testudo graeca Linnaeus, 1758, collected in 

Settat, Morocco. In addition, 2 new species, Thaparia carlosfeliui sp. n. and Thaparia bourgati sp. n. from the 

cecum of Testudo hermanni Gmelin, 1789, collected in Catalonia, Spain, are described. Scanning electron 

microscopy studies revealed substantial differences in the structure of the mouth and the caudal end, which 

enabled us to differentiate the 2 new species from the others and from each other. 

KEY WORDS: Thaparia thapari thapari, Thaparia carlosfeliui sp. n., Thaparia bourgati sp. n., Nematoda, 

Pharyngodonidae, Testudo graeca, spur-thighed tortoise, Testudo hermanni, Hermann's tortoise, Morocco, Spain. 

The genus Thaparia was erected by Ortlepp 

(1933) for Thaparia macrospiculum Ortlepp, 

1933, a parasite of the tent tortoise, Psammobates 

tentorius (Bell, 1828). Ortlepp (1933) gave 

the following diagnosis: Medium-sized worms 

possessing 3 lips and a relatively short esophagus 

consisting of an anterior muscular portion, 

a middle glandular portion, and a posterior bulb; 

excretory pore post-bulbar; lateral alae absent. 

Vulva approximated to anus; vagina very long; 

2 uteri and 2 ovaries. Caudal extremity of male 

cut ventrally and continued backward to form a 

short truncated and alate tail. Four pairs of caudal 

papillae, 3 pairs circumcloacal and 1 pair 

toward tip of tail. Single spicule very long and 

stout, extending to or even anterior of the esophageal 

bulb. Type species T. macrospiculum from 

P. tentorius. 

Walton (1942) described a second species, 

Thaparia contortospicula, a parasite of the Galapagos 

tortoise, Geochelone nigra (Quoy and 

Gaimard, 1824) from the Galapagos Islands. 

Fitzsimmons (1961) described Thaparia capensis 

from the South African bowsprit tortoise, 

Chersina angulata (Schweigger, 1812), in South 

Africa. 

Fetter (1966) described Thaparia domerguei, 

from the common spider tortoise, Pyxis arachnoides 

Bell, 1827, and the radiated tortoise, 

Geochelone radiata (Shaw, 1802), from Mada- 

Corresponding author. 

169 

gascar. She also transferred the species Tachygonetria 

thapari Dubinina, 1949, described from 

the Central Asian tortoise, Testudo horsfieldii 

Gray, 1844, in Afghanistan and from other Palearctic 

tortoises, the spur-thighed tortoise Testudo 

graeca Linnaeus, 1758, and Hermann's tortoise, 

Testudo hermanni Gmelin, 1789, to the 

genus Thaparia. Fetter (1966) also modified the 

diagnosis of the genus, which is now: Pharyngodoninae—mouth 

with 3 lips; short esophagus 

divided into 2 equal parts; 4 pairs of caudal papillae: 

3 pairs at the level of the cloaca and 1 

near tail extremity. Caudal alae present or absent 

in males. Type species: T. macrospiculum Ortlepp, 

1933. 

Petter and Douglass (1976) described 2 additional 

species, T. rnacrocephala and T. microcephala, 

from the Bolson tortoise, Gopherus flavomarginatus 

Legler, 1959, in Mexico. 

Baker (1987) cited the 7 species above plus 3 

subspecies of T. thapari, following Petter (1966) 

(Thaparia thapari thapari (Dubinina, 1949), 

Thaparia thapari australis (Petter, 1966), and 

Johnson (1973a) (Thaparia thapari lysavyi 

Johnson, 1973, from Testudo hermanni from Albania). 

The 7 species of Thaparia are distributed 

in 5 biogeographical regions. 

In this study, we describe 2 new species of 

Thaparia from a testudinid species and emend 

the diagnosis of the genus. 


A first collection of nematode parasites from 18 

specimens of Testudo graeca from Settat, Morocco 



(deposited at the Institut Agronomique et Veterinaire 

Hassan II, Rabat, Morocco), was made by one of us 

(S.B.). The second collection, from a single specimen 

of T. hennanni from Catalonia, Spain, was made by 

C. Feliu, Barcelona, Spain, and deposited at the Barcelona 

Zoo, Spain. Nematodes were preserved in 70% 

ethanol before being cleared with lactophenol for 

study. Figures were made with the aid of a drawing 

tube. Nematodes were dehydrated by passage through 

progressive ethanol concentrations to absolute ethanol 

and critical-point-dried (M scope 500, Hitachi, Japan). 

The scanning electron microscope used was a Hitachi 

S 520, Hitachi, Japan at 20 kV. Measurements given 

are for the holotype male and the allotype female. 

Measurements in parentheses are the ranges of paratype 

males and females. All measurements are in micrometers. 

Results 

Thaparia thapari thapari (Dubinina, 1949) 

(Figs. 1-10) 

Redescription 

GENERAL: The material examined consisted 

of 6 males and 15 females. Body medium-sized, 

stout. Mouth triangular, with 3 transparent lips. 

Buccal cavity with denticles. Cephalic sense organs 

consisting of inner circle of 6 nerve endings, 

papillae not pedunculate (Figs. 2, 3, 6, 7), 

the outer circle not observed, and amphids present. 

Esophagus divided into 2 portions: anterior 

muscular part, and comparatively longer posterior 

glandular part terminating in valvular bulb; 

17 chitinoid pieces surrounding anterior end of 

esophagus (Figs. 1, 3). Excretory pore postesophageal. 

MALE: Length 2,754-3,169; maximum 

thickness 191-229. In worm measuring 2,773, 

esophagus 388: corpus 180 and isthmus plus 

bulb 208. Nerve ring and excretory pore 161 

and 889, respectively, from anterior end. Posterior 

extremity truncated. Tail 67 long. Spicule 

needle-shaped, 100 long. Gubernaculum Vshaped. 

Three pairs of caudal papillae: 2 circumcloacal 

(1 pair preanal and 1 pair postanal) and 

1 pair at tail end. Preanal membrane, present 

with 6 lobes (Figs. 4, 5, 8, 9, 10), and caudal 

alae absent. 

FEMALE: Length 4,282-4,716; maximum 

thickness 356-378. In a worm measuring 4,600, 

esophagus 615: corpus 240 and isthmus plus 

bulb 375. Nerve ring, excretory pore, and vulva 

at 180, 1,282, and 2,264, respectively, from anterior 

end. Tail 270 long. 


HOST: Spur-thighed tortoise, Testudo graeca 

Linnaeus, 1758. 

SITE IN HOST: Cecum. 

TYPE LOCALITY/COLLECTION DATE: Settat, 

Morocco, 32°30'45"N, 7°45'30"W, 22 July 1999, 

by S.B. 

SPECIMENS DEPOSITED: Museum National 

d'Histoire Naturelle, Paris, France. Number 825 

HE 

Remarks 

Thapar (1925) described the female of this 

species from Testudo graeca, as Oxyuris sp. Dubinina 

(1949) studied males, the structures of 

which made it possible to include the species in 

Tachygonetria Wedl, 1862. Fetter (1966) redescribed 

and transferred this species to the genus 

Thaparia and divided it in 2 subspecies: T. thapari 

thapari and T. thapari australis. 

The emended diagnosis characterizes T. thapari 

thapari with 17 chitinoid pieces, whereas 

Petter (1961, 1966) cited 6 chitinoid pieces. Cephalic 

sense organs consist of an inner circle of 

6 nerve endings (papillae not pedunculate), 4 

submedian and 2 lateral close to amphids; outer 

papillae were not observed. 

Description 


Thaparia carlosfeliui sp. n. 

(Figs. 11-23) 


of 20 males and 40 females. Nematoda, Oxyuroidea, 

Pharyngodonidae, Thaparia. Robust 

worms of small size. Mouth surrounded by 3 

lips. Esophagus in 2 parts, with elongated isthmus. 

Amphids prominent. Differs from the diagnosis 

of the genus in the number of caudal 

papillae. 

MALE: (holotype and 3 paratypes): Mouth 

surrounded by 3 V-shaped cut lips (Figs. 13, 21). 

Six oral papillae, arranged in 3 pairs (Fig. 13). 

Buccal cavity without denticles. Esophagus 

lobes visible. No chitinoid pieces visible. Tail 

without alae. Structure of caudal region complex 

(Figs. 15, 16, 22, 23). One pair of preanal papillae, 

1 pair of large postanal elongated papillae. 

Posterior lip of anus with central nipple. 

Surrounding ventral membrane present lateral 

and anterior to anus, and second preanal membrane 

situated posterior to first membrane. Anterior 

lip of anus with 2 lobes, extremity of spic-

BOUAMER AND MORAND—OXYUROID GENUS THAPARIA 171 

Figures 1-5. Thaparia thapari thapari, male. 1. Anterior end. 2. Cephalic end, en face view. 3. Cephalic 

end, deeper en face view. 4. Posterior end, lateral view. 5. Posterior end, ventral view. All scale lines = 50 |un. 



Figures 6-9. Thaparia thapari thapari, scanning electron micrographs: 6. Cephalic end of female. 7. 

Cephalic end of male. 8. Caudal end of male, ventral view (a = preanal papillae, b = postanal papillae, 

c = caudal papillae). 9. Caudal end of male, lateral view (a = preanal papillae, b = postanal papillae, c 

= caudal papillae, e = preanal membrane). Scale lines: Fig. 6 = 17.6 (xm; Fig. 7 = 20 urn; Fig. 8 = 20 

fjim; Fig. 9 = 20 (xm. 


Figure 10. Thaparia thaparia thaparia, scanning 

electron micrograph: Cloacal view of male (d = anterior 

lip, e = preanal membrane, f = posterior lip, g 

= ventral lobe of preanal membrane, h = subventral 

lobe of preanal membrane, i = spicule). Scale line = 

5 (xm. 

ule visible between them. U-shaped gubernaculum. 

Third pair of papillae lateral at end of tail. 

Length 1,797 (1,324-1,943) with maximal width 

180 (150-255). Nerve ring 112 (70-116) from 

anterior end. Excretory pore 586 (435-640) 

from anterior end. Esophagus 416 (351-421) 

long, corpus 165 (139-167) long, isthmus plus 

bulb 251 (212-254) long. Tail 45 (45-49) long. 

Spicule 123 (109-123) long. 

FEMALE (allotype and 3 paratypes): Mouth 

surrounded by 3 V-shaped cut lips (Figs. 19, 20). 

Six oral papillae arranged in 3 pairs as in male 

(Fig. 19). Buccal cavity without denticles. No 

chitinoid pieces visible. Esophagus lobes visible. 

Length 3,471 (2,867-3,848), maximum width 

426 (426-482). Nerve ring corpus 114 (114- 

188) from anterior end. Excretory pore 814 

(575-814) from anterior end. Esophagus: 615 

(482-634) long with corpus 244 (192-252) long 

and isthmus plus bulb 371 (290-382). Vulva 

1,622 (1,528-2,056) from anterior end. Tail 244 

(232-283) long. Eggs asymmetrical, measuring 

89 X 147 (69 X 126-89 X 147). 



TYPE HOST: Hermann's tortoise, Testudo hermanni 

Gmelin, 1789. 

SITE IN HOST: Cecum. 

TYPE LOCALITY/COLLECTION DATE: South Catalonia, 

Spain, 41°23'14"N, 2°11'17"E, 17 December 

1993 by Dr. Carlos Feliu. 



HE 

ETYMOLOGY: The species is named in honor 

of Professor Carlos Feliu (University of Barcelona, 

Spain). 

Remarks 

Thaparia carlosfeliui sp. n. differs from T. 

macrospiculum in the size of the spicule and 

from T. domerguei in the absence of caudal alae. 

Thaparia capensis differs from the new species 

in the presence of caudal alae and the length of 

the body. 

Thaparia contortospicula resembles T. carlosfeliui 

in the size of the spicule but differs in 

the presence of caudal alae and the length of the 

body. Thaparia macrocephala and T. microcephala 

differ in the presence of caudal alae, the 

number of lips, and the length of the body. 

Thaparia carlosfeliui differs from T. thapari 

(Dubinina, 1949) in the absence of teeth, the arrangement 

of labial papillae, and the absence of 

6 chitinoid pieces around the mouth. Thaparia 

carlosfeliui differs from T. thapari australis in 

the arrangement of labial papillae, the lack of a 

tip at the extremity of the male tail, and the absence 

of chitinoid pieces. 

Thaparia bourgati sp. n. 

(Figs. 24-31) 

Description 


of 10 males and 1 female. Nematoda, Oxyuroidea, 

Pharyngodonidae, Thaparia. Robust 

worms of small size. Mouth surrounded by 3 

lips. Esophagus in 2 parts, with an elongated 

isthmus. Amphids prominent. Differs from the 

diagnosis of the genus in the number of caudal 

papillae. 

MALE (holotype and 3 paratypes): Labial papillae 

conspicuous, arranged as in T. carlosfeliui 

sp. n. (Figs. 20, 28). Three projections visible 

inside mouth, forming diaphragm under lip. 

Buccal cavity without denticles. No chitinoid 



15 



Figures 17-19. Thaparia carlosfeliui sp. n. female allotype. 17. Entire specimen, lateral view. 18. Anterior 

end. 19. Cephalic end, en face view. Scale lines: Fig. 17 = 400 (Jim; Figs. 18, 19 = 50 (xm. 

pieces visible. Tail without alae. Three pairs of 

caudal papillae. Ventral membrane surrounding 

anus anteriorly, showing on each side 2 small 

submedian lobes and 2 large lateral lobes (Figs. 

26, 27, 29—31). Anterior lip of anus composed 

of 2 submedian lobes congruent at distal extremity. 

Posterior lip of anus with central nipple. Ushaped 

gubernaculum. Length 3,122 (2,204- 

3,207), maximum width 221 (143-255). Nerve 

ring 98 (68-104) from anterior end. Excretory 

pore 756 (705-851) from anterior end. Esophagus 

439 (251-416) long, corpus 169 (97-169) 

long, isthmus plus bulb 270 (154-270) long. Tail 

53 (45-53) long. Spicule 162 (112-225) long. 

FEMALE (allotype): A single female has been 

found, which closely resembles females of the 

other species of the genus. Nerve ring not visible. 

Length 2,226, maximum width 166. Excre- 

Figures 11-16. Thaparia carlosfeliui sp. n. male holotype. 11. Entire specimen, lateral view. 12. Anterior 

end. 13. Cephalic end, en face view. 14. Cephalic end, deeper en face view. 15. Posterior end, lateral 

view. 16. Posterior end, ventral view. Scale lines: Fig. 11 = 100 jim; Figs. 12-16 = 50 jim. 



zz Z3 

Figures 20-23. Thaparia carlosfeliui sp. n., scanning electron micrographs. 20. Cephalic end of female. 

21. Cephalic end of male. 22. Caudal end of male, lateral view (a = preanal papillae, b = postanal papillae, 

c = caudal papillae, d = anterior lip, e = preanal membrane). 23. Caudal end of male, ventral view (a = 

preanal papillae, b = postanal papillae, c = caudal papillae, d = anterior lip, e = preanal membrane, 

f = posterior lip). Scale lines: Fig. 20 = 30 urn; Figs. 21—23 = 25 jjim. 

tory pore 549. Esophagus 502, corpus 225 long, 

isthmus plus bulb 277 long. Vulva 804 from anterior 

end. Tail 131. Eggs asymmetrical, measuring 

49 X 90. 


TYPE HOST: Hermann's tortoise, Testudo hermanni 

Gmelin, 1789. 


INFECTION SITE: Cecum. 

TYPE LOCALITY/COLLECTION DATE: South Catalonia, 

Spain, 41°23'14°N, 2°11'17"E, 17 December 

1993, by Dr. Carlos Feliu. 



HE 

ETYMOLOGY: The species is named in honor 

c


Figures 24-27. Thaparia bourgati sp. n. male holotype. 24. Cephalic end, en face view. 25. Cephalic 

end, deeper en face view. 26. Caudal end, lateral view. 27. Caudal end, ventral view. All scale lines = 50 

|xm. 



Figures 28—31. Thaparia bourgati sp. n. male, scanning electron micrograph. 28. Cephalic end. 29. 

Caudal end, lateral view (a = preanal papillae, b = postanal papillae, c = caudal papillae). 30. Caudal 

end, ventral view (a = preanal papillae, b = postanal papillae, c = caudal papillae, d = anterior lip, e = 

preanal membrane). 31. Cloacal view (d = anterior lip, e = preanal membrane, f = posterior lip). Scale 

lines: Fig. 28 = 10 fjim; Fig. 29 = 17.6 jjim; Fig. 30 = 15 u.m; Fig. 31 = 7.5 fjim. 


of Professor Robert Bourgat (University of Perpignan, 

France). 

Remarks 

Thaparia bourgati sp. n. differs from all other 

species of the genus in the same characters as 

T. carlosfeliui. Thaparia bourgati differs from 

T. carlosfeliui in the shape of the preanal membrane 

in the male—9 lobes in T. bourgati and 4 

lobes in T. carlosfeliui—and in the size of the 

eggs (smaller in T. bourgati). 

Discussion 

Thaparia bourgati sp. n. and T. carlosfeliui 

sp. n. differ from all other species of the genus 

Thaparia, except T. thapari, in the lack of caudal 

alae. Both species differ from the subspecies 

T. thapari australis in the shape and the disposition 

of labial papillae, the lack of apical chitinoid 

pieces, the lack of a tip at the end of the 

male tail, and the presence of a longer spicule 

compared with the length of the tail. They differ 

from the subspecies T. thapari thapari in the 

disposition of labial papillae, the lack of esophageal 

teeth and the lack of apical chitinoid pieces, 

the shape of the preanal membrane, and the 

shape of the gubernaculum in the male, and 

from T. thapari rysavyi in the arrangement of 

labial papillae and in the shape of the adanal 


tudinidae. Medium-sized, lateral alae present or 

absent. Mouth with 3 or 6 slightly bilobed lips. 

Esophagus rather short, divided into 2 parts of 

about equal length: an anterior muscular corpus 

and a posterior glandular isthmus terminating in 

a valvulated bulb. Excretory pore bulbar or postbulbar. 

MALE: Tail truncated, spiked, or simple. 

Caudal alae present or absent. Spicule simple or 

contorted, may be very long. Gubernaculum U-, 

V-, or Y-shaped. Caudal papillae in 3 pairs: 2 

circumcloacal, 1 pair at or near tail end. 

FEMALE: Tail tapering to sharp point. Vulva 

postequatorial, sometimes very close to anus. 

Vagina long; ovijector present. Eggs thinshelled, 

relatively few. 

TYPE SPECIES: Thaparia macrospiculum Ortlepp, 

1933, in Psammobates tentorius; South 

Africa. 

Key to the Species and Subspecies of the 

Genus Thaparia 

This key follows Johnson (1973b) and includes 

T. capensis Fitzsimmons, 1961, T. macrocephala 

Petter and Douglass (1976), T. microcephala 

Petter and Douglass (1976), and T. 

carlosfeliui sp. n. and T. bourgati sp. n., both 

described herein. 

membrane in the male. Finally, T. carlosfeliui 1. Caudal alae in male present 2 

sp. n. resembles T. bourgati sp. n. in the arrange- Caucal alae in male absent ..... 7 

ment of the labial papillae, but it is distinguished 2' Iai! in ma!e spiked " I 

^ r Tail in male truncated 5 

by the shape of the preanal membrane in the 3 Spicule contorted; vulva away from anus 

male and by the size of the eggs. T. contortospicula Walton, 1942 

The genus Thaparia shows a wide geograph- Spicule simple; vulva far away from anus ..... 4 

ical distribution, with 3 Palearctic species (T. 4- Len§th of sPicule 1/2 of body 

..... _, , . . „, , T. macrocephala Fetter and Douglass (1976) 

carlosfelim sp. n., T. bourgati sp. n., and T. tha- Length of spicule 1/3 of body 

pan), 2 Nearctic species, 2 South African spe- T. micmcephala Fetter and Douglass (1976) 

cies, and 1 species from the Galapagos Islands. 5. Spicule simple; vulva far (more than 1,450 [Jim) 

The question remains open concerning the pres- from anus T. capensis Fitzsimmons, 1961 

ence of this genus in other members of the Pa- Splcule simPle; vulva near (less than 20° ^m) f 

^ anus 6 

learctic tortoises (T. horsfieldii, T. graeca, T. 6 Spicule less than 1 mm in length 

hermanni, the Egyptian tortoise, Testudo klein- T. domerguei Fetter, 1966 

manni Lortet, 1883, and the marginated tortoise, Spicule more than 2.5 mm in length 

Testudo marginata Schoepff, 1792). T macrospiculum Ortlepp, 1933 

7. Buccal cavity with 6 teeth 

Emended diagnosis of the genus Thaparia , , . T: ll*apari th«pari Dubinina (1949) 

to l Buccal cavity without teeth 8 

The use of a scanning electron microscope al- g. Spicule less than 90 jxm; tail more than 90 jxm 

lowed verification that the previously described in length T. thapari australis Fetter, 1966 

adanal papillae are simple lobes. The lack of ter- Spicule more than 90 jun; tail less than 90 ^.m 

. , . c j u * - , u in length 9 

mmal nerves is confirmed by optical observa- 9 preana, ^embrane absent 

tion. The new diagnosis of the genus is: T. thapari rysavyi Johnson, 1973 

Pharyngodonidae: Intestinal parasites of Tes- Preanal membrane present 10 



10. Preanal membrane with 4 lobes 

T. carlosfeliui sp. n. 

Preanal membrane with 9 lobes 

T. bourgati sp. n. 

Acknowledgment 

We thank Dr. Annie Fetter for helpful comments 

on an earlier version of the manuscript. 


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

in amphibians and reptiles. Memorial University 

of Newfoundland, Occasional Papers in 

Biology 11:1-325. 

Dubinina, M. H. 1949. (Ecological studies on the parasite 

fauna of the Testudo horsfieldii Gray from 

Tadjikistan). Parazitologicheskii Sbornik Zoologicheskogo 

Instituta AN SSSR 11:61-97. (In Russian.) 

Fitzsimmons, W. M. 1961. Thaparia capensis n. sp., 

an oxyuroid parasite of Testudo angidata. British 

Journal of Herpetology 3:7-12. 

Johnson, S. 1973a. Some oxyurid nematodes of the 

genera Mehdiella and Thaparia from the tortoise 

Testudo hermani. Folia Parasitologica 20:141— 

148. 


MARION M. FARR 

1903-2000 

. 1973b. The first record of the nematode Thaparia 

thapari thapari (Dubinina, 1949) in Afghanistan, 

with remarks on the genus Thaparia 

Ortlepp, 1933. Folia Parasitologica 20:178. 

Ortlepp, R. J. 1933. On some South African reptilian 

oxyurids. Onderstepoort Journal of Veterinary 

Science and Animal Industry 1:9-114. 

Fetter, A. J. 1961. Redescription et analyse critique 

de quelques especes d'Oxyures de la tortue grecque 

(Testudo graeca L.). Diversite des structures cephaliques. 

Annales de Parasitologie Humaine et 

Comparee 10:648-671. 

. 1966. Equilibre des especes dans les populations 

de Nematodes parasites du colon des Tortues 

terrestres. Memoires du Museum National 

d'Histoire Naturelle, Paris, Nouvelle Serie, Serie 

A, Zoologie 39:1-252. 

, and J. F Douglass. 1976. Etude des populations 

d'Oxyures du colon des Gopherus (Testudinidae). 

Bulletin du Museum National d'Histoire 

Naturelle, Paris, Serie 3, No. 389, Zoologie 271: 

731-768. 

Thapar, G. S. 1925. Studies on the oxyurid parasites 

of the reptiles. Journal of Helmintology 3:83-150. 

Walton, A. C. 1942. Some oxyurids from a Galapagos 

tortoise. Proceedings of the Helminthological Society 


Elected to Membership, 1938 

Executive Committee Member at Large, 1943-1945 

21st Recording Secretary, 1946 

Vice President, 1946 

34th President, 1951 

Assistant Secretary-Treasurer, 1964 

Society Representative to the Washington Academy of 

Sciences, 1964-1965 

Elected to Life Membership, 1979 

Published in the Proceedings from 1939-1963 on 

Eimeria and Histomonas 




Parasites of Farm-Raised Trout in Michigan, U.S.A. 

PATRICK M. MUZZALL 

Department of Zoology, Natural Science Building, Michigan State University, East Lansing, Michigan 48824, 

U.S.A. (e-mail: rnuzzall@pilot.msu.edu) 

ABSTRACT: A total of 635 trout (366 rainbow trout, Oncorhynchus mykiss Richardson, 1836; 166 brook trout, 

Salvelinus fontinalis Mitchill, 1814; 103 brown trout, Salmo trutta Linnaeus, 1758; Salmonidae) collected in 

March-July 1996, 1997, and 1998 from 12 trout farms in Michigan, U.S.A., was examined for parasites. Twelve 

parasite species (1 Acanthocephala, Acanthocephalus dims (Van Cleave, 1931) Van Cleave and Townsend, 1936; 

1 Monogenea, Gyrodactylus sp.; 2 Cestoda, Eubothrium salvelini (Schrank, 1790), Proteocephalus sp.; 1 Nematoda, 

Truttaedacnitis sp.; 1 Copepoda, Salmincola edwardsii (Olsson, 1869); 1 Myxozoa, Myxobolus cerebralis 

(Hofer, 1903); 4 Ciliophora, Capriniana sp. [ = Trichophrya sp.], Chilodonella sp., Ichthyophthirus multifiliis 

(Fouquet, 1876), Trichodina sp.; and 1 Mastigophora, Ichthyobodo sp. [ = Costia sp.]) were found. Rainbow trout 

were infected with A. dims, Truttaedacnitis sp., E. salvelini, Prutcocephalus sp., Gyrodactylus sp., M. cerebralis, 

Ichthyobodo sp., Capriniana sp., Chilodonella sp., /. multifiliis, and Trichodina sp. Brook trout were infected 

with A. dirus, S. edwardsii, E. salvelini, M. cerebralis, and Trichodina sp. Acanthocephalus dims was the only 

parasite infecting brown trout. Eubothrium salvelini, A. dirus, and Trichodina sp. infected trout from 9, 8, and 

8 farms, respectively. Acanthocephalus dirus in all trout species and S. edwardsii on brook trout had the highest 

prevalences, mean intensities, and mean abundances. 

KEY WORDS: trout, rainbow trout, Oncorhynchus mykiss, brook trout, Salvelinus fontinalis, brown trout, Salmo 

trutta, Salmonidae, helminths, protozoans, parasites, aquaculture, Michigan, U.S.A. 

Newman and Kevern (1994) reported that 

more than half of the fish growers in the state 

of Michigan, U.S.A., raise rainbow, brook, and 

brown trout. These growers produce trout for 

sale: 1) to individuals or groups for stocking, 2) 

to retail stores or restaurants, and 3) through 

their own fee-fishing ponds. In 1996, predators 

and diseases were the leading causes of death 

for trout in culture conditions in Michigan, accounting 

for 53% and 13% of all fish lost, respectively 

(Anonymous, 1997). Except for a few 

reports in local newspapers of diseases of trout 

in the Michigan Department of Natural Resources 

hatcheries and the studies by Sawyer et al. 

(1974) and Yoder (1972), little has been published 

on the parasites of trout raised in culture 

in Michigan. The present study reports on the 

parasites infecting rainbow, brook, and brown 

trout from 12 privately owned farms in Michigan. 

The emphasis of this study was on the 

metazoan parasites of trout, but observations and 

comments are also made on protozoans. Furthermore, 

information is presented on the life 

cycles of some of the parasites, their pathogenicity, 

and factors influencing their occurrence. 


Trout were collected by dip net or seine from ponds 

or raceways (hereinafter referred to as ponds) in 

March-July 1996, 1997, and 1998 from 12 farms in 

181 

Michigan. These trout farms are in an area of the lower 

peninsula between 42.0° and 45.5°N and 84.0° and 

86.5°W. Specific information on the locations of the 

trout farms, however, cannot be provided because of 

conditions of confidentiality imposed by the growers. 

Fish were either brought to the laboratory alive and 

necropsied within 48 hr of collection or were put on 

ice at the farm, brought to the laboratory, frozen, and 

examined later. Total length (mm) and sex were recorded 

during necropsy. The fins, external surface, 

buccal cavity, gills, brain, eyes, gonads, swim bladder, 

gastrointestinal tract, liver, spleen, and muscles (left or 

right side of each fish) were examined from all fish. 

In fish collected in 1997 and 1998, the skull bones and 

cartilage and 2 or more gill arches (without the filaments) 

were macerated separately into a slurry and examined 

with a compound microscope at 20 X. During 

the entire study, rainbow trout were examined from 23 

ponds, brook trout from 13 ponds, and brown trout 

from 7 ponds. These totals include some of the same 

ponds sampled on different dates and in different 

years. Parasite prevalence is defined as the percentage 

of fish infected, mean intensity as the mean number of 

metazoan parasites in infected fish, and mean abundance 

as the mean number of metazoan parasites in 

examined fish. Population numbers of each metazoan 

species at 1 facility were estimated as (prevalence) X 

(mean abundance) X (estimated number of trout) at 

the facility when the fish were sampled. It should be 

emphasized that the results of examination of frozen 

trout may not accurately reflect the occurrence and (or) 

numbers of protozoans and monogeneans. Therefore, 

mean numbers were not calculated for these parasite 

groups. Protozoan taxonomy follows that of Lom and 

Dykova (1992). Voucher specimens have been deposited 

in the United States National Parasite Collection 



Table 1. Numbers and mean lengths of Oncorhynchus 

mykiss, Salvelinus fontinalis, and Salmo trutta 

examined in 1996, 1997, and 1998. 

Species* 

OM, 1996 

OM, 1997 

OM, 1998 

SF, 1996 

SF, 1997 

SF, 1998 

ST, 1996 

ST, 1997 

No. 

examined 

184 

50 

132 

112 

27 

27 

88 

15 

Mean length ± SD 

(range, 95% confidence 

intervals) 

162 ± 71 (62-338, 152-172) 

177 ± 59 (93-279, 161-195) 

215 ± 64 (94-409, 204-226) 

154 ± 72 (56-378, 141-168) 

222 ± 45 (153-328, 204-239) 

200 ± 26 (160-260, 190-210) 

146 ± 69 (43-283, 131-161) 

208 ± 25 (147-244, 194-222) 

* OM = Oncorhynchus mykiss; SF = Salvelinus fontalis', ST 

= Salmo trutta. 

(USNPC), Beltsville, Maryland, U.S.A., with the following 

accession numbers: Eubothrium salvelini 

(89483), Acanthocephalus dims (89484), Salmincola 

edwardsii (89485). 

Results 

Totals of 366 rainbow trout, 166 brook trout, 

and 103 brown trout were examined for parasites 

from 12 Michigan farms in March—July 

1996, 1997, and 1998. All farms did not raise 

all species; thus, unequal numbers of each species 

were examined. The mean lengths of the 

trout species examined each year are in Table 1. 

Rainbow trout in 1998 were significantly larger 

than those in 1996 and 1997 (analysis of variance, 

F = 24.4, P < 0.0001). Brook trout in 

1996 were significantly smaller that those examined 

in 1997 and 1998 (analysis of variance, 

F = 15.6, P < 0.0001). Brown trout in 1997 

were significantly larger than those examined in 

1996 (Student's f-test, t = -6.34, P < 0.0001). 

Twelve parasite species were found in trout in 

this study (Table 2). Eleven parasite species infected 

rainbow trout, 5 species infected brook 

trout, and only 1 species infected brown trout. 

The prevalences, mean intensities, and mean 

abundances of the parasites found in trout in 

ponds varied dramatically. Of the metazoan parasite 

species found, Eubothrium salvelini 

(Schrank, 1790), Acanthocephalus dims (Van 

Cleave, 1931) Van Cleave and Townsend, 1936, 

and Salmincola edwardsii (Olsson, 1869) were 

gravid. The sites (in parentheses) where the parasites 

were found in trout were: E. salvelini (pyloric 

ceca, small intestine); Proteocephalus sp. 

(intestine); Gyrodactylus sp. (gills), Tmttaedac- 


nitis sp. (small intestine); A. dims (intestine); S. 

edwardsii (primarily gills, inner operculum, base 

of fins); Myxobolus cerebral is Hofer, 1903 (head 

bones and cartilage, gill arches); Ichthyobodo 

sp., Capriniana sp., Chilodonella sp., Ichthyophthirus 

multifiliis (Fouquet, 1876), and Trichodina 

sp. (gills and [or] head area). 

Acanthocephalus dims was the most common 

parasite species occurring in the gastrointestinal 

tract of each trout species. When S. edwardsii 

occurred, it commonly infested brook trout. Although 

several small immature S. edwardsii 

were seen on many brook trout, mean intensities 

and mean abundances of 5. edwardsii reflect 

only gravid females counted. Trichodina sp. was 

the most common external protozoan found on 

rainbow and brook trout. Myxobolus cerebralis 

infected 13 of 22 rainbow trout at 1 farm, 2 of 

15 rainbow trout at another farm, and 1 of 20 

brook trout at a third facility. Capriniana sp., 

Chilodonella sp., /. multifiliis, and Ichthyobodo 

sp. each infected fish in only 1 pond. 

All farms had trout that were infected with at 

least 1 parasite species. Eubothrium salvelini, A. 

dims, and Trichodina sp. infected trout from 9 

(75%), 8 (67%), and 8 (67%) farms, respectively, 

of the 12 farms from which trout were examined. 

Acanthocephalus dims and Trichodina 

sp. each infected rainbow trout from 52% of the 

23 ponds examined (Table 3). Eubothrium salvelini, 

A. dims, and 5. edwardsii each infected 

brook trout from 31% of the 13 ponds. Acanthocephalus 

dims infected brown trout from 2 

of 7 ponds. 

Trout were examined for parasites from 1 

farm in March and July 1997. Prevalences, mean 

abundances, and estimated numbers of helminths 

varied dramatically between these 

months (Table 4). In March, 4 parasite species 

infected trout, and A. dims and 5. edwardsii 

were common. Acanthocephalus dims had the 

highest mean abundances and estimated number 

of helminths. In July, only A. dims infrequently 

infected trout. 

There were no significant differences in the 

prevalence (chi-square analysis, P > 0.05) and 

intensity (Mann-Whitney test, P > 0.05) of E. 

salvelini, A. dims, and S. edwardsii between female 

and male trout of each species. At the 

farms where these 3 parasite species were common, 

examined trout did not vary enough in 

length to determine if these parasite infections 

had a significant relationship with length.

Discussion 

Twelve parasite species (2 Cestoda, 1 Monogenea, 

1 Nematoda, 1 Acanthocephala, 1 Copepoda, 

1 Myxozoa, 1 Mastigophora, 4 Ciliophora) 

were found in 635 trout examined from 12 

farms in the present study. Parasites of these 

trout and their infection values varied between 

ponds and years, a variability characteristic of 

wild trout populations as well. Most if not all of 

these parasite species have been found in wild 

trout from Michigan environments (Muzzall, 

1984, 1986; Hernandez and Muzzall, 1998). Digenetic 

trematodes, however, found in wild trout 

by Muzzall (1984, 1986), did not infect trout 

from culture ponds. 

Of the parasitic species found in the present 

study, E. salvelini, A. dims, S. edwardsii, and 

M. cerebralis had prevalences of 50% or more 

in at least 1 pond and deserve further discussion. 

Eubothrium salvelini was a common parasite of 

rainbow and brook trout. It utilizes copepods as 

intermediate hosts and commonly infects wild 

salmonids in inland waters (Hernandez and 

Muzzall, 1998) as well as in the Great Lakes 

(Muzzall, 1993, 1995a, b). Muzzall (1984) 

found immature Eubothrium sp. in brook trout 

from a Michigan creek. Boyce (1969) reported 

that E. salvelini reduced the growth, swimming 

performance, and survival of salmon. Smith and 

Margolis (1970) suggested that this cestode 

caused indirect damage to young salmonids. 

Hendee in 1980 believed it reduced the growth 

of brook trout in the state of New Hampshire, 

U.S.A. (in Hoffman, 1999). 

Acanthocephalus dirus had the highest prevalences, 

mean intensities, and mean abundances 

of all parasites found. It is widespread in Michigan 

trout farms and is common in some natural 

environments of Michigan (Muzzall, 1984). 

Muzzall (1984) also reported that the isopod, 

Caecidotea intermedius Forbes, 1876, was the 

intermediate host for this parasite in the Rogue 

River. In the present study, some individuals of 

all 3 trout species infected with 100 worms or 

more from 3 farms appeared emaciated, and the 

head appeared large for the size of the fish. Bullock 

(1963) demonstrated that the most pronounced 

effects of A. dirus (=A. jacksoni) in 

rainbow and brook trout in a New Hampshire 

hatchery were damage to the intestinal epithelium 

and proliferation of connective tissue, leading 

to malnutrition and emaciation. Furthermore, 

MUZZALL—PARASITES OF TROUT 183 

he stated (p. 33) that "this worm seriously impairs 

the health of the fish." Allison (1954) discussed 

the advancements in prevention and 

treatment of parasitic diseases of fish and listed 

13 parasitic genera that warranted discussion. 

However, A. dirus was not listed, and acanthocephalans 

in general receive little attention in 

hatchery manuals about fish diseases. It is not 

known if A. dirus was common when Allison 

wrote his article or if it has become increasingly 

common in Michigan. 

The presence of S. edwardsii on brook trout 

and its absence from rainbow and brown trout 

were not unexpected, because it parasitizes only 

the former species (Kabata, 1969). Some brook 

trout infested with S. edwardsii had 1 or both 

opercula folded underneath itself, and the distal 

portions of many gill filaments showed hyperplasia 

and clubbing. These characteristics also 

occurred on uninfected trout, suggesting previous 

infestation by this parasite. This copepod 

has a direct life cycle and is a common parasite 

of brook trout in Michigan (Allison and Latta, 

1969; Muzzall, 1984, 1986). The mean intensities 

of S. edwardsii are higher than those found 

on trout in Michigan lotic environments but are 

comparable to the high mean intensities found 

on brook trout in Michigan lakes by Allison and 

Latta (1969). Many studies on Salmincola spp. 

suggest that they debilitate their hosts but may 

not be direct causes of trout mortality. Allison 

and Latta (1969) found no relationship between 

S. edwardsii and brook trout mortality in Michigan 

lakes. 

Owners of 2 Michigan farms told me that they 

had seen a parasitic copepod on the gills of rainbow 

trout in their ponds. However, in this study, 

none was found infesting rainbow trout. A copepod 

that infests rainbow trout is Salmincola 

californiensis (Dana, 1853), which is native to 

streams in the Pacific Northwest, U.S.A., and 

Canada (Kabata, 1969). Hoffman (1984) reported 

on its eastward movement in North America, 

being transferred on live fish and with shipments 

of trout eggs. Sutherland and Wittrock (1985) 

believed that 5. californiensis entered central 

Iowa through the importation of infested rainbow 

trout from a Missouri farm. They found a 

mean intensity of 4.6 adults and suggested that 

this copepod may be responsible for host mortality 

if fish are sufficiently stressed. Perhaps this 

species has made its way to Michigan but is infrequent 

on rainbow trout in this state. 



Table 2. Prevalences, mean intensities, mean abundances of parasites found in Oncorhynchus mykiss, 

Salvelinus fontinalis, and Salmo trutta from farms in 1996, 1997, and 1998. 

Parasite 

Cestoda 

Eiibothrium salvelini 

Proteocephalus sp. 

Monogenea 

Gyrodactylus sp. 

Nematoda 

Truttaedacnitis sp. 

Acanthocephala 

Acanthocephalus dims 

Copepoda 

Salmincola edwardsii 

Myxozoa 

Myxobolus ccrebralis 

Ciliophora 

Capriniana sp. 

Chilodonella sp. 

Ichthyophthirus multifiliis 

Farm 

number, 

year 

4, 96 

4, 96 

5, 96 

5, 97 

6, 98 

8, 98 

9, 98 

11, 98 

3, 96 

12, 96 

1, 98 

12, 98 

4, 96 

1, 97 

11, 98 

11, 98 

5, 96 

2, 

4, 

4, 

4, 

5, 

5, 

5, 

5, 

7, 

8, 

10, 

11, 

5, 

5, 

5, 

8, 

5, 

5, 

5, 

96 

96 

96 

96 

96 

97 

98 

98 

98 

98 

98 

98 

96 

96 

97 

98 

96 

96 

97 

1, 96 

5, 96 

1, 97 

5, 97 

5, 97 

11, 98 

5, 97 

8, 98 

2, 96 

3, 96 

Trout 

species* 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

SF 

SF 

SF 

SF 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

SF 

SF 

SF 

SF 

ST 

ST 

ST 

SF 

SF 

SF 

SF 

OM 

OM 

SF 

OM 

OM 

OM 

No. examined 

25 

13 

15 

15 

16 

20 

18 

22 

14 

4 

19 

2 

25 

5 

22 

22 

15 

12 

13 

13 

25 

15 

15 

5 

16 

20 

20 

20 

22 

20 

16 

15 

6 

20 

15 

15 

10 

20 

12 

15 

15 

22 

20 

20 

12 

20 

Prevalence 

No. infected 

(%) 

1 (4) 

1 (8) 

3 (20) 

4(27) 

1 (6) 

14 (70) 

13 (72) 

5 (23) 

1 (7) 

1 (25) 

1 (5) 

1 (50) 

2(8) 

2(40) 

4(18) 

1 (5) 

3 (20) 

4(33) 

1 (8) 

1 (8) 

10 (40) 

15 (100) 

15 (100) 

1 (20) 

9(56) 

7(35) 

16 (80) 

1 (5) 

5(23) 

20 (100) 

3(19) 

15 (100) 

6 (100) 

20 (100) 

3 (20) 

15 (100) 

10 (100) 

20 (100) 

7(58) 

14 (93) 

2 (13) 

13 (59) 

1 (5) 

8 (40) 

3 (25) 

8 (40) 

Mean intensity 

±SD (max) 

1.7 ± 

2.0 ± 

2.4 ± 

20.7 ± 

2.6 ± 

1.5 ± 

93.5 ± 

2.3 ± 

44.7 ± 

56.5 ± 

3.2 ± 

17.9 ± 

98.1 ± 

5.6 ± 

42.7 ± 

2.0 ± 

46.3 ± 

11.7 ± 

36.0 ± 

3.3 ± 

75.7 ± 

1.0 

1.0 

1.2 (3) 

2.0 (5) 

1.0 

1.5 (5) 

44.9 (163) 

1.8 (5) 

2.0 

1.0 

1.0 

1.0 

1.0 

1.0 

0.6 (2) 

— 

1.0 

107.5 (209) 

1.0 

2.0 

1.6(5) 

43.0 (127) 

110.5 (432) 

22.0 

3.3 (11) 

39.4 (107) 

137.8 (490) 

1.0 

9.2 (22) 

48.6 (172) 

1.4 (4) 

43.8 (116) 

6.9 (20) 

24.3 (87) 

4.0 (8) 

80.6 (298) 

39.5 ± 20.9 (71) 

3.6 ± 3.1 (11) 

6.6 ± 6.4 (20) 

45.4 ± 31.2 (88) 

— 

— 

— 


— 

— - 

Mean abundance 

±SD 

0.04 ± 0.20 

0.08 ± 0.27 

0.33 ± 0.82 

0.53 ± 1.30 

0.06 ± 0.25 

1,65 ± 1.66 

14.94 ± 6.63 

0.59 ± 1.37 

0.14 ± 0.54 

0.25 ± 0.50 

0.05 ± 0.23 

0.50 ± 0.71 

0.08 ± 0.28 

0.40 ± 0.55 

0.27 ± 0.63 

— 

0.20 ± 0.41 

31.20 ± 72.6 

0.08 ± 0.28 

0.15 ± 0.56 

0.92 ± 1.49 

44.70 ± 43.00 

56.50 ± 110.50 

4.40 ± 9.80 

1.81 ± 2.93 

6.25 ± 23.87 

78.40 ± 128.90 

0.05 ± 0.22 

1.27 ± 4.67 

42.70 ± 48.60 

0.38 ± 1.03 

46.3 ± 43.8 

11.7 ± 6.9 

36.0 ± 24.3 

0.67 ± 2.05 

75.7 ± 80.6 

39.5 ± 20.9 

3.6 ± 3.1 

3.8 ± 5.8 

42.4 ± 32.2 

— 

— 

— 

— 

— 

—

Table 2. Continued. 

Parasite 

Trichodina sp. 

Mastigophora 

Ichthyobodo sp. 

Farm 

number, 

year 

1, 96 

1, 96 

2, 96 

4, 96 

4, 96 

1, 97 

1, 97 

1, 97 

7, 98 

9, 98 

11, 98 

12, 98 

1, 96 

1, 96 

1, 97 

1, 98 

8, 98 

12, 98 

3, 96 

Trout 

species* 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

OM 

SF 

SF 

SF 

SF 

SF 

SF 

OM 

No. examined 

10 

24 

12 

13 

25 

10 

20 

5 

20 

18 

22 

11 

20 

10 

12 

19 

6 

2 

20 


Prevalence 

No. infected Mean intensity 

(%) ±SD (max) 

10 (100) — 

14 (58) 

3 (25) 

2(15) 

7(28) 

3 (30) 

2(10) 

1 (20) 

4 (20) 

10 (56) 

4(18) 

2(18) 

5 (25) 

10 (100) 

5 (42) 

5 (26) 

2(33) 

1 (50) 

2 (10) — 

OM = Oncorhynchus mykiss; SF = Salvelinus fontinalis; ST = Salmo trutta. 

Myxobolus cerebralis has been present in 

Michigan waters since at least 1968, when it was 

discovered in 3 commercial trout hatcheries. 

Yoder (1972) discussed the spread of M. cerebralis 

into native trout populations in the Tobacco 

River, Michigan, from 1 of these hatcheries. 

The protozoan spread down the first 6 mi of water, 

and factors involved in this spread were the 

high incidence of disease at the hatchery, abundance 

of susceptible trout, and trout movement. 

In 1998 and 1999, M. cerebralis was reported 

from at least 6 privately owned trout farms in 

Michigan. It has been suggested that it was endemic 

in 1 or more facilities and transferred to 

other facilities with infected fish or by piscivorous 

birds that ate infected fish. In the present 

study, M. cerebralis-infected trout were detected 

in 3 farms. Infected trout, however, did not 

have clinical symptoms. Furthermore, Sutherland 

(1999) reported that M. cerebralis has been 

found in fish from the Au Sable and Manistee 

rivers in lower Michigan. 

In the present study, parasites and their numbers 

infecting trout in a pond may dramatically 

change the next time the fish are sampled and 

examined. One reason for this is that trout are 

moved into and out of facilities during the year. 

An example of this was evident at 1 farm, when, 

Mean abundance 

±SD 

— 

— 

— 

— 

— 

— 

— 

— 

— 

— 

— 

— 

— 

— 

— 

— 

— 

in March 1997, 4 parasite species infected trout 

and 2 were common (Table 4). In July, only 1 

species infrequently infected trout after the infected 

trout were moved out and uninfected ones 

were moved in. Also, informing the owner of 

the facility on the parasites found can affect infection 

levels from 1 sampling date to the next. 

After an owner was informed that brook trout 

were infested with S. edwardsii, he told me that 

"a treatment had been done to the pond." Approximately 

2 months later, the prevalence and 

mean intensity of S. edwardsii on trout from the 

same pond were dramatically reduced. Another 

suggestion for these differences is that parasite 

species may exhibit a seasonal cycle in their occurrence. 

Hare and Frantsi (1974) found 12 parasite 

species, 10 parasite species, and 1 parasite species 

infecting, respectively, Atlantic salmon, 

Salmo solar Linnaeus, 1758; brook trout; and 

rainbow trout, in 13 Canadian hatcheries in the 

Maritime provinces. Hexamita salmonis (Moore, 

1923) Wenyon, 1926; Trichophyra piscium 

Buetschli, 1889; Diplostomum spathaceum (Rudolphi, 

1819) Braun, 1893; Acanthocephalus later 

alls (Leidy, 1851); and S. edwardsii were 

considered to be serious fish pathogens, based 

on the work of other authors. Buchmann and 


—


Table 3. Parasites of Oncorhynchus mykiss examined 

from 23 ponds, Salvelinus fontinalis from 13 

ponds, and Salmo trutta from 6 ponds in 1996,1997, 

and 1998. 

Cestoda 

No. (%) of 

ponds where 

Trout parasites 

Parasite species* occurred 

Eubothrium salvelini 

Proteocephalus sp. 

Monogenea 

Gyrodactylus sp. 

Nematoda 




Copepoda 


Myxozoa 

Myxobolus cerebralis 

Ciliophora 

Capriniana sp. 

Chilodonella sp. 

Ichthyophthirus multifiliis 

Trichodina sp. 

Mastigophora 

Ichthyobodo sp. 

OM 

SF 

OM 

OM 

OM 

OM 

SF 

ST 

SF 

OMf 

OMt 

SFt 

OM 

OM 

OM 

OM 

SF 

OM 

9(39) 

4(31) 

3 (13) 

1 (4) 

1 (4) 

12 (52) 

4(31) 

2(33) 

4(31) 

2(17) 

1 (8) 

1 (25) 

1 (4) 

1 (4) 

1 (4) 

12 (52) 

6 (46) 

1 (4) 

* OM = Oncorhynchus mykiss; SF = Salvelinus fontinalis; 

ST = Salmo trutta. 

t OM examined from 12 ponds and SF from 4 ponds in 1997 

and 1998. 

Bresciani (1997) listed investigations performed 

on the parasites of farmed salmonids, and reported 

22 parasite species (12 protozoans and 10 

metazoans) infecting 805 pond-reared rainbow 

trout from 5 freshwater farms in Denmark. 

Based on these and the present studies, there is 

a relationship between the number of trout examined 

and number of parasite species found. 

As the number of fish examined increases, so 

does the number of parasite species found. 

Hnath (1993) suggested that a sample size of 60 

individuals should be examined from a population 

of 2,000 fish or more in a pond in order to 

detect a pathogen. The numbers of parasite species 

found in rainbow and brook trout in the present 

study are low compared with the numbers 

found by Hare and Frantsi (1974) and by Buchman 

and Bresciani (1997). More rainbow and 

brook trout were examined in those studies than 

in the present one. 

The total numbers of parasite species found 

in rainbow, brook, and brown trout in this study 

are low compared with the numbers for each 

species listed by Hoffman (1999) in North 

America. This may be explained by the artificial 

conditions in trout farms, which harbor very few 

potential intermediate invertebrate hosts. In most 

ponds, snails, which serve as intermediate hosts 

for digenetic trematodes, were never collected. 

In contrast, protozoans with direct life cycles are 

easily introduced and spread between fish. In 

conversations with several farmers, it was apparent 

that they used several "antiparasitic" 

drugs to treat against ectoparasitic infections 

when they were aware that their fish exhibited 

signs of infection. This treatment regime also 

explains the paucity of parasites. The current 

and increasing use of well water and spring wa- 

Table 4. Prevalence (P), mean abundance (MA), and estimated number (EN) of parasites from 1 farm 

in March and July 1997. 

Parasite 


Eubothrium salvelini 



Trout 

species 

(»)* 

SF (20) 

ST (20) 

OM (15) 

OM (15) 

OM (15) 

SF (20) 

P 

100 

100 

100 

20 

7 

100 

March 

42.7 

36.0 

44.7 

0.33 

0.20 

3.60 

MA ± SD 

(Max.) 

± 49 (172) 

± 24 (87) 

± 43 (127) 

± 0.82 (3) 

± 0.41 

± 3 (11) 

EN 

106,750 

90,000 

312,900 

462 

98 

9,000 

Trout 

species 

(/>)* 

SF(16) 

ST (15) 

OM (15) 

OM (15) 

OM (15) 

SF (16) 

SF = Salvelinus fontinalis; ST = Salmo trutta; OM = Oncorhynchus mykiss. (no. examined) 


P 

19 

20 

0 

0 

0 

0 

July 

MA ± SD 

(Max.) EN 

0.38 ± 1.03 (4) 14 

0.67 ± 2.05 (8) 27 

— — 

— — 

— — 

—

ter and fiberglass or concrete ponds and tanks 

will also reduce the incidence of parasites. 

The effects of natural bodies of water and the 

fish in them serving as a source of parasites in 

culture should be addressed. This involves facilities 

with "flow-through" systems (those that 

receive water from lentic or lotic environments). 

Eggs, other infective stages, and hosts can be 

carried with water that flows into and through 

facilities. Obviously this facilitates infection of 

fish. In general, trout in the present study being 

raised in flow-through systems had more parasite 

species and more individuals of the species 

present in comparison with the other systems. 

Similarly, Valtonen and Koskivaara (1994), 

studying the relationships between parasites of 

wild and cultured fishes in 2 lakes and a fish 

farm in Finland, reported that the source of parasites 

in the fish farm was the water-supplying 

lake. 

Cone and Cusack (1988) reported on the occurrence 

of 2 monogeneans, Gyrodactylus colemanensis 

Mizelle and Kritsky, 1967, and Gyrodactylus 

salmonis Yin and Sproston, 1948, on 

brook and rainbow trout, and Atlantic salmon, 

Salmo salar Linnaeus, 1758, in a farm in Nova 

Scotia, Canada, and discussed the origins of infection 

and their dispersal in the farm. Sources 

of infection with G. salmonis were stocks of infected 

rainbow trout brought into the facility 

from another farm, as well as wild infected Atlantic 

salmon and brook trout gaining access to 

the hatchery. Parasite dispersal in the farm involved 

infected fish jumping and wriggling from 

one pond to the next and the workers using 

transfer nets and buckets that contained live parasites. 

Also, brood stocks were infected and constituted 

internal reservoirs of infection. 

The effects of fish culture on natural waters 

receiving water from the farms has been a subject 

of increasing debate. If surveillance of parasites 

in the water above and below the fish facility 

is not continuous, little will be known 

about where the parasite really originated or 

how long it has been present. Regarding M. cerebralis, 

fish known to have whirling disease 

were imported into a commercial trout farm in 

Michigan in 1968. The receiving stream (a 

brook and brown trout stream) of this facility 

yielded M. cerefera/zs—infected rainbow trout escapees 

directly below the positive facility effluent. 

Valtonen and Koskivaara (1994) suggested 

that the farm itself was unlikely to affect the fish 


parasite fauna of the water-recipient lake, although 

some ectoparasites could originate from 

the farm. 

Muzzall (1995c), studying the parasites of 

pond-reared yellow perch, Perca flavescens 

(Mitchill, 1814) in Michigan, suggested that 

conditions of a pond associated with producing 

a good crop of fish also support a good crop of 

helminths that infect fish. Later, Muzzall (1996) 

referred to this as "the good fish crop-good helminth 

crop" relationship. Based on the results 

of the parasites infecting trout in the present 

study, this relationship does not occur. Of the 

parasite species found by Muzzall (1995c) infecting 

perch, 8 were represented as only larval 

stages, 6 of which were digenetic trematodes. 

Only 2 genera (generalist protozoans, Trichodina 

sp., Capriniana sp.) infesting perch were also 

found infesting trout. The dramatic differences 

in parasites found in yellow perch and trout from 

culture conditions can be explained by many 

factors. Probably the most important are the 

types of ponds used to culture the particular species, 

water temperatures, water sources, whether 

ponds are periodically drained, the surroundings 

of the ponds, and animals associated with the 

ponds. 

The state of control and prevention of parasites 

and diseases of fishes in culture in Michigan 

is difficult to assess. I refer to it as "crisis 

fisheries health," which can be defined as follows: 

"Some state and university officials, extension 

specialists, aquaculture centers, and trout 

farmers are not apparently concerned with fish 

health in aquaculture and in nature unless there 

is a crisis health problem, then action takes 

place." This approach is understandable with so 

many interested parties having different motives 

and the low priority of funding for parasite and 

disease work. I suggest that more studies on fish 

parasites and diseases in Michigan be encouraged 

and supported by the interested groups. 

Surveillance and surveys are needed to determine 

what parasites are infecting trout in culture 

conditions and in the surrounding waters. 

As mentioned earlier, growers in Michigan are 

involved in 3 activities in producing and selling 

trout. In regard to the first, the sale of infected 

trout for stocking could transfer some parasites 

to other fish directly or contaminate the watershed 

with other parasites. However, most if not 

all parasites reported in this study have been 

found infecting trout in the wild. Second, no par- 



asites were found that could infect humans if 

poorly cooked infected meat was eaten. Third, 

the sale of infected fish in fee-fishing ponds 

should not play a role in transmitting parasites, 

unless these fish are placed in other environments 

or the ponds have effluents to public waters. 

Obviously it should be emphasized that if 

trout are not routinely examined, light infections 

will not be noticed; when the infections do become 

evident, it may be too late to help the diseased 

fish. 


I thank the trout farmers in Michigan who 

generously provided fish for this study, and Liz 

Osmer, Chris Henderson, Mindy Place, and Amy 

Hawkins for their technical assistance. I gratefully 

acknowledge Bob Baldwin, president of 

the Michigan Aquaculture Association, for making 

this study possible, and John Hnath for reviewing 

an early draft of the manuscript and 

sharing information with me on parasites. 


Allison, L. N. 1954. Advancements in prevention and 

treatment of parasitic diseases of fish. Transactions 

of the American Fisheries Society 83:221-228. 

, and W. C. Latta. 1969. Effects of gill lice 

(Salmincola edwardsii) on brook trout (Salvelinus 

fontinalis) in lakes. Michigan Department of Natural 

Resources, Research and Development Report 

No. 189. 32 pp. 

Anonymous. 1997. Michigan agricultural statistics 

1996—97. Trout. Commercial Fisheries Newsline 

(Michigan Sea Grant Extension)(December, 1997) 

16(2): 14. 

Boyce, N. P. J. 1969. Parasite fauna of pink salmon 

(Oncorhynchus gorbuscha) of the Bella Coola 

River, central British Columbia, during their early 

sea life. Journal of the Fisheries Research Board 

of Canada 26:813-820. 

Buchmann, K., and J. Bresciani. 1997. Parasitic infections 

in pond-reared rainbow trout Oncorhynchus 

mykiss in Denmark. Diseases of Aquatic Organisms 

28:125-138. 

Bullock, W. L. 1963. Intestinal histology of some salmonid 

fishes with particular reference to the histopathology 

of acanthocephalan infections. Journal 

of Morphology 112:23-44. 

Cone, D. K., and R. Cusack. 1988. A study of Gyrodactylus 

colemanensis Mizelle and Kritsky, 

1967 and Gyrodactylus salmonis (Yin and Sproston, 

1948) (Monogenea) parasitizing captive salmonids 

in Nova Scotia. Canadian Journal of Zoology 

66:409-415. 

Hare, G. M., and C. Frantsi. 1974. Abundance and 

potential pathology of parasites infecting salmonids 

in Canadian Maritime hatcheries. Journal of 


the Fisheries Research Board of Canada 31:1031- 

1036. 

Hernandez, A. D., and P. M. Muzzall. 1998. Seasonal 

patterns in the biology of Eubothrium salvelini 

infecting brook trout in a creek in lower 

Michigan. Journal of Parasitology 84:1119—1123. 

Hnath, J. G., ed. 1993. Great Lakes fish disease control 

policy and model program (supersedes September 

1985 edition). Great Lakes Fishery Commission 

Special Publication 93-1:1-38. 

Hoffman, G. L. 1984. Salmincola californiensis continues 

the march eastward. Fish Health Section, 

American Fisheries Newsletter 12:4. 

. 1999. Parasites of North American Freshwater 

Fishes. Cornell University Press, Ithaca, New 

York. 539 pp. 

Kabata, Z. 1969. Revision of the genus Salmincola 

Wilson, 1915 (Copepoda: Lernaeopodidae). Journal 

of the Fisheries Research Board of Canada 26: 

2987-3041. 

Lorn, J., and I. Dykova. 1992. Protozoan Parasites of 

Fishes. Developments in Aquaculture and Fisheries 

Science, 26. Elsevier Science Publishers, New 

York. 315 pp. 

Muzzall, P. M. 1984. Parasites of trout from four lotic 

localities in Michigan. Proceedings of the Helminthological 


••—. 1986. Parasites of trout from the Au Sable 

River, Michigan, with emphasis on the population 

biology of Cystidicoloides tenuissima. Canadian 


. 1993. Parasites of parr and lake age chinook 

salmon, Oncorhynchus tshawytscha, from the Pere 

Marquette River and vicinity, Michigan. Journal 

of the Helminthological Society of Washington 

60:55-61. 

. 1995a. Parasites of pacific salmon, Oncorhynchus 

spp., from the Great Lakes. Journal of Great 

Lakes Research 21:248-256. 

. 1995b. Parasites of lake trout, Salvelinus namaycush, 

from the Great Lakes: a review of the 

literature 1874-1994. Journal of Great Lakes Research 

21:594-598. 

. 1995c. Parasites of pond-reared yellow perch 

from Michigan. Progressive Fish-Culturist 57: 

168-172. 

. 1996. Parasites and diseases of pond-reared 

walleye and yellow perch. Aquaculture, November/December:49-61. 

Newman, J. R., and N. R. Kevern. 1994. Production 

of Michigan Aquacultural Products. Research Report 

526. Michigan Agricultural Experiment Station, 

Michigan State University, East Lansing. 78 

pp. 

Sawyer, T. K., J. G. Hnath, and J. F. Conrad. 1974. 

Thecamoeba hoffmani sp. n. (Amoebida: Thecamoebidae) 

from gills of fingerling salmonid fish. 

Journal of Parasitology 60:677-682. 

Smith, H. D., and L. Margolis. 1970. Some effects 

of Eubothrium salvelini (Schrank, 1790) on sockeye 

salmon, Oncorhynchus nerka (Walbaum), in 

Babine Lake, British Columbia. Journal of Parasitology 

56 (4, section 2, part l):321-322. (Abstract.)

Sutherland, D. R. 1999. Assessing the risk of whirling 

disease becoming established in the Great Lakes. 

Commercial Fisheries Newsline 18:10. 

, and D. D. Wittrock. 1985. The effects of 

Salmincola californiensis (Copepoda: Lernaeopodidae) 

on the gills of farm-raised rainbow trout, 

Salmo gairdneri. Canadian Journal of Zoology 63: 

2893-2901. 


Valtonen, E. T., and M. Koskivaara. 1994. Relationships 

between the parasites of some wild and culture 

fishes in two lakes and a fish farm in central 

Finland. International Journal for Parasitology 24: 

109-118. 

Yoder, W. G. 1972. The spread of Myxosoma cerebralis 

into native trout populations in Michigan. 

Progressive Fish-Culturist 34:103-106. 

Museums for Depositing of Specimens 

It is the policy of Comparative Parasitology to require the deposit of type and voucher specimens to document 

survey or taxonomic papers. Moreover, the value of any paper is enhanced by the deposit of reference specimens. The 

following museum collections in the United States will accept such specimens, provide professional curatorial services 

for their preservation, provide accession numbers for inclusion in your publication, and make the deposited materials 

available for study by researchers worldwide. If other museum collections are used, they must provide comparable 

services, and information for contacting the museum must be provided in your publication. Materials designated as type 

specimens often carry the implication that they have been placed under such curatorial care; therefore, specimens retained 

in private collections should not carry type designations. 

The Editors would appreciate receiving additional information regarding other institutions that provide equivalent 

services. This information will be published in a future number of Comparative Parasitology and added to the directory 

that we are compiling for publication on the Society's website. 

Collection and domestic and international shipment of wildlife, including invertebrates, are governed by the laws 

and regulations of the countries of origination and destination. Detailed information for the United States is available 

through the U.S. Fish and Wildlife Service website, http://www.fws.gov. It is also wise to contact the appropriate curator/ 

collections manager of the receiving institution for special instructions before sending specimens for deposit. 

Helminths and Protozoans 

U.S. National Parasite Collection 

Biosystematics & National Parasite Collection Unit 

USDA, ARS, LPSI, BARC East No. 1180 

Beltsville, MD 20705-2350 

Curator: Dr. Eric P. Hoberg 

e-mail: ehoberg@lpsi.barc.usda.gov 

Telephone: (voice) 301-504-8444; (fax) -8979 

Homepage: http://www.lpsi.barc.usda.gov/bnpcu 

Harold W. Manter Laboratory of Parasitology 

University of Nebraska State Museum 

W-529 Nebraska Hall, 

University of Nebraska 

Lincoln, NE 68588-0514 

Curator: Dr. Scott Lyell Gardner 

e-mail: slg@unl.edu 


Homepage: http://lamarck.unl.edu/research/parasitology 

Ticks 

U.S. National Tick Collection 

Institute of Arthropodology and Parasitology 

Georgia Southern University 

Statesboro, GA 30460-8056 

Curator: Dr. James E. Keirans 

e-mail: jkeirans@gasou.edu 


Homepage: http://www2.gasou.edu/iap/ 

Mites 

J. Ralph Lichtenfels and Janet W. Reid 

National Mite Collection 

Systematic Entomology Laboratory 

USDA, ARS, BARC West No. 047 

Beltsville, MD 20705-2350 

Curator: Dug Miller 

e-mail: dmiller@sel.barc.usda.gov 


Homepage: http://www.sel.barc.usda.gov/Selhome/ 

selhome.htm 

Crustaceans, Hirudineans 

Department of Invertebrate Zoology 

National Museum of Natural History 

Smithsonian Institution 

Washington, D.C. 20560-0163 

Collections Manager: Cheryl Bright 

e-mail: bright.cheryl@nmnh.si.edu 


Homepage: http://www.nmnh.si.edu/iz/collect.html 

Insects and Their Allies, Including Spiders 

Department of Entomology 

National Museum of Natural History 

Smithsonian Institution 

Washington, D.C. 20560-0165 

Collections Manager: Dr. David Furth 

e-mail: furth.david@nmnh.si.edu 

Telephone: (voice) 202-357-3146; (fax) 202-786-2894 

Homepage: http://entomology.si.edu 




Six New Host Records and an Updated List of Wild Hosts for 

Neobenedenia melleni (MacCallum) (Monogenea: Capsalidae) 

STEPHEN A. BuLLARD,1-5 GEORGE W. BENZ,2 ROBIN M. OVERSTREET,' 

ERNEST H. WILLIAMS, JR.,3 AND JAY HEMDAL4 

1 Gulf Coast Research Laboratory, Department of Coastal Sciences, University of Southern Mississippi, 703 

East Beach Drive, Ocean Springs, Mississippi 39564, U.S.A. (e-mail: ash.bullard@usm.edu; 

Robin.Overstreet@usm.edu), 

2 Tennessee Aquarium and Southeast Aquatic Research Institute, 1 Broad Street, RO. Box 1 1048, 

Chattanooga, Tennessee 37401, U.S.A. (e-mail: gwb@sari.org), 

3 Department of Marine Sciences, University of Puerto Rico, RO. Box 908, Lajas, Puerto Rico 00667-0908, 

U.S.A. (e-mail: bert@rmocfis.uprm.edu), and 

4 Toledo Zoo, 2700 Broadway, Toledo, Ohio 43609, U.S.A. (e-mail: jay.hemdal@toledozoo.org) 

ABSTRACT: Six new host records and an updated list of wild hosts for Neobenedenia melleni (MacCallum) 

(Monogenea: Capsalidae) are provided. We report specimens of N. melleni from the skin of a whitefin sharksucker 

(Echeneis neucratoides Zuieuw [Echeneidae]) caught off Mayagiiez, Puerto Rico; from the skin of a 

mosquitofish (Gambusia xanthosoma Greenfield [Poeciliidae]) caught in Little Salt Creek, Grand Cayman Island, 

British West Indies; from a freshwater immersion bath of red grouper (Epinephelus morio (Valenciennes) [Serranidae]) 

caught in the Gulf of Mexico off Sarasota, Florida, U.S.A.; from the skin of a garden eel (Heteroconger 

hassi (Klausewitz and Eibl-Eibesfeldt) [Congridae]) in the Toledo Zoo, Toledo, Ohio, U.S.A.; from the skin of 

a raccoon butterflyfish (Chaetodon liinula (Cuvier) [Chaetodontidae]) in the Fort Wayne Children's Zoo, Fort 

Wayne, Indiana, U.S.A.; and from the gill cavity of a red snapper (Lutjanus campechanus (Poey) [Lutjanidae]) 

in holding facilities at the Gulf Coast Research Laboratory, Ocean Springs, Mississippi, U.S.A. Neobenedenia 

melleni had not been reported previously from a suspected wild host in the Gulf of Mexico (i.e., E. morio) or 

from a member of Echeneidae, Atheriniformes, or Anguilliformes. Published host records indicate that N. melleni 

exhibits a relatively low degree of host specificity among captive and wild hosts; in nature, N. melleni infests 

predominantly shallow-water or reef teleosts. 

KEY WORDS: Neobenedenia melleni, Echeneis neucratoides, Gambusia xanthosoma, Epinephelus morio, Heteroconger 

hassi, Chaetodon lunula, Lutjanus campechanus, Monogenea, Capsalidae, host specificity, zoogeography, 

public aquaria, aquaculture, U.S.A., Puerto Rico, British West Indies, Florida, Mississippi, Gulf of Mexico. 

The capsalid Neobenedenia melleni (Mac- accounts of TV. melleni infesting wild hosts (see 

Callum, 1927) is relatively unusual among references in Table 1) are relatively scarce, and 

members of Monogenea in that it has been re- little is known about the breadth of host speciported 

from a wide range of hosts. This capsalid ficity exhibited by this parasite in nature. Thereinfests 

the eyes, fins, gill cavity, nasal cavity, fore, reports of TV. melleni from wild hosts are 

and skin of over 100 species of marine teleosts significant because they offer insight into the 

(Whittington and Horton, 1996). Most of these natural geographic distribution and host range of 

records are from fishes in aquaria and aquacul- this parasite. We report 6 new host records for 

ture systems where the parasite is identified as N. melleni: 3 from wild fishes and 3 from capa 

lethal pathogen (e.g., MacCallum, 1927; Jahn 

and Kuhn, 1932; Nigrelli and Breder, 1934; 

tive fishes, 

Mueller et al., 1994). However, there is no report Materials and Methods 

of disease associated with infestations of TV. me/leni 

among wild fishes. Neobenedenia melleni 

Worms were fixed in 10% neutral buffered formalin, 

70% ethanol or Bouin,s fixadve Eight worms were 

had been reported previously from wild hosts in stained in Van Cleave's hematoxylin containing sevthe 

Caribbean Sea, Gulf of California, and east- eral additional drops of Ehrlich's hematoxylin and 

ern Pacific Ocean off the coasts of Chile, Mex- were then dehydrated to 70% ethanol. Several drops 

ico, and 

j *u 

the 

TT 

United 

-4- j o«. 

States 

« /T 

(Table 

ui i\ 

1). 

ui- 

Published 

u j of aqueous 

. J' . 

saturated 

, , 

lithium 

. 

carbonate 

. „ ,_. 

were 

, 

then 

. 

add- 

ed, followed by several drops of 6% butylamme 

. 

solution. 

Stained worms were dehydrated in an ethanol 

series, cleared in clove oil, and mounted permanently 

5 Corresponding author. on glass slides using neutral Canada balsam. Five 

190 


Table 1. Wild hosts for Neobenedenia melleni (MacCallum, 1927). 

Host Site Lo 

ATHERINIFORMES 

Poeciliidae 

Gambusia xanthosoma Greenfield. 1983 

Skin Little Salt Creek, Gra 

ish West Indies 

SCORPAENIFORMES 

Skin Southeast Pacific Oc 

Gills Northeast Pacific Oce 

Washington. U.S.A 

Mouth and skin Northeast Pacific Oc 

California, U.S.A. 

Scorpaenidae 

Sebastes capensis (Gmelin. 1789) 

Sebastes melanops Girard, 1856 (as Sebastodes melanops) 

Sebastes serranoides (Eigenmann and Eigenmann, 1890) 

Hexagrammidae 

Hexagrammos decagrammus (Pallas, 1810) 

Gills Northeast Pacific Oce 

Washington. U.S.A 

Cottidae 

Leptocottus armatus Girard, 1854 

Not indicated Northeast Pacific Oce 

nia, U.S.A. 

Gills Caribbean Sea off La 

PERCIFORMES 

Serranidae 

Epinephelus guttatus (Linnaeus, 1758) 

Not indicated Gulf of Mexico off S 

Epinephelus morio (Valenciennes, 1828) 

Not indicated Caribbean Sea off Bi 

Epinephelus striatus (Bloch, 1792) 

Gills Gulf of California of 

Mycteroperca rosacea (Gilbert, 1892) (as Mycteroperca pardalis) 

Skin Caribbean Sea off M 

Echeneidae 

Echeneis neucratoides Zuieuw. 1789 

Not indicated Caribbean Sea off Bi 

Lutjanidae 

Lutjanus apodus (Walbaum, 1892) (as Lutianus apodus) 


Table 1. Continued. 

Host Site Loca 

Sparidae 

Archosargus probatocephalus (Walbaum 1792)j 

Northeast Pacific Ocea 

nia, U.S.A. 

Eyes and skin 

Caribbean Sea off Bim 

Not indicated 

Chaetodontidae 

Chaetodon capistratus Linnaeus, 1758 


Not indicated 

Chaetodon ocellatits Bloch, 1787 


Not indicated 

Chaetodon striatus Linnaeus, 1758 


Not indicated 

Pomacanthidae 

Holocanthus ciliaris (Linnaeus, 1758) 


Not indicated 

Holocanthus tricolor (Bloch, 1795) 


Not indicated 

Pomacanthus arcuatux (Linnaeus, 1758) 


Not indicated 

Pomacanthus paru (Bloch, 1787) 

Skin 

Kyphosidae 

Cirella nigricans (Ayres, 1860) 

Northeast Pacific Ocean 

nia, U.SA. 


land, California, U.S 


land, California, U.S 

Fins and skin 

Skin 

Medialuna californiensis (Steindachner, 1876) 

Exterior 

Embiotocidae 

Embiotoca jacksoni Agassiz, 1853 


ta Barbara, Californi 







Exterior of head 

Embiotoca lateralis Agassiz, 1854 

Exterior 

Exterior 

Rhacochilus vacca (Girard, 1855) (as Damalichthys vacca) 


CB < 

•£ a 

o '= 

2 I 

BULLARD ET AL.—HOSTS OF NEOBENEDENIA MELLENl 193 

worms intended for study using Nomarski illumination 

were dehydrated, cleared in clove oil, and mounted 

unstained in neutral Canada balsam. Worms were identified 

using the original description of N. melleni (as 

Epibdella melleni MacCallum, 1927), the redescription 

of N. melleni contained in a recent revision of Neobenedenia 

Yamaguti, 1963, and the key to the species 

of Neobenedenia (see Whittington and Horton, 1996). 

We primarily used 1) anterior attachment organs circular 

and not bipartite; 2) anterior hamuli recurved, 

nonserrated (i.e., smooth), and robust (i.e., width usually 

greater than that of both accessory sclerites and 

posterior hamuli and with root of consistent width 

along total length [i.e., root not tapered, constricted, or 

pinched]); 3) glands of Goto not evident, and 4) other 

specific features indicated by Whittington and Horton 

(1996). Nomenclature used herein for members of 

Neobenedenia follows that of Whittington and Horton 

(1996). Specimens of N. melleni from Gambusia xanthosoma 

(Poeciliidae) and Lutjanus campechanus 

(Poey, 1860) (Lutjanidae) were deposited in the United 

States National Parasite Collection (USNPC) at Beltsville, 

Maryland, U.S.A. (USNPC Nos. 089159 and 

089160), and specimens from Echeneis neucratoides 

(Echeneidae), Epinephelus morio (Serranidae), Heteroconger 

hassi (Klausewitz and Eibl-Eibesfeldt, 1959) 

(Congridae), and Chaetodon lunula (Cuvier, 1831) 

(Chaetodontidae) were deposited there (USNPC Nos. 

089161, 089162, 089163, and 089164) and in the helminth 

collections of the H. W. Manter Laboratory 

(HWML) of the University of Nebraska State Museum 

at Lincoln, Nebraska, U.S.A. (HWML Nos. 15063, 

15064, 15065, and 15066). 

Results and Discussion 

Regarding our new host records, 2 specimens 

of N. melleni were collected from the skin of a 

whitefin sharksucker (E. neucratoides), a rernora 

that was attached to a West Indian manatee (Trichechus 

manatus Linnaeus, 1758 [Trichechidae]) 

off Mayagiiez, Puerto Rico. This is the first report 

of N. melleni from a remora and may help 

to explain in part the wide geographic distribution 

of N. melleni. Although carriers of infested 

remoras may not travel between oceans, infested 

remoras may transfer infestations of N. melleni 

among fish, mammalian, and turtle species and 

individuals with which they associate. In addition, 

remoras can attached to or mingle with 

their carriers for prolonged periods of time. This 

habit may provide N. melleni opportunity to infest 

the remora's carrier host or other fishes in 

close proximity to the infested remora. Various 

cleaner fishes (e.g., bluehead wrasse, Thalassoma 

bifasciatum (Bloch, 1791) [Labridae]; neon 

goby, Gobiosoma oceanops (Jordan, 1904) [Gobiidae]; 

and cleaning goby, Gobiosoma genie 

Bohlke and Robins, 1968) were effective in controlling 

infestations of N. melleni among aquar- 



ium-kept fish (see Cowell et al., 1993). Some 

species of remora feed on ectoparasites (Cressey 

and Lachner, 1970), and, because of this, aquaculturists 

eventually may use remoras to control 

infestations of N. melleni on large hosts. However, 

as previously suggested, remoras may 

transport worms to adjacent groups of fishes. 

A specimen of TV. melleni was collected from 

the skin of a mosquitofish (G. xanthosoma) from 

Little Salt Creek (western shore of North Sound, 

Grand Cayman, British West Indies). Neobenedenia 

melleni had not been reported previously 

from a member of Atheriniformes or from the 

western Caribbean Sea. The specimen of N. melleni 

was conspicuous, 3 mm in total length, and 

attached to the dorsal surface of the head at the 

level of the eyes of a mosquitofish that was 33 

mm in total length. Gambusia xanthosoma is apparently 

endemic to the high salinity mangrove 

habitats throughout North Sound (Abney and 

Heard, personal communication); therefore, it is 

of ecological interest to report on the occurrence 

of nonendemic parasites, such as N. melleni, that 

infest a wide range of hosts and that are identified 

as lethal pathogens among confined fishes. 

Nigrelli (1947) reported several wild hosts for 

N. melleni in the Caribbean Sea off Bimini (see 

Table 1). Robinson et al. (1992) and Hall (1992) 

reported heavy infestations of N. melleni among 

cultured, seawater-acclimated red hybrid tilapia 

in floating cages off southern Jamaica. Cowell 

et al. (1993) reported infestations of TV. melleni 

on Florida red tilapia (descendants of an original 

cross between Oreochromis urolepis hornorum 

(Norman, 1922) [Cichlidae] and Oreochromis 

mossambicus (Peters, 1852)) in aquaria at the 

Caribbean Marine Research Center (CMRC), 

Lee Stocking Island, Exuma Cays, Bahamas. 

However, because N. melleni has a broad host 

range and wide geographic distribution and 

heavily infests some hosts in aquaculture, we 

cannot determine if, when, or how it was introduced 

to the endemic population of G. xanthosoma. 

At least 3 specimens of TV. melleni infested 

the red grouper (E. morio); they were caught off 

Sarasota, Florida, U.S.A., in January 1993. Material 

of TV. melleni was collected from a freshwater 

immersion bath at the Mote Marine Laboratory 

(MML), Sarasota, Florida, when the fish 

were initially treated after being captured from 

the Gulf of Mexico. Nevertheless, TV. melleni later 

became established in culture facilities at the 


MML. Neobenedenia melleni was previously reported 

from E. morio and Mycteroperca tnicrolepis 

(Goode and Bean, 1879) (Serranidae) in 

recirculating-seawater tanks in northwestern 

Florida (Florida State University Marine Laboratory, 

Turkey Point, Florida, U.S.A.) by Mueller 

et al. (1994) and from other members of the 

sea bass family in the Caribbean Sea and the 

Gulf of California (see Table 1); however, this 

is the first report of TV. melleni from a suspected 

wild host in the Gulf of Mexico. 

We also report numerous adult and juvenile 

specimens of TV. melleni from the skin of a garden 

eel (H. hassi) from the Toledo Zoo, Toledo, 

Ohio, U.S.A. This is the first report of TV. melleni 

from any member of Anguilliformes, and to the 

best of our knowledge, it is also the first report 

of TV. melleni from a host that lacks scales. 

Whereas the exact geographic origin of the eel 

was not known, we suspect that it became infested 

while confined in a compartmentalized 

quarantine system at the Toledo Zoo. One of us 

(J.H.) observed a cream angelfish (Apolemichthys 

xanthurus (Bennett, 1832) [Pomacanthidae]) 

in this same water system that harbored 

numerous specimens of a platyhelminth on its 

skin that were probably TV. melleni. Nigrelli and 

Breder (1934) reported that some angelfishes 

were foci for epidemics of TV. melleni in the New 

York Aquarium. Specimens of TV. melleni have 

yet to be reported from A. xanthurus. However, 

because the aforementioned worms from this 

host were not available for identification, we did 

not report this fish as a host for TV. melleni. 

Numerous specimens of TV. melleni were also 

collected from a raccoon butterflyfish (C. lunula) 

that died while in quarantine at the Fort 

Wayne Children's Zoo, Fort Wayne, Indiana, 

U.S.A. We are not certain of the exact geographic 

origin of that wild-caught fish or whether it 

was infested in the wild. However, C. lunula is 

a reef species that ranges from East Africa to 

Polynesia (Randall et al., 1990), and that raccoon 

butterflyfish most likely came from there 

(i.e., Indo-Pacific Region). Neobenedenia melleni 

has been reported from 3 members of Chaetodon 

in the Caribbean Sea (Nigrelli, 1947; Table 

1). 

A single specimen of TV. melleni was collected 

from the gill cavity of a red snapper (L. campechanus) 

caught in the northern Gulf of Mexico 

and maintained in an aquaculture tank with 

other red snapper at the Gulf Coast Research

Laboratory (GCRL), Ocean Springs, Mississippi, 

U.S.A. The tank and filtration system that 

supported this host had been sanitized before 

adding any fish, and no other fishes shared the 

water of this system. There was no history of 

infestation by this monogenean in culture facilities 

at GCRL. Therefore, it is likely that this red 

snapper was infested with N. melleni in the wild. 

In addition, 3 juvenile red snapper (each approximately 

120 mm in total length) that were 

spawned and reared at the GCRL aquaculture 

facility and then transferred to the GCRL Marine 

Education Center (MEC), Biloxi, Mississippi, 

U.S.A., became heavily infested with TV. melleni. 

These red snapper were maintained in a 7,700liter 

aquarium with a spadefish (Chaetodipterus 

faber (Broussonet, 1782) [Ephippidae]) that was 

also heavily infested with the monogenean. Nigrelli 

(1947) reported Lutjanus apodus (as Lutianus 

apodus) as a wild host for N. melleni (as 

Benedenia melleni) in the West Indies. We suspect 

that L. campechanus also may be a wild 

host of N. melleni in the northern Gulf of Mexico. 

However, it does not seem to be a common 

host, because we have yet to observe a specimen 

of N. melleni on a red snapper directly from the 

wild, in spite of examinations of at least 276 

such fish. 

The most recent list of captive and wild hosts 

for N. melleni was presented by Lawler (1981). 

Whittington and Horton (1996) subsequently 

provided a list of hosts for N. melleni; however, 

that list did not distinguish between captive and 

wild hosts. Because a list identifying wild hosts 

for N. melleni has not been presented in 18 

years, we consider Table 1 a useful update. 

Rarely does a monogenean species, let alone 

a capsalid, occur in more than 1 ocean and infest 

more than 1 host species, and if so, those hosts 

are usually closely related species (e.g., Byrnes 

and Rohde, 1992; Whittington, 1998). Neobenedenia 

melleni has now been reported from 27 

species comprising 18 genera, 14 families, and 

3 orders of wild hosts (Table 1). These records 

suggest that N. melleni is a parasite of predominantly 

shallow-water or reef-dwelling marine 

teleosts. Neobenedenia melleni exhibits a relatively 

low degree of host specificity among both 

captive and wild hosts. Nigrelli and Breder 

(1934) studied the host-parasite relationship between 

N. melleni and several fishes held at the 

New York Aquarium. However, the factors that 

allowed it to infest a broad array of hosts in 

BULLARD ET AL.—HOSTS OF NEOBENEDENIA MELLENI 195 

captivity and in the wild were not clearly understood. 

In some cases, horizontal transfer and 

levels of infestation may be limited initially only 

by the physical distance between parasite and 

potential host. This could, in part, explain the 

apparent abundance of N. melleni among reef 

fishes that live in close proximity to one another 

in the wild and among those and other fishes 

held in public aquaria and aquaculture systems. 

Further study of this unique monogenean utilizing 

molecular techniques could possibly reveal 

population differences. 


We thank Reg Blaylock for commenting on 

the manuscript; Nate Jordan, Jason Sleekier, 

Jody Peterson, and Casey Nicholson (all of 

GCRL) for providing red snapper for examination; 

Joyce Shaw (GCRL) for requesting some 

of the pertinent literature via interlibrary loan; 

Alex Schesny (MEC) for providing juvenile 

specimens of L. campechanus infested with N. 

melleni; Michael Abney (University of Kentucky) 

and Richard Heard (GCRL) for providing 

the specimen of G. xanthosoma infested with TV. 

melleni; Pamela Phelps (MML) for providing 

specimens of N. melleni from E. morio; David 

Miller (Fort Wayne Children's Zoo) for providing 

specimens of N. melleni from C. lunula; and 

the Cayman Islands National Trust and the Cayman 

Islands Department of the Environment for 

allowing and facilitating collection of G. xanthosoma 

on Grand Cayman. This study was supported 

in part from National Oceanic and Atmospheric 

Administration, National Marine 

Fisheries Service, award No. NA86FL0476 and 

NA96FL0358. 


Bravo-Hollis, M. 1957. Trematodos de peces marinos 

de aguas mexicanas. XIV. Cuatro monogeneos de 

la familia Capsalidae Baird, 1853, de las costas 

del Pacifico, incluyendo una especie nueva. Anales 

del Institute de Biologia, Mexico 28:195-216. 

, and J. C. Deloya. 1973. Catalogo de la coleccion 

helmintologica del Institute de Biologia. 

Institute de Biologia, Universidad Nacional Autonoma 

de Mexico, Publicaciones Especiales 2. 

137 pp. 

Brooks, D. R., and M. A. Mayes. 1975. Phyllodistomum 

scrippsi sp. n. (Digenea: Gorgoderidae) 

and Neobenedenia girellae (Hargis, 1955) Yamaguti, 

1963 (Monogenea: Capsalidae) from the California 

sheephead, Pimelometopon pulchrum (Ayers) 

(Pisces: Labridae). Journal of Parasitology 61: 

407-408. 



Byrnes, T., and K. Rohde. 1992. Geographical distribution 

and host specificity of ectoparasites of 

Australian bream, Acanthopagrus spp. (Sparidae). 

Folia Parasitologica 39:249-264. 

Cowell, L. E., W. O. Watanabe, W. D. Head, J. J. 

Grover, and J. M. Shenker. 1993. Use of tropical 

cleaner fish to control the parasite Neobenedenia 

melleni (Monogenea: Capsalidae) on seawater-cultured 

Florida red tilapia. Aquaculture 

113:189-200. 

Cressey, R. F., and E. A. Lachner. 1970. The parasitic 

copepod diet and life history of diskfishes 

(Echeneidae). Copeia 1970:310-318. 

Dyer, W. G., E. H. Williams, and L. Bunkley-Williams. 

1992. Neobenedenia pargueraensis n. sp. 

(Monogenea: Capsalidae) from the red hind, Epinephelus 

giittatus, and comments about Neobenedenia 

melleni. Journal of Parasitology 78:399- 

401. 

Gaida, I. H., and P. Frost. 1991. Intensity of Neobenedenia 

girellae (Monogenea: Capsalidae) on 

the halfmoon, Mediahma californiensis (Perciformes: 

Kyphosidae), examined using a new method 

for detection. Journal of the Helminthological Society 


Goldberg, J. L., R. Millar, and S. Sanchez. 1991. 

The ontogenic acquisition of infestation of the 

trematode ectoparasite Neobenedenia girellae on 

the marine teleost Girella nigricans. Bulletin of 

the Southern California Academy of Sciences 90: 

83-85. 

Gonzalez, M. T., and E. Acuna. 1998. Metazoan parasites 

of the red rockfish Sebastes capensis off 

northern Chile. Journal of Parasitology 84:783- 

788. 

Hall, R. N. 1992. Preliminary investigations of marine 

cage culture of red hybrid tilapia in Jamaica. Proceedings 

of the Gulf and Caribbean Fisheries Institute 

42:440. 

Hargis, W. J. 1955. A new species of Benedenia 

(Trematoda: Monogenea) from Girella nigricans, 

the opaleye. Journal of Parasitology 41:48-50. 

Jahn, T. L., and L. R. Kuhn. 1932. The life history 

of Epibdella melleni MacCallum 1927, a monogenetic 

trematode parasitic on marine fishes. Biological 

Bulletin 62:89-111. 

Lawler, A. R. 1981. Zoogeography and host-specificity 

of the superfamily Capsaloidea Price, 1936 

(Monogenea: Monopisthocotylea): an evaluation 

of the host-parasite locality records of the superfamily 

Capsaloidea Price, 1936, and their utility 


in determinations of host-specificity and zoogeography. 

Special Papers in Marine Science No. 6 

(Virginia Institute of Marine Science, Gloucester 

Point, Virginia, U.S.A.). 650 pp. 

Love, M. S., K. Shriner, and P. Morris. 1984. Parasites 

of olive rockfish, Sebastes serranoides, 

(Scorpaenidae) off central California. Fishery Bulletin 

82:530-537. 

MacCallum, G. A. 1927. A new ectoparasitic trematode. 

Zoopathologica 1:291-300. 

Moser, M., and L. Haldorson. 1982. Parasites of two 

species of surfperch (Embiotocidae) from seven 

Pacific coast locales. Journal of Parasitology 68: 

733-735. 

Mueller, K. W., W. O. Watanabe, and W. D. Head. 

1994. Occurrence and control of Neobenedenia 

melleni (Monogenea: Capsalidae) in cultured tropical 

marine fish, including three new host records. 

Progressive Fish-Culturist 56:140—142. 

Nigrelli, R. F. 1947. Susceptibility and immunity of 

marine fishes to Benedenia ( = Epibdella) melleni 

(MacCallum), a monogenetic trematode. III. Natural 

hosts in the West Indies. Journal of Parasitology 

33:25. 

, and C. M. Breder, Jr. 1934. The susceptibility 

and immunity of certain marine fishes to 

Epibdella melleni, a monogenetic trematode. Journal 

of Parasitology 20:259-269. 

Randall, J. E., G. R. Allen, and R. C. Steene. 1990. 

Fishes of the Great Barrier Reef and Coral Sea. 

University of Hawaii Press, Honolulu, Hawaii. 

507 pp. 

Robins, C. R., G. G. Ray, and J. Douglas. 1986. A 

Field Guide to Atlantic Coast Fishes of North 

America. Houghton Mifflin Company, Boston, 

Massachusetts. 354 pp. 

Robinson, R. D., L. F. Khalil, R. N. Hall, and R. D. 

Steele. 1992. Infection of red hybrid tilapia with 

a monogenean in coastal waters off southern Jamaica. 

Proceedings of the 42nd Annual Gulf and 

Caribbean Fisheries Institute 42:441-447. 

Whittington, I. D. 1998. Diversity "down under": 

monogeneans in the antipodes (Australia) with a 

prediction of monogenean biodiversity worldwide. 

International Journal for Parasitology 28:1481- 

1493. 

, and M. A. Horton. 1996. A revision of Neobenedenia 

Yamaguti, 1963 (Monogenea: Capsalidae) 

including a redescription of N. melleni 

(MacCallum, 1927) Yamaguti, 1963. Journal of 

Natural History 30:1113-1156.



Hymenolepis nana in Pet Store Rodents 

LAURA M. DUCLOS AND DENNIS J. RICHARDSON' 

Department of Biological Sciences, Box 138, 275 Mount Carmel Avenue, Quinnipiac University, Hamden, 

Connecticut 06518, U.S.A. (e-mail: dennis.richardson@quinnipiac.edu) 

ABSTRACT: The rodent tapeworm, Hymenolepis nana, is a zoonotic pathogen transmissible through the ingestion 

of eggs in feces or cysticercoids in arthropods. Since data addressing the potential for acquiring human infections 

of H. nana from pet rodents are lacking, a survey of pet stores in southern Connecticut, U.S.A., was conducted. 

Fecal flotation analysis revealed 9.1% overall prevalence in 110 samples collected weekly from cages holding 

group-housed small animals, from 3 stores for 4 weeks. Of 11 species, only cages holding rats (3 of 22 samples), 

mice (6 of 30 samples), and prairie dogs (1 of 2 samples) were positive. Necropsies of 38 rats, 72 domestic 

mice, and 39 golden hamsters purchased from 9 stores showed prevalences of 31.6%, 22.2%, and 10.3%, 

respectively. Mean intensity was 66 worms per rat, 14 worms per mouse, and 15 worms per hamster. Overall, 

75% of surveyed pet stores were selling infected rats, mice, or hamsters, indicating that pet store rodents pose 

a potential threat to public health. 

KEY WORDS: Hymenolepis nana, pets, rodents, survey, zoonosis. 

Hymenolepis nana Siebold, 1852, infects 75 

million people worldwide (Crompton, 1999), of 

whom the majority are children (Little, 1985; 

Markell et al., 1999). Hymenolepis nana has a 

cosmopolitan distribution, with human, Old 

World monkey, and rodent definitive hosts becoming 

infected through ingestion of infective 

cysticercoids within beetle or flea intermediate 

hosts or through ingestion of eggs in feces. In 

the latter route, cysticercoids develop in intestinal 

villi, with worms later reemerging and attaching 

to the mucosa (Roberts and Janovy, 

2000). Direct transmission through the ingestion 

of eggs may be the most common route of infection 

in humans (Turner, 1975). 

It has been suggested that 2 morphologically 

identical subspecies of H. nana exist, yet this 

tapeworm is typically classified as a zoonotic 

and is capable of horizontal transmission between 

human and nonhuman animals (Fox et al., 

1984; Jacoby and Fox, 1984; Chomel, 1992). 

Human infections "produce either no symptoms 

or vague abdominal disturbances. In fairly heavy 

infections, children may show lack of appetite, 

abdominal pain with or without diarrhea, anorexia, 

vomiting, and dizziness" (Neva and 

Brown, 1994). Such nondescript symptoms may 

account for the low number of reported clinical 

cases, and many subclinical infections may go 

undiagnosed. Available prevalence data regarding 

human populations were obtained in most 

cases from fecal surveys conducted in develop- 


197 

ing nations. For example, H. nana was found in 

20.5% of Australian aborigines (Meloni et al., 

1993), 8-10% of oncology patients in Mexico 

(Guarner et al., 1997), 16% of Egyptian school 

children (Khalil et al., 1991), and 0.6% of Thai 

laborers (Wilairatana et al., 1996). In developed 

regions such as Western Europe and North 

America, human infections are seldom identified 

or acknowledged, and survey data are often 

patchy and scarce (Croll and Gyorkos, 1979; Jacobs, 

1979; Seaton, 1979; Cooper et al., 1981). 

Still, H. nana is estimated to be an important 

cause of cestodiasis in the southeastern United 

States, with infections found in approximately 

1% of school children (Roberts and Janovy, 

2000) and 4% of pediatric clinic patients (Flores 

et al., 1983). In 1987, 34 state diagnostic laboratories 

identified H. nana in collected stool 

samples (0.4%), with Connecticut reporting a 

prevalence of 0.8%, Massachusetts 0.4%, and 

Rhode Island 1.6% (Kappus et al., 1991). 

Most studies focus on human infection, but 

fail to adequately address potential zoonotic 

sources of the infection. In Turkey, 5.6% of surveyed 

wild mice and rats harbored H. nana (Sahin, 

1979), and in Saudi Arabia, H. nana was 

reported from baboons living in close proximity 

to humans (Ghandour et al., 1995). In the United 

States, Stone and Manwell (1966) reported infection 

in 21% of mice and 9% of hamsters from 

Syracuse University, Syracuse, New York, 

U.S.A., animal rooms and various commercial 

vendors. In the same study, pet mice and ham- 


198 uuMKAKAiIVt, rAKAsuOLOuY, 6/(2), JULY 2000 

Table 1. Results of cage sampling for Hymenolepis nana using fecal flotation. 

Host species 

Domestic spiny mouse (Heteromyidae) 

Long-tailed chinchilla (Chinchilla lanigcra Molina, 1782) 

Black-tailed prairie dog (Cynomys ludovicianus Ord, 1815) 

Guinea pig (Cavia porcellus Linnaeus, 1758) 

Domestic mouse (Mus tnusculus Linnaeus, 1758) 

Ferret (Mustelaputoriusfa.ro Linnaeus, 1758) 

Mongolian gerbil (Meriones unguiculatus Milne-Edwards, 1867) 

European rabbit (Oryctolagux cuniculus Linnaeus, 1758) 

Siberian hamster (Phodopus sungorus Pallus, 1773) 

Norway rat (Rattus norvegicus Berkenhout, 1769) 

:i: Overall prevalence of infected cages was 9.1%. 

t A total of 5 individual prairie dogs was surveyed from the 2 cage samples. 

:i: Hymenolepis dirninuta eggs were also detected in the cage sample. 

sters showed prevalences of 66% and 44% respectively. 

Pet stores are traditionally implicated as potential 

sources of human parasite infections, but 

emphasis is centered upon feline, canine, or avian 

species rather than rodents. However, H. 

nana is a common zoonosis of pet rodents (Chomel, 

1992), and in 1969 infection was detected 

in Mongolian gerbils purchased as pets from a 

department store (Lussier and Loew, 1970). Given 

that children have less than optimal hygiene 

habits, and immune-compromised individuals, 

such as those with the acquired immunodeficiency 

syndrome (AIDS) or undergoing cancer 

treatment, are at greater risk for disease (Gerba 

et al., 1996), pet rodent infections raise obvious 

public health concerns. Additionally, there is a 

lack of survey data addressing the assumption 

that golden hamsters are more often parasitized 

with H. nana than are other rodents (Chomel, 

1992; Teclaw et al., 1992). The purpose of this 

study was to assess health risks associated with 

human and rodent interaction as they pertain to 

H. nana, through a survey of small animals sold 

by pet stores in southern Connecticut. 


Once a week for 4 weeks beginning in July 1999, a 

fecal survey was conducted on all small animal cages 

from 3 pet stores. Samples of 5-10 fecal pellets were 

collected from the bedding of cages housing grouped 

animals and analyzed by fecal flotation (Hendrix, 

1998). A total of 110 cage samples was obtained from 

representatives of 11 domesticated small animal species 

(Table 1). 

Based on the findings from fecal analysis of small 

No. of cage 

samples 

3 

6 

2 

19 

18 

30 

3 

7 

11 

1 

22$ 


Samples ( + ) 

for H. nana 

0 

0 

1 

0 

0 

6 

0 

0 

0 

0 

3 

Cage 

prevalence 

(%)* 

0 

0 

sot 

0 

0 

20 

0 

0 

0 

0 

14 

animal cages, individual rodents were purchased from 

9 different pet stores not included in the fecal survey, 

and a postmortem examination of the intestinal tract 

of each rodent was performed. Necropsy was conducted 

on a total of 38 rats, 39 golden hamsters, and 

72 domestic mice. Animals were killed by CO2 narcosis, 

and the small intestine, from the pyloric sphincter 

to the ileocecal juncture, was removed, placed in a 

Petri dish of tap water, and opened longitudinally. 

Worms were removed and counted. Representative 

specimens were stained, mounted, and deposited in the 

United States National Museum Parasite Collection in 

Beltsville, Maryland, U.S.A. (USNPC No. 089330.00). 

Results 

Fecal analysis showed that 9.1% of cages 

housed infected animals, with animals from 1 

pet store testing positive for 3 of 4 weeks. Domestic 

mice and Norway rats exhibited prevalences 

of 30.0% and 13.6%, respectively. One 

of 2 black-tailed prairie dog cage samples revealed 

H. nana. All other species, including 

golden hamsters, were negative by fecal flotation 

analysis (Table 1). 

Necropsy results of purchased animals revealed 

that 7 of 9 pet stores were selling infected 

rats, domestic mice, and/or golden hamsters. 

Prevalence was highest in rats (31.6%), with 

mean intensity (MI) of 66 worms per host. 

Mouse prevalence was lower at 22.2% (MI = 

15), and only 4 golden hamsters (10.3%) were 

infected (MI = 15). One rat was infected with 

Hymenolepis dirninuta Rudolphi, 1819 (Table 2). 

Discussion 

Rodents typically remained in pet stores approximately 

7-10 days. The prepatent period for

Table 2. Presence of Hymenolepis nana in necropsied 

rats, mice, and hamsters. 

No. of No. (%) of 

individuals individuals Mean 

Host species necropsied infected intensity 

Golden hamster 

Domestic mouse 

Norway rat 

39 

72 

38 

4 (10.3) 

16 (22.2) 

12 (31.6)* 

* One rat was infected with Hymenolepis dimimita. 

DLJCLOS AND RICHARDSON—HYMENOLEPIS NANA IN PETS 199 

15 

14 

66 

H. nana is approximately 25 days (Hunninen, 

1935; Jacoby and Fox, 1984), leading to the conclusion 

that animals arrived infected from commercial 

vendors or private breeders, rather than 

acquiring infection through exposure at pet 

stores. Because of the high demand for and 

quick turnover rate of rats, mice, and hamsters, 

the majority of pet stores surveyed purchased 

their rodents from various vendors rather than 

relying on in-house breeding programs. In this 

study, 7 of the 12 pet stores purchased animals 

from 5 different vendors, while the other 5 

stores relied on in-house breeding programs or 

various suppliers, either private or commercial 

sources. Rodents in those 7 vendor-supplied 

stores tested positive for H. nana, while only 3 

of the other 5 stores revealed positive rodents 

(Table 3). 

Evidence for direct transmission of H. nana 

as the common route of infection in rodents can 

be derived from the concomitant presence of H. 

dimimita and H. nana within the same rat cage. 

Transmission of H. diminuta requires an arthropod 

intermediate host (Roberts and Janovy, 

2000). Since the same arthropods may serve as 

intermediate hosts for both tapeworms, lower 

prevalence of H. diminuta and higher prevalence 

of H. nana indicate transmission through direct 

rather than indirect routes. If H. nana were using 

an intermediate host, one would assume the 

prevalences of the 2 tapeworms to be nearly 

equivalent. 

Traditionally, H. nana is considered a tapeworm 

of mice (Markell et al., 1999). However, 

the public health significance of the higher prevalence 

in rats becomes apparent when the type 

of pet most often purchased for children is considered. 

According to pet store owners, mice are 

usually sold as feeder animals, but hamsters and 

Table 3. Summary of survey data on Hymenolepis nana in rodents from pet stores in southern Connecticut, 

U.S.A. 

Pet store Sample 

location size 

Necropsy animals:]: 

Hamden 

Wallingford 

Meriden 

East Haven 

North Branford 

Fairrield 

Orange 

Stratford 

Naugatuck (a) 

Fecal sample|| 

Naugatuck (b) 

Seymore (a) 

Seymore (b) 

20 

17 

10 

17 

15 

21 

13 

18 

18 

37 

23 

9 

Store prevalence 


rats are more often purchased as pets. Surprisingly, 

our results indicate that pet store rats constitute 

a more important reservoir for H. nana 

than do mice or hamsters. Also, this is the first 

report of H. nana from a pet prairie dog, a nontraditional 

or exotic animal that is becoming 

common in pet stores (Storer and Watson, 1997). 

Improper hygiene following handling of all rodents, 

including feeders, may lead to transmission. 

A pet ownership profile (Teclaw et al., 

1992) showed that approximately 50% of households, 

primarily those with children between the 

ages of 6-17 years, owned some type of pet. 

Further, 2% of Florida AIDS patients interviewed 

owned pet rodents, but most health care 

providers failed to advise them of possible zoonoses 

from their companion animals (Conti et 

al., 1995). 

Overall, 75% of surveyed pet stores were selling 

animals infected with H. nana. Despite the 

fact that H. nana infection in rodents is easily 

treatable with praziquantel (Harkness and Wagner, 

1995), none of the pet stores reported practicing 

antihelmintic treatment and control measures. 

The combination of high prevalence and 

absence of control measures demonstrates that 

pet rodents pose a zoonotic threat to pet store 

personnel, animal care workers, and customers. 

Surveys of human populations, together with 

further epidemiological information, are needed 

to assess the extent to which this potential health 

threat is actually being realized. 


This work was funded in part by an Interdisciplinary 

Research Grant from Quinnipiac College. 

Robin LePardo assisted in collecting fecal 

samples. Kristen E. Richardson, Quinnipiac College, 

assisted in the preparation of the manuscript. 


Chomel, B. B. 1992. Zoonoses of house pets other 

than dogs, cats, and birds. Pediatric Infectious 

Disease Journal 11:479-487. 

Conti, L., S. Lieb, T. Liberti, M. Wiley-Bayless, K. 

Hepburn, and T. Diaz. 1995. Pet ownership 

among persons with AIDS in three Florida counties. 

American Journal of Public Health 85:1559- 

1561. 

Cooper, B. T., H. J. Hodgson, and V. S. Chadwick. 

1981. Hymenolepiasis: an unusual cause of diarrhea 

in Western Europe. Digestion 21:115-116. 

Croll, N. A., and T. W. Gyorkos. 1979. Parasitic dis- 


ease in humans: the extent in Canada. Canadian 

Medical Association Journal 120:310-312. 

Crompton, D. W. T. 1999. How much human helminthiasis 

is there in the world? Journal of Parasitology 

85:397-403. 

Flores, E. C., S. C. Plumb, and M. C. McNeese. 

1983. Intestinal parasitosis in an urban pediatric 

clinic population. American Journal of Diseases 

of Children 137:754-756. 

Fox, J. G., C. E. Newcomer, and H. Ruzmiarek. 

1984. Selected zoonoses and other health hazards. 

Pages 614-643 in J. G. Fox, B. J. Cohen, and F. 

M. Loew, eds. Laboratory Animal Medicine. Academic 

Press, Orlando, Florida. 

Gerber, C. P., J. B. Rose, and C. N. Haas. 1996. 

Sensitive populations: who is at the greatest risk? 

International Journal of Food Microbiology 30: 

113-123. 

Ghandour, A. M., N. Z. Zahid, A. A. Banaja, K. B. 

Karnal, and A. I. Bouq. 1995. Zoonotic intestinal 

parasites of hamadryas baboons Papio hamadryas 

in the western and northern regions of Saudi Arabia. 

Journal of Tropical Medicine and Hygiene 98: 

431-439. 

Guarner, J., T. Matilde-Nava, R. Villasenor-Flores, 

and G. Sanchez-Mejorada. 1997. Frequency of 

intestinal parasites in adult cancer patients in 

Mexico. Archives of Medical Research 28:219- 

222. 

Harkness, J. E., and J. E. Wagner. 1995. The Biology 

and Medicine of Rabbits and Rodents, 4th 

ed. Williams & Wilkins, Media, Pennsylvania. 

372 pp. 

Hendrix, C. M. 1998. Diagnostic Veterinary Parasitology, 

2nd ed. Mosby, Inc., St. Louis, Missouri. 

321 pp. 

Hunninen, A. V. 1935. Studies on the life history and 

host-parasite relations of Hymenolepis fraterna 

(H. nana, van fraterna, Stiles) in white mice. 

American Journal of Hygiene 20:414-443. 

Jacobs, D. E. 1979. Man and his pets. Pages 174-200 

in R. J. Donaldson, ed. Parasites and Western 

Man. University Park Press, Baltimore, Maryland. 

Jacoby, R. O., and J. G. Fox. 1984. Biology and 

diseases of mice. Pages 31-88 in J. G. Fox, B. J. 

Cohen, and F. M. Loew, eds. Laboratory Animal 

Medicine. Academic Press, Orlando, Florida. 

Kappus, K. K., D. D. Juranek, and J. M. Roberts. 

1991. Results of testing for intestinal parasites by 

state diagnostic laboratories, United States, 1987. 

Morbidity and Mortality Weekly Reports Intestinal 

Parasite Surveillance Summary 40 (No. SS- 

4): 24-45. 

Khalil, H. M., S. el Shimi, M. A. Sarwat, A. F. 

Fawzy, and A. O. el Sorougy. 1991. Recent 

study of Hymenolepis nana infection in Egyptian 

children. Journal of the Egyptian Society of Parasitologists 

21:293-300. 

Little, M. D. 1985. Cestodes (tapeworms). Pages 110- 

126 in P. C. Beaver and R. C. Jung, eds. Animal 

Agents and Vectors of Human Disease, 5th ed. 

Lea and Febiger, Philadelphia, Pennsylvania. 

Lussier, G., and F. M. Loew. 1970. Natural Hymenolepis 

nana infection in Mongolian gerbils (Mer-

tones ungiiiculatus). Canadian Veterinary Journal 

11:105-107. 

Markell, E. K., D. T. John, and W. A. Krotoski. 

1999. Markell and Voge's Medical Parasitology, 

8th ed. W. B. Saunders Co., Philadelphia, Pennsylvania. 

501 pp. 

Meloni, B. P., R. C. A. Thompson, R. M. Hopkins, 

J. A. Reynoldson, and M. Gracey. 1993. The 

prevalence of Giardia and other intestinal parasites 

in children, dogs and cats from Aboriginal 

communities in the Kimberly. Medical Journal of 

Australia 158:157-159. 

Neva, F. A., and H. W. Brown. 1994. Basic Clinical 

Parasitology, 6th ed. Appleton and Lange, Norwalk, 

Connecticut. 356 pp. 

Roberts, L. S., and J. Janovy, Jr. 2000. Gerald D. 

Schmidt and Larry S. Roberts' Foundations of 

Parasitology, 6th ed. McGraw-Hill, Boston, Massachusetts. 

670 pp. 

Sahin, I. 1979. Parasitosis and zoonosis in mice and 

rats caught in and around Beytepe Village. Mikrobiyoloji 

Bulteni 13:283-290. 

Seaton, J. R. 1979. Cestodes and trematodes. Pages 

114-132 in R. J. Donaldson, ed. Parasites and 

DUCLOS AND RICHARDSON—HYMENOLEPIS NANA IN PETS 201 

NEW BOOK AVAILABLE 

Western Man. University Park Press, Baltimore, 

Maryland. 

Stone, W. B., and R. D. Manwell. 1966. Potential 

helminth infections in humans from pet or laboratory 

mice and hamsters. Public Health Reports 

31:647-653. 

Storer, P., and L. Watson. 1997. Prairie Dog Primer. 

Country Storer Enterprises, Columbus, Texas. 76 

pp. 

Teclaw, R., J. M. Mendlein, P. Garbe, and P. Mariolis. 

1992. Characteristics of pet populations and 

households in the Purdue Comparative Oncology 

Program catchment area, 1988. Journal of the 

American Veterinary Medical Association 210: 

1725-1729. 

Turner, J. A. 1975. Other cestode infections. Pages 

708-744 in J. A. Hubbert, W. F. McCulloch, and 

P. R. Schnurrenberger, eds. Diseases Transmitted 

from Animals to Man, 6th ed. Charles C. Thomas, 

Springfield, Illinois. 

Wilairatana, P., B. Radomyos, R. Phraevanich, W. 

Plooksawasdi, P. Chanthavanich, C. Viravan, 

and S. Looareesuwan. 1996. Intestinal sarcocystosis 

in Thai laborers. Southeast Asian Journal of 

Tropical Medicine and Public Health 27:43-46. 

Echinostomes as Experimental Models for Biological Research. Edited by Bernard Fried and 

Thaddeus K. Graczyk. 2000. Kluwer Academic Publications, Dordrecht, The Netherlands. 284 pp. 

ISBN 0-7923-6156-3. Hardcover. US$ 150.00/NLG 250.00/GBP 88.00. Abstract: ". . . Considerable 

but scattered literature has been published on the subject of echinostomes and a synthesis of 

this wide range of topics has now been achieved with the publication of this book, which represents 

a wide range of topics in experimental biology related to the use of echinostomes as laboratory 

models. It will have a special appeal to advanced undergraduates and graduate students in parasitology 

and should also appeal to professional parasitologists, physicians, veterinarians, wildlife 

disease biologists, and any biomedical scientists interested in new model systems for studies in 

experimental biology." 




Seasonal Occurrence and Community Structure of Helminth 

Parasites from the Eastern American Toad, Bufo americanus 

americanus., 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 September 1996, 47 American toads, Bufo americanus americanus Holbrook, were 

collected from Waukesha County, Wisconsin, U.S.A., and examined for helminth parasites. Forty-six (98%) of 

47 toads were infected with 1 or more helminth species. The component community consisted of 6 species, 3 

direct-life-cycle nematodes, and 3 indirect-life-cycle helminths (2 trematodes and 1 metacestode). Totals of 2,423 

individual nematodes (92%), 45 trematodes (2%), and 155 cestodes (6%) were found, with infracommunities 

being dominated by skin-penetrating nematodes. A significant correlation existed between wet weight and overall 

helminth abundance, excluding larval platyhelminths. Helminth populations and communities were seasonally 

variable but did not show significant differences during the year. However, a number of species showed seasonal 

variations in location in the host, and these variations were related to recruitment period. 

KEY WORDS: Bufo americanus, American toad, Cosmocercoides variahilis, Rhabdias americanus, Oswaldocruzia 

pipiens, Mesocestoidex sp., Gorgoderina sp., Trematoda, Nematoda, Cestoda, echinostome metacercariae, 

seasonal study, Wisconsin, U.S.A. 

American toads, Bufo americanus americanus 

Holbrook, 1836, are large, thick-bodied terrestrial 

anurans found in North America near 

marshes, oak savannas, semiopen coniferous and 

deciduous forests, and agricultural areas. They 

range from Labrador and Hudson Bay to eastern 

Manitoba, south to eastern Oklahoma and the 

coastal plains, and are distributed throughout 

Wisconsin (Vogt, 1981). Toads are active foragers, 

differing from most anurans that are sitand-wait 

predators (Scale, 1987). Although a 

number of surveys and natural history studies 

on the helminths and ecology of toads exists 

(Bouchard, 1951; Odlaug, 1954; Ulmer, 1970; 

Ulmer and James, 1976; Williams and Taft, 

1980; Coggins and Sajdak, 1982; Joy and Bunten, 

1997), no studies have used measures of 

helminth communities. Here we report on the 

helminth infracommunity and component community 

structure in American toads from southeastern 

Wisconsin. 


American toads were collected from April to November 

of 1996 by driving 4.0-km sections of highways 

N and 67 (42°54'N, 88°29'W) in Eagle, Wau- 

1 Corresponding author. Current address: Department 

of Veterinary Pathobiology, Purdue University, 

West Lafayette, Indiana 47907, U.S.A. (e-mail: 

mgb @ vet.purdue.edu). 

202 


kesha County, Wisconsin, U.S.A., during the night and 

collecting individuals as they crossed roads. Animals 

were placed in plastic containers, transported to the 

laboratory, stored at 4°C, and killed in MS-222 (ethyl 

m-aminobenzoate methane sulfonic acid) within 72 hr 

of capture. Snout-vent length (SVL) and wet weight 

(WW) were recorded for each individual. Toads were 

individually toe-clipped and frozen. At necropsy, the 

digestive tracts, limbs, body wall musculature, and internal 

organs were examined for helminth parasites. 

Each organ was individually placed in a Petri dish and 

examined under a stereomicroscope. The body cavity 

was 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. Echinostome 

metacercariae were badly damaged during 

necropsy, and these were identified but not retained. 

Prevalence, mean intensity, and abundance are according 

to Bush et al. (1997). Mean helminth species richness 

is the sum of helminth species per individual amphibian, 

including noninfected individuals, divided by 

the total sample size. All values are reported as the 

mean ± 1 SD. Undigested stomach contents were 

identified to class or order following Borror et al. 

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

the number of arthropods in a given class or order, 

divided by the total number of arthropods recovered 

X 100. Voucher specimens have been deposited in the 

helminth collection of the H. W. Manter Laboratory 

(HWML), University of Nebraska State Museum, Lincoln, 

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

BOLEK AND COGGINS—HELMINTH COMMUNITIES IN TOADS 203 

Table 1. Prevalence, mean intensity, mean abundance, and total numbers of helminths found in 47 specimens 

of Bufo americanus americanus. 

Species 

Trematoda 

Echinostome metacercariae* 

Gorgoderina sp. 

Cestoda 

Mesocestoides sp.* 

Nematoda 

Oswaldocruzia pipiens 

Cosmocercoides variabilis 

Rhahdias americanus 

Underestimate. 

Prevalence, Mean intensity 

No. (%) ± 1 SD (range) 

3 (6.3) 

1 (2.1) 

13 ± 19 (1-35) 

6 ± 0 (6) 

6 (12.7) 25.8 ± 22 (12-70) 

15051, male Oswaldocruzia pipiens; 15052, male Cosmocercoides 

variabilis; 15053, Rhabdias americanus; 

15054, Gorgoderina sp.; 15055, Mesocestoides sp.). 

The chi-square test for independence was calculated 

to compare differences in prevalence among host sex, 

seasonal differences in prevalence, and seasonal differences 

in location of nematodes in the host. Yates' 

adjustment for continuity was used when sample sizes 

were low, and a single-factor, independent-measures 

analysis of variance was used to compare among seasonal 

differences in mean intensity and mean helminth 

species richness (Sokal and Rohlf, 1981). Student's ttest 

was used to compare differences in mean intensity 

and mean helminth species richness between sex of 

hosts. Approximate f-tests 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. 

Pearson's con-elation was calculated for host SVL and 

WW and helminth species richness per individual amphibian. 

Because WW gave a stronger correlation than 

SVL in each case, it is the only parameter reported. 

Because of low sample sizes during certain collection 

periods, data were pooled on a bimonthly basis to form 

samples of 15 to 16 toads per season. Larval platyhelminths 

were not included in the seasonal analysis, 

because they can accumulate throughout an amphibian's 

life. 

Results 

A total of 47 American toads, 28 males and 

19 females, was collected. The overall mean 

SVL and WW of toads was 56.6 ± 12.5 mm 

(range = 26.2-72.6 mm) and 26.6 ± 13.5 g 

(range = 2.19-55.5 g), respectively. No significant 

difference existed in numbers of male 

toads and female toads collected throughout the 

No. of 

Mean worms 

abundance recov- 

± 1 SD ered Location 

0.8 ± 5.1 

0.1 ± 0.9 

41 (87) 8.5 ± 7 (1-31) 7.4 ± 7.1 

43(91) 32.3 ± 31.5 (1-135) 29.6 ±31.5 

39 Kidneys, body cavity 

6 Bladder 

155 Leg muscles 

349 Small intestine 

1,392 Lungs, body cavity, 

large and small intestine 

43(91) 15.8 ± 17.9 (1-75) 14.5 ± 17.7 682 Lungs, body cavity 

year (x2 = 1.72, P > 0.05). Although female 

toads were larger (58.2 ± 15.4 mm) and heavier 

(30.5 ± 17.3 g) than males (55.5 ± 10.2 mm, 

23.9 ± 9.7 g), these differences were not significant 

(t = 0.73, P > 0.05, t's = 1.50, P > 0.05). 

Stomach contents analyses revealed that the 

toads fed mostly on ants (98%), with beetles and 

other terrestrial arthropods representing a small 

portion of the diet (2%). 

Forty-six (98%) of 47 toads were infected 

with 1 or more species of helminths. The component 

community consisted of 6 species, 3 direct-life-cycle 

nematodes, and 3 indirect-life-cycle 

helminths (2 trematodes and 1 metacestode). 

Overall mean helminth abundance, excluding 

larval platyhelminths, was 55.7 ± 45.3 worms 

per toad infracommunity (range = 0-180). Prevalence 

was highest for nematodes, ranging from 

91% for Cosmocercoides variabilis Harwood, 

1930, and Rhabdias americanus Baker, 1978, to 

87% for Oswaldocruzia pipiens Walton, 1929. 

Prevalence for indirect life cycle parasites was 

generally low, being highest for the cestode Mesocestoides 

sp. (12.7%) and lowest for Gorgoderina 

sp. (2.1%) (Table 1). No significant differences 

existed in prevalence between male and 

female toads for any of the 6 helminth species 

recovered. Mean intensity differed significantly 

only in Mesocestoides sp., being higher in male 

(32.7 ± 32) than in female toads (19 ± 4.6, /; 

= 4.70, P < 0.05). 

Mean helminth species richness was 2.9 ± 0.9 


20-1 COMPARATIVE PARASITOLOGY, 67(2), JULY 2000 

.1 u 

ft, 

cc 

"e 

1 

"S 

ffi 

o 

L. 

a> 

ja 

S 

3 

O - 

5 - 

4 - 

Figure 1. 

3 - -*• • 4*MHN»*++ •» * ** * 

2 - 

** «•* • 

1 - •*• •*• 

n . 

* 

* ++ * • 

, A .4,.- 4 1 

20 40 

Wet Weight in Grams 

Figure 3. 

S Body Cavity B Lungs 

100 -r D Small Intestine • Large Intestine 

= 405 

Aug-Sep 

60 

0 

Number of ¥ 

1.80-1 

160- 

140- 

120- 

100 - 

80- 

60- 

40- 

Figure 2. n 

• Males 

D Females + D 

• D 

& U 

• 

- * DV** 

^ • 

20- 

"P 

A . ctk 

^ •* tl * 

* * n i 

20 40 

Wet Weight in Grams 

w 

1 

100-r 

Figure 4. 

I Body Cavity D Lungs 

60 

= 261 

Figures 1—4. 1. Wet weight versus number of helminth species per individual of the American toad, 

Bufo americanus americanus, r = 0.31, P < 0.05. 2. Wet weight versus total helminth abundance, excluding 

larval platyhelminths, in American toads, B. a. americanus, overall r = 0.47, P < 0.01; female toads r = 

0.57, P < 0.01; and male toads r = 0.20, P > 0.05. 3. Seasonal distribution of the relative proportions of 

Cosmocercoides variabilis recovered in the body cavity, lungs, small intestine, and large intestine of B. a. 

americanus. N equals number of nematodes recovered in each sampling period. 4. Seasonal distribution 

of the relative proportions of Rhabdias americanus recovered in the body cavity and lungs of B. a. americanus. 

N equals number of nematodes recovered in each sampling period. 

species per toad. Infections with multiple species 

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

occurring in 1, 2, 6, 30, 7, and 1 host, respectively. 

No statistically significant differences in 

mean helminth species richness were found between 

male (3.0 ± 0.86) and female toads (2.8 


± 0.85, t = 0.83, P > 0.05). There was a significant 

positive correlation between WW and 

helminth species richness per toad (r = 0.31, P 

< 0.05, Fig. 1). However, this relationship became 

insignificant when a single uninfected toad 

was removed (r = 0.20, P > 0.05). A significant


Table 2. Seasonal prevalence and mean intensity of 3 species of nematodes in Bufo americanus americanus. 

Species Apr Jun-Jul Aug-Sep Statistic 

Cosmocercoides variahilis 

Oswaldocruzia pipiens 

Rhabdias americanus 

Prevalence 

Mean intensity ± 

Prevalence 


Prevalence 


1 SD 

1 SD 

1 SD 

93% 

30.4 

93% 

7 

93% 

17.8 

positive correlation existed between WW and 

overall helminth abundance, excluding larval 

platyhelminths (r = 0.47, P < 0.01, Fig. 2), although 

this correlation was only significant for 

female toads (r = 0.57, P < 0.01) and not for 

males (r = 0.20, P > 0.05). Similar results were 

obtained for C. vanabilis (r = 0.41, P < 0.01) 

and O. pipiens (r = 0.43, P < 0.01), whereas 

no significant correlation was observed for R. 

americanus (r = 0.27, P > 0.05). When these 

analyses were performed separately for male and 

female hosts, significant positive correlations occurred 

only in female toads (C. variabilis, r — 

0.52, P < 0.05; O. pipiens, r = 0.58, P < 0.01; 

R. americanus, r = 0.54, P < 0.05). 

Only 1 male toad was infected with 6 Gorgoderina 

sp. during the early spring (April) collection. 

There was no significant seasonal difference 

in prevalence or mean intensity for any 

140 

Number of Osn>aldocruzia pipiens 

Figure 5. Number of Oswaldocruzia pipiens versus 

Cosmocercoides variabilis in Bufo americanus 

americanus, overall r = 0.54, P < 0.01; female toads 

r = 0.58, P < 0.01; and male toads r = 0.51, P < 

0.01. 

(14/15) 

± 28.5 

(14/15) 

± 6.7 

(14/15) 

± 21.6 

88% (14/16) 

40 ± 31.7 

88% 

10 

100% 

10.7 

(14/16) 

± 8.1 

(16/16) 

±11.2 

94% 

27 

81% 

8.3 

81% 

20 

(15/16) 

± 34.6 

(13/16) 

± 6.1 

(13/16) 

± 19.8 

X2 

F 

X2 

F 

X2 

F 

= 0.5 

= 0.65 

= 1.1 

= 0.63 

= 3.7 

= 1.12 

P > 0.05 

P > 0.05 

P > 0.05 

P > 0.05 

P > 0.05 

P > 0.05 

of the nematodes recovered (Table 2). Prevalence 

was highest during early spring for O. pipiens 

(93%), early summer (June-July) for R. 

americanus (100%), and late summer-early fall 

(August-September) for C. vanabilis (94%). 

Mean intensities were higher during early summer 

for C. variabilis and O. pipiens and in late 

summer-early fall for R. americanus. Seasonally, 

there was a significant difference in location 

of C. variabilis (x2 = 556, P < 0.01), and R. 

americanus (x2 = 232, P < 0.01). Cosmocercoides 

variabilis occurred in greater numbers in 

the body cavity and lungs during the early spring 

and in the small and large intestines during early 

summer and late summer-early fall collections 

(Fig. 3). Individuals of Rhabdias americanus 

were found primarily in the lungs during early 

spring and progressively increased in numbers 

in the body cavity during early summer and late 

summer-early fall collections (Fig. 4). The nematode 

O. pipiens showed no seasonal variation 

in location and was found in the small intestine 

throughout the year. Oswaldocruzia pipiens was 

significantly correlated with C. variabilis (r = 

0.54, P < 0.01) in both male (r = 0.51, P < 

0.01) and female toads (r = 0.58, P < 0.01) 

(Fig. 5) but was not significantly correlated with 

R. americanus (r = 0.23, P > 0.05) in either 

male (r = 0.23, P > 0.05) or female toads (r = 

0.24, P > 0.05). 

Mean species richness varied throughout the 

year, being highest (3.26 ± 0.79) in early spring, 

intermediate (2.88 ± 0.71) in early summer, and 

lowest during the late summer-early fall collection 

(2.62 ± 0.95), although these differences 

were not statistically significant (F = 2.3, P > 

0.05). Wet weight of toads showed similar seasonal 

variability, but these differences were also 

not significant (F = 2.99, P > 0.05). 

Discussion 

The component community of Wisconsin 

toads was similar to those of other published re- 



ports (Bouchard, 1951; Odlaug, 1954; Ulmer, 

1970; Ulmer and James, 1976; Williams and 

Taft, 1980; Coggins and Sajdak, 1982; Joy and 

Bunten, 1997; Yoder, 1998). Toad helminth infracommunities 

were dominated by 3 species of 

skin-penetrating nematodes with high overall 

prevalence and mean intensities and with few 

toads infected by indirect-life-cycle parasites. 

Bladder flukes of the genus Gorgoderina are 

common parasites of amphibians, but few lifecycle 

studies exist (Prudhoe and Bray, 1982). 

The life cycles of Gorgoderina attenuata Stafford, 

1902, and Gorgoderina vitelliloba Olsson, 

1876, have been determined. For these species, 

amphibians acquire infection by feeding on 

semiaquatic insect larvae or tadpoles, and the 

worms excyst in the stomach and migrate to the 

kidneys and bladder (Rankin, 1939; Smyth and 

Smyth, 1980). The low prevalence (2.1%) and 

intensity (6) of Gorgoderina sp. observed in our 

study were not surprising, since analyses of 

stomach contents revealed few kinds of arthropods. 

This is characteristic of actively foraging 

species such as toads. Ants made up the largest 

portion of the diet, with beetles and other terrestrial 

arthropods being found less frequently. 

Kirkland (1904) also found that ants and beetles 

made up the greatest portion of the diet of 149 

toads from New England, U.S.A. These results 

are similar to other investigations on diet of species 

ofBufo (Toft, 1981; Collins, 1993; Indraneil 

and Martin, 1998), where ants and beetles appeared 

to be an important food item in the diet 

of toads. Toft (1981) reported that ants made up 

64% to 91% of the arthropods consumed by 3 

South American toad species, while Collins 

(1993) mentioned that ants and beetles were important 

items in the diet of 5 species of Bufo 

from Kansas, U.S.A. Because most trematodes 

of amphibians use aquatic or semiaquatic arthropods 

as intermediate hosts (Prudhoe and Bray, 

1982), these observations may indicate why 

toads usually have low species richness and 

prevalence of adult trematodes (Williams and 

Taft, 1980; Coggins and Sajdak, 1982; McAllister 

et al., 1989; Goldberg and Bursey, 1991a, 

1991b, 1996; Goldberg et al., 1995; Bursey and 

Goldberg, 1998). 

The most commonly occurring nematode was 

C. variabilis, with a total of 1,392 worms recovered. 

Vanderburgh and Anderson (1987) 

studied the seasonal transmission of this species 

in American toads from Ontario, Canada. They 


observed J4 larvae in the lungs during the breeding 

season and adult worms in the rectum of 

toads throughout the year. They suggested that 

toads may acquire C. variabilis soon after 

emerging in the spring and that transmission 

may decline during summer and fall. However, 

they stated that this may have been an artifact 

of sampling, because all toads collected after the 

breeding season were from another location and 

may have had a lower prevalence and mean intensity 

of C. variabilis. Vanderburgh and Anderson 

(1987) also observed larvae in the lungs 

of 5 toads collected in October of the following 

year and concluded that transmission probably 

occurs throughout the year. 

Data from the present study suggest that the 

breeding period may be important in transmission 

of C. variabilis in adult toads. All our toads 

were collected from the same general location 

and had high prevalence and mean intensities 

throughout the year. Although the differences 

were not significant, mean intensity increased 

after the breeding season and decreased during 

the late summer-early fall collection. Ten percent 

of the worms recovered during April were 

located in the body cavity, 37% were located in 

the lungs, and 28% and 24% were located in the 

small and large intestine, respectively. Subsequent 

sampling revealed that only 1 toad collected 

during June had 6 larvae in the lungs, 

with all other worms being recovered from the 

small and large intestine. Baker's (1978a) study 

on the life cycle of Cosmocercoides dukae Holl, 

1928 (=C. variabilis} in toads revealed that 8- 

10 days are required for larvae to reach the 

lungs, and more than 30 days at 14-18°C to migrate 

to the rectum and develop to a gravid 

stage. Our observations suggest that toads became 

infected during the breeding season, and 

there appeared to be a decline during summer 

and early fall. Unfortunately, no toads were collected 

during October, and therefore it is not 

known if infection may occur during the fall (but 

see below). All adult female worms recovered 

throughout the year were gravid, indicating that 

eggs were being produced from April through 

September. 

The second most frequently recovered nematode 

was R. americanus, primarily a parasite of 

toads (Baker, 1979a, 1987). Few studies exist on 

the seasonal occurrence of species of Rhabdias 

in amphibians (Lees, 1962; Plasota, 1969; Baker, 

1979b). Lees (1962) studied the seasonal occur-

ence of Rhabdias bufonis Schrank, 1788, in its 

host, Rana temporaria Linnaeus, 1758, in England, 

and Baker (1979b) studied the seasonal occurrence 

of Rhabdias ranae Walton, 1929, in the 

wood frog, Rana sylvatica Le Conte, 1825, in 

Canada. Prevalence and intensities in these species 

were lowest during summer and highest in 

spring and early fall. Baker (1979b) observed 

many subadult worms in the body cavity of 

wood frogs during late summer and early fall, 

with no worms being found in the body cavity 

during early spring and few worms in the fall. 

Worms occurred in the lungs during early spring 

and fall, while few were found in the lungs during 

late summer and early fall. Baker (1979b) 

concluded that transmission of R. ranae occurs 

throughout the summer and early fall, with 

worms maturing in the lungs during the fall and 

overwintering in their hosts. 

During this study, no significant differences in 

prevalence or mean intensity were observed 

throughout the collection period, although the 

location (lungs or body cavity) of worms varied 

during the year. Most R. americanus recovered 

in April occurred in the lungs, with numbers of 

worms in the lungs decreasing during early summer 

(June-July) and late summer-early fall (August-September). 

In contrast, the number of 

worms in the body cavity increased during the 

June-July and August-September collections. 

Therefore, transmission of this species occurred 

during the summer and early fall, with worms 

overwintering in their host. These results are 

consistent with earlier studies of seasonal distribution 

of other species of Rhabdias (Lees, 1962; 

Baker, 1979b). 

Oswaldocruzia pipiens also had high prevalence 

but lower mean intensities than the other 

2 species of nematodes recovered. Because of 

its fast migration, reaching the stomach and 

small intestine within 1 to 3 days of infection 

(Baker, 1978b), differences in location of these 

worms within the host were not observed. Prevalence 

and mean intensity were variable but not 

significant over the course of this study. Baker 

(1978b), in a seasonal study of O. pipiens in 

wood frogs, observed peak prevalence and intensity 

during spring (May-June) and early fall 

(September-October) in Ontario, Canada. He 

stated that worms overwintered in the host and 

that transmission occurred in early spring, with 

an initial decline during early summer and continued 

transmission during summer and early 


fall. A significant positive correlation existed for 

this species and C. variabilis but not for R. 

americanus, which suggested that toads became 

infected with O. pipiens and C. variabilis during 

the same time and in the same places, while infection 

with R. americanus occurred at a later 

time during the summer. Hosts that contain different 

species of adult parasites that co-occur in 

an infracommunity may contaminate an area 

when they release eggs in their hosts' feces. 

Therefore, acquisition of a certain parasite species 

by a host may often be coupled with the 

acquisition of other species into the infracommunity. 

The spring breeding period may also be 

important in the recruitment of O. pipiens in 

toads at our study site. Because toads were not 

collected during October, it is not known if recruitment 

occurs during this time. If infection 

occurs during the fall, as Baker's (1978b) data 

for wood frogs imply, C. variabilis may also be 

acquired in the fall. 

Significant, positive relationships between 

WW and species richness and abundance were 

observed in toad helminth communities. However, 

significant relationships between WW and 

abundance were observed only in female toads. 

Interestingly, the largest toads collected were females, 

and these had the highest intensities of 

helminths; therefore, they may provide a greater 

surface area for colonization by skin-penetrating 

nematodes. Similar observations were reported 

by Me Alpine (1997) for female leopard frogs, 

which were significantly larger than males. Although 

species richness also showed a significant 

positive relationship with WW, once the 

single noninfected individual was excluded from 

the calculation, the relation became nonsignificant. 

Therefore, no conclusions can be drawn 

from this relationship. Seasonal variance in species 

richness was not significant in B. a. americanus, 

with toad infracommunities being dominated 

by 3 skin-penetrating nematodes throughout 

the year. The toad's terrestrial habitat and 

diet of ants and beetles may be important in excluding 

transmission of adult and larval trematodes, 

unlike other anurans such as semiaquatic 

species of Rana, which spend more time in an 

aquatic environment and have a broader diet of 

semiaquatic invertebrates (Muzzall, 1991; 

McAlpine, 1997; Bolek, 1998). 


We thank Melissa Ewert and Luke Bolek for 

help in collecting toads. We also thank 2 anon- 



ymous reviewers and the editors, Drs. W. A. 

Reid and J. W. Reid, for improvements on an 

earlier draft of the manuscript. 


Baker, M. R. 1978a. Transmission of Cosmocercoides 

dukae (Nematoda: Cosmocercoidea) to amphibians. 

Journal of Parasitology 64:765-766. 

. 1978b. Development and transmission of Oswaldocruzia 

pipiens Walton, 1929 (Nematoda: 

Trichostrongylidae) in amphibians. Canadian 


. 1979a. The-free living and parasitic development 

of Rhabdias spp. (Nematoda: Rhabdiasidae) 

in amphibians. Canadian Journal of Zoology 57: 

161-178. 

. 1979b. Seasonal population changes in Rhabdias 

ranae Walton, 1929 (Nematoda: Rhabdiasidae) 

in Rana sylvatica of Ontario. Canadian Journal 

of Zoology 57:179-183. 

-. 1987. Synopsis of the Nematoda Parasitic in 

Amphibians and Reptiles. Occasional Papers in 

Biology, Memorial University of Newfoundland, 

St. John's, Newfoundland, Canada. 325 pp. 

Bolek, M. G. 1998. A seasonal and comparative study 

of helminth parasites in nine Wisconsin amphibians. 

M.S. Thesis, University of Wisconsin-Milwaukee. 

134 pp. 

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. 875 pp. 

Bouchard, J. L. 1951. The platyhelminths parasitizing 

some northern Maine Amphibia. Transactions of 

the American Microscopical Society 70:245-250. 

Bursey, C. R., and S. R. Goldberg. 1998. Helminths 

of the Canadian toad, Bufo hemiophrys (Amphibia: 

Anura), from Alberta, Canada. Journal of Parasitology 

84:617-618. 

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 


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 


Collins, J. T. 1993. Amphibians and Reptiles in Kansas, 

3rd ed. The University of Kansas Museum of 

Natural History, Lawrence, Kansas, Public Education 

Series 13. 397 pp. 

Goldberg, S. R., and C. R. Bursey. 199la. Helminths 

of the red-spotted toad, Bufo punctatus (Anura: 

Bufonidae), from southern Arizona. Journal of the 

Helminthological Society of Washington 58:261- 

269. 

, and . 199 Ib. Helminths of three toads, 

Bufo alvarius, Bufo cognatus (Bufonidae), and 

Scaphiopus couchii (Pelobatidae), from southern 

Arizona. Journal of the Helminthological Society 


, and . 1996. Helminths of the oak toad 


(Bufo quercicus, Bufonidae) from Florida 

(U.S.A.). Alytes 14:122-126. 

-, 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 


Indraneil, D., and G. N. Martin. 1998. Bufo fergusonii. 

Ecology. Herpetological'Review 29:164. 

Joy, J. E., and C. A. Bunten. 1997. Cosmocercoides 

variabilis (Nematoda: Cosmocercoidea) populations 

in the eastern American toad, Bufo a. americanus 

(Salienta: Bufonidae), from western West 

Virginia. Journal of the Helminthological Society 


Kirkland, A. H. 1904. Usefulness of the American 

toad. Farm Bulletin, U.S. Department of Agriculture, 

Washington, D.C., 196:1-16. 

Lees, E. 1962. The incidence of helminth parasites in 

a particular frog population. Parasitology 52:95— 

102. 

McAllister, C. T., S. J. Upton, and D. B. Conn. 

1989. A comparative study of endoparasites in 

three species of sympatric Bufo (Anura: Bufonidae), 

from Texas. Proceedings of the Helminthological 

Society of Washington 56:162-167. 

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

(Rana catesbeiana), green frogs (Rana 

clamitans), and leopard frogs (Rana pipiens) from 

New Brunswick, Canada. Canadian Journal of Zoology 

75:1883-1890. 

Muzzall, P. M. 1991. Helminth infracommunities of 

the frogs Rana catesbeiana and Rana clamitans 

from Turkey Marsh, Michigan. Journal of Parasitology 

77:366-371. 

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

Ohio Journal of Science 54:126-128. 

Plasota, K. 1969. The effect of some ecological factors 

on the parasitofauna of frogs. Acta Parasitologica 

Polonica 16:47-60. 

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

Parasites of the Amphibia. British Museum 

(Natural History)/Oxford University Press, London, 

England. 217 pp. 

Rankin, J. S. 1939. The life cycle of the frog bladder 

fluke, Gorgoderina attenuata Stafford, 1902 

(Trematoda: Gorgoderidae). American Midland 

Naturalist 21:476-488. 

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

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

Volume 2. Academic Press, San Diego, California. 

Smyth, J. D., and M. M. Smyth. 1980. Frogs As 

Host-Parasite Systems. 1. Macmillan, London, 

England. 212 pp. 

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

ed. W. H. Freeman and Company, New York. 859 

pp. 

Toft, C. A. 1981. Feeding ecology of Panamanian litter 

Anurans: patterns in diet and foraging mode. 

Journal of Herpetology 15:139-144. 

Ulmer, M. J. 1970. Studies on the helminth fauna of

Iowa I. Trematodes of amphibians. American 

Midland Naturalist 83:38-64. 

-, and H. A. James. 1976. Studies on the helminth 

fauna of Iowa II. Cestodes of amphibians. 

Proceedings of the Helminthological Society of 


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

liminary observations on seasonal changes in 

prevalence and intensity of Cosmocercoides variabilis 

(Nematoda: Cosmocercoidea) in Bufo 

americanus (Amphibia). Canadian Journal of Zoology 

65:1666-1667. 


Meeting Notices 

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

Reptiles of Wisconsin. The Milwaukee Public 

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

Wisconsin. 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. 

Yoder, H. R. 1998. Community ecology of helminth 

parasites infecting molluscan and amphibian 

hosts. Ph.D. Thesis, University of Wisconsin-Milwaukee. 

138 pp. 

The 8th European Multicolloquium of Parasitology (EMOP-8) will be held on 10-14 September, 

2000 in Poznari, Poland. "Parasitology at the Turn of the Millenium" is the theme of the EMOP- 

8, and the program will consist of: 6 symposia (Malaria in European Travelers; Congenital Toxoplasmosis; 

Toxocara and Toxocariasis; Tapeworm Zoonoses; Human and Animal Trematode Infections; 

and the Status of Food-Born Parasites at the Dawn of the Millenium), 8 scientific sessions 

(Biology and Taxonomy; Molecular Biology; Immunology, Including Vaccines; Epidemiology and 

Control; Parasitic Infections and Diseases; Antiparasitic Drugs and Drug Resistance; The Parasites 

of Fishes and Other Hosts from the Aquatic Environment; and General Parasitology, including 

SNOPAD), and 5 technical workshops (Microscopy Updated; New Methods in Serological Diagnosis 

of Parasitic Infections; Molecular Methods in Diagnosis of Parasitic Infections; Education 

in Parasitology on CD-ROM; and Parasitology on the Internet). Contact information: Organizing 

Committee (Prof. Z. S. Pawlowski and Prof. K. Boczofi), Department of Biology and Medical 

Parasitology, Karl Marcinkowski University of Medical Sciences, Fredry Street 10, 61-701 Poznari, 

Poland. Telephone/Fax (48-61) 852-71-92, e-mail: emop8@eucalyptus.usoms.poznan.pl. Internet: 

http//www.emop8.am.poznan.pl. 

The 4th International Symposium on Monogenea will be held on 9 -13 July 2001 at the Women's 

College of the University of Queensland, Brisbane, Queensland, Australia. A local committee has 

been established to organize details for the various scientific sessions, social events, excursions and 

accompanying persons program. Subject to sufficient demand, there will be a post-symposium 

workshop on Heron Island on the Great Barrier Reef. Calls for expressions of interest to attend the 

symposium, and details regarding registration, submission of abstracts, and a preliminary scientific 

program will be available later in 2000. Contact information: Dr. Ian D. Whittington and Dr. 

Leslie A. Chisolm, Department of Microbiology and Parasitology, The University of Queensland, 

Brisbane, Queensland 4072, Australia. Fax +61 7 3365 4620, e-mail: i.wmttmgton@mailbox.uq. 

edu.au or l.chisolm@mailbox.uq.edu.au. For further information and notices see the Internet web 

page at: http://www.biosci.uq.edu.au/micro/academic/ianw/ism4.htm. 




Ecological Aspects of Endohelminths Parasitizing Cichla monoculus 

Spix, 1831 (Perciformes: Cichlidae) in the Parana River near Porto 

Rico, State of Parana, Brazil 

PATRICIA MIYUKI MACHADO,1 SILVIA CAROLINA DE ALMEIDA,1 

GlLBERTO CEZAR PAVANELLI,2'3 AND RlCARDO MASSATO TAKEMOTO2 

1 Postgraduate Course in Ecology of Continental Aquatic Environments and 

2 Center for Research in Limnology, Ichthyology and Aquaculture (DBI/NUPELIA), State University of 

Maringa, Avenida Colombo 5790, Bloco G-90, 87020-900 Maringa, Parana, Brazil (e-mail: 

gcpavanelli@uem.br) 

ABSTRACT: We examined 136 specimens of Cichla monoculus Spix, 1831, collected in the Parana River near 

Porto Rico, State of Parana, Brazil, from July 1996 through October 1997. Of the total number of fish, 133 

(97.8%) were infected with at least 1 species of helminth. A total of 8 helminth species was recorded: 3 Digenea, 

Clinostomum sp., Diplostomum (Austrodiplostomuni) compactum (Lutz, 1928), and Diplostomum sp.; 3 Cestoda, 

Proteocephalus microscopicus Woodland, 1935, Proteocephalus rnacrophallus (Diesing, 1850), and Sciadocephalus 

megalodiscus Diesing, 1850; 1 Nematoda, Contracaecum sp.; and 1 Acanthocephala, Quadrigyrus machadoi 

Fabio, 1983. Proteocephalus microscopicus and P. macrophallus showed the highest values of prevalence 

and intensity of infection, followed by Contracaecum sp. In the endoparasite community of C. monoculus, the 

cestodes are both dominant and codominant species. The typical pattern of overdispersion or aggregation was 

observed for P. microscopicus, P. macrophallus, S. megalodiscus, Q. machadoi, and Contracaecum sp. Prevalence 

and total host length were positively correlated in fish parasitized by P. microscopicus, P. macrophallus, 

and S. megalodiscus. Infection intensity and host length were positively correlated only for P. microscopicus. 

There were significant differences in the prevalence of P. macrophallus and Q. machadoi in males and females 

of C. monoculus. Clinostomum sp., Diplostomum sp., D. (A.) compactum, and Q. machadoi were found for the 

first time in C. monoculus. 

KEY WORDS: ecology, endohelminths, Digenea, Cestoda, Nematoda, Acanthocephala, freshwater fish, tucunare, 

Cichla monoculus, Cichlidae, Teleostei, Parana River, Brazil. 

Of the main factors influencing the composition 

of endoparasite communities, the feeding 

habits of the hosts are most important, since diverse 

animals that serve as intermediate hosts 

for the hosts' parasites are found in their diets 

(Dogiel, 1970). Changes in the diet and feeding 

habits of fishes also influence the composition 

of their parasite fauna (Dogiel, 1970) and account 

for the differences in the parasite faunas 

of young and adults (Burn, 1980; Scott, 1982; 

Moser and Hsieh, 1992). It is also well understood 

that the fluctuations in water level characteristic 

of floodplains may modify the feeding 

habits of fish because of changes in the quantity 

and quality of available food (Junk, 1980; Lowe- 

McConnell, 1987; Brasil-Sato and Pavanelli, 

1999). The influence of the sex of the hosts is 

another important factor responsible for the variation 

in the composition of their parasitofauna 

and may be related to behavioral, biological, and 

physiological differences between male and fe- 


210 


male fish (Paling, 1965; Muzzall, 1980; Fernandez, 

1985; Moser and Hsieh, 1992; Takemoto et 

al., 1996; Machado et al., 1994). The number of 

studies in Brazil on the ecology of helminth parasites 

of fish, especially in floodplain environments, 

is still small. 

The tucunare, Cichla monoculus Spix, 1831, 

the object of this investigation, is an important 

commercial and sport fish in the Upper Parana 

River. It is a native of the Amazon Basin and 

was first recorded in the Parana River in 1986 

(Agostinho et al., 1994). It is a predator (Lowe- 

McConnell, 1969), considered piscivorous because 

of the predominance of fish in its diet 

(FUEM/CIAMB/PADCT, 1995) and carnivorous 

because it eats shrimp (Fontenele and Peixoto, 

1979; Gery, 1984; Bonetto and Castello, 1985) 

and benthopelagic fish (Ortega and Vari, 1986). 

This study was intended to extend our existing 

knowledge of the helminth fauna parasitizing 

the fishes of the Parana River floodplain, to 

show the structure and diversity of the endoparasitic 

infrapopulations of the tucunare, and to

MACHADO ET AL.—ECOLOGY OF ENDOHELMINTHS OF CICHLA MONOCULUS 211 

Table 1. Prevalence (P%), mean intensity of infection (Mil), mean abundance (MA), range of variation 

(Rx), and sites of infection of the endohelminths of 136 specimens of Cichla monoculus collected in Pau 

Veio Bayou near Porto Rico, state of Parana, Brazil, from July 1996 through October 1997.* 

Parasite Ni Np P (%) Mil ± SD MA ± SD Rx Site of infection 

Digenea 

Clinostomum sp. (M) 

D. (A.) compactum (M) 

Diplostomum sp. (M) 

Cestoda 

Proteocephalus 

microscopicus (A) 


macrophallus (A) 

Sciadocephahts 

megalodiscus (A) 

Nematoda 

Contracaecum sp. (L) 


Quadrigyrus machadoi (L) 

2 18 1.5 9.0 ± 9.9 0.13 ± 1.4 2-16 Branchial cavity and 

stomach 

7 19 5.2 2.7 ± 1.7 0.1 ± 0.7 1-5 Vitreous humor (eye) 

12 16 8.8 1.3 ± 0.5 0.1 ± 0.4 1-2 Vitreous humor (eye) 

128 36,863 94.1 288.0 ± 793.0 27 1.1 ±772.1 1-8,594 Stomach and intestine 

61 1,121 44.9 18.4 ± 74.6 8.2 ± 50.6 1-573 Stomach and intestine 

18 154 13.2 8.6 ±11.0 1.1+4.9 1-42 Stomach and intestine 

96 1,034 70.6 10.8 ± 32.1 7.6 ± 27.3 1-309 Mesentery 

30 76 22.1 2.5 ± 2.0 0.6 ±1.4 1-7 Mesentery (encysted 

L); stomach and intestine 

(free L) 

Ni = number of infected fish; Np = number of parasites; M = metacercaria; A = adults; L = larvae. 

analyze the possible influences of sex and length 

of the hosts on these infrapopulations. 


The fish were collected monthly in Pau Veio Bayou 

on Mutum Island in the floodplain of the Upper Parana 

River, state of Parana, Brazil (22°45'00"S, 

53°16'50"W) from July 1996 through October 1997. 

After capture and identification, the fish were measured, 

weighed, and sexed. They were eviscerated, and 

the visceral cavity, eyes, digestive tract and associated 

organs, kidney, urinary and reproductive tracts, and 

gonads were removed. The organs were separated and 

placed in Petri dishes containing 0.65% physiological 

solution and examined individually with a stereomicroscope. 

The digenetic trematodes were compressed 

between slides and/or coverglasses and fixed in cold 

AFA. The cestodcs and nematodes were fixed in warm 

formol. The acanthocephalans were killed in distilled 

water in Petri dishes under refrigeration and fixed unpressed 

in AFA. All of the worms were preserved in 

70% alcohol. The digenetic trematodes, cestodes, and 

acanthocephalans were stained with acetic carmine or 

Delafield's hematoxylin. All of the worms were dehydrated 

in a graded ethanol series, cleared with 

beechwood creosote, and mounted in Canada balsam. 

For identification of the parasites, the following works 

were used: Diesing (1850), LaRue (1914), Woodland 

(1933, 1935), Yamaguti (1963), Freze (1965), Travassos 

et al. (1969), Schmidt and Hugghins (1973), Moravec 

(1994), Rego (1994), Takemoto and Pavanelli 

(1996), Sholtz et al. (1996), and Silva-Souza (1998). 

Helminths were deposited in the Helminthological 

Collection of the Institute Oswaldo Cruz (FIOCRUZ), 

Rio de Janeiro, state of Rio de Janeiro, Brazil, under 

the following accession numbers: Clinostomum sp. 

34235, Diplostomum (Austrodiplostomwri) compactum 

34233, Diplostomum sp. 34232, Proteocephalus microscopicus 

34234, Proteocephalus macrophallus 

34230, S. megalodiscus 33951, 33952, and 33953 ac, 

Contracaecum sp. 34231, and Quadrigyrus machadoi 

34236. 

Parasite diversity was evaluated by the Shannon diversity 

index (H'). The possible variation in parasite 

diversity was analyzed in relation to sex of the hosts 

by Student's Mest, and in relation to the total length 

of the hosts by the Spearman rank correlation coefficient 

(rs) (Ludwig and Reynolds, 1988). The importance 

value (I) proposed by Bush, according to Thul 

et al. (1985), was used to classify the parasite community 

components. Species in the larval stage were 

not considered in this classification. The dispersion index 

was used to determine the distribution of the infrapopulation 

in the sample. The degree of overdispersion 

or aggregation was calculated using Green's 

index (Ludwig and Reynolds, 1988). These tests were 

applied only to the endohelminth species present at 

prevalences higher than 10%. The correlation between 

total host length and the intensity of infection of the 

parasite species was evaluated by Spearman rank correlation 

coefficient (rs) (Zar, 1996). The existence of a 

correlation between total host length and prevalence of 

infection was tested using Pearson's correlation coefficient 

(r) (9 length classes between 13.1 and 49 cm 

were established) after angular transformation of the 

prevalence data (arc sinVx)(Zar, 1996). Student's r-test 

was used to compare the total lengths of male and 


Table 2. Monthly values of prevalence (P%) and mean intensity of infection (Mil) of the endohelminths 

in Pau Veio Bayou near Porto Rico, state of Parana, Brazil, from July 1996 through October 1997 (N = 

Proteocephalus S 

macrophallus m 

P(%) Mil P 


rnicroscopicus 

Diplostomurn 

sp. 

Diplostornum 

Clinostomum (A.) 

sp. compactum 

Mil 

P(%) 

Mil 

P(%) 

N P(%) Mil P(%) Mil 

Month 

4.8 6 

2.0 

1.0 

12.8 5 

179.5 

53.0 3 

50.0 

50.0 

100.0 

71.4 

80.0 

33.3 

195.6 

37.0 

42.0 

448.9 

2,562.5 

240.0 

87.5 

50.0 

100.0 

100.0 

80.0 

100.0 

1.5 

2.0 

25.0 

50.0 

8 12.5 16.0 — — 

2 — — — — 

2 

1.3 

1.0 

— 

42.9 

20.0 

— 

5 — — — — 

3 — — — — 

1996 

Jul 

Aug 

Sep 

Oct 

Nov 

Dec 

4.0 

1.0 

9.8 

6.0 

1.8 

3.3 

1.0 

2.5 

1.0 

12.0 

100.0 

12.5 

66.7 

46.2 

40.0 

37.5 

42.9 

22.2 

100.0 

100.0 

1,074.0 

1.0 

293.8 

335.7 

180.2 

98.4 

131.3 

112.4 

1 34.0 

531.7 

100.0 

84.4 

100.0 

100.0 

100.0 

100.0 

100.0 

100.0 

100.0 

100.0 

— 

— 

1.5 

1.0 

— 

1.0 

— 

— 

— 

— 

— 

— 

8.3 

15.4 

— 

12.5 

— 

— 

— 

— 

2 — — 100.0 3.5 

32 — — 3.1 1.0 

24 4.2 2.0 8.3 2.5 

13 — — 7.7 5.0 

10 — — 10.0 1.0 

8 — — — — 

7 — — — — 

9 _ _ _ __ 

1 _ _ _ __ 

3 — — — — 

1997 

Jan 

Feb 

Mar 

Apr 

May 

Jun 

Jul 

Aug 

Sep 

Oct 


COMPARATIVE PARASITOLOGY, 67(2;

40- 

35- 

30- 

25- 

20- 

15- 

10- 

5- 

n . 

18.1 

27.1 


39.1 

1 2 3 4 5 

No. parasite species 

Figure I. Parasite richness in 136 specimens of 

Clchla monoculus collected in Pau Veio Bayou near 

Porto Rico, state of Parana, Brazil from July 1996 

through October 1997. 

female hosts. The effect of sex of the host on the prevalence 

of each parasite species was evaluated by the 

log-likelihood G test using a 2 X 2 contingency table 

(Zar, 1996), and the intensities of infection of each 

species of parasite in the male and female hosts were 

compared using the Mann-Whitney (/-test (Siegel, 

1975). In the data analysis, only values with significance 

levels of P •& 0.05 were considered significant. 

Ecological terms are based on Bush et al. (1997). 

Results 

Of the 136 hosts examined, 133 (97.8%) were 

parasitized by 1 or more species of endohelminth, 

of which 39,301 specimens were collected. 

The endohelminths included 3 species of digeneans 

(Clinostomum sp., Diplostomum (Austrodiplostomum) 

compactum (Lutz, 1928), and 

Diplostomum sp.); 3 species of cestodes (Proteocephalus 

microscopicus Woodland, 1935, 

Proteocephalus macrophallus (Diesing, 1850), 

and Sciadocephalus megalodiscus Diesing, 

1850); 1 species of nernatode (Contracaecum 

sp.); and 1 species of acanthocephalan (Quadrigyrus 

machadoi Fabio, 1983) that were found 

free in the intestine or encysted in the mesentery 

in the same stage of development (Tables 1 and 

2). Parasite richness varied from 1 to 5 species, 

and 52 fish were infected by 3 species of parasites 

(Fig. 1). 

The cestodes were the most frequently encountered 

parasites, corresponding to 97% of the 

helminths collected, and were present in 130 

hosts. Proteocephalus microscopicus showed 

the highest percentages of parasitism, followed 

by Contracaecum sp. (Table 1). Proteocephalus 

microscopicus also was the species that presented 

the highest monthly values of prevalence and 

mean intensity (Table 2). The acanthocephalan 

Q. machadoi made up 0.2% of the parasite spec- 

11.3 

1.5 

Table 3. Classification and Bush's Importance values 

(I) of the species of endohelminth parasites of 

136 specimens of Cichla monoculus collected in Pau 

Veio Bayou near Porto Rico, state of Parana, Brazil 

from July 1996 through October 1997. 

Parasite I 

Dominant species 

Proteocephalus microscopicus 98.50 

Proteocephalus macrophallus 1.40 

Co-dominant species 

Sciadocephalus megalodiscus 0.06 

imens collected, while the digenetic trematodes 

represented 0.14%. 

The mean parasite diversity according to the 

Shannon index (H') was 0.1329 (SD = 0.1879), 

and the maximum diversity was 1.3716. Parasite 

diversity did not differ significantly between 

male and female hosts (t = 0.6004, P = 0.5492), 

and was not correlated with total length of the 

hosts (rs = 0.03124, P = 0.7180). Total host 

length varied from 13.5 to 45.7 cm (mean 25.1 

cm). 

Using the importance value (I) proposed by 

Bush, 2 species were classified as dominants and 

1 as co-dominant (Table 3). The parasites of C. 

monoculus showed the typical pattern of overdispersion 

or aggregation of the parasite populations. 

Proteocephalus macrophallus and S. 

megalodiscus showed the highest values of 

Green's index of aggregation (Table 4). There 

was no significant difference in length between 

the 61 male and 75 female tucunares examined 

(t = 0.6130, P = 0.5409). 

There was a positive correlation between total 

length of the hosts and prevalence for fish parasitized 

by P. microscopicus, P. macrophallus, 

and S. megalodiscus. A positive correlation be- 

Table 4. Dispersion Index (DI) and Green's Index 

of Aggregation (GI) for the endohelminth parasite 

species of 136 specimens of Cichla monoculus collected 

in Pau Veio Bayou near Porto Rico, state of 

Parana, Brazil from July 1996 through October 

1997. 

Parasite 

Proteocephalus microscopicus 

Proteocephalus macrophallus 

Sciadocephalus megalodiscus 

Quadrigyrus machadoi 

Contracaecum sp. 


DI 

2,198.96 

312.23 

21.83 

3.27 

98.06 

GI 

0.059 

0.278 

0.136 

0.030 

0.094

. JI 

Table 5. Values of Spearman's rank correlation coefficients (rs) and Pearson's correlation coefficients 

(r), to evaluate the relationship between intensity and prevalence of infection, respectively, of the endohelminth 

fauna with the total length of 136 specimens of Cichla monoculus collected in Pau Veio Bayou 

near Porto Rico, state of Parana, Brazil from July 1996 through October 1997. 

Parasite 






Level of significance. 

rs 

0.3596 

0.2175 

0.1943 

-0.0755 

-0.2954 

tween the total length of the hosts and the intensity 

of parasite infection was seen only for P. 

microscopicus (Table 5). 

In the cestode P. macrophallus and the acanthocephalan 

Q. machadoi, prevalence and intensity 

of infection were influenced by sex of the 

host (Table 6). For P. macrophallus these indices 

were higher in the males, and for Q. machadoi 

in the females (Table 6). In addition to these 

species, S. megalodiscus showed a significant 

difference in intensity of infection according to 

sex of the host, with higher intensity in males 

(Tables 6 and 7). 

Discussion 

In the tucunare, the proteocephalid cestodes 

P. microscopicus and P. macrophallus showed 

the highest values of mean intensity of infection. 

This is explainable by the feeding habits of the 

tucunare, which includes in its diet fish species 

that act as intermediate or paratenic hosts of 

these parasites. The cestodes found in the present 

work appear to be exclusive to the tucunare, 

never having been recorded in other fish (Rego, 

1994). 

For the acanthocephalans, the main factor regulating 

the prevalence and intensity of the par- 

P* 

P < 0.0001 

P = 0.0922 

P = 0.4397 

p = 0.4648 

P = 0.1130 

r 

0.7501 

0.9048 

0.7603 

0.2509 

0.5697 

P 

P = 0.0199 

P = 0.0008 

P = 0.0174 

P = 0.5149 

P = 0.1093 

asitoses is also predation by the fish on the intermediate 

or paratenic hosts (Amin and Burrows, 

1977). In the specimens of C. monoculus 

studied, only larvae of Q. machadoi in the same 

stage of development were found free in the intestine 

or encysted in the mesentery, which may 

indicate that these fish are intermediate or paratenic 

hosts of this parasite. The natural predators 

of tucunare in the study region are carnivorous 

fish or piscivorous birds. The small number 

of fish infected by Clinostomum sp., together 

with the fact that the worms were found free in 

the stomach, may indicate that they were ingested 

accidentally with prey. 

Only 2 individuals of C. monoculus showed 

the maximum parasite richness found, i.e., 5 

species of endoparasites. Most of the population 

was parasitized by 3 species of helminths, of 

which P. microscopicus was always present. 

Holmes (1990) pointed out that parasite richness 

is higher in fishes of intermediate trophic levels, 

since they harbor both adult and larval stages of 

parasites. 

The relationship between body length or age 

of the host and parasite diversity is based on the 

process of temporal accumulation and on the increase 

in the dimensions of the sites of infection 

Table 6. Results of the log-likelihood test (G) to compare prevalence between males and females and of 

the Mann-Whitney l/-test to compare intensity of infection between males and females of 136 specimens 

of Cichla monoculus collected in Pau Veio Bayou near Porto Rico, state of Parana, Brazil from July 1996 

through October 1997. 

Parasite 






G 

1.067 

5.321 

2.210 

0.111 

5.410 

P 

0.5 > P > 0.25 

0.025 > P > 0.01 

0.25 > P > 0.10 

0.75 > P > 0.50 

0.025 > P > 0.01 

P = level of significance; Z = value of normal approximation of {/-test. 

Z 

0.16 

7.32 

3.18 

1.05 

5.71 


P 

P > 0.25 

P < 0.0005 

0.01 > P > 0.005 

0.25 > P > 0.10 

P < 0.005


Table 7. Prevalence and mean intensity of infection by helminth parasites of Cichla monoculus collected 

in Pau Veio Bayou near Porto Rico, state of Parana, Brazil from July 1996 through October 1997.* 

Parasite Nm Nf Nmi Nfi Pm (%) Pf (%) Mim Mif 

Digenea 

Clinostomum sp. 

Diplostomum (A.) compactum 

Diplostomiim sp. 

Cestoda 




Nematoda 




61 

61 

61 

61 

61 

61 

61 

61 

75 

75 

75 

75 

75 

75 

75 

75 

0 

2 

6 

56 

34 

11 

44 

8 

2 

5 

6 

72 

27 

7 

52 

22 

0.0 

3.3 

9.8 

91.8 

55.7 

18.0 

70.5 

13.1 

2.7 

6.7 

8.0 

96.0 

36.0 

9.3 

70.7 

29.3 

— 

4.0 ± 0.0 

1.2 ± 0.4 

434.7 ± 1,160.2 

26.4 ± 99.0 

9.3 ± 12.0 

14.5 ± 46.4 

2.0 ± 

1.1 

9.0 ± 9.9 

2.2 ± 1.8 

1.5 ± 0.6 

173.9 ± 227.6 

8.4 ± 15.3 

7.4 ± 10.0 

7.7 ± 8.8 

2.7 ± 2.2 

* Nm = number of males examined; Nf = number of females examined; Nmi = number of males infected; Nfi = number of 

females infected; Pm and Pf = prevalence of males and females, respectively; Mim and Mif = mean intensity of infection of 

males and females, respectively. 

as a function of growth (Luque et al., 1996). 

Such a relationship has been shown not to exist 

in other species of freshwater fishes (Adams, 

1986; Janovy and Hardin, 1988; Machado et al., 

1996). The lack of this relationship in the tucunare 

may indicate a homogeneity in their 

feeding habits during ontogenetic development. 

The independence of diversity in relation to 

the sex of C. monoculus may constitute evidence 

that the occupation of the habitat and the diet 

are similar in males and females. Adams (1986), 

Janovy and Hardin (1988), and Machado et al. 

(1996) obtained similar results for other species 

of freshwater fishes. 

The cestodes P. microscopicus, P. macrophallus, 

and S. megalodiscus must be considered 

as basic components of the parasite community 

of C. monoculus. The first 2 species were classified 

as dominants, and the third as codominant 

(Thul et al., 1985). The dominant and codominant 

species of endohelminths showed a pattern 

of spatial aggregation, in agreement with the 

typical pattern of endoparasitism demonstrated 

by other investigators (Skorping, 1981; Janovy 

and Hardin, 1987; Oliva et al., 1990; Takemoto, 

1993; Machado et al., 1996). 

The positive correlation observed between the 

total length of the hosts and the prevalence of 

P. microscopicus, P. macrophallus, and S. megalodiscus 

indicates the occurrence of a cumulative 

process. A positive correlation between 

the standard length of hosts and the prevalence 

and/or intensity of infection was observed also 

by Conneely and McCarthy (1986), Machado et 

al. (1994), and Takemoto and Pavanelli (1994); 

the latter 2 investigations were also carried out 

in the Parana River. 

Esch et al. (1988) pointed out that the sex of 

the hosts may also be one of the factors that 

influence levels of parasitism. The influence of 

physiological factors (hormones, mucosity) was 

demonstrated by Paling (1965) and Moser and 

Hsieh (1992), who hypothesized that certain 

species of parasites possess a greater facility to 

infect male or female hosts. Muzzall (1980) and 

Takemoto and Pavanelli (1994) found no influence 

of the sex of the host on the parasite fauna, 

showing that the ecological relationships (behavior, 

habitat, and diet) of males and females 

are similar. In C. monoculus, the sex influences 

the prevalence and intensity of infection of P. 

macrophallus and Q. machadoi and only the intensity 

of S. megalodiscus. This can be explained 

by the fact that some species of fish become 

more susceptible in the breeding season 

because of physiological and behavioral chang- 


We are grateful to Drs. Janet W. Reid and Willis 

A. Reid, Jr., who assisted in translating the 

text into English. 




Adams, A. M. 1986. The parasite community on the 

gills of Fundulus kansae (Garman) from the South 

Platte River, Nebraska (USA). Acta Parasitologica 

Polonica 31:47-54. 

Agostinho, A. A., H. F. Julio, Jr., and M. Petrere, 

Jr. 1994. Itaipu Reservoir (Brazil): impacts of the 

impoundment on the fish fauna and fisheries. Pages 

171-184 in I. G. Cowx, ed. Rehabilitation of 

Freshwater Fisheries. Hull International Fisheries 

Institute, University of Hull, U.K. 

Amin, O. M., and J. M. Burrows. 1977. Host and 

seasonal associations of Echinorhynchus salmonis 

(Acanthocephala, Echinorhynchidae) in Lake 

Michigan fishes. Journal of the Fisheries Research 

Board of Canada 34:325-331. 

Bonetto, A. A., and H. P. Castello. 1985. Pesca y 

Piscicultura en Aguas Continentales de America 

Latina. Secretaria General de la Organizacion de 

los Estados Americanos, Programa Regional de 

Desarrollo Cientifico y Tecnologico, Washington, 

D.C. Monografia 31. 118 pp. 

Brasil-Sato, M. C., and G. C. Pavanelli. 1999. Ecological 

and reproductive aspects of Neoechinorhynchus 

pimelodi Brasil-Sato & Pavanelli (Eoacanthocephala, 

Neoechinorhynchidae) of Pirnelodus 

maculatus Lacepede (Siluroidei, Pimelodidae) 

of the Sao Francisco River, Brasil. Revista Brasileira 

de Zoologia 16:73-82. 

Burn, P. R. 1980. The parasites of smooth flounder, 

Liopsetta putnami (Gill), from the Great Bay Estuary, 

New Hampshire. Journal of Parasitology 

66:532-541. 





Conneely, J. J., and T. K. McCarthy. 1986. Ecological 

factors influencing the composition of the parasite 

fauna of the European eel, Anguilla anguilla 

(L.), in Ireland. Journal of Fish Biology 28:207- 

219. 

Diesing, K. M. 1850. Systema Helminthum. Vol. 1. 

Wilhelm Braumuller, Vindobonae. 679 pp. 

Dogiel, V. A. 1970. Ecology of the parasites of freshwater 

fishes. Pages 1-47 in V. A. Dogiel, G. K. 

Petrushevsky, and Yu. I. Polyanski, eds. Parasitology 

of Fishes. T. F. H. Publications, Inc. Ltd., 

Hong Kong. 

Esch, G. W., C. R. Kennedy, A. O. Bush, and J. M. 

Aho. 1988. Patterns in helminth communities in 

freshwater fish in Great Britain: alternative strategies 

for colonization. Parasitology 96:519-532. 

Fernandez, J. 1985. Estudio parasitologico de Merlucclus 

australis (Hutton, 1872) (Pisces: Merluccidae): 

aspectos sistematicos, estadisticos y zoogeograficos. 

Boletin de la Sociedad de Biologfa 

de Concepcion 56:31-41. 

Fontenele, O., and J. T. Peixoto. 1979. Aprecia9ao 

sobre os resultados da introdu9ao do tucunare 

comum, Cichla ocellaris (Bloch & Schneider, 

1801), nos acudes do nordeste brasileiro, atraves 

da pesca comercial. Boletim Tecnico DNOCS, 

Fortaleza 37:109-134. 


Freze, V. I. 1965. Principles of Cestodology. K. I. 

Skrjabin, ed. Proteocephalata Cestodes of Fishes, 

Amphibians and Reptiles. Moscow: Translated 

from Russian, Israel Program for Scientific Translations, 

Jerusalem. 538 pp. 

FUEM/CIAMB/PADCT. 1995. Estudos ambientais 

da plamcie de inunda£ao do Rio Parana no trecho 

compreendido entre a foz do rio Paranapanema e 

o reservatorio de Itaipu. Final Report 2:411—426. 

Gery, J. 1984. The fishes of Amazonia. In H. Sioli, 

ed. The Amazon: Limnology and Landscape Ecology 

of a Mighty Tropical River and Its Basin. 

Monographiae Biologicae 56, Dr. W. Junk Publishers, 

Dordrecht, The Netherlands. 

Holmes, J. C. 1990. Competition, contacts, and other 

factors restricting niches of parasitic helminths. 

Annales de Parasitologie Humaine et Comparee 

65:69-72. 

Janovy, J., and E. L. Hardin. 1987. Population dynamics 

of the parasites in Fundulus zebrinus in 

the Platte River of Nebraska. Journal of Parasitology 

73:689-696. 

, and . 1988. Diversity of the parasite 

assemblage of Fundulus zebrinus in the Platte 

River of Nebraska. Journal of Parasitology 74: 

207-213. 

Junk, W. J. 1980. Areas inundaveis—um desafio para 

a limnologia. Acta Amazonica 10:775-796. 

LaRue, G. R. 1914. A revision of the cestode family 

Proteocephalidae. III. Biological Monographs 1: 

1-350. 

Lowe-McConnell, R. H. 1969. Speciation in tropical 

freshwater fishes. Biological Journal of the Linnean 

Society 1:51-75. 

. Ecological Studies in Tropical Fish Communities. 

Cambridge University Press, Cambridge, 

U. K. 382 pp. 

Ludwig, J. A., and J. F. Reynolds. 1988. Statistical 

Ecology: Primer on Methods and Computing. Wiley-Interscience 

Publications, New York. 337 pp. 

Luque, J. L., J. F. R. Amato, and R. M. Takemoto. 

1996. Comparative analysis of the communities of 

metazoan parasites of Orthopristis ruber and Haemulon 

steindachneri (Osteichthyes: Haemulidae) 

from the Southeastern Brazilian littoral: II. Diversity, 

interspecific associations, and distribution of 

gastrointestinal parasites. Revista Brasileira de 

Biologia 56:293-302. 

Machado, M. H., G. C. Pavanelli, and R. M. Takemoto. 

1994. Influence of host's sex and size on 

endoparasitic infrapopulations of Pseudoplatystoma 

corruscans and Schizodon borelli (Osteichthyes) 

of the High Parana River, Brazil. Revista 

Brasileira de Parasitologia Veterinaria 3:143-148. 

, , and . 1996. Structure and diversity 

of endoparasitic infracommunities and the 

trophic level of Pseudoplatystorna corruscans and 

Schizodon borelli (Osteichthyes) of High Parana 

River. Memorias do Institute Oswaldo Cruz 97: 

441-448. 

Moravec, F. 1994. Parasitic Nematodes of Freshwater 

Fishes of Europe. Kluwer Academic Publishers. 

Ceske Budejovice. 473 pp. 

Moser, M., and J. Hsieh. 1992. Biological tags for


stock separation in Pacific herring Clupea harengus 

pallasi in California. Journal of Parasitology 

78:54-60. 

Muzzall, P. M. 1980. Population biology and hostparasite 

relationships of Triganodistomum attenuatum 

(Trematoda: Lissorchiidae) infecting the 

white sucker, Catostornus commersoni (Lacepede). 


Oliva, M., J. L. Luque, and J. A. lannacone. 1990. 

The metazoan parasites of Stellifer minor (Tschudi, 

1844): an ecological approach. Memorias do 

Instituto Oswaldo Cruz 85:271-274. 

Ortega, H., and P. R. Vari. 1986. Annotated checklist 

of the freshwater fishes of Peru. Smithsonian Contributions 

to Zoology 437:1-25. 

Paling, J. E. 1965. The population dynamics of the 

monogenean gill parasite Discocotyle sagittata 

Leuckart on Windermere trout, Salmo trutta L. 

Parasitology 55:667-694. 

Rego, A. A. 1994. Order Proteocephalidea Mola, 

1928. Pages 257-293 in L. F. Khalil, A. Jones, 

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

of Vertebrates. CAB International, Wallingford, 

U.K. 

Schmidt, G. D., and E. J. Hugghins. 1973. Acanthocephala 

of South American fishes. Part 1. Eoacanthocephala. 


Scott, J. S. 1982. Digenean parasite communities in 

flatfishes of the Scotia Shelf and southern Gulf of 

St. Lawrence. Canadian Journal of Zoology 60: 

2804-2811. 

Scholz, T., A. Chambrier, A. Prouza, and R. Royero. 

1996. Redescription of Proteocephalus macrophallus, 

a parasite of Cichla ocellaris (Pisces: 

Cichlidae) from South America. Folia Parasitologica 

43:287-291. 

Siegel, S. 1975. Estatfstica Nao Parametrica (Para 

Ciencias do Comportamento). McGraw-Hill do 

Brasil, Sao Paulo, Brazil. 350 pp. 

Silva-Souza, A. T. 1998. Estudo do Parasitismo de 

Plagioscion squamosissimus (Heckel, 1840) (Perciformes, 

Scianidae) por Diplostomum (Austrodiplostomum) 

compactum (Lutz, 1928) (Trematoda, 

Digenea) no Rio Tibagi, PR. Ph.D. Thesis, Universidade 

Federal de Sao Carlos. 125 pp. 

Skorping, A. 1981. Seasonal dynamics in abundance, 

development and pattern of infection of Bunodera 

luciopercae (Miiller) in perch, Perca fluviatilis L. 

from an oligotrophic lake in Norway. Journal of 

Fish Biology 18:401-410. 

Takemoto, R. M. 1993. Estudo taxonomico, sistematico 

e ecologico dos metazoarios parasites de 

Oligoplites palometa, O. saums e O. saliens (Osteichthyes: 

Carangidae) do literal do Estado do 

Rio de Janeiro. M.S. Dissertation, Universidade 

Federal Rural do Rio de Janeiro. 157 pp. 

, J. F. R. Amato, and J. L. Luque. 1996. 

Comparative analysis of the metazoan parasite 

communities of leather] ackets, Oligoplites palometa, 

O. saums, and O. saliens (Osteichthyes: 

Carangidae) from Sepetiba Bay, Rio de Janeiro, 

Brazil. Revista Brasileira de Biologia 56:639— 

650. 

, and G. C. Pavanelli. 1994. Ecological aspects 

of proteocephalidean cestodes parasites of 

Paulicea luetkeni (Steindachner) (Osteichthyes: 

Pimelodidae) from the Parana River, Parana, Brazil. 

Revista Unimar 16:17-26. 

, and . 1996. Proteocephalidean cestodes 

in the freshwater fish Cichla monoculus 

from the Parana River, Brazil. Studies on Neotropical 

Fauna and Environment 31:123-127. 

Thul, J. E., D. J. Forrester, and C. L. Abercrombie. 

1985. Ecology of parasitic helminths of wood 

ducks Aix sponsa, in the Atlantic Fly way. Proceedings 

of the Helminthological Society of 


Travassos, L., J. F. T. Freitas, and A. Kohn. 1969. 

Trematodeos do Brasil. Memorias do Instituto Oswaldo 

Cruz 67:1-886. 

Woodland, W. N. F. 1933. On the anatomy of some 

fish cestodes described by Diesing from the Amazon. 

Quarterly Journal of Microscopical Science 

76:175-208. 

. 1935. Some new Proteocephalids and a Pythobothriid 

(Cestoda) from the Amazon. Proceedings 

of the Zoological Society of London 3:619- 

624. 

Yamaguti, S. 1963. Systema Helminthum. V. Acanthocephala. 

Interscience Publishers, New York. 

860 pp. 

Zar, J. H. 1996. Biostatistical Analysis, 3rd ed. Prentice-Hall, 

Inc., Upper Saddle River, New Jersey. 

662 pp. 




Prevalence of Hookworms (Uncinaria lucasi Stiles) in Northern Fur 

Seal (Callorhinus ursinus Linnaeus) Pups on St. Paul Island, Alaska, 

U.S.A.: 1986-1999 

EUGENE T. LYONS,u TERRY R. SPRAKER,2 KIMBERLY D. OLSON,2 SHARON C. TOLLIVER,' 

AND HEATHER D. BAIR' 

1 Department of Veterinary Science, University of Kentucky, Gluck Equine Research Center, Lexington, 

Kentucky 40546-0099, U.S.A. (e-mail: elyonsl@pop.uky.edu; sctolll@pop.uky.edu) and 

2 Department of Pathology, College of Veterinary Medicine, Colorado State University, Ft. Collins, Colorado 

80523, U.S.A. (e-mail: tspraker@vth.colostate.edu) 

ABSTRACT: Prevalence of hookworms (Uncinaria lucasi) was studied in northern fur seal (Callorhinus ursinus) 

pups at necropsy on St. Paul Island, Alaska, U.S.A. Gross examination of 2,121 pups during the period 1986- 

1998 revealed that only 13 had a light hookworm infection and 2 other pups had hookworm disease (moderate 

hookworm infection, enteritis, and anemia). Specific parasitologic inspections of 230 pups in 1988 and 1996- 

1999 included examinations of feces for eggs or intestines for adult worms. Hookworm eggs were present in 

fecal samples of 1 (2%) of 48 pups in 1988 and none of 19 pups in 1996. Specimens of U. lucasi, found mostly 

during qualitative examinations, were in the intestines of 4 (5%) of 77 pups in 1997, 4 (9%) of 47 in 1998, and 

1 (3%) of 39 in 1999. The 9 infected pups harbored 1-8 hookworms each. Observations in this study indicate 

a dramatic decline in hookworm prevalence in C. ursinus pups on St. Paul Island compared with that of several 

years previously. 

KEY WORDS: Uncinaria lucasi, adult hookworms, prevalence, northern fur seal pups, Callorhinus ursinus, St. 

Paul Island, Alaska, U.S.A. 

Uncinaria lucasi Stiles, 1901, and Uncinaria 

hamiltoni Baylis, 1933, are the only 2 species of 

hookworms described from pinnipeds (Baylis, 

1933, 1947; Stiles, 1901). However, there are 

types with measurements intermediate between 

U. lucasi and U. hamiltoni (Baylis, 1933; Dailey 

and Hill, 1970). Specimens of Uncinaria are 

common in otariids (eared seals) and rare in 

phocids (earless or true seals) (George-Nascimento 

et al., 1992). Classification of hookworms 

in pinnipeds remains uncertain. George-Nascimento 

et al. (1992) considered all hookworms in 

otariids to be the same species, U. lucasi. In the 

present paper, the hookworms are designated as 

U. lucasi because they were first described and 

named from northern fur seals (Callorhinus ursinus 

Linnaeus, 1758). This research was done 

to compare the current prevalence of adult U. 

lucasi in northern fur seal pups on St. Paul Island, 

Alaska, U.S.A., with prevalences found in 

earlier studies. 


Dead fur seal pups were collected from 2 rookeries, 

Northeast Point and Reef, on St. Paul Island, Alaska 

(57°09'N, 170°13'W), for pathologic studies, from 


218 


1986 to 1999. Some of these pups were selected for 

specific research on hookworms. The pups were gathered 

daily from early July through the first 2 weeks of 

August of each year. Pups selected had not been dead 

for more than about 24 hr. Collection was by a person 

working on a catwalk over a rookery and using a 5m-long 

pole equipped with a noose. Pup carcasses recovered 

from rookeries were taken directly to the research 

laboratory on St. Paul Island for necropsy. 

Gross examination of 2,121 dead pups, for the overall 

period from 1986 to 1998, included observations 

for hookworms in the opened ileocecal area. About 

95% of the pups were from Northeast Point and 5% 

from Reef rookeries. The infections were separated 

into 2 categories: 1) light hookworm infections, when 

only a few parasites were noted, and 2) hookworm 

disease, defined as moderate hookworm infection, 

when a number of parasites, enteritis, and anemia were 

evident. 

Special parasitologic examinations of 230 pups were 

conducted. Fecal samples were obtained from the colons 

of 48 pups in 1988 and 19 pups in 1996, placed 

in glass vials or plastic bags containing 5% formalin, 

and examined for hookworm eggs (Lyons et al., 1976). 

Determinations (Lyons, 1963; Olsen and Lyons, 1962, 

1965) for presence of adult hookworms in the intestines 

were made for 77 pups in 1997, for 47 in 1998, 

and for 39 in 1999. Parts of the intestines examined 

were 1) approximately 60-100 mm of the ileum, the 

entire cecum, and about 45-60 mm of the proximal 

end of the colon for all 77 pups in 1977; 2) up to about 

300 mm of the ileum, the entire cecum, and approximately 

150 mm of the colon, including the proximal 

end, for 36 pups, and the entire intestinal tract for 11

Table 1. Summary of results of gross examination 

of the ileocecal area of the intestines of northern 

fur seal pups (n = 2,121) for hookworm (Uncinaria 

lucasi) infection and hookworm disease at necropsy 

from 1986 through 1998 on St. Paul Island, Alaska, 

U.S.A. 

Year 

1986 

1987 

1988 

1989 

1990 

1991 

1992 

1993 

1994 

1995 

1996 

1997 

1998 

Total 

Examined 

39 

90 

91 

113 

364 

248 

227 

141 

251 

130 

172 

165 

90 

2,121 

No. of pups 

With light 

hookworm 

infection 

0 

5 

1 

2 

0 

1 

2 

0 

0 

0 

0 

1 

1 

13 

LYONS ET AL.—HOOKWORMS IN NORTHERN FUR SEAL PUPS 219 

With 

hookworm 

disease* 

0 

0 

0 

1 

0 

0 

0 

0 

0 

0 

1 

0 

0 

2 

Table 2. Results of examination of intestines* of 

northern fur seal pups at necropsy on St. Paul Island, 

Alaska, U.S.A., for the hookworm, Uncinaria 

lucasi, in 1997, 1998, and 1999. 

Year 

1997 

1998 

1999 

Total 

No. of pups 

Examined 

77 

47 

39 

163 

Infected 

(%) 

4(5) 

4 (9) 

1 (3) 

9 (6) 

No. (mean) of hookworms/ 

infected pups 

Male Female Total 

0.50 

0.75 

1.00 

0.67 

0.75 

2.00 

0.00 

1.38 

1.25 

2.75 

1.00 

1.89 

* Ileocecal areas for 113 pups and complete intestinal tracts 

for 50 pups. 

* Moderate hookworm infection, enteritis, and anemia. 

Discussion 

In this study, prevalence of U. lucasi in northern 

fur seal pups on St. Paul Island was low. 

This was determined mostly from gross examination 

of the ileocecal area of the pups' intestines. 

The observed prevalence might have been 

higher if the entire small and large intestine had 

been scrutinized for every pup. However, Olsen 

(1958) demonstrated that U. lucasi concentrate 

in the ileum, cecum, and proximal colon; these 

pups in 1998; and 3) the entire intestinal tract for 39 

pups in 1999. 

areas were examined in all fur seal pups in the 

present study. 

Current rates of infection of adult U. lucasi in 

Results 

northern fur seal pups appear to be much lower 

Results of gross examination of the entire 

sample of 2,121 pups at necropsy from 1986 to 

1998 are summarized in Table 1. Only 13 pups 

had light hookworm infections, and 2 other pups 

were considered to have hookworm disease. 

Examinations of fecal samples revealed hookworm 

eggs in only 1 (2%) of 48 pups in 1988 

and none of 19 pups in 1996. Prevalence of U. 

lucasi, based on recovery of adult specimens in 

the intestines of pups at necropsy, was


Table 3. Prevalence of Uncinaria lucasi in intestines of northern fur seal pups at necropsy in some 

previous surveys on St. Paul Island, Alaska, U.S.A. 

Reference Year of study 

Lucas (1899)* 1897 

Olsen (1954) 1952 

Olsen (1954) 1953 

Olscn (1956, 1958) 1955 

Lyons and Olsen (1960) 1960 

Rookery 

Gorbatch 

Kitovi 

Lagoon 

Lukanin 

Northeast Point 

Polovina 

Reef 

Tolstoi 

Zapadni 

Total 

Unknown 

Polovina 

Kitovi 

Polovina 

Reef 

Tolstoi 

Vostochnif 

Zapadni 

Total 

Little Polovina 

Polovina 

Reef 

Vostochnit 

Zapadni 

Total 

No. of pups 

Examined Infected (%) 

33 

17 

4 

12 

10 

10 

57 

109 

93 

345 

42 

26 

28 

164 

4 

100 

112 

100 

553 

30 

112 

63 

30 

17 

252 

* These data apparently are for causes of death of pups due to U. lucasi rather than actual prevalence. 

t A rookery on Northeast Point. 

rain. The ground where fur seals now breed on 

the 2 rookeries (Northeast Point and Reef) in the 

present study is generally rocky. Previously, 

when populations of fur seals were much higher, 

breeding animals were more widely dispersed to 

include sandy surfaces on these rookeries 

(E.T.L., personal observation). Possibly the decline 

in numbers of animals breeding on sandy 

areas has contributed to the dramatic decrease in 

prevalence of U. lucasi. 

Table 4 summarizes literature on prevalences 

of adult U. lucasi in northern fur seal pups in 

some localities other than St. Paul Island. On the 

Commander Islands (Bering Island and Medny 

Island), Russia, and the Channel Islands (San 

Miguel Island), California, U.S.A., the hookworm 

prevalence is currently high, based on examinations 

of relatively small numbers of dead 

pups. 

No definite cause has been determined for the 

spectacular decline of hookworm infections in 

northern fur seal pups on St. Paul Island. Per- 


15 (45) 

7 (41) 

0(0) 

7 (58) 

7 (70) 

6 (60) 

12 (21) 

52 (48) 

38(41) 

144 (42) 

38 (90) 

24 (92) 

5 (18) 

120 (73) 

13 (27) 

73 (73) 

89 (79) 

67 (67) 

367 (66) 

20 (67) 

86 (77) 

35 (56) 

22 (73) 

1 1 (65) 

174 (69) 

haps it is related to one or more unknown factors 

in combination with a corresponding decline in 

the herd. Numbers of fur seals in the 20th century 

peaked in the 1950s and 1960s and began 

to decline in the 1970s (Trites, 1992). Estimated 

size of the fur seal population on the Pribilof 

Islands (St. Paul Island and St. George Island) 

was about 1.5 million in the 1960s (Baker, 1957; 

Riley, 1961). This population is now estimated 

at about 973,000 (York et al., 2000). The overall 

population decreased about 40% in the last 40 

years, but the number of pups born declined 

much more; i.e., about 60% or from about 

500,000 to 200,000 (York et al., 2000). 

Although it is known that parasitic third-stage 

larvae (L3) of U. lucasi can live for many years 

in the tissues of northern fur seals, there is no 

definite information on the time period or number 

of lactations for clearance of these larvae 

through the mammary system. In experimental 

infections of the nematode Strongyloides ransomi 

Schwartz and Alicata, 1930, in pigs, colos-


Table 4. Prevalence of Vncinaria lucasi in intestines of northern fur seal pups at necropsy in some studies 

in Russia and in California, U.S.A. 

Location Reference 

Russia 

Commander Islands 

Bering Island 

Northern rookery 

Northwestern rookery 

Medny Island 

Southeastern rookery 

Urilie rookery 

Bering Island 

Unknown rookery 


Channel Islands 

San Miguel Island 

West Cove rookery 

Adams Cove rookery 

Year of 

study 

Kolevatova et al. (1978) 1997 

Mizuno (1997) 

Lyons et al. (1997) 

trum samples were collected for 4 consecutive 

lactations from 6 individual sows (Stewart et al., 

1976). There was an exponential decline of parasitic 

L3 of S. ransomi with each lactation. The 

mean number of larvae/ml of colostrum was 1.1 

for the first lactation and 0.06 for the fourth lactation. 

Several aspects of the life cycle of U. lucasi, 

as studied in northern fur seals on St. Paul Island 

(Lyons, 1994), may have a role in the decline in 

prevalence: 1) the only source of adult hookworms 

in pups is from parasitic L3 passed in the 

mother's milk for


ever, at the present time on St. Paul Island, 

hookworm infections and numbers of infected 

fur seals are so low that these parasites appear 

to be at a minimal or subminimal level for continued 

existence. If older fur seals had infections 

of adult hookworms contributing eggs to the environment, 

continued success of the hookworms 

presumably would be more likely. Possibly the 

highly evolved, exclusive manner of transmammary 

transmission of U. lucasi has become a 

detriment for the species under the present conditions 

on St. Paul Island. 


This investigation was made in connection 

with a project of the Kentucky Agricultural Experiment 

Station and is published with the approval 

of the director as paper No. 99-14-5. The 

research was conducted under Marine Mammal 

Protection Act Research Permit No. 837, issued 

to the National Marine Mammal Laboratory, Seattle, 

Washington, U.S.A. 


Baker, R. C. 1957. Fur Seals of the Pribilof Islands. 

Conservation in Action, Number 12. Fish and 

Wildlife Service, U.S. Department of the Interior, 

Washington, D.C. 23 pp. 

Baylis, H. A. 1933. A new species of the nematode 

genus Uncinaria from a sea-lion, with some observations 

on related species. Parasitology 25: 

308-316. 

. 1947. A redescription of Uncinaria lucasi 

Stiles, a hookworm of seals. Parasitology 38:160- 

162. 

Dailey, M. D., and B. L. Hill. 1970. A survey of 

metazoan parasites infecting the California (Zalophus 

californianus) and Steller (Eumetopias jubatus) 

sea lion. Bulletin of the Southern California 

Academy of Science 69:126-132. 

George-Nascimento, M., M. Lima, and E. Ortiz. 

1992. A case of parasite-mediated competition? 

Phenotypic differentiation among hookworms Uncinaria 

sp. (Nematoda: Ancylostomatidae) in 

sympatric and allopatric populations of South 

American sea lions Otaria byronia, and fur seals 

Arctocephalus australis (Carnivora: Otariidae). 

Marine Biology 112:527-533. 

Kolevatova, A. I., le. la. Serebriannikov, V. V. 

Fornin, and L. A. Safronova. 1978. Uncinaria 

lucasi in fur seals. Prophylaxis and treatment of 

agricultural animal diseases. Trudy Kirovskogo 

Sel'skokhoziaistvennogo Instituta 61:49-54. (In 

Russian.) 

Lucas, F. A. 1899. The causes of mortality among seal 

pups. Pages 75—98, Plates 16—21 in The Fur Seals 

and Fur Seal Islands of the North Pacific Ocean 

(David Starr Jordan Report, 1899) Part 3. Washington, 

D.C. 


Lyons, E. T. 1963. Biology of the hookworm, Uncinaria 

lucasi Stiles, 1901, in the northern fur seal, 

Callorhinus ursinus Linn, on the Pribilof Islands, 

Alaska. Ph.D. Dissertation, Colorado State University, 

Fort Collins. 87 pp., 5 pis. 

. 1994. Vertical transmission of nematodes: emphasis 

on Uncinaria lucasi in northern fur seals 

and Strongyloides westeri in equids. Journal of the 

Helminthological Society of Washington 61:169- 

178. 

, R. L. DeLong, S. R. Melin, and S. C. Tolliver. 

1997. Uncinariasis in northern fur seal and 

California sea lion pups from California. Journal 

of Wildlife Diseases 33:848-852. 

, J. H. Drudge, and S. C. Tolliver. 1976. 

Studies on the development and chemotherapy of 

larvae of Parascaris equorum (Nematoda:Ascaridoidea) 

in experimentally and naturally infected 

foals. Journal of Parasitology 62:453-459. 

, M. C. Keyes, and J. Conlogue. 1978. Activities 

of dichlorvos or disophenol against the hookworm 

{Uncinaria lucasi) and sucking lice of 

northern fur seal pups (Callorhinus ursinus) on St. 

Paul Island, Alaska. Journal of Wildlife Diseases 

14:455-464. 

, and O. W. Olsen. 1960. Report on the seventh 

summer of investigations on hookworms, 

Uncinaria lucasi Stiles, 1901, and hookworm diseases 

of fur seals, Callorhinus ursinus Linn, on 

the Pribilof Islands, Alaska, from 15 June to 3 

October 1960. U. S. Department of the Interior, 

Fish and Wildlife Service, Washington, D.C. 26 

pp. 

Mizuno, A. 1997. Ecological study on the hookworm, 

Uncinaria lucasi, of northern fur seal, Callorhinus 

ursinus, in Bering Island, Russia. Japanese Journal 

of Veterinary Research 45:109-110. (English 

summary of a thesis presented to the School of 

Veterinary Medicine, Hokkaido University, Japan.) 

Olsen, O. W. 1954. Report on the third summer of 

investigations on hookworms, Uncinaria lucasi 

Stiles, 1901, and hookworm disease of fur seals, 

Callorhinus ursinus Linn., on the Pribilof Islands, 

Alaska, from May 21 to September 17, 1953. U.S. 

Department of the Interior, Fish and Wildlife Service, 


. 1956. Report on the fifth summer of investigations 

on the hookworm, Uncinaria lucasi Stiles, 

1901 and hookworm disease in fur seals, Callorhinus 

ursinus Linn., on the Pribilof Islands, Alaska, 

from June 9 to August 20, 1955. U.S. Department 

of the Interior, Fish and Wildlife Service, 


. 1958. Hookworms, Uncinaria lucasi Stiles, 

1901, in fur seals, Callorhinus ursinus (Linn.), on 

the Pribilof Islands. Pages 152-175 in Transactions 

of Twenty-third North American Wildlife 

Conference, Wildlife Management Institute, 

Washington, D.C. 

, and E. T. Lyons. 1962. Life cycle of the 

hookworm, Uncinaria lucasi Stiles, of northern 

fur seals, Callorhinus ursinus on the Pribilof Is-

lands in the Bering Sea. Journal of Parasitology 

48 (supplement):42-43. 

-, and . 1965. Life cycle of Uncinaria 

lucasi Stiles, 1901 (Nematoda: Ancylostomatidae) 

of fur seals, Callorhinus ursinus Linn., on the 

Pribilof Islands, Alaska. Journal of Parasitology 

51:689-700. 

Riley, F. 1961. Fur seal industry of the Pribilof Islands, 

1786-1960. Fishery Leaflet 516, U.S. Department 

of the Interior, Fish and Wildlife Service, 


Stewart, T. B., W. M. Stone, and O. G. Marti. 1976. 

Strongyloides ransomi: prenatal and transmammary 

infection of pigs of sequential litters from 


Editors' Acknowledgments 

dams experimentally exposed as weanlings. 

American Journal of Veterinary Research 37:541- 

544. 

Stiles, C. W. 1901. Uncinariosis (Anchylostomiasis) 

in man and animals in the United States. Texas 

Medical News 10:523-532. 

Trites, A. W. 1992. Northern fur seals: why have they 

declined? Aquatic Mammals 18:3-18. 

York, A. E., R. G. Towell, R. R. Ream, J. D. Baker, 

and B. W. Robson. 2000. Population assessment, 

Pribilof Islands, Alaska. Pages 7-26 in B. W. Robson, 

ed. Fur Seal Investigations, 1998. U.S. Department 

of Commerce, NOAA Technical Memorandum 

NMFS-AFSC-113. 101 pp. 

We would like to acknowledge, with thanks, the following persons for providing their valuable 

help and insights in reviewing manuscripts for the Journal of the Helminthological Society of 

Washington and Comparative Parasitology: Omar Amin, Roy Anderson, Hisao Arai, Ann Barse, 

Diane Barton, Jeffrey Bier, Walter Boeger, Matthew Bolek, Daniel Brooks, Richard Buckner, Charles 

Bursey, Albert Bush, Rebecca Cole, Gary Conboy, Murray Dailey, Raymond Damian, David Dean, 

Donald Duszynski, Barbara Doughty, Tommy Dunagan, Larry Duncan, Ralph Eckerlin, William 

Font, Chris Gardner, Tim Goater, Stephen Goldberg, Richard Heckmann, Sherman Hendrix, Eric 

Hoberg, Jane Huffman, Renato Inserra, John Janovy, Hugh Jones, James Joy, Il-Hoi Kim, Michael 

Kinsella, Delane Kritsky, Ralph Lichtenfels, Donald Linzey, Austin Maclnnis, David Marcogliese, 

Lillian Mayberry, Chris McAllister, Robert Miller, Serge Morand, Michael Moser, Patrick Muzzall, 

Steve Nadler, Kazuya Nagasawa, Ronald Neafie, Paul Nollen, John Oaks, David Oetinger, Kazuo 

Ogawa, Niels 0rnbjerg, Wilbur Owen, Michael Patrick, Danny Pence, Gary Pfaffenberger, Oscar 

Pung, Dennis Richardson, Larry Roberts, Klaus Rohde, Wesley Shoop, Allen Shostak, Scott Snyder, 

Robert Sorensen, Jane Starling, Mauritz ("Skip") Sterner, Vernon Thatcher, Dennis Thoney, John 

Ubelaker. 




Life History of Spiroxys hanzaki Hasegawa, Miyata, et Doi, 1998 

(Nematoda: Gnathostomatidae) 

HIDEO HASEGAWA,1'3 TOSHIO Doi,2 AKIKO FunsAKi,1 AND AKIRA MIYATA' 

1 Department of Biology, Oita Medical University, Hasama, Oita 879-5593, Japan 

(e-mail: hasegawa@oita-med.ac.jp) and 

2 Suma Aqualife Park, Wakamiya, Suma, Kobe, Hyogo 654-0049, Japan 

ABSTRACT: The life history of Spiroxys hanzaki Hasegawa, Miyata, et Doi, 1998 (Nematoda: Gnathostomatidae), 

a stomach parasite of the Japanese giant salamander, Andrias japonicus (Temminck, 1836) (Caudata: Cryptobranchidae), 

was studied. The eggs developed in water to liberate sheathed second-stage larvae with a cephalic 

hook. They were ingested by the cyclopoid copepods, Mesocyclops dissimilis Defaye et Kawabata, 1993, and 

Macrocyclops albidus (Jurine, 1820) and developed to infective third-stage larvae in the hemocoel. Natural 

infections with third-stage larvae were also found in the cobitid loaches, Misgurnus anguillicaudatus (Cantor, 

1842) and Cobitis biwae (Jordan et Snyder, 1901). The largest third-stage larva from A. japonicus had almost 

the same body size as the smallest immature adult. 

KEY WORDS: Spiroxys hanzaki, Nematoda, Gnathostomatidae, life history, Andrias japonicus, Japanese giant 

salamander, Caudata, Cryptobranchidae, Copepoda, Mesocyclops, Macrocyclops, Japan. 

The Japanese giant salamander, Andrias japonicus 

(Temminck, 1836) (Cryptobranchidae), 

is an endangered amphibian distributed only in 

West Japan and protected by Japanese national 

law. From this salamander, a new nematode, Spiroxys 

hanzaki Hasegawa, Miyata, et Doi, 1998 

(Gnathostomatidae), was described recently 

(Hasegawa et al., 1998). Although it was suggested 

that the salamander acquired the infection 

by ingesting freshwater fish harboring the infective 

stage of S. hanzaki (Hasegawa et al., 1998), 

there is insufficient evidence for this. Recently, 

viable eggs of S. hanzaki were unexpectedly 

available, allowing attempts to experimentally 

infect copepods as intermediate hosts. In addition, 

freshwater fish captured in the rivers where 

the giant salamanders live were examined for 

larvae of S. hanzaki. The larval stages were also 

compared with those observed in the definitive 

host. We present herein the results of these observations, 

with a discussion on the developmental 

stages of gnathostomatoid nematodes. 


Experiments on embryonic and larval 

development 

On 4 July 1998, 1 A. japonicus reared in the Suma 

Aqualife Park, Kobe, Hyogo Prefecture, Japan, vomited 

a half-digested loach, Misgurnus anguillicaudatus 

Cantor, 1842, that had been given on the previous day 

as food. Many individuals of S. hanzaki at various de- 


224 


velopmental stages were found invading the skin, muscles, 

and viscera of the loach. The loach was kept at 

4°C and transported to the Department of Biology, Oita 

Medical University, for further examination. On arrival 

(6 July 1998), all the worms were still alive. Eggs were 

obtained by tearing the uteri of 2 gravid females. 

Meanwhile, the remaining worms were fixed with 70% 

ethanol at 70°C for routine morphological examination 

or were stored at — 25°C for future biochemical analysis. 

The eggs were incubated in distilled water in a Petri 

dish (9 cm in diameter) at 15°C for 11 days, and then 

the temperature was raised to 20°C to facilitate hatching. 

When larvae hatched, 1 or 2 were transferred by 

a capillary pipette to each of several small Petri dishes 

(3 or 4 cm in diameter) containing about 5 ml of pond 

water. Copepods were collected in a nearby pond or 

paddy with a plankton net and were introduced to the 

dishes containing S. hanzaki larvae. Each copepod was 

observed daily thereafter under a stereomicroscope to 

examine the development of 5. hanzaki larvae inside 

the body. Identification of copepods was based on 

Ueda et al. (1996, 1997). 

Some newly hatched larvae were fixed by slight 

heating to observe their morphology. Infected copepods 

were dissected in physiological saline at various 

days of infection, and recovered larvae were killed by 

slight heating or by placing them in 70% ethanol at 

70°C. Heat-killed larvae were examined immediately, 

whereas those fixed in 70% ethanol were cleared in 

glycerol-alcohol solution by evaporating the alcohol, 

mounted on a glass slide with 50% glycerol aqueous 

solution, and observed under a Nikon Optiphot microscope 

equipped with a Nomarski differential interference 

apparatus. Measurements are in micrometers unless 

otherwise stated. 

Larvae parasitic in naturally infected fish 

Between May and November 1998, the following 

fish were netted in the Hatsuka River and the Okuyama

River, Kobe, Hyogo Prefecture, Japan, and were examined 

for larvae of Spiroxys: from the Hatsuka River: 

47 Zacco temmincki (Temminck et Schlegel, 1846) 

(Cyprinidae) (body length 37-137 mm); 2 Morokojouyi 

(Jordan et Snyder, 1901) (Cyprinidae) (54-58 mm); 

2 Pungutungia herzi Herzenstein, 1892 (Cyprinidae) 

(90-95 mm); 4 Misgurnus anguillicaudatus (Cantor, 

1842) (Cobitidae) (46-80 mm); 13 Cobitis biwae Jordan 

et Snyder, 1901 (Cobitidae) (38-95 mm); 4 Rhinogobius 

flumineus (Mizuno, 1960) (Gobiidae) (33-45 

mm); and 1 Odontobutis obscura (Temminck et Schlegel, 

1845-1846) (Gobiidae) (96 mm); from the Okuyama 

River: 10 Z. temmincki (41-75 mm). Their viscera 

were pressed between 2 thick glass plates and 

observed under a stereomicroscope with transillumination 

to detect Spiroxys larvae. The remaining portions 

of the fish were minced and digested with artificial 

gastric fluid for 3 hr at 37°C. The residues were 

transferred to a Petri dish and examined for nematode 

larvae under a stereomicroscope. Larvae detected were 

processed as described above for morphological observation. 

Scientific names of the fishes follow those 

adopted by Masuda et al. (1984). 

The third-stage larvae of Spiroxys japonica Morishita, 

1926, from the Asian pond loach, M. anguillicaudatus, 

and frogs, Rana nigromaculata Hallowell, 

1860, and Rana rugosa Schlegel, 1838, captured in 

Niigata and Akita Prefectures, northeastern Japan, 

where A. japonicus does not occur, were examined for 

comparison. 

Voucher nematode specimens were deposited in the 

United States National Parasite Collection (USNPC), 

Beltsville, Maryland, U.S.A., Nos. 89629-89638. 

Results 

Embryonic development 

When the culture started, the nematode eggs 

contained 1- to 4-cell-stage embryos. After 2 

days of culture, they developed to 16-cell to 

morula stage. On days 7 and 8 of culture, tadpole-stage 

embryos were seen. On day 10, firststage 

larvae showed movement within the eggshell, 

and some larvae began to molt to become 

second stage. On day 11, molted larvae were 

observed. On day 18, eggs began to hatch (Fig. 

1), and hatched second-stage larvae were still 

enclosed in a sheath, adhered by the tips of their 

tails to the bottom of the culture dish. They seldom 

swam in the water. 

MORPHOLOGY OF HATCHED SECOND-STAGE LAR- 

VAE (n = 4): Stumpy worm with tapered posterior 

portion (Fig. 1). Enclosed within doublelayered 

sheath: outer layer lacking striations, 

and inner layer with reticular markings (Figs. 2, 

3). Length 330-435, maximum width 25-32. 

Anterior end with dorsal sclerotized hooklet 

with elongated base (Fig. 2). Esophagus 118- 

173 long, widened posteriorly and narrowed at 

HASEGAWA ET AL.—LIFE HISTORY OF SPIROXYS HANZAKI 225 

level of nerve ring. Nerve ring 65-85 from anterior 

extremity. Intestinal wall with brown granules. 

Excretory pore, genital primordium, and 

anus indiscernible. 

Development in intermediate host 

Several species of copepods were used for experimental 

infection. Preliminary trials revealed 

that Mesocyclops dissimilis Defaye et Kawabata, 

1993, Macrocyclops albidus (Jurine, 1820), and 

3 species of unidentified cyclopoids readily ingested 

the hatched larvae, but infection was established 

only in the former 2 species. The other 

species could not tolerate the infection and soon 

died. Hence, the following results were based on 

the experiments using M. dissimilis and M. albidus 

as intermediate hosts. 

After being ingested by the copepods, the larvae 

soon migrated to the hemocoel of the host 

(Fig. 4). The sheath was not observed in the larvae 

that had migrated to the hemocoel. Among 

31 M. dissimilis challenged, 15 were found to 

ingest the larvae, whereas worm uptake was not 

confirmed in the remaining individuals. The larvae 

disappeared from the hemocoel of 3 M. dissimilis 

by day 7 after infection. The copepods 

harboring S. hanzaki larvae became emaciated, 

5 of them died by day 10, and 6 more died by 

day 20. The larvae recovered by dissecting these 

dead copepods showed little development, still 

possessing the cephalic hooklet (Fig. 5). In 1 M. 

dissimilis, disseminated fatal infection with unidentified 

flagellates was caused after migration 

of S. hanzaki larvae. Ultimately, only 1 M. dissimilis 

survived for more than 25 days. When 

dissected on the 35th day of infection, this copepod 

harbored 1 living third-stage larva and 1 

dead second-stage larva. 

Among 10 M. albidus challenged, only 2 were 

found to harbor the larvae in the hemocoel on 

day 2 after infection, but 1 of them died by day 

10. The remaining individual died on day 24, 

but 1 third-stage larva was recovered from it by 

dissection. The control copepods, 36 M. dissimilis 

and 10 M. albidus, were not observed to 

be infected with any nematode throughout the 

experiment. 

MORPHOLOGY OF THE SECOND-STAGE LARVAE 

COLLECTED FROM THE INFECTED COPE- 

PODS: Identical with that of the hatched larvae 

but lacking sheaths; size gradually increased as 

the duration of infection lengthened. On day 8 

after infection, length 313-333, maximum width 



Figures 1-3. Second-stage larva of Spiroxys hanzaki. 1. Hatching larva (scale bar = 50 urn). 2. Anterior 

portion, lateral view, showing cephalic hooklet (arrow) and double-layered cuticle (darts) (scale bar = 25 

(Am). 3. Inner layer of cuticle showing reticulated nature (darts) (scale bar = 25 jxm). 

Figure 4. Spiroxys hanzaki larva (arrow) in the hemocoel of Mesocyclops dissimilis on day 1 after 

exposure (scale bar = 200 u.m). 

Figure 5. Second-stage larva collected from the hemocoel of M. dissimilis at 15 days after infection. 

Arrow indicates cephalic hooklet (scale bar = 50 u.m). 

Figures 6-8. Third-stage larvae collected from the infected copepod, lateral view (scale bars = 50 u,m). 

6. Anterior portion. 7. Genital primordium (arrow). 8. Posterior portion. 

Figures 9-10. Third-stage larva of S. hanzaki in naturally infected sand loach, Cobitis biwae, lateral 

view (scale bars = 50 (xm). 9. Anterior portion. 10. Posterior portion. 


HASEGAWA ET AL.—LIFE HISTORY OF SPIROXYS HANZAKl 227 

Table 1. Measurements of third-stage larvae of Spiroxys hanzaki collected from experimentally-infected 

copepods, naturally infected fish, and salamanders (measurements in micrometers unless stated otherwise). 

No. measured 

Body length, mm 

Maximum width 

Nerve ringt 

Excretory poret 

Deiridst 

Esophagus length, mm 

Esophagus width 

Genital primordium, mm:|: 

Tail length 

* Advanced third-stage larvae. 

t Distance from cephalic extremity. 

+ Distance from caudal extremity. 

Mesocyclops 

dissimilis and 

Macrocyclops 

albidus 

2 

1.39-1.80 

61-80 

140-198 

175-219 

220-296 

0.50-0.70 

32-34 

0.46-0.47 

45-56 

18—19 at posterior esophagus level, nerve ring 

69—72 from anterior extremity and esophagus 

115-143 long (n = 2). On day 15, length 345, 

maximum width 18, nerve ring 75 from anterior 

extremity and esophagus 148 long (n =1). 

MORPHOLOGY OF THE THIRD-STAGE LARVAE 

COLLECTED FROM THE INFECTED COPEPODS: Body 

slender. Cuticle with fine transverse striations. 

Lateral alae absent. Anterior extremity with lateral 

pseudolabia with trilobed internal sclerotized 

structure of which dorsal and ventral lobes 

much smaller than median lobe, directed anteriorly 

(Figs. 6, 17). Large submedian papillae 

and amphidial pore present (Fig. 17). Esophagus 

club-shaped. Intestinal Wall densely packed with 

brown granules. Genital primordium with elongated 

2 branches extending anteriorly and posteriorly 

(Fig. 7). Tail conical, with prominent 

phasmidial pores and blunt extremity (Fig. 8). 

Measurements are presented in Table 1. 

Natural infection of fish with Spiroxys larva 

A total of 83 individuals of 7 fish species belonging 

to 3 families was examined during the 

Misgtirnus 

anguillicaudatus and 

Cobitis biwae 

3 

1.33-2.01 

46-56 

118-144 

149-205 

226-304 

0.41-0.58 

28-38 

0.43-0.77 

56-69 

Andrias 

japonicus 

2 

1.76-2.00 

56-90 

176-143 

214-190 

293-296 

0.57-0.65 

28-32 

0.54-0.65 

54-64 

Andrias 

japonicus 

5~'~ 

6.70-9.00 

208-286 

384-455 

462-539 

666-813 

1.83-2.16 

78-102 

2.36-3.02 

150-183 

period from May to November 1998. Only 1 M. 

anguillicaudatus and 2 sand loach, Cobitis biwae 

(Jordan et Snyder, 1901), were found to be 

infected each with 1 larva of Spiroxys. Two of 

the larvae were found encysted on the stomach 

wall and liver surface, whereas the remaining 

larva was recovered by artificial digestion. The 

morphology was identical with that of the larvae 

recovered from the experimentally infected copepods 

(Figs. 9, 10, 18). Measurements are also 

comparable with those of the third-stage larvae 

from the experimentally infected copepods as 

shown in Table 1. 

The third-stage larva of S. hanzaki is readily 

distinguished from that of S. japonica, because 

the latter has inwardly curved dorsal and ventral 

lobes of the internal sclerotized structure in the 

pseudolabium (Figs. 11, 19). 

Morphology of 5. hanzaki larvae and 

immature adults vomited from A. japonicus 

THIRD-STAGE LARVAE (Figs. 12-14): Morphology 

comparable with those from the exper- 

Figure 11. Third-stage larva of Spiroxys japonica collected from pond loach, Misgurnus anguillicaudatus 

from Hachiro-gata, Akita Prefecture, Japan, lateral view, showing inwardly bent dorsal and ventral 

lobes of the sclerotized structure in pseudolabium (scale bar = 50 |xm). 

Figures 12-14. Smallest third-stage larva of S. hanzaki collected from Andrias japonicus, lateral view 

(scale bars = 50 jxm). 12. Anterior portion. 13. Genital primordium (arrow). 14. Posterior portion. 

Figures 15, 16. Advanced third-stage larva vomited by A. japonicus, lateral view (scale bars = 50 |xm). 

15. Anterior extremity. 16. Posterior extremity. 



Figures 17-19. Cephalic extremities of third-stage larvae of Spiroxys spp., lateral view (scale bars = 

25 (Jim). 17, 18. Spiroxys hanzaki collected from experimentally infected copepod, Mesocyclops dissimilis 

(17), and naturally infected sand loach, Cobitis biwae (18). 19. Spiroxys japonica collected from naturally 

infected Rana rugosa (tadpole) in Akita, Akita Prefecture, Japan. 

imentally infected copepods or naturally infected 

fish. Measurements are stated in Table 1. 

ADVANCED THIRD-STAGE LARVAE (Figs. 15, 16): 

Morphology identical with that of the third-stage 

larvae described above but with much larger 

body (Table 1). 

IMMATURE ADULTS: Morphology identical 

with that of mature adults described in Hasegawa 

et al. (1998), but much smaller in size: males 

10.2-13.3 mm long (« = 5) and females 10.2- 

15.5 mm long (n = 5). 

Discussion 

Although only a few third-stage larvae were 

recovered from the experimentally infected copepods, 

it is apparent that 5. hanzaki utilizes copepods 

as its intermediate host, like most gnathostomatoids 

for which life histories have been 

elucidated (cf. Anderson, 1992). Compared with 

Spiroxys contortus (Rudolphi, 1819) and S. japonica 

(cf. Hedrick, 1935; Hasegawa and Otsuru, 

1978), S. hanzaki shows some different 

features in the life history. Hatched larvae are 

much larger (175-294 long in S. contortus, and 

148-207 long in S. japonica). The hatched larva 

attaches at the bottom, like that of 5. contortus, 

whereas the larva of S. japonica often swims in 

the water. Moreover, the period to attain the third 

stage in the copepods is much longer (10 to 14 

days and 6 to 8 days in S. contortus and S. japonica, 


The large size of the hatched larva and the 

slow development may be responsible for the 

high mortality rate of the infected copepods. 

Penetration of such a large larva through the alimentary 

canal wall of the copepod may result 

in perforation, through which pathogenic organisms 

could easily invade the hemocoel, as shown 

by the disseminated infection with flagellates in 


the present experiment. Meanwhile, it is also 

probable that the larva in the hemocoel would 

stimulate some defense mechanism of the copepods 

to eliminate the invader, because the larvae 

often died without further development. 

The worm size and morphology of the thirdstage 

larvae from the copepods are similar to 

those of the smallest third-stage larva found in 

the salamander. This suggests that the third-stage 

larva developed in copepods could be infective 

to the final host. However, most of the gnathostomatoids 

require the second intermediate or 

paratenic hosts, often fish, in which the thirdstage 

larva becomes the so-called advanced third 

stage, that shows significant gain in body size 

but without essential morphological change (cf. 

Anderson, 1992). A similar pattern was also 

postulated for S. hanzaki (Hasegawa et al., 

1998). Because the low tolerance of the copepods 

to the infection in our experiments prevented 

experimental infection of fish with raised 

larva, it remains unknown whether the larvae 

grow to the advanced third stage in fish. 

In most parasitic nematodes, the fourth stage 

exists between the infective third stage and fifth 

(adult) stage. However, the presence of the 

fourth-stage larva in Spiroxys is doubtful. Hedrick 

(1935) stated that the third and fourth molts 

of S. contortus were observed in the definitive 

host turtles but did not present the morphology 

of the fourth stage. In the life history study of 

S. japonica, Hasegawa and Otsuru (1978) could 

not find any larva that was distinguishable morphologically 

from the third-stage larva in the experimentally 

infected definitive host, frogs. Berry 

(1985) described the larvae of Spiroxys chelodinae 

Berry, 1985, collected from the stomach 

ulcers of Australian chelonians, as fourth stage, 

but the morphology resembles that of third-stage

larvae of 5. contortus or S. japonica. In the present 

study, the largest third-stage larva from the 

salamander had nearly the same body size as 

that of the smallest immature adult. These facts 

suggest that Spiroxys lacks a fourth larval stage. 

The presence of the fourth larval stage is also 

unclear- for other gnathostomatoids. In Gnathostorna 

spp., there has been no description of the 

fourth-stage larva. In the life history study of 

Gnathostoma procyonis Chandler, 1942, Ash 

(1962) termed the larva of which a cross-section 

was presented as the fourth stage in the figure 

caption. However, he did not use this term in the 

text or describe a stage morphologically different 

from both the third stage and the adult stage. 

Moreover, we recently observed that Gnathostoma 

doloresi Tubangui, 1925, larvae in molting 

to the adult stage had the cuticle with typical 

arrangement of cephalic booklets of the third 

stage (specimens courtesy of Dr. J. Imai). Further 

careful study is required to determine 

whether gnathostomatoids molt only once in the 

definitive host. 


Sincere thanks are rendered to Dr. J. Imai, Miyazaki 

Medical College, Dr. M. Koga, Kyushu 

University School of Medicine, and Dr. H. Akahane, 

Fukuoka University School of Medicine, 

for their kindness in providing invaluable information 

on the development of Gnathostoma spp. 

Thanks are also extended to Dr. T. Yoshino, University 

of the Ryukyus, for his kindness in verifying 

the scientific names of the fish. This study 

HASEGAWA ET AL.—LIFE HISTORY OF SPIROXYS HANZAKI 229 

was partly supported by the grant-in-aid from 

the Ministry of Education, Science and Culture, 

Japanese Government, No. 11640700. 


Anderson, R. C. 1992. Nematode Parasites of Vertebrates. 

Their Development and Transmission. 

C.A.B International, Wallingford, U.K. 578 pp. 

Ash, L. R. 1962. Migration and development of Gnathostoma 

procyonis Chandler, 1942, in mammalian 

hosts. Journal of Parasitology 48:306-313. 

Berry, G. N. 1985. A new species of the genus Spiroxys 

(Nematoda: Spiruroidea) from Australian 

chelonians of the genus Chelodina (Chelidae). 

Systematic Parasitology 7:59-68. 

Hasegawa, H., A. Miyata, and T. Doi. 1998. Spiroxys 

hanzaki n. sp. (Nematoda: Gnathostomatidae) collected 

from the giant salamander, Andrias japonicus 

(Caudata: Cryptobranchidae), in Japan. Journal 


, and M. Otsuru. 1978. Notes on the life cycle 

of Spiroxys japonica Morishita, 1926 (Nematoda: 

Gnathostomatidae). Japanese Journal of Parasitology 

27:113-122. 

Hedrick, L. R. 1935. The life history and morphology 

of Spiroxys contortus (Rudolphi); Nematoda: Spiruridae). 

Transactions of the American Microscopical 

Society 54:307-335. 

Masuda, H., K. Amaoka, C. Araga, T. Uyeno, and 

T. Yoshino, eds. 1984. The Fishes of the Japanese 

Archipelago. Tokai University Press, Tokyo. 456 

pp. 

Ueda, H., T. Ishida, and J. Imai. 1996. Planktonic 

cyclopoid copepods from small ponds in Kyushu, 

Japan. I. Subfamily Eucyclopinae with descriptions 

of micro-characters on appendages. Hydrobiologia 

333:5=56. 

, , and . 1997. Planktonic cyclopoid 

copepods from small ponds in Kyushu, Japan. 

II. Subfamily Cyclopinae. Hydrobiologia 

356:61-71. 




Inducible Nitric Oxide Synthase in the Muscles of Trichinella sp.- 

Infected Mice Treated with Glucocorticoid Methylprednisolone 

KRYSTYNA BOCZON' AND BARBARA WARGIN 

Department of Biology and Medical Parasitology, Karol Marcinkowski University of Medical Sciences, 

61-701 Poznari, Poland (e-mail: kboczon@eucalyptus.usoms.poznan.pl) 

ABSTRACT: The dynamics of inducible nitric oxide synthase (iNOS) activity in mice infected with Trichinella 

spiralis larvae were followed between the first and tenth week postinfection (p.L). During infection with T. 

spiralis, a bimodal stimulation of iNOS activity to 371% of the control value by day 21 p.i. and to 285% by 

day 70 p.i. was observed. The first increase in iNOS activity was abolished by glucocorticoid treatment. In T. 

pseudospiralis infection, the dynamics of iNOS stimulation differed from that in mice infected with T. spiralis: 

a constant but much weaker stimulation of iNOS starting on day 21 p.i. lasted until the end of the study. The 

results suggest that nitric oxide synthase activity is induced in muscle of the mouse during trichinellosis and 

that nitric oxide may participate in the host's biochemical defense mechanism. 

KEY WORDS: iNOS, inducible nitric oxide, Trichinella spiralis, Trichinella pseudospiralis, muscle, mouse, 

glucocorticoid treatment, methylprednisolone. 

The past decade has witnessed an increase in 

the number of papers devoted to the role of nitric 

oxide (NO) synthase in the pathogenesis of 

many diseases. Part of this surge in interest is 

related to the discovery of a role in both signal 

transduction and cell toxicity for NO. Induction 

of inducible nitric oxide synthase (iNOS) has 

been observed in the course of many human diseases. 

The parasitic infections investigated until 

now include malaria (Tsuji et al., 1995); leishmaniasis 

(Stenger et al., 1996); and toxoplasmosis 

(Holscher et al., 1998). The role of NO in 

killing protozoans of the genus Leishmania was 

studied in greater detail as early as 1993 (Callahan 

et al., 1993), when it was established that 

the course of the disease is dependent to a considerable 

extent on the type of lymphokines generated 

by T lymphocytes. During infection with 

such protozoans as Trypanosoma cruzi Chagas, 

1909 (Rottenberg et al., 1996), or Toxoplasma 

gondii Nicolle et Manceaux, 1908 (Hayashi et 

al., 1996), NO has both antiparasitic and immunosuppressive 

effects. Recent publications 

have also reported modulation of the expression 

of messenger RNA responsible for tumor necrosis 

factor—and prostaglandin E2-independent 

synthesis of iNOS and production of NO in Entamoeba 

histolytica Schaudinn, 1903, infection 

(Wang et al., 1994). The type of free radicals 

contributing to pathogenesis in specific parasitic 

invasion depends on the developmental stage of 


230 


the parasite, and the protective function of NO 

seems to be tissue-specific (Scharton-Kersten et 

al., 1997). 

Nitric oxide generated by nitrogen free radicals 

(RNI), specifically one generated in inflammatory 

conditions by the inducible form of NOS 

(iNOS), is associated with macrophages and 

plays a fundamental role in killing or suppressing 

various pathogens (Gross and Wolin, 1995). 

The mechanism whereby NO influences the cell 

includes, among others, an effect on both respiration 

and oxygen potential in mitochondria 

and Fe-S proteins engaged in the Krebs cycle 

and in electron transport (Kroncke et al., 1995). 

In the case of NO overproduction, the concentration 

of oxygen in the environment plays an 

important role in regulating the functions of mitochondria. 

The balance between RNI and oxygen 

free radicals (ROS) is of special importance. 

Nitric oxide also participates in modulating 

enteritis during the intestinal phase of infection 

with Trichinella spiralis Owen, 1935, since inflammatory 

changes in the intestine of animals 

infected with T. spiralis were eliminated with a 

specific iNOS inhibitor. This suggests that iNOS 

may participate in the disease process associated 

with intestinal invasion by adult forms of T. 

spiralis (Hogaboam et al., 1996) and may 

through its influence on enteritis play an important 

role in rejection of adult worms. 

Our laboratory proposed a hypothesis that 

RNI may also play a role in protective mechanisms 

during the muscular phase of trichinellos-

is. Using histochemical methods our group demonstrated 

NOS in basophilically transformed 

muscle fibers in T. spiralis—infected mice (Hadas 

et al., 1999). In a separate paper we reported on 

the participation of ROS in the biochemical protective 

mechanisms in host muscle infected with 

T. spiralis larvae (Wandurska-Nowak et al., 

1998). In the same paper we demonstrated that 

administration of the glucocorticoid methylprednisolone 

had a profound effect on the activity of 

antioxidant enzymes that were examined (superoxide 

dismutase [SOD] and peroxidase). According 

to Connors and Moncada (1991), glucocorticoid 

also inhibits iNOS. 

The initiation of research on the participation 

of iNOS in biochemical defense mechanisms of 

the host in T. spiralis infection was also important 

from the point of view of its possible participation 

in the mechanism of uncoupling of oxidative 

phosphorylation, which can be observed 

in the mitochondria of tissue infected with helminths 

(Michejda and Boczori, 1972; Van den 

Bosche et al., 1980; Boczori and Bier, 1986; 

Ruble et al., 1989). It was shown that the expected 

temporal correlation between the increase 

in the activity of SOD and peroxidase and the 

peaks in trichinellosis phosphorylation uncoupling 

did not occur (Wandurska-Nowak et al., 

1998). 

The objectives of the present investigation 

were to determine 1) quantitative changes in the 

activity of iNOS in muscles from hosts infected 

with T. spiralis or Trichinella pseudospiralis 

Garkavi, 1976, and 2) if glucocorticoid prevents 

changes in the quantity of NO generated in infected 

tissues. 


Experimental tissue consisted of muscles removed 

from uninfected mice (2-mo-old female mice, strain 

BALB/C) and from mice infected per os with 700-800 

infective larvae of either T. spiralis (strain MSUS/PO/ 

60/ISS3) or T. pseudospiralis (strain MPRO/US/72/ 

ISS13). The infective larvae obtained after pepsin-HCl 

digestion after about 2 hr for T. spiralis larvae and 

about 1-1.5 hr for T. pseudospiralis were administered 

per os to mice anesthetized with ether. The mice were 

killed by decapitation. The amount of larvae per 1 g 

of muscle tissue obtained after pepsin-HCl digestion 

at 6-8 wk post-infection (p.i.) were 10,000-12,000 and 

5,000 for T. spiralis and T. pseudospiralis, respectiveiy. 

Mice were bred and housed in the animal laboratory, 

which ensured approximately constant temperature, 

humidity, and ad libitum access to LMS Labofeed B 

BOCZON AND WARGIN—iNOS IN MOUSE MUSCLE 231 

(Feed and Concentrates Production Plant) granulated 

food and water. 

Only 1 group of animals infected with T. spiralis 

larvae was treated with methylprednisolone (Depomedrol 

[Jelfa, Poland], a drug with prolonged action) 

administered on day 7 p.i. by subcutaneous injection 

at a dose of 20 mg/kg of body weight. Quadriceps 

muscles from hind legs were removed and homogenized 

for 15 to 30 sec in a sucrose medium of the 

following content (in final concentration): 0.25 M sucrose, 

0.002 M EGTA, 0.01 M Tris HC1 buffer (pH 

7.3), and 20 jxl heparin with a concentration of 500 

units/g per 10 ml medium. The homogenate was centrifuged 

for 10 min at 4,500 rpm, and the resulting 

supernatant was centrifuged for 12 min at 15,000 rpm. 

The activity of iNOS was measured in the latter supernatant 

spectrophotometrically by Green's method as 

modified by Lepoivre (Lepoivre et al., 1989), using the 

following solutions: A) Griess' reagent containing 

0.5% sulphanilamide dissolved in 1 N HC1 and 0.15% 

/V-(l-napthyl) ethylendiamine mixed in a ratio of 1:1 

and B) consisting of (in final concentrations) 40 mM 

Tris HC1 buffer (pH 8.0), 2 mM NADPH, and 7 mM 

arginine. Enzyme activity was measured in 140 JJL! of 

supernatant after 30 min of incubation (to induce the 

enzyme activity) at 1-wk intervals at a wavelength of 

X = 540 nm in a cuvette containing 1,200 u,l of solution 

A and 100 u.1 of solution B. In some pilot experiments 

1.5 mM CaCl2 was added. Absorption readings 

were taken after a 30-min incubation period at a 

temperature of 24 °C, and NO concentration was determined 

using a NaNO2 standard curve. Protein was 

measured applying Lowry's method (Lowry et al., 

1951). 

The measurements were carried out in 4 groups of 

animals: for T. spiralis-infected mice (I+NaCl), T. 

spiral is-infected mice under treatment (I+D), and also 

for 2 control groups (C+NaCl and C+D). Both infected 

and untreated mice (I+NaCl) and those from 

the respective control group (C + NaCl) were given intramuscular 

injections of 0.9% NaCl. Activity measured 

in the respective control groups was taken as 

100% for the calculation of percentage changes in such 

activity during T. spiralis infection and treatment. 

Analysis of variance or the Mann—Whitney test was 

used for statistical comparison between groups; P < 

0.01 (very significant) or


Table 1. Activity of iNOS (in nmoles/mg of protein/min) in T. spiralis-infected (I+NaCl) and infected + 

methylprednisolone-treated mouse muscles (I+D). 

Days 

postinfection 

(d.p.i.) 

Controls 

7 

14 

21 

34 

42 

49 

70 

90 

Activity (±SEM) 

0.14 ± 0.01 (8) 

0.08 ± 0.007 (2) 

(P < 0.05) 

0.15 ± 0.007 (4) 

0.52 ± 0.1 (5) 

(P < 0.01) 

0. 1 1 ± 0.003 (6) 

0.07 ± 0.003 (5) 

(P < 0.05) 

0.07 ± 0.003 (6) 

(P < 0.05) 

0.40 ± 0.063 (6) 

(P < 0.01) 

0.04 ± 0.003 (3) 

(P < 0.05) 

I + NaCl* 

% of control 

57 

107 

371 

79 

50 

50 

285 

29 

I + D| 

Activity (±SEM) 

0.11 ± 0.003 (3) 

0.20 ± 0.026 (4) 

0.11 ± 0.053 (5) 

0.16 ± 0.003 (3) 

(P < 0.05) 

0.14 ± 0.056 (11) 

0.06 ± 0.003 (5) 

0.07 ± 0.003 (6) 

(P < 0.05) 

0.37 ± 0.254 (6) 

0.04 ± 0.001 (3) 

% of control 

* The statistically significant differences in column I+NaCl when the values of activity in infected mice were compared with 

normal mice. The number in parentheses is the number of measurements. 

t The comparison of the results from infected and infected and treated by glucocorticoid methylprednisolone mice was carried 

out using a Mann-Whitney test. The number in parentheses is the number of measurements. 

infected and treated, between 7 and 70 days p.i., 

are presented in Table 1. 

The investigations of iNOS activity in muscles 

of mice infected with T. spiralis larvae 

showed a 2-stage increase in enzyme activity 

during the course of trichinellosis. An initial 

peak of activity was seen at 21 days p.i., and a 

second rise in activity occurred at 70 days pi 

(285% of the activity in the control group). Statistically 

the changes in activity during the stages 

of trichinellosis mentioned above varied significantly 

from controls (P < 0.01 or P < 0.05). 

The investigation of muscle enzyme activity 

was carried out on mice infected with T. spiralis 

and treated simultaneously with glucocorticoid 

methylprednisolone (I+D) at the same intervals 

as those for the group of infected and untreated 

animals. The I+D group mice had higher enzyme 

activity than those of the I+NaCl group 

on day 7 p.i. (182% of the control values) and 

at 70 days p.i. (up to 336% of the control value) 

with a statistically significant result compared 

with the control group (P < 0.01). At 90 days 

p.i. enzyme activity fell in a manner similar to 

that seen in T. spiralis-infected and untreated 

animals. Therefore, it may be assumed that 

methylprednisolone exerted a normalizing influence 

on iNOS activity only in the initial stage 

of the muscular phase, i.e., at day 21 p.i. 


182 

100 

145 

127 

56 

64 

336 

The comparison of changes in iNOS activity 

in 2 infections, 1 caused by nonencysting T. 

pseudospiralis larvae and the other by encysting 

T. spiralis larvae, as presented in Figure 1, revealed 

a totally different dynamic for iNOS 

changes in muscles of infected mice. A statistically 

significant difference existed between the 

activity of the enzyme examined in mice infected 

with the T. spiralis and T. pseudospiralis invasion 

during all phases of trichinellosis (P < 

0.01). In general, unlike T. spiralis, infection 

with T. pseudospiralis was characterized by an 

absence of iNOS stimulation during the first 

weeks, while considerable stimulation (up to approximately 

280% of the control) lasted 

throughout and continued up to the end of the 

muscle phase (from 42 to 70 days p.i.). 

Discussion 

The biochemical defense mechanisms for killing 

T. spiralis larvae by eosinophils are mediated 

by peroxidase (POX), with inclusion of the 

process in which both neutrophils and eosinophils 

produce hypochlorous acid, which is toxic 

to the larvae of the parasite (Buys et al., 1981). 

The quantitative results of research on the activity 

of the inducible NOS isoform (iNOS) presented 

in this paper clearly indicate that this enzyme, 

supplying NO lethal to many parasites, is 

37

1.2 n 

1.0- 

|T. spiralis 

|T. pseudospiralis 


49 70 dpi 

Figure. 1. Comparison of the changes of the iNOS activity (mean ± SEM) during T. spiralis and T. 

pseudospiralis infection between 0-70 days postinfection. Values with * or ** (P

234 COMPARATIVE PARASITOLOGY, 67(2), JULY 2(X)0 

lymphocytes and suppression of NO production. 

In the present study, the immunosuppressive 

drug had no effect on iNOS induction in T. spira/zs-infected 

mice muscles. On the other hand, 

during the muscular phase of trichinellosis, 

methylprednisolone exerts a suppressive influence 

on the subpopulation of CDS lymphocytes 

(Boczori et al., unpublished data) with the degree 

of change dependent on the doses of larvae 

used to infect the host. 

Similar investigations on T. pseudospiralis 

were not performed because human cases of T. 

pseudospiralis have not been reported. Nevertheless, 

in severe cases of human trichinellosis 

caused by T. spiralis, glucocorticoid chemotherapy 

is popular, although still disputable. 

It is well known that larvae of T. pseudospiralis 

are less immunogenic than those of T. spiralis 

(Flockhart, 1986). Irrespective of the fact 

that they share 60% of antigens with T. spiralis 

larvae, they still possess 8 different proteins. In 

the intestinal phase, the adult form of T. pseudospiralis 

causes a much weaker inflammatory 

reaction than that of T. spiralis (Flockhart, 

1986). In the late muscular phase (70 days p.i-X 

the presence of T. pseudospiralis larvae resulted 

in a level of iNOS stimulation similar to that 

caused by T. spiralis larvae (approximately 

240%). 

Taking into account that the intensity of the 

T. pseudospiralis infection was 2 times lower 

than that of T. spiralis, we can assume that despite 

the lower immunogenicity of the former, it 

induced much stronger RNI generation. For example, 

on day 7 p.i. the iNOS activity recalculated 

per 1,000 muscle larvae of T. pseudospiralis/g 

of tissue was about 0.08 nmoles/mg of 

protein/min, and during the muscular phase 

measured on day 42 p.i. was 0.16 nmoles/mg of 

protein/min. In T. spiralis infection, the respective 

values for iNOS activity were 0.007 and 

0.006 nmoles/mg of protein/min, being about 10 

and 26 times lower, respectively, than in T. pseudospiralis 

infection. 

Thus, the continuous migration of T. pseudospiralis 

larvae causes a much greater degree 

of damage, but instead of an immunological response, 

a set of biochemical defense reactions, 

including RNI generation, take place. 

The agents present in excretory-secretory 

products of larvae induced iNOS for almost 3 

months. Still, in order to establish the possible 

duration of this effect, it would be necessary to 


carry out a long-term study for at least 6 to 8 

mo p.i. 


Boczori, K., and J. W. Bier. 1986. Anisakis simplex: 

uncoupling of oxidative phosphorylation in the 

muscle of infected fish. Experimental Parasitology 

61:270-279. 

, E. Hadas, E. Wandurska-Nowak, and M. 

A. Derda. 1996. Stimulation of antioxidants in 

muscles of Trichinella spiralis infected rats. Acta 

Parasitologica 41:136-138. 

Brown, G. C. 1995. Nitric oxide regulates mitochondrial 

respiration and cell functions by inhibiting 

cytochrome oxidase. Federation of European Biochemical 

Societies Letters 369:136-139. 

Butterworth, A. E. 1980. Eosinophils and immunity 

to parasites. Transactions of the Royal Society of 

Tropical Medicine and Hygiene 74 (supplement): 

38-43. 

Buys, J., R. Wever, R. Van Stigt, and E. J. Ruitenberg. 

1981. The killing of newborn larvae of 

Trichinella spiralis by eosinophils peroxidase in 

vitro. European Journal of Immunology 11:843— 

845. 

Callahan, H. L., E. R. James, and R. K. Crouch. 

1993. Oxidants and anti-oxidants in trypanosomiasis 

and leishmaniasis. Pages 81 — 110 in O. I. 

Aruoma, ed. Free Radicals in Tropical Diseases. 

Harwood Academic Publishers GmbH, Switzerland. 

Connors, K. J., and S. Moncada. 1991. Glucocorticoids 

inhibit the induction of nitric oxide synthase 

and the related cell damage in adenocarcinoma 

cells. Biochimica et Biophysica Acta 1097:227- 

231. 

Derda, M. 1998. Biochemical aspects of defence in 

experimental trichinellosis. Ph.D. Thesis, K. Marcinkowski 

University of Medical Sciences, Poznari, 

Poland. 51 pp. 

Despommier, D. D. 1990. The worm that would be 

virus. Parasitology Today 6:193—196. 

Flockhart, H. A. 1986. Trichinella speciation. Parasitology 

Today 2:1-3. 

Gross, S. S., and M. S. Wolin. 1995. Nitric oxide 

pathophysiological mechanism. Pages 737—769 in 

J. F. Hoffman and P. De Weer, eds. Annual Review 

of Physiology 57. Annual Review, Inc., Palo Alto, 

California. 

Hadas, E., L. Gustowska, K. Boczori, and D. Janczewska. 

1999. Histochemical investigations of 

the biochemical defence mechanism in experimental 

trichinellosis. II. Nitric oxide synthase activity. 

Wiadomosci Parazytologiczne 45:5—15. 

Hayashi, S., C. C. Chan, R. Gazzinelli, and F. G. 

Roberge. 1996. Contribution of nitric oxide to the 

host parasite equilibrium in toxoplasmosis. Journal 

of Immunology 156:1476-1481. 

Hogaboam, C. M., S. M. Collins, and M. G. Blennerhassett. 

1996. Effects of oral L-NAME during 

Trichinella spiralis infection in rats. American 

Journal of Physiology 271 (2, part l):G338-346. 

Holscher, C., G. Kohler, U. Muller, H. Mossmann,

G. A. Schaub, and F. Brombacher. 1998. Defective 

nitric oxide effector functions lead to extreme 

susceptibility of Trypanosoma crwzz-infected 

mice deficient in gamma interferon receptor or 

inducible nitric oxide synthase. Infective Immunology 

66:1208-1215. 

Kroncke, K. D., K. Fehsel, and V. Kolb-Bachofen. 

1995. Inducible nitric oxide synthase and its product 

nitric oxide, a small molecule with complex 

biological activities. Biological Chemistry 376: 

327-343. 

Lepoivre, M., H. Boudbid, and J. F. Petit. 1989. 

Antiproliferative activity of gamma-interferon 

combined with lipopolysaccharide on murine adenocarcinoma: 

dependence on an L-arginine metabolism 

with production of nitrite and citrulline. 

Cancer Research 9:1970-1976. 

Lowry, O. H., W. J. Rosenbrough, A. Z. Faar, and 

R. J. Randall. 1951. Protein measurments with 

the Folin phenol reagent. Journal of Biological 

Chemistry 193:265-275. 

Michejda, J. W., and K. Boczori. 1972. Changes in 

bioenergetics of skeletal muscle mitochondria during 

experimental trichinosis in rats. Experimental 


Rottenberg, M. E., E. Castanos-Velez, R. Mesquita, 

O. G. Laguardia, P. Bibcrfeld, and A. 6rn. 

1996. Intracellular co-localization of T. cruzi and 

inducible nitric oxide synthase: Evidence for a 

dual pathway of iNOS induction. European Journal 

of Immunology 26:3203-3213. 

Ruble, C. J., C. A. Boehm, and F. L. Bygrave.1989. 

Aberrant energy-linked reactions in mitochondria 

isolated from the livers of rats infected with the 

liver fluke Fasciola hepatica. Biochemical Journal 

260:517-523. 


Scharton-Kersten, T. M., G. Yap, J. Magram, and 

A. Sher. 1997. Inducible nitric oxide is essential 

for host control of persistent but not acute infection 

with the intracellular pathogen Toxoplasma 

gondii. Journal of Experimental Medicine 185: 

1261-1273. 

Stenger, S., N. Donhauser, H. Thuring, M. Rollinghoff, 

and C. Bogdan. 1996. Reactivation of 

latent leishmaniasis by inhibition of inducible nitric 

oxide synthase. Journal of Experimental Medicine 

183:1501-1514. 

Tsuji, M., Y. Miyahira, R. S. Nussenzweig, M. 

Aguet, M. Reichel, and F. Zavala. 1995. Development 

of antimalarial immunity in mice lacking 

IFN-gamma receptor. Journal of Immunology 154: 

5338-5344. 

Wandurska-Nowak, E., K. Boczori, M. A. Derda, E. 

Hadas, and A. Matias-Guzek. 1998. The effect 

of methylprednisolone on the dynamics of oxygen 

uptake and peroxidase and superoxide dismutase 

activities in skeletal muscles of mice infected with 

Trichinella spimlis larvae. Acta Parasitologica43: 

54-56. 

Wang, W., K. Keller, and K. Chadee. 1994. Entamoeba 

histolytica modulates the nitric oxide synthase 

gene and nitric oxide production by macrophages 

for cytoxicity against amoebae and tumour 

cells. Immunology 83:601-610. 

Van den Bosche, H., H. Verhoeven, and H. Lauwers. 

1980. Uncoupling of liver mitochondria associated 

with fascioliasis in rats: normalization by 

closantel. Pages 699-704 in H. Van den Bosche, 

ed. The Host-Invader Interplay. Elsevier/North- 

Holland Biomedical Press, Amsterdam, New 

York, and Oxford. 




The Expulsion of Echinostoma trivolvis: Worm Kinetics and 

Intestinal Cytopathology in Jirds, Meriones unguiculatus 

TAKAHIRO FUJINO, K5 TOMONORI SHINOHARA,' KOICHI FuKUDA,2 HIDETAKA ICHIKAWA,S 

TOMOYUKI NAKANO,' AND BERNARD FRIED4 

1 Department of Biology, Faculty of Science, Yamagata University, 990-8560 Yamagata, Japan 

(e-mail: tfuji@sci.kj.yamagata-u.ac.jp), 

2 Center for Laboratory Animal Science, National Defense Medical College, Tokorozawa 359-8513, Japan 

(e-mail: kfukuda@cc.ndmc.ac.jp), 

3 Department of Medical Zoology, Kanazawa Medical University, Ishikawa 920-0293, Japan 

(e-mail: ichikawa@kanazawa-med.ac.jp), and 

4 Department of Biology, Lafayette College, Easton, Pennsylvania 18042, U.S.A. 

(e-mail: fried@lafvax.lafayette.edu) 

ABSTRACT: Worm kinetics and cytopathology of jirds, Meriones unguiculatus Milne-Edwards, 1967, infected 

with Echinostoma trivolvis (Cort, 1914) Kanev, 1985, were reported and compared with previous studies on 

echinostome infections in murine hosts. Seven jirds were each infected with 40 metacercarial cysts, and the 

worms were recovered at days 5, 8, 10, 12, 15, and 17 postinfection (p.i.). Worm recoveries were 35.4, 10.7, 

and 0.4 at days 5, 10, and 15 p.i., respectively. Worm expulsion occurred on about day 10 p.i., corresponding 

to the peak increase in the number of goblet cells at 24.3 ± 0.6/villus-crypt unit (VCU) at day 10 p.i. These 

data showed that worm expulsion of E. trivolvis in jirds occurred earlier than that in C3H and BALB/c mouse 

hosts. The difference in expulsion times and rates reflects differences in the peak number of goblet cells in the 

host intestines of jirds versus mice. The number of mucosal mast cells increased slightly and peaked at 1.1 ± 

0.32/10 VCU at day 10 in jirds. An increase in mucosal mast cells occurred earlier and was smaller in jirds 

than in BALB/c mice. Scanning electron microscope observations showed an irregular arrangement of microvilli 

in the small intestine of infected jirds. Transmission electron microscope observations also showed damage in 

the distal parts of villi in infected jird intestines and the appearance of numerous vesicles in the infected 

epithelium. 

KEY WORDS: Echinostoma trivolvis, worm expulsion, worm kinetics, intestine, cytopathology, jird, Meriones 

unguiculatus, SEM, TEM. 

Echinostoma trivolvis (Cort, 1914) Kanev, contained sulfomucins, whereas those in ham- 

1985, is expelled within several weeks of infec- sters contained sialomucins. Probably differention 

from the intestines of various strains of ces in mucin characteristics account in part for 

mice (Mus musculus Linnaeus, 1758): ICR (Ho- differences in infectivity of E. trivolvis in these 

sier and Fried, 1986; Weinstein and Fried, hosts. Thus, differences in infectivity depend in 

1991), BALB/c (Fujino et al., 1993), C3H (Fu- part on genetic differences in murine strains 

jino and Fried, 1993a; Fujino et al., 1996), and used as hosts of echinostomes. 

Swiss Webster (Hosier and Fried, 1986) mice, Studies on E. trivolvis in jirds have not been 

whereas this echinostome species is retained for done. However, Christensen et al. (1990) demore 

than 15 weeks in the intestines of golden scribed survival and fecundity of an allopatric 

hamsters (Mesocricetus auratus Waterhouse, echinostome species, Echinostoma caproni Ri- 

1839) (Huffman et al., 1986; Franco et al., chard, 1964, in hamsters and jirds (Meriones un- 

1986). Fujino et al. (1993, 1996) noted that guiculatus Milne-Edwards, 1867). Mahler et al. 

worm expulsion is mainly caused by an in- (1995) noted considerable differences in the recreased 

secretion of mucins by hyperplastic gob- productive capacity of E. caproni in hamsters 

let cells. Fujino and Fried (1993b; 1996) exam- versus jirds. The hamster was more susceptible 

ined glycoconjugates in intestinal mucins in to E. caproni infection than was the jird. 

C3H mice versus golden hamsters infected with The purpose of the present study was to report 

E. trivolvis and reported that goblet cells in mice information on the infection, growth, and distribution 

of E. trivolvis in jirds and to compare our 

data with previous studies on this species in 

5 Corresponding author. mouse strains and in the golden hamster. Path- 

236 


Table 1. Infectivity and distribution of Echinostoma trivolvis in jirds. 

Group 

A 

B 

C 

D 

E 

F 

Day 

postinfection 

5 

8 

10 

12 

15 

17 

No. of 

exposed 

(infected) 

jirds 

7 (7) 

7 (7) 

7 (7) 

7 (5) 

7 (1) 

7 (0) 

* I = anterior; II = middle; III = posterior. 

t Not significant. 

Mean (± SE) 

No. (%) of worms 

recovered 

14.1 ± 2.4 (35.4) 

13.6 ± 2.2 (33.9)t 

4.3 ± 0.9 (10.7) 

1.7 ± 1.8 (4.3) 

0. 1 ± 0.4 (0.4) 

0 

ological, histochemical, and electron microscopical 

studies of the intestines of jirds infected 

with E. trivolvis were also carried out. 


Metacercarial cysts of Echinostoma trivolvis were 

obtained from the kidney and pericardial sac of laboratory-infected 

Biomphalaria glabrata (Say, 1816) 

snails. The worm strain was previously described by 

Fujino and Fried (1993a). Forty cysts were fed via a 

stomach tube to each jird, and 7 jirds were lightly 

anesthetized with ether and killed by cervical dislocation 

at days 5, 8, 10, 12, 15, and 17 postinfection 

(p.i.). Six groups of untreated control jirds, 7 per 

group, were also killed on the same days as the infected 

hosts. The jirds were starved for about 12 hr 

prior to necropsy to avoid food residue in the intestine. 

The intestine was removed and opened longitudinally 

to determine worm location. The worms were counted, 

and their distribution was recorded in the small intestine, 

which was divided equally into anterior, middle, 

and posterior regions, and in the cecum and colon plus 

rectum. Where applicable, Student's /-test was used to 

analyze differences between means, and P < 0.05 was 

considered statistically significant. 

For histological samples, pieces of intestine (2 cm 

long) located 10 cm anterior to the cecum, corresponding 

to the middle jejunum to ileum, were excised and 

fixed for 3 hr in Carney's fixative. The samples were 

dehydrated with an ethanol series and embedded in 

paraffin. Histological sections 5ixm thick were stained 

with periodic-acid Schiff for goblet cell mucins. Mucosal 

mast cells were stained with alcian blue (pH 0.3) 

and safranin O. All counts were expressed as the number 

of cells per villus-crypt unit (VCU) (Miller and 

Jarrett, 1971) for goblet cells and cells per 10 VCU 

for mast cells. Thirty to 50 VCUs were analyzed per 

host. Logarithmic transformation of data: geometric 

means (antilog of mean log of data) was performed. 

For comparison of the cell counts, goblet cell numbers 

were multiplied 10 times as for the mast cells. This 

transformation tends to stabilize variance and to normalize 

such data. 

For scanning electron microscopy (SEM), the intestinal 

tissue from jirds infected with E. trivolvis at 8 

FUJINO ET AL.—EXPULSION OF ECHINOSTOMES 237 

No. of worms located in the: 

Small intestine 

Total (I II III)1 Cecum 

97 ( 4 49 44) 

90 (30 21 39) 

30(13 8 9) 

10 ( 3 4 3) 

1(1 0 0) 

0 

1 

4 

0 

1 

0 

0 

Colon + 

rectum 

and 10 days p.i. and control tissues were excised from 

the upper ileum, opened longitudinally with fine needles, 

and pinned on small rubber boards in physiological 

saline. The intestinal debris was removed by gentle 

flow of saline forced over the surface with a pipette. 

After a brief rinse in 0.1M sodium cacodylate buffer 

(pH 7.4), the specimens were fixed for 3 hr with 3% 

glutaraldehyde, postfixed for 3 hr in 0.1 M osmium 

tetroxide (pH 7.4), and then dehydrated in an ethanol 

series. The material was dried in a carbon dioxide critical-point 

drying apparatus (Hitachi HCP-2, Tokyo, Japan), 

coated with palladium in a Hitachi E 1030, Tokyo, 

Japan ion sputter, and examined in a Hitachi s- 

450 SEM, Tokyo, Japan at 10 kV. For transmission 

electron microscopy (TEM), the material was prepared 

as described for SEM procedures in Fujino and Fried 

(1993a). Ultrathin sections stained with uranyl acetate 

and lead acetate were viewed in a JEOL JEM 1210 

electron microscope operating at 80 kV. 

Results and Discussion 

Infectivity and worm recovery data are presented 

in Table 1. All jirds were infected with 

E. trivolvis at days 5, 8, and 10 p.i., and this was 

confirmed by fecal examination under the microscope. 

By day 12 p.i., 5 of 7 jirds were infected, 

but only 1 was infected by day 15 p.i. 

Worm recoveries were 35.4% and 33.9% at days 

5 and 8 p.i., respectively, and this difference was 

not statistically significant. The recovery data 

dropped to 10.7% at day 10 p.i., fell markedly 

to 4.3% by day 12 p.i., and finally to 0.4% by 

day 15 p.i. Most worms were expelled between 

days 10 and 15 p.i., and all were expelled by 

day 17 p.i. Most worms were found in the middle 

to posterior part of the small intestine at day 

5 p.i. The worms moved mainly anteriad to the 

middle of the small intestine by day 10 p.i. Such 

an anteriad worm shift was reported previously 

in ICR mice infected with E. trivolvis at day 21 

p.i. (Weinstein and Fried, 1991). In the present 


1 

0 

0 

1 

0 

0


500 

0.01 

8 10 

Day post-infection 

Figure 1. Goblet cell (mean ± SE)/VCU (•) and mast cell (•) (mean ± SE)/VCU numbers of the 

anterior section of the ileum of jirds, each of which was infected with Echinostoma trivolvis metacercarial 

cysts. A logarithmic transformation was performed on the number of the cells for normalizing the data. 

For comparison of the cell counts, goblet cell numbers were multiplied 10 times as for the mast cells. 

study, the worms moved posteriad to the cecum 

and colon plus rectum by day 12 p.i. In BALE/ 

c mice infected with E. trivolvis, the recovery 

rate of the worms was over 44% for days 6-10 

p.i. and worm expulsion occurred from day 10 

to 12 p.i., corresponding to the peak increase in 

goblet cells (Fujino et al., 1996). Those worm 

recovery rates were much higher than what is 

seen in the present study on jirds, i.e., 35.4% 

and 33.9% at days 5 and 8 p.i., respectively. 

Therefore, worm expulsion occurred from days 

8 to 12 p.i. These data showed that worm expulsion 

in jirds occurred earlier than in murine 

hosts, probably reflecting a difference in the 

peak number of goblet cells in jirds and mice. 

Christensen et al. (1990) examined the establishment, 

survival, and fecundity in E. caproni and 

the allopatric species of E. trivolvis in hamsters 

and jirds. They noted that the jird exhibited an 

overall low susceptibility to E. caproni infection. 

The jird's low susceptibility to E. caproni 

is different from that of E. trivolvis. According 

to Ellerman and Morrison-Scott (1951), the jird 


(M. unguiculatus) belongs to the subfamily Gerbillinae 

of the family Muridae and differs both 

taxonomically and genetically from the golden 

hamster (M. auratus) of the subfamily Cricetinae 

and also from various mouse strains of Mus 

musculus of the subfamily Murinae. It is known 

that Gerbillinae is genetically closer to Murinae 

than Cricetinae (Ellerman and Morrison-Scott, 

1951). The present infection data on E. trivolvis 

in jirds generally correspond to the above-noted 

taxonomic and genetic differences in murine 

hosts. In conclusion, the recoveries of E. trivolvis 

from jirds were lower than those from mice 

and much lower than those from golden hamsters. 

It is possible that these differences in recoveries 

reflect the genetic differences among 

these 3 hosts, jirds, mice, and hamsters. 

Kinetic changes in the number of goblet cells/ 

VCU at the anterior sections (n = 50) of the 

ileum with or without the parasites present are 

shown in Figure 1. The number of goblet cells 

in infected jirds increased markedly, peaked at 

24.3 ± 0.6/VCU at day 10 p.i. and then de- 

12

FUJINO ET AL.—EXPULSION OF ECHINOSTOMES 239 

Figure 2. SEM of the control and infected intestinal surface of jirds. a. Control (normal) intestine, 

with round villi having regularly arranged microvilli. Scale bar = 5.0 (Jim. b. Intestine infected for 10 

days with Echinostoma trivolvis. The intestinal microvilli appear irregularly arranged (arrows) and partly 

peeled off (arrowheads). Scale bar = 5.0 fjim. 

clined. The number of goblet cells in the control 

was 9.6 ± 0.4/VCU. The numbers of mucosal 

mast cells were so small that their kinetic changes 

were examined in 10 VCU, although the 

number of mast cells in infected jirds increased 

gradually from 0.2 ± 0.42/10 VCU at day 5 p.i. 

to reach a peak of 1.1 ± 0.32/10 VCU at day 

10 p.i. and then declined gradually. For comparison 

of the cell counts, goblet cell numbers 

were multiplied 10 times as for the mast cells. 

The numbers of cells were log-transformed to 

normalize the data in Figure 1. Fujino et al. 

(1993) examined the worm kinetics and intestinal 

cytopathology in conventional and congenitally 

athymic BALB/c mice and noted that worm 

rejection was caused by goblet cell hyperplasia 

and not by mast cells. 

The SEM observations of the surface of the 

intestinal villi infected with E. trivolvis and the 

control showed a rough and irregular arrangement 

of microvilli in the infected intestine compared 

with the regular microvilli arrangement in 

the control intestine (Fig. 2). The intestine infected 

with worms was partly damaged by day 

10 p.i., and its epithelial surface was eroded. The 

TEM observations showed that the intestinal epithelium 

appeared more electron-dense than that 

in the control (not shown). The distal ends of 

the villi were partly broken and the microvilli 

were partially eroded. Numerous vesicles of various 

sizes appeared in the intestinal epithelium. 

The appearance of these vesicles was also re- 

ported in BALB/c and C3H mice infected with 

E. trivolvis by Fujino et al. (1993) and Fujino 

and Fried (1993a), respectively. Matrices of 

many small rounded mitochondria were granular 

and irregularly condensed. Elongate nuclei with 

an irregular peripheral margin had heterochromatin 

arranged in small patches and randomly 

distributed. Fujino and Fried (1996) noted histopathological 

differences in mouse versus hamster 

small intestine infected with E. trivolvis and 

showed no marked histopathological and histochemical 

changes in the hamster intestines. They 

suggested that the response of the hamster to E. 

trivolvis infection was relatively weak and that 

this host showed only a limited capacity to expel 

E. trivolvis. 


Christensen, N. 0., P. E. Simonsen, A. B. Odaibo, 

and H. Mahler. 1990. Establishment, survival 

and fecundity in Echinostoma caproni (Trematoda) 

infections in hamsters and jirds. Journal of the 

Hclminthological Society of Washington 57:104- 

107. 

Ellerman, J. R., and T. C. S. Morrison-Scott. 1951. 

Checklist of Palaearctic and Indian Mammals, 

1758 to 1946. British Museum (Natural History), 

London, England. 810 pp. 

Franco, J., J. E. Huffman, and B. Fried. 1986. In 

fectivity, growth, and development of Echinostoma 

revolutum (Digenea: Echinostomatidae) in the 

golden hamster, Mesocricetus auratus. Journal of 


Fujino, T., and B. Fried. 1993a. Expulsion of Echinostoma 

trivolvis (Cort, 1914) Kanev, 1985 and 



retention of E. caproni Richard, 1964 (Trematoda: 

Echinostomatidae) in C3H mice: pathological, ultrastructural, 

and cytochemical effects on the host 

intestine. Parasitology Research 79:286-292. 

, and . 1993b. Echinostoma caproni 

and E. trivolvis alter the binding of glycoconjugates 

in the intestinal mucosa of C3H mice as determined 

by lectin histochemistry. Journal of Helminthology 

67:179-188. 

, and . 1996. The expulsion of Echinostoma 

trivolvis from C3H mice: differences in 

gycoconjugates in mouse versus hamster small intestinal 

mucosa during infection. Journal of Helminthology 

70:115-121. 

-, H. Ichikawa, and I. Tada. 1996. 

Rapid expulsion of the intestinal trematodes Echinostoma 

trivolvis and E. caproni from C3H mice 

by trapping with increased goblet cell mucins. International 

Journal for Parasitology 26:319-324. 

-, and I. Tada. 1993. The expulsion of 

Echinostoma trivolvis: worm kinetics and intestinal 

cytopathology in conventional and congenitally 

athymic BALB/c mice. Parasitology 106: 

297-304. 

Hosier, D. W., and B. Fried. 1986. Infectivity, 

growth, and distribution of Echinostoma revolutum 

in Swiss Webster and ICR mice. Proceedings 

of the Helminthological Society of Washington 

53:173-176. 

Huffman, J. E., C. Michos, and B. Fried. 1986. Clinical 

and pathological effects of Echinostoma revolutum 

(Digenea: Echinostomatidae) in the golden 

hamster, Mesocricetus auratus. Parasitology 

93:505-515. 

Mahler, H., N. 0. Christensen, and 0. Hindsbo. 

1995. Studies on the reproductive capacity of 

Echinostoma caproni (Trematoda) in hamsters and 

jirds. International Journal for Parasitology 25: 

705-710. 

Miller, H. R. P., and W. F. H. Jarret. 1971. Immune 

reactions in mucous membranes. I. Intestinal mast 

cell response during helminth expulsion in the rat. 

Immunology 20:277-288. 

Weinstein, M. S., and B. Fried. 1991. The expulsion 

of Echinostoma trivolvis and retention of Echinostoma 

caproni in the ICR mouse: pathological 

effects. International Journal for Parasitology 21: 

255-257. 

2000-2001 MEETING SCHEDULE OF THE 

HELMINTHOLOGICAL SOCIETY OF WASHINGTON 

11 October 2000 

15 November 2000 

17 January 2001 

14 March 2001 

5 May 2001 

George Washington University, Washington, DC (Contact 

Person: Ralph Eckerlin, 703-323-3234). 

Anniversary Dinner, Location to be announced. 

Nematology Laboratory, Beltsville Agricultural Research 

Service, USDA, Beltsville, MD (Contact Person: Lynn 

Carta, 301-504-8787). 

Naval Medical Research Center, 503 Robert Grant Avenue, 

Silver Spring, MD (Walter Reed Forest Glen Annex 

Bldg. 503) (Contact Person: Eileen Franke-Villasante, 

301-319-7667). 

Joint Meeting with the New Jersey Society for Parasitology 

at the New Bolton Center, University of Pennsylvania, 

Kennett Square, PA (Contact Person: Jay Ferrell, 

215-898-8561). 




Effects of a High-Carbohydrate Diet on Growth of Echinostoma 

caproni in ICR Mice 

MARK R. DARAS, SUSAN SISBARRO, AND BERNARD FRIED' 

Department of Biology, Lafayette College, Easton, Pennsylvania 18042, U.S.A. (e-mail: friedb@lafayette.edu) 

ABSTRACT: The effects of a high-carbohydrate diet (HCD) on the host-parasite relationship of Echinostoma 

caproni Richard, 1964, in ICR mice were studied. The experimental diet was a customized HCD containing 

63% carbohydrates, 14% protein, 4% fat, and 19% cellulose. The control diet, a standard laboratory diet, 

contained 31% carbohydrate, 20% protein, 7% fat, and 42% cellulose. Thirty-six mice were each infected with 

35 metacercarial cysts; 18 mice were fed the HCD and the remaining mice received the control diet. Equal 

numbers of experimental and control mice were necropsied at 2, 3, and 4 weeks postinfection (p.i.). Comparisons 

of worm body area in uniformly fixed and stained worms were made at 2, 3, and 4 weeks p.i. There was no 

significant difference in body area in worms from each group at 2 and 3 weeks p.i. At 4 weeks p.i. the body 

area of worms from hosts on the HCD was significantly greater than that of worms from hosts on the control 

diet. The findings suggest that the HCD contributes to growth enhancement of E. caproni in ICR mice. 

KEY WORDS: trematodes, high-carbohydrate diet, Echinostoma caproni, ICR mice, growth. 

Previous studies in our laboratory have examined 

the effects of various experimental diets 

of hosts on growth and development of Echinostoma 

caproni Richard, 1964, in Institute for 

Cancer Research (ICR) mice. Sudati et al. (1996, 

1997) used this model to study the effects of 

high-lipid and high-protein diets, respectively, in 

ICR mice. Rosario and Fried (1999) examined 

the effects of a protein-free host diet on growth 

and development of E. caproni in ICR mice. 

Although studies are available on the effects 

of a high-carbohydrate host diet on gastrointestinal 

trematodes, this topic has been studied extensively 

in rats infected with hymenolipid cestodes 

(e.g., Read; 1959; Read and Simmons, 

1963). It is clear from the literature that hymenolipids 

thrive best in rodent hosts maintained on 

high-carbohydrate diets (see Von Brand, 1973, 

for review). Because of the lack of information 

on gastrointestinal trematodes maintained in rodent 

hosts fed a high-carbohydrate diet (HCD), 

we initiated this study to examine the effects of 

such a diet on worm recovery, growth, and distribution 

of E. caproni in ICR mice. Echinostoma 

caproni now is a well-established model 

for conducting such studies of intestinal trematode 

infections in nutritionally altered hosts. 

Gracyzk and Fried (1998) examined the recent 

literature on human echinostomiasis and 

noted that it is a common but forgotten foodborne 

disease. Because echinostomiasis may oc- 


241 

cur in people from socioeconomic groups that 

have relatively high-carbohydrate, low-protein 

diets, studies on the effects of HCD on the model 

echinostome, E. caproni, seemed appropriate. 


Metacercarial cysts of Echinostoma caproni were 

removed from the kidney/pericardial region of experimentally 

infected Biomphalaria glabrata (Say, 1818) 

snails and fed by stomach tube (35 cysts per mouse) 

to 36, 6 to 8-week-old, female ICR mice (Manger and 

Fried, 1993). The experimental mice were fed a customized 

HCD in pellet form containing 63% cornstarch 

as a source of carbohydrate, 14% protein, 4% 

fat, and 19% cellulose (Dyets Inc., Bethlehem, Pennsylvania, 

U.S.A.). The control mice were fed a standardized 

rat-mouse-hamster (RMH) 3000 diet in pellet 

form containing 31% carbohydrate, 20% protein, 7% 

fat, and 42% cellulose (US Biochemicals Co., Cleveland, 

Ohio, U.S.A.). Both diets contained essential vitamins 

and minerals as described previously. The HCD 

was about 1.3 times more calorific than the normal diet 

(Rosario and Fried, 1999). 

A total of 36 mice was used in the experiment; 18 

mice were maintained on the HCD, and the remainder 

on the RMH diet. On the day of infection, the mice 

were weighed and maintained 6 per cage on either the 

HCD or the RMH diet. Food and water were provided 

ad libitum. Six mice on the HCD and 6 mice on the 

RMH diet were each necropsied at 2, 3, and 4 weeks 

post infection (p.i.). Mice were weighed on the day 

they were fed cysts and at necropsy. At that time, the 

small intestine was removed from the pyloric sphincter 

to the ileocecal valve and divided into 5 equal sections 

numbered 1-5, beginning with the pylorus. Worms 

were removed from the small intestine, and their location 

and number in each section were recorded. 

Worms were rinsed in Locke's solution and fixed in 

hot (85°C) alcohol-formalin-acetic acid. Twenty 



0 1 2 3 4 

Weeks postinfection 

Figure 1. Mean (± SE) weights of mice on high 

carbohydrate (diamonds) versus control (squares) 

diet at 0—4 weeks postinfection. 

worms at each data point were selected at random from 

mice on the HCD and RMH diets and stained in Gower's 

carmine, dehydrated in ethanol, cleared in xylene, 

and mounted in Permount (Kaufman and Fried, 

1994). Length and maximum width measurements of 

worms were made with the aid of a calibrated ocular 

micrometer to give body area in mm2 for control and 

experimental worms at 2, 3, and 4 weeks p.i. Length 

and width measurements were also made on the gonads 

and suckers to determine if there were significant 

differences in organ sizes between worms on HCD 

versus RMH diet (Sudati et al., 1997). Whenever possible, 

differences in means between groups were determined 

using Student's f-test, with P < 0.05 being 

considered significant. 

Results 

Mean weights of mice on both the HCD and 

RMH diet are shown in Figure 1. Mouse weight 

in both groups increased rapidly until 2 weeks 

p.i. and then less rapidly until 4 weeks p.i. Although 

the weights of mice on the RMH diet 

were slightly higher than those of mice on the 

HCD diet, there was no significant difference in 

mouse weight between groups at any week p.i. 

There was no apparent difference in food consumption 

in mice on either diet. 

From 2 to 4 weeks p.i., the small intestines of 

hosts on the HCD were yellow compared to the 

tan-colored intestines of hosts on the RMH diet; 

the intestines of mice on the HCD were thinner, 

more translucent, and more brittle than those of 

hosts on the RMH diet. All worms from hosts 

on both diets were ovigerous at 2 to 4 weeks p.i. 


2 3 4 

Weeks postinfection 

Figure 2. Effects of diet on mean (± SE) E. caproni 

worm body area; control diet (closed bar) and 

high-carbohydrate diet (open bar). 

Eggs taken at random from some worms maintained 

on the HCD, when incubated in artificial 

spring water, produced miracidia that were capable 

of infecting B. glabrata. 

The mean body areas of worms from the hosts 

on the RHM diet and on the HCD are shown in 

Figure 2. At 2 and 3 weeks p.i., there were no 

significant differences in the body areas of 

worms from either group. However, a significant 

increase in body areas was seen in worms from 

the experimental hosts at 4 weeks p.i. compared 

with that of worms from the control hosts. There 

was a significant difference at 4 weeks p.i. in 

the length of the anterior and posterior testes and 

in the diameter of the acetabulum and oral sucker 

of worms from the HCD group compared 

with those on the RMH diet. 

The percent worm recovery is shown in Figure 

3 and was similar in control and experimental 

mice at all 3 sampling points with about 50% 

recovery in both groups at all data points. More 

worms from hosts on the RMH diet were located 

in segments 3 and 4 than those from hosts on 

the HCD at sampling points. Considerably more 

worms on the HCD were located in segment 5, 

compared with worms on the RMH diet at all 

sampling points. Worms from the HCD group 

were more widely dispersed in their hosts than 

those from hosts on the RMH diet; worms from 

HCD hosts were also located more posteriad 

than those from hosts on the RMH diet.

0) 

0> 

O 20- 

O 

2 3 4 

Weeks post infect ion 

Figure 3. Effects of diet on E. caproni worm recovery 

in mice exposed to 35 cysts/hosts; control 

diet (closed bar) and high-carbohydrate diet (open 

bar). 

Discussion 

Worms from hosts on the HCD, when compared 

with those from mice on the RMH diet, 

showed a marked increase in body area at 4 

weeks p.i. This is the first report that documents 

enhanced growth of a digenean maintained in an 

experimental vertebrate host fed an HCD. Echinostomes 

on the HCD showed greater body area 

by 4 weeks p.i. Reasons for the increase in 

worm body area are not readily apparent from 

the findings in this study. We have no way of 

knowing if the HCD had a direct effect on worm 

growth (i.e., if the worms consumed more carbohydrates 

from the HCD than the RMH diet) 

or had an indirect effect by altering gut constituents. 

Our results suggest that the intestines of 

hosts on the HCD showed a loss of normal integrity. 

The HCD at 4 weeks p.i. could have 

contributed to the altered gut that allowed for a 

large number of mucosal epithelial cells to be 

sloughed off, thereby increasing the food supply 

available to the echinostomes in hosts on the 

HCD. Perhaps such an increased food supply 

DARAS ET AL.—GROWTH OF ECH1NOSTOMA CAPRONI 243 

was a factor in the enhanced worm growth. 

Since there were no control uninfected mice on 

the HCD, there is also no way of knowing if 

some of the changes in host guts may not have 

been caused by interactions between diet and 

worms. 

Distribution data are interesting in that, in 

hosts on the HCD, worms were more spread out 

and also located more posteriad than worms 

from hosts on the RMH diet. The disparate arrangement 

of the worms in the HCD hosts is 

similar to previous observations on mice infected 

with E. caproni and maintained on diets with 

altered amounts of fats and proteins (Sudati et 

al., 1996, 1997; Rosario and Fried, 1999). 


Graczyk, T. K., and B. Fried. 1998. Echinostomiasis: 

a common yet forgotten food borne disease. 

American Journal of Tropical Medicine and Hygiene 

58:501-504. 

Kaufman, A. R., and B. Fried. 1994. Infectivity, 

growth, distribution and fecundity of a six versus 

twenty-live metacercarial cyst inoculum of Echinostoma 

caproni in ICR mice. Journal of Helminthology 

68:203-206. 

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. 

Read, C. P. 1959. The role of carbohydrates in the 

biology of cestodes. VIII. Some conclusions and 

hypotheses. Experimental Parasitology 8:365- 

382. 

, and J. E. Simmons. 1963. Biochemistry and 

physiology of tapeworms. Physiological Reviews 

43:263-305. 

Rosario, C., and B. Fried. 1999. Effects of a proteinfree 

diet on worm recovery, growth and distribution 

of Echinostoma caproni in ICR mice. Journal 

of Helminthology 73:167-169. 

Sudati, J. E., A. Reddy, and B. Fried. 1996. Effects 

of high fat diets on worm recovery, growth and 

distribution of Echinostoma caproni in ICR mice. 

Journal of Helminthology 70:351-354. 

, F. Rivas, and B. Fried. 1997. Effects of a 

high protein diet on worm recovery, growth and 

distribution of Echinostoma caproni in ICR mice. 

Journal of Helminthology 71:351-354. 

Von Brand, T. 1973. Biochemistry of Parasites, 2nd 

ed. Academic Press, New York. 499 pp. 




Surface Ultrastructure of Larval Gnathostoma cf. binucleatum from 

Mexico 

MASATAKA KoGA,1-6 HIROSHIGE AKAHANE,2 RAFAEL LAMOTHE-ARGUMEDO,3 

DAVID OSORIO-SARABIA,3 LUIS GARC1A-PRIETO,3 JUAN MANUEL MARTINEZ-CRUZ,4 

SYLVIA PAz DiAZ-CAMACHO,5 AND KANAMI NOD A1 

1 Department of Microbiology (Parasitology), Graduate School of Medical Sciences, Kyushu University, 

Fukuoka 812-8582, Japan (e-mail: masakoga@linne.med.kyushu-u.ac.jp), 

2 Department of Parasitology, School of Medicine, Fukuoka University, Fukuoka 814-0180, Japan, 

3 Laboratorio de Helmintologia, Departamento de Zoologia, Institute de Biologia, Universidad Nacional 

Autonoma de Mexico 04510 D.F., Mexico, 

4 Pedro Garcia No. 918, Tierra Blanca, Veracruz, Mexico, and 

5 Facultad de Ciencias Quirnico-Biologicas, Universidad Autonoma de Sinaloa, Culiacan, Sinaloa, Mexico 

ABSTRACT: We examined the morphology of gnathostome larvae obtained in Temazcal and Sinaloa, Mexico, 

mainly using scanning electron microscopy. The mean body length was 4.67 mm. The head had 4 transverse 

rows of hooklets, and the mean number of each row was 40, 44, 47, and 50. The bodies were wholly covered 

with minute cuticular spines along their transverse striations. The mean number of striations varied from 227 to 

275. The cervical papillae were situated between the 13th and 17th transverse striations, and most specimens 

had them between the 14th and 15th transverse striations. An excretory pore was also located between the 24th 

and 28th transverse striations. We identified this Mexican gnathostome as Gnathostoma cf. binucleatum Almeyda-Artigas, 

1991. 

KEY WORDS: Gnathostoma cf. binucleatum, scanning electron microscopy, morphology, Mexico. 

Gnathostomiasis is an important parasitic zoonosis, 

mainly endemic in such countries as Japan, 

Thailand, and Vietnam, where people often 

eat raw freshwater fish. For this reason, this 

food-borne disease was thought to be limited to 

Southeast Asian countries. In 1970, however, a 

case of human gnathostomiasis was reported in 

Mexico (Pelaez and Perez-Reyes, 1970). The patient 

was neither a traveler nor an immigrant 

from Southeast Asia. After this initial discovery, 

the number of gnathostomiasis patients increased 

drastically; more than 1,000 cases have 

been diagnosed in Mexico. The endemic area in 

Mexico includes 6 states, which are roughly divided 

into 3 regions, including the Pacific coast 

(Culiacan), Atlantic coast areas (Tampico), and 

regions (Veracruz) adjacent to Central American 

countries (Ogata et al., 1998). Lamothe-Argumedo 

et al. (1989) and Almeyda-Artigas (1991) 

examined the morphology of gnathostome larvae 

from fish in Oaxaca-Veracruz. Later Akahane 

et al. (1994) examined by light microscopy 

the morphology of the larvae collected from pelicans 

in the same area. 

We herein report the morphology of specimens 

of Gnathostoma cf. binucleatum Almeyda- 


244 


Artigas, 1991, from Mexico, which were examined 

using scanning electron microscopy 

(SEM). The results were compared with our previous 

SEM study of larvae of Gnathostoma spinigerum 

Owen, 1836, Gnathostoma doloresi 

Tubangui, 1925, and Gnathostoma hispidum 

Fedtschenko, 1872, obtained in Japan, China, 

and Thailand (Koga et al., 1987, 1988, 1994). 


Three American white pelicans (Pelecanus erythrorhynchos 

Gmelin, 1789) were collected in the Presidente 

Miguel Aleman Reservoir in Temazcal, Oaxaca, 

Mexico, and their muscles were examined for gnathostome 

larvae. The muscles were removed, chopped 

into small pieces, and then cut into thin slices. The 

slices were then placed between 2 glass plates (10 X 

10 cm, 2 mm thick), pressed by hand, and examined 

under a dissecting microscope. The muscle remnants 

were then digested in artificial gastric juice (0.2 g pepsin 

in 0.7 ml HC1/100 ml distilled water) for 3 hours 

at 37°C to collect any larvae that might have been 

overlooked. The muscles of another ichthyophagous 

bird, a great egret (Egrctta alba Linnaeus, 1758), captured 

at a dike of the San Lorenzo River in Culiacan, 

were also examined. These larvae were processed for 

morphological examination by both light microscopy 

and SEM. Paraffin sections of specimens were prepared 

by conventional methods and stained with Mayer's 

hematoxylin and eosin. 

For the SEM specimen preparations, 10 viable lar-

vae from Temazcal and 3 from Culiacan were washed 

in distilled water and stored in a refrigerator until the 

worms relaxed completely. They were then fixed in 

10% formalin for 7 days. The larvae then were washed 

overnight in running tap water to remove the fixative 

and were transferred to distilled water. The specimens 

were rinsed twice in Millonig's phosphate buffer and 

postfixed overnight in 0.5% OsO4 in the same buffer. 

All specimens were then carefully and gradually dehydrated 

in an ascending series of ethanol, since such 

specimens often shrink or have surface wrinkles because 

of rapid dehydration. They were transferred into 

amyl acetate and CO2 critical-point dried with a Hitachi 

HCP-2 dryer (Tokyo, Japan). The specimens 

were sputter-coated with gold and examined with a 

JEOL JSM-U3 SEM (Tokyo, Japan) operated at 15 kV. 

Results 

As many as 570 larvae were obtained from 

the 3 pelicans in Temazcal. Only 3 larvae were 

found in 5 egrets in Culiacan. The mean body 

length (10 larvae) was 4.67 mm, measured in a 

relaxed state after natural death in cold distilled 

water. The heads had 4 transverse rows of hooklets 

(Fig. 1), and the mean number in each row 

was 40, 44, 47, and 50 booklets. The typical 

hooks on the head bulb had sharp tapering points 

composed of hard keratin that emerged from an 

oblong chitinous base (Fig. 2). The bodies were 

wholly covered with minute cuticular spines 

along their transverse striations. The mean number 

of striations varied from 227 to 275. A pair 

of cervical papillae was laterally situated between 

the 13th and 17th transverse striations 

(Fig. 3). In most specimens, the papillae were 

located between the 14th and 15th striations. A 

ventral excretory pore was located between the 

24th and 28th transverse striations (Fig. 4). A 

wide terminal anal opening was visible on the 

ventral surface, and the transverse striations on 

the body were limited to the extent of this opening 

(Fig. 5). Both ends of the larva had a pair 

of lateral phasmidial pores (Fig. 6). 

The intestinal cells had multiple nuclei in the 

larvae from Temazcal (Fig. 7). The larvae from 

Sinaloa had 2 to 7 nuclei in each intestinal cell 

(Fig. 8). 

Discussion 

Lamothe-Argumedo et al. (1989) determined 

their larval gnathostome specimens obtained 

from Temazcal to be Gnathostoma sp. However, 

based on our observations, their specimens 

seemed to be the same as those reported by Almeyda-Artigas 

(1991); both specimens of larvae 

were from both fish and waterfowl in the same 

KOGA ET AL.—SURFACE ULTRASTRUCTURE OF GNATHOSTOMA 245 

endemic area of human gnathostomiasis, and the 

descriptions of the larval morphology were quite 

similar. We attributed this specimen as G. binucleatum. 

Lamothe-Argumedo et al. (1989) 

had previously observed larvae in Oaxaca, Temazcal, 

Mexico. We think that their SEM observations 

were insufficient, especially regarding 

the location of excretory pores and numbers of 

the transverse striations on the larval bodies. We 

reexamined the Temazcal specimens using SEM 

and made some new observations. We also examined 

the surface structures of the specimens 

from Sinaloa, Culiacan. Previously, 5 specimens 

from Sinaloa were examined by Camacho et al. 

(1998) using SEM. They mentioned the numbers 

of booklets of 4 rows on the head bulb as 39, 

42, 44, and 49. Furthermore, they recognized 1 

pair of cervical papillae located between the 

13th and 15th striations of the cuticular spines 

on a single larva. The number of transverse striations 

on the body was more than 200. There 

were no descriptions regarding the location of 

the excretory pore. The locations of the cervical 

papillae, the excretory pore, and the number of 

transverse striations are very important for the 

identification of species of gnathostome larvae. 

As shown in Table 1, the number of transverse 

striations is more than 200 in G. spinigerum. 

However, the number is less than 200 in most 

specimens of G. doloresi. On the other hand, the 

cervical papillae and excretory pores of G. hispidum 

were situated more anteriorly than those 

of the other 2 species. 

In the present study, we compared the larvae 

from 2 districts in Mexico, Temazcal and Culiacan, 

and found no differences between them in 

the larval morphology (Table 1). In particular, 

the surface ultramorphologies were very similar. 

However, when our findings were compared 

with those of G. spinigerum in Thailand (Table 

1), they were the same, including the shape of 

the larval hooks, which had oblong chitinous bases 

and are known to be one of the characteristic 

structures of G. spinigerum (Miyazaki, 1960). 

Akahane et al. (1994) also compared the number 

of booklets in each row on the head bulb of the 

Temazcal larvae and the larvae of G. spinigerum 

in Thailand by light microscopy and concluded 

that the numbers of booklets in Temazcal larvae 

were slightly less than those of G. spinigerum. 

The intestinal epithelium of Temazcal specimens 

consisted of a single layer of intestinal 

cells, and each columnar cell had 2 to 5 nuclei 



Figures 1-4. Scanning electron micrographs of Gnathostoma cf. binucleatum. 1. Lateral view of the 

head bulb of Temazcal specimen. The arrow indicates the cervical papilla. Scale = 50 |xm. 2. An enlarged 

view of the booklets. The base of each booklet has an oblong shape. Sharp keratin hooks armed posteriorly. 

Scale = 10 u.m. 3. A mammary form of the cervical papilla (CP) protruding from the tegument. Scale = 

3 jxm. 4. The oval-shaped opening of the excretory pore (EP), which opens ventrally. Scale = 3 u.m. 

(Akahane et al., 1994). This feature closely resembles 

that of the Sinaloan specimen. Once 

again, no differences were observed in intestinal 

cells between the larvae from Temazcal and Sinaloa, 

and we conclude that both should be included 

in the same species (G. binucleatum). 

Further, we could not differentiate G. binucleatum 

from G. spinigerum based on the number of 

nuclei in the intestinal cells. Most specimens of 

G. spinigerum from Thailand also had 2 to 4 


nuclei in the intestinal cells. On the other hand, 

the number of nuclei in the intestinal cells of 

other Asian species, e.g., G. hispidurn and G. 

doloresi, have only 1 nucleus per cell (Akahane 

et al., 1994). Almeyda-Artigas' light microscopic 

observations of the larvae were limited regarding 

the number of booklets in each row and 

the number of nuclei in the intestinal cells. Recently 

Koga et al. (1999) experimentally obtained 

the adults of this Mexican gnathostome


Figures 5, 6. Scanning electron micrographs of Gnathostoma cf. binucleatum. 5. The terminal end of 

a larva where the anal opening (AP) is clearly visible on the ventral surface of the larva, which has a 

crescent shape. Scale = 3.5 (Jim. 6. The terminal extremity of a larva, showing a lateral phasmidial pore 

(PH). Scale = 12 urn. 

and found that the eggs have no surface pits. 

Furthermore, Kuramochi et al. (unpublished 

data, 1999) found arrangement differences in the 

mitochondrial DNA of adult Thai specimens of 

G. spinigerum and the adult Mexican gnathostome. 

Although our SEM observations did not 

show typical differences in larval stages between 

these 2 species, we think that this Mexican 

gnathostome may be a separate species. 

Such designation must, however, await a more 

detailed analysis. 

Gnathostoma spinigerum was reported in Ecuador 

in 1981 (Ollague et al., 1981), yet their 

description remains unclear. The adult of this 

species should be re-examined more precisely. 

We assume that this human-infecting Latin 

American gnathostome may be the same as that 

of G. binucleatum. 

Figures 7, 8. Cross-sections of the larval intestines of Gnathostoma cf. binucleatum. 7. A cross section 

of the Temazcal larva. Multiple nuclei are evident in 1 cell. Scale = 20 |xm. 8. A cross-section of the 

Sinaloan larva. Arrows indicate the cells with 5 nuclei each. Scale = 20 urn. 



Table 1. Morphological dimensions of the advanced third-stage larvae of species of Gnathostoma (data 

obtained by SEM).* 

No. of booklets 

on head bulb 

Gnathostoma 

species: 

Locality I II III IV 

G. binucleatunr. 

Temazcal 40 43 46 49 

Present specimens: 

Temazcal 

Sinaloa 

G. spinigerum: 

Thailand 

G. doloresi: 

Japan 

G. hispidum: 

China 

G. procyonis: 

U.S.A. 

39 

40 

40 

39 

40 

33 

44 

44 

43 

39 

41 

* ND = not described. 

37 

46 

45 

46 

36 

47 

41 

50 

49 

50 

38 

48 

45 

Location 

between transverse 

striations of transverse 

striations (No. 

Cervical Excretory of larvae 

papillae pore examined) References 

12th-13th about 30th 260(12) Lamothe-Argumedo et al. (1989) 

12th- 15th 

12th- 15th 

llth- 16th 

15th-19th 

10th- 13th 

ND* 


The authors would like to thank Professor 

Isao Tada, Department of Microbiology (Parasitology), 

Graduate School of Medical Sciences, 

Kyushu University, for reviewing the manuscript. 

Thanks are due to Associate Professor 

Brian T. Quinn, Division of Applied Linguistics, 

Kyushu University, for final revision of the English. 

This work was supported by a Grant-in- 

Aid for International Scientific Research (Field 

Research No. 08041187) from the Ministry of 

Education, Science, Sports, and Culture, Japan. 


Akahane, H., R. Lamothe-Argumedo, J. M. Martinez-Cruz, 

D. Osorio-Sarabia, and L. Garcia- 

Prieto. 1994. A morphological observation of the 

advanced third-stage larvae of Mexican Gnathostoma. 

Japanese Journal of Parasitology 43:18-22. 

Almeyda-Artigas, R. J. 1991. Hallazgo de Gnathostoma 

binucleatum n. sp. (Nematoda: Spirurida) en 

felinos silvestres y el papel de peces dulceacuicolas 

y oligohalinos como vectores de la gnathostomiasis 

humana en la cuenca baja del rio Papaloapan, 

Oaxaca-Veracruz, Mexico. Anales del Institute 

de Ciencias del Mar y Limnologia, Universidad 

Nacional Autonoma de Mexico 18:137- 

155. 

Ash, L. R. 1962. Development of Gnathostoma procyonis 

Chandler, 1942, in the first and second intermediate 

hosts. Journal of Parasitolology 48: 

298-305. 

24th-28th 

23rd-24th 

22nd-28th 

25th-28th 

19th-20th 

ND 

227-225 (10) 

228-256 (3) 

225-256 (8) 

176-211 (10) 

202-216 (10) 

ND (15) 


This report 

This report 

Koga et al. (1994) 

Koga and Ishii (1987) 

Koga et al. (1988) 

Ash (1962) 

Camacho, S. P. D., M. Zazueta-Ramos, E. Ponce- 

Torrecillas, I. Osuna-Ramirez, R. Castro-Velazquez, 

A. Elores-Gaxiola, J. Baquera-Heredia, 

K. Willms, H. Akahane, K. Ogata, and Y. 

Nawa. 1998. Clinical manifestations and immunodiagnosis 

of gnathostomiasis in Culiacan, Mexico. 

American Journal of Tropical Medicine and 

Hygiene 59:908-915. 

Koga, M., H. Akahane, Y. Ishii, and S. Kojima. 

1994. External morphology of the advanced thirdstage 

larvae of Gnathostoma spinigerum: a scanning 

electron microscopy. Japanese Journal of 


, , K. Ogata, R. Lamothe-Argumedo, 

D. Osorio-Sarabia, L. Garcia-Prieto, and J. M. 

Martinez-Cruz. 1999. Adult Gnathostoma cf. 

binucleatum obtained from dogs experimentally 

infected with larvae as an etiological agent in 

Mexican gnathostomiasis: external morphology. 


66:41-46. 

, J. Ishibashi, Y. Ishii, and T. Nishimura. 

1988. Scanning electron microscopic comparisons 

among the early and advanced third-stage larvae 

of Gnathostoma hispidum and the gnathostome 

larvae obtained from loaches. Japanese Journal of 


and Y. Ishii. 1987. Surface morphology of 

the advanced third-stage larvae of Gnathostoma 

doloresi: an electron microscopic study. Japanese 


Lamothe-Argumedo, R., R. L. Medina-Vences, S. 

Lopez-Jimenez, and L. Garcia-Prieto. 1989. 

Hallazgo de la forma infectiva de Gnathostoma 

sp., en peces de Temazcal, Oaxaca, Mexico. An-

ales del Institute de Biologfa de la Universidad 

Nacional Autonoma de Mexico, Series Zoologia 

60:311-320. 

Miyazaki, I. 1960. On the genus Gnathostorna and 

human gnathostomiasis, with special reference to 

Japan. Experimental Parasitology 9:338-370. 

Ogata, K., Y. Nawa, H. Akahane, S. P. Camacho, 

R. Lamothe-Argumedo, and A. Cruz-Reyes. 

1998. Gnathostomiasis in Mexico. American Jour- 


nal of Tropical Medicine and Hygiene 58:316- 

318. 

Ollague, W., J. Ollague, A. Guevara de Veliz, S. 

Penaherrera, C. Von Buchwald, and J. Mancheno. 

1981. Gnathostomiasis humana en el Ecuador 

(larva migrans profunda). Nuestra Medicina 

6:9-23. 

Pelaez, D., and R. Perez-Reyes. 1970. Gnathostomiasis 

humana en America. Revista Latino-Americana 

de Microbiologia 12:83-91. 

Report on the Brayton H. Ransom Memorial Trust Fund 

The Brayton H. Ransom Memorial Trust Fund was established in 1936 to "Encourage and 

promote the study and advancement of the Science of Parasitology and related sciences." Income 

from the Trust currently provides token support of Comparative Parasitology and limited support 

for publication of meritorious manuscripts by authors lacking institutional or other backing. Donations 

or memorial contributions may be directed to the Secretary-Treasurer. Information about 

the Trust may be found in the following articles: Proceedings of the Helminthological Society of 

Washington (1936) 3:84-87; (1983) 50:200-204 and Journal of the Helminthological Society of 

Washington (1993) 60:144-150. 

Financial Report for 1999 

Balance on hand, January 1, 1999 $23,289.69 

Receipts: $1,286.37 

Contributions from Members of the Helminthological Society of Washington: 

1998 $84.00 

1999 (in memory of Dr. Francis G. Tromba) $50.00 

Interest received in 1999 $1,152.37 

Disbursements ($117.00) 

Grant to the Helminthological Society of Washington for 1999 ($50.00) 

Membership in the American Association for Zoological Nomenclature ($50.00) 

Service charges on accounts ($17.00) 

On hand, December 31, 1999 $24,459.69 

Robin N. Huettel 

Secretary-Treasurer 

USDA, CSREES, Agri-Box 2220, 

1400 Independence Avenue 

Washington, DC 20250 

Trustees of the Brayton H. Ransom Memorial Trust Fund 

J. Ralph Lichtenfels, President 

Robin N. Huettel, Secretary-Treasurer 

A. Morgan Golden 

Harley G. Sheffield 

Nancy D. Pacheco 




Research Note 

Helminth Parasites in Six Species of Shorebirds (Charadrii) from 

Bristol Bay, Alaska, U.S.A. 

ALBERT G. CANARis1-3 AND JOHN M. KiNSELLA2 

1 University of Texas at El Paso, P.O. Box 717, Hamilton, Montana 59840, U.S.A. (e-mail: 

acanaris@bitterroot.net), and 

2 Department of Pathobiology, College of Veterinary Medicine, University of Florida, Gainesville, Florida 

32611, U.S.A. (e-mail: wormdwb@aol.com) 

ABSTRACT: Nineteen species of gastrointestinal helminth 

parasites were recovered from 6 species of charadriid 

shorebirds (Aves: Charadriiformes) from Bristol 

Bay, Alaska: the surfbird Aphriza virgata, the western 

sandpiper Calidris mauri, the rock sandpiper Calidris 

ptilocnemis, the whimbrel Numenius phaeopus, the 

northern phalarope Phalaropus lobatus, and the blackbellied 

plover Pluvialis squatarola. Cestode species 

were dominant (N = 14), followed by trematode species 

(N = 4) and an acanthocephalan (N = 1). No 

nematodes were observed. Only the cestode Aploparaksis 

daviesi infected more than 1 species of host, the 

surfbird Aphriza virgata and the northern phalarope 

Phalaropus lobatus. All species of helminths have 

been reported from birds on other continents, particularly 

Eurasia. 

KEY WORDS: Helminth parasites, Aves, Charadrii, 

surfbird, Aphriza virgata, western sandpiper, Calidris 

mauri, rock sandpiper, Calidris ptilocnemis, whimbrel, 

Numenius phaeopus, northern phalarope, Phalaropus 

lobatus, black-bellied plover, Pluvialis squatarola, littoral 

zone, Bristol Bay, Alaska, U.S.A. 

Bristol Bay, Alaska, and adjacent tundra provide 

significant habitat to nesting and postbreeding 

shorebirds (Suborder Charadrii) (Gill and 

Handel, 1981). The bay is shallow and shorebirds 

are easily observed foraging on extensive 

sand—mud habitats exposed at low tide. Observations 

and counts (A.G.C.) near the mouth of 

the Egegik River, Bristol Bay, over the summers 

of 1991, 1993, 1996, and 1997 indicated arrival 

and utilization of the littoral zone by large numbers 

of postbreeding shorebirds. There is no 

published information on helminth parasites of 

shorebirds from this important area and very little 

from the other localities on Bristol Bay. 

Schmidt and Neiland (1968) recorded 19 species 

of cestodes and trematodes from 5 species of 

shorebirds collected on Kvichak Bay, Bristol 


250 


Bay. Deblock and Rausch (1968) reported Aploparaksis 

leonovi Spasskii, 1962, and Aploparakris 

stricta Spasskii, 1962, from the western 

sandpiper Calidris mauri Cabanis, 1857, from 

the watersheds of Bristol Bay. Van Cleave and 

Rausch (1950), in the only previous report of 

helminths from the surfbird Aphriza virgata 

Gmelin, 1789, recovered 2 immature specimens 

of the acanthocephalan Arythmorhynchus comptus 

Van Cleave and Rausch, 1950, near Juneau, 

Alaska. We examined these 2 host species and 

4 additional species from Bristol Bay and herein 

report our findings. 

Birds were obtained from a 6.0-km-long section 

of littoral zone just north of the mouth of 

the Egegik River, Bristol Bay, Alaska, between 

Bishop Creek (58°14'31"N, 157°29'43"W) and 

Big Creek (58°17'01"N, 157°32'25"W). All species 

of hosts were common except the surfbird 

and rock sandpiper Calidris ptilocnemis Coues, 

1873, which were rare. Five surfbirds A. virgata, 

5 western sandpipers C. mauri, 4 whimbrels Numenius 

phaeopus Linnaeus, 1758, and 1 C. ptilocnemis, 

were collected in 1996, and 5 A. virgata, 

10 black-bellied plovers Pluvialus squatarola 

Linnaeus, 1758, 10 northern phalaropes 

Phalaropus lobatus Linnaeus, 1758, in 1997. 

Birds were collected with a shotgun between 

July 2 and July 23 and examined within 6 hr. 

All internal organs were examined. The koilon 

of the ventriculus was removed, and both the 

ventriculus and proventriculus tissues were 

teased apart. Skin and blood were not examined 

for parasites. 

Acanthocephalans, cestodes, and trematodes 

were fixed and preserved in alcohol-formalinacetic 

acid, stained in Ehrlich's hematoxylin, 

cleared in methyl salicylate, and mounted in 

Canada balsam. Voucher specimens were deposited 

in the United States National Helminthol-

CANARIS AND KINSELLA—RESEARCH NOTES 251 

Table 1. Helminth parasites of 6 species of shorebirds (Charadrii) from Bristol Bay, Alaska, U.S.A. 

Number 

Host and parasite infected 

Black-bellied plover, Pluvialis squatarola Linnaeus, 1758 (N = 

Anornotacnia ericetorum (Krabbe, 1869) 

Aploparaksis diagonalis Spasskii and Bobova, 1961 

Liga brevis (Linstow, 1884) 

Proterogynotaenia variabilix Belopol'skya, 1954 

Schistocephalus solidiix (Mueller, 1776) 

Wardium squatarolae Kornyushin, 1970 

Echinoparyphium reciirvatum (Linstow, 1873) 

Polymorphic magnus (Southwell, 1927) 

Aploparaksis daviesi Deblock and Rausch, 1968 

Echinocotyle tennis Clerc, 1906 

Trichocephaloides megalocephala (Krabbe, 1869) 

Plagiorchis morosovi Sobolev, 1946 

Surfbird Aphriza virgata Gmelin, 1789 (N = 10) 

Aploparaksis diagonalis Spasskii and Bobova, 1961 

Dictymetra nymphaea (Schrank, 1790) 

Lacunovermes sp. Ching, 1965 

Western sandpiper Calidris mauri Cabanis, 1857 (A' = 5) 

Aploparaksis leonovi Spasskii, 1961 

Kowalewskiella cingulifera (Krabbe, 1869) 

Whimbrel Numenius phaeopus Linnaeus, 1758 (A' = 4) 

Brachylaima fuscatum (Rudolphi, 1819) 

Rock sandpiper Calidris ptilocnemis Coues, 1873 (N = I) 

Wardium ampliitricha (Rudolphi, 1819) 

ogical Collection, Beltsville, Maryland, U.S.A., 

accession numbers 89038-89055. 

Nineteen species of helminths were recovered 

from the 6 species of hosts. Cestode species 

were dominant (TV = 14), followed by trematode 

species (TV = 4) and an acanthocephalan (TV = 

1). No nematodes were observed. Each of the 6 

species of host was parasitized by at least 1 helminth 

species. Only the cestode Aploparaksis 

daviesi Deblock and Rausch, 1968, infected 

more than 1 species of host—the surfbird A. virgata 

and northern phalarope P. lobatus (Table 

1). All are new host records for Alaska. All species 

of helminths were previously reported from 

birds on other continents, particularly from Eurasia 

(Table 1). 

Generally, trematode species are dominant in 

marine habitats, and cestodes are dominant in 

freshwater environments (Bush, 1990; Canaris 

and Kinsella, 1998). In both our study and that 

10) 

7 

1 

5 

5 

1 

4 

6 

1 

t)\ 

1 

2 

1 

3 

6 

8 

1 

2 

2 

1 

1 

Mean 

intensity 

14.9 

0.1 

1 1.7 

129.5 

0.1 

4.6 

62.0 

0.2 

0.2 

0.4 

0.1 

0.7 

10.5 

22.4 

89.3 

7.4 

2.6 

0.25 

2.0 

Range 

1-72 

— 

6-55 

1-1,155 

— 

1-34 

1-468 

— 

1-3 

— 

1-3 

1-56 

2-66 

— 

1-36 

1-12 

— 

— 

Other localities 

Europe 

Russia 

Eurasia 

Russia 

Eurasia, North America 

Eurasia 

Cosmopolitan 

Russia 

Alaska, U.S.A. 

Russia 

Eurasia 

Russia 

Africa, Eurasia 

Eurasia 

British Columbia, 

Canada 

Russia 

Eurasia, Guadeloupe 

Australia, Europe, 

North America, 

Russia 

Europe, North 

America, Russia 

by Schmidt and Neiland (1968), cestode species 

were dominant (72% and 79%, respectively). 

This may reflect the hosts' recent association 

with the terrestrial (freshwater) nesting area, an 

absence of proper intermediate molluscan hosts 

for trematodes in Bristol Bay, or both. Also, it 

may reflect early summer season examination of 

hosts in both studies. In this study, the bulk of 

the trematodes was obtained later in July. Trematodes 

obtained earlier in July were often immature 

or recently mature, as indicated by the 

presence of small numbers of eggs and lack of 

pigmentation of the eggshell. Small numbers or 

absence of species of acanthocephalans and 

nematodes in Bristol Bay have also been found 

in studies done in Canada on 3 species of shorebirds: 

the long-billed curlew Numenius americanus 

Bechstein, 1812 (Goater and Bush, 1988); 

the American avocet Recurvirostra americana 

Gmelin, 1789 (Edwards and Bush, 1989); and 



the whimbrel Catoptrophorus semipalmatus 

Gmelin, 1789 (Bush, 1990). The absence of 

nematodes from the upper digestive tract in this 

study is also somewhat puzzling. Anderson et al. 

(1996) reviewed records for these nematodes in 

shorebirds from North and South America. As 

in the present study, they found no species of 

the genus Skrjabinoclava Sobolev, 1943, or ventricular 

nematodes in 15 surfbirds (A. virgatd) 

or 44 northern phalaropes (P. lobatus). However, 

species of Skrjabinoclava were common in 83 

black-bellied plovers (P. squatarold), 93 western 

sandpipers (C. mauri), and 8 whimbrels (N. 

phaeopus). It is possible that the intermediate 

hosts of these nematodes are absent in Bristol 

Bay or that they were not detected in our relatively 

small sample sizes. Part of the explanation, 

at least in our study, was that skin and 

blood were not examined for nematodes. 

Most helminths reported herein are not host 

specific (Baer, 1962; Deblock and Rausch, 1968; 

Schmidt and Neiland, 1968). Low overlap in 

helminth species may be attributed to small sample 

size, but it may also be influenced by specialized 

feeding habits of shorebirds (Storer, 

1971) prior to arrival from the nesting grounds 

and in Bristol Bay. Natural sorting out of shorebird 

species into preferred feeding habitats as 

the tide recedes, as reported by Ehrlich et al. 

(1988), is easily observed (A.G.C.) on the sandmud 

habitats of Bristol Bay. 

All species of shorebirds nesting at Bristol 

Bay migrate to more distant southern wintering 

localities, many to other continents. We expect 

that further studies will reveal more relationships 

of helminth species of shorebirds on Bristol 

Bay to those from distant localities. 

The littoral zone of Bristol Bay is an important 

postbreeding locality for many species of 

shorebirds. Studies of helminth communities 

need to be extended, in both time and location, 

to understand the dynamics of the helminth 

communities and their interactions among the 

many species of shorebirds during the long days 

but relatively short summer. 

We wish to thank Hilda Ching for her opinion 

on Lacunovermis sp. and Jerry Solie and Jerry 


Lang for their very able assistance, support, and 

long friendship with A.G.C. 


Anderson, R. C., P. L. Wong, and C. M. Bartlett. 

1996. The acuarioid and habronematoid nematodes 

(Acuarioidea, Habronematoidea) of the upper 

digestive tract of waders. A review of observations 

on their host and geographic distributions 

and transmissions in marine environments. Parasite 

4:303-312. 

Baer, J. G. 1962. Cestoda. Pages 1-63 in Zoology of 

Iceland. Vol. 2, Part 12. Ejnat Munksgaard, Copenhagen. 

Bush, A. O. 1990. Helminth communities in avian 

hosts: determinants of pattern. Pages 197-232 in 

G. W. Esch, A. O. Bush, and J. M. Aho, eds. Parasite 

Communities: Patterns and Processes. Chapman 

and Hall, New York. 

Canaris, A. G., and J. M. Kinsella. 1998. Helminth 

parasites in four species of shorebirds (Charadriidae) 

on King Island, Tasmania. Papers and Proceedings 

of the Royal Society of Tasmania 132: 

49-57. 

Deblock, S., and R. L. Rausch. 1968. Dix Aploparaksis 

(Cestoda) de Charadriiformes d'Alaska et 

quelques autres d'ailleurs. Annales de Parasitologie 

Humaine et Comparee 43:429-448. 

Edwards, D. D., and A. O. Bush. 1989. Helminth 

communities in avocets: importance of the compound 

community. Journal of Parasitology 75: 

225-238. 

Ehrlich, P. R., D. S. Dobkin, and D. Wheye. 1988. 

The Birder's Handbook: A Field Guide to the Natural 

History of North American Birds. Simon & 

Schuster Inc., New York. 785 pp. 

Gill, Jr., R. E., and C. M. Handel. 1981. Shorebirds 

of the eastern Bering Sea. Pages 719-738 in W. 

Hood and J. A. Calder, eds. The Eastern Bering 

Sea Shelf: Oceanography and Resources. Vol. 2D. 

University of Washington Press, Seattle. 

Goater, C. P., and A. O Bush. 1988. Intestinal helminth 

communities in long-billed curlews: the importance 

of cogeneric host-specialists. Holarctic 

Ecology 11:40-145. 

Schmidt, G. D., and K. A. Neiland. 1968. Hymenolepis 

(Hym.) deblocki sp. n., and records of other 

helminths from charadriiform birds. Canadian 


Storer, R. W. 1971. Adaptive radiation of birds. Pages 

149-188 in D. S. Earner, J. R. King, and K. C. 

Parkes, eds. Avian Biology. Vol. 1. Academic 

Press, New York. 

Van Cleave, H. J., and R. Rausch. 1950. A new species 

of the acanthocephalan genus Aiythmorhynchus 

from sandpipers of Alaska. Journal of Parasitology 

36:278-283.



Research Note 

RESEARCH NOTES 253 

Colobomatus embiotocae (Copepoda: Philichthyidae) from Shiner 

Perch, Cymatogaster aggregata (Osteichthyes: Embiotocidae) in 

Canadian Waters 

SHELLEY F. JEPPS AND TIMOTHY M. GOATER' 

Biology Department, Malaspina University-College, Nanaimo, British Columbia, Canada V9R 5S5 (e-mail: 

goatert@mala.bc.ca) 

ABSTRACT: During an examination of the parasitic 

crustacean fauna of shiner perch, Cymatogaster aggregata 

(Embiotocidae) from eastern Vancouver Island in 

Nanaimo, British Columbia, Canada, the copepod, Colobomatus 

embiotocae Noble, Collard, and Wilkes, 

1969 (Philichthyidae), was noted in the sensory ducts 

of the preopercular cephalic canals. Prevalence and 

mean intensity of C. embiotocae were 59.2% and 1.36 

± 0.57, respectively. This parasite was also recovered 

from 68.4% (mean intensity = 1.62 ± 0.65) of shiner 

perch sampled near Bamfield Marine Station on the 

western coast of Vancouver Island. The high prevalence 

of C. embiotocae probably reflects increased 

transmission resulting from the aggregation behavior 

of the fish host. These results establish a range extension 

for C. embiotocae in C. aggregata to include Canadian 

Pacific waters. 

KEY WORDS: Colobomatus embiotocae, Copepoda, 

shiner perch, Cymatogaster aggregata, British Columbia, 

Canada. 

Members of the poecilostome family Philichthyidae 

are endoparasitic copepods that occupy 

the subcutaneous spaces associated with 

the sensory canals of the skull bones and lateral 

line of marine fishes (Kabata, 1979). They are 

highly specialized parasitic copepods, with pronounced 

sexual dimorphism and females exhibiting 

reduced organs of attachment, reduced appendages, 

and bizarre morphological processes 

projecting from their bodies. 

The richest genus of this family, Colobomatus, 

is recorded from a diversity of marine teleosts 

and elasmobranchs (Kabata, 1979; West, 

1992). Colobomatus embiotocae Noble, Collard, 

and Wilkes, 1969, was first described from shiner 

perch, Cymatogaster aggregata Gibbons, 

1854, and was found infecting several other spe- 


cies of embiotocid fishes in California and 

Oregon in the United States and in Mexico (Noble 

et al., 1969). Samples were not collected 

from Canadian waters, though the range of C. 

aggregata, among the most widely distributed 

embiotocid fish species, extends from Port 

Wrangel, Alaska, U.S.A., to Quintin Bay, Baja 

California, Mexico (Odenweller, 1975). Arai et 

al. (1988) did not find C. embiotocae during 

their study of metazoan parasites of C. aggregata 

from British Columbia. To date, the only 

species of Colobomatus recorded from Canadian 

waters is Colobomatus kyphosus Sekerak, 1970, 

from Sebastodes alutus Gilbert, 1890, and several 

species of Sebastes (Sekerak, 1970; Sekerak 

and Arai, 1977; Kabata, 1988). 

Females and males of C, embiotocae have 11 

body segments; in the female the fourth and fifth 

are fused. The average length for females and 

males is approximately 3.7 mm and 1.2 mm, respectively 

(Noble et al., 1969). Diagnostic morphological 

features distinguishing the female 

parasite from other species of Colobomatus include 

the caudal furcae with a spine on their 

inside lateral surfaces, the egg-laying apparatus 

with a bulbous structure equipped with a flagellate 

seta, and 3 eyes arranged in a compact cluster. 

Males of C. embiotocae are distinguished on 

the basis of their 6-segmented first antennae and 

1-segmented mandibles (Noble et al., 1969). 

During an investigation of the parasitic crustacean 

fauna of C. aggregata from Piper's Lagoon, 

Nanaimo, British Columbia, males and females 

of C. embiotocae were noticed infecting 

the sensory canals of the skull. A total of 76 C. 

aggregata was seined from the littoral region 

during March 1996, returned to the laboratory, 

and killed in concentrated anesthetic (MS-222), 

and their cephalic sensory canals and lateral 



lines were carefully examined. Live males and 

females of C. embiotocae were teased out of the 

canals with fine needles. The prevalence and 

mean ± SD intensity were 59.2% and 1.36 ± 

0.57, respectively. There was no association between 

host size and copepod intensity (n = 45, 

r = 0.09, P = 0.557). Females were less prevalent 

(22.4% females vs. 48.7% males) and 

abundant (mean intensity of 1.0 ± 0 females vs. 

1.19 ± 0.46 males) than males. There was no 

significant difference in the intensity of males 

compared with the intensity of females in infected 

hosts (F,,54 = 2.99, P = 0.09). 

Colobomatus embiotocae were also present in 

C. aggregata caught in a trawl (March 1996) in 

Trevor Channel on the western coast of Vancouver 

Island near the Bamfield Marine Station, 

British Columbia. Nineteen fish were necropsied 

for the presence of C. embiotocae in the cephalic 

canals. The prevalence was 68.4% and the mean 

intensity was 1.62 ± 0.65. The aggregating behavior 

characteristic of this fish species may be 

one factor explaining the high prevalence of this 

parasite, because host aggregation likely increases 

contact with C. embiotocae larvae. 

Only 1 of the 95 fish examined from both localities 

had 2 female C. embiotocae sharing the 

same canal. The presence of a gravid female 

within the cephalic sensory canals may prevent 

or inhibit other females from establishing themselves 

within such a space-constrained microhabitat. 

The 2 females were found aligned head 

to furca in the left preopercular canal. All females 

were recovered from either the left or 

right preopercular canals. Males were found in 

all of the skull's sensory canals. Only 1 male 

was recovered from the lateral line, and several 

males were observed exiting the fish via the 

pores associated with the sensory canals. 

These observations establish the first definitive 

record of C. embiotocae in Canadian eastern 

Pacific waters and add another species to the diverse 

list of shiner perch parasites in Canada 

(Margolis and Arthur, 1979; McDonald and 

Margolis, 1995). Very little is known about the 

population biology of philichthyid copepods, 

probably because they are endoparasites inhabiting 

a unique and seldom studied microhabitat 

(Kabata, 1988) and are mostly found in fish species 

that are of limited commercial importance 

(West, 1992). We urge other investigators to include 

the sensory canals and lateral line system 


of marine fish as sites to routinely examine for 

these copepods. The site and host specificity of 

these unusual parasites might inspire experimental 

and field-based studies examining how seasonality 

and aspects of the fish host's behavior 

and ecology interact to influence the parasite's 

transmission dynamics. 

Voucher specimens have been deposited in 

the United States National Parasite Collection, 

Bethesda, Maryland, U.S.A. (USNPC accession 

No. 87634). We thank Jason Lewis for assisting 

with fish collections and Cameron Weighill for 

necropsy assistance. The manuscript benefited 

from constructive comments by Bob Kabata and 

Cam Goater. 


Arai, H. P., Z. Kabata, and D. Noakes. 1988. Studies 

on seasonal changes and latitudinal differences in 

the metazoan fauna of the shiner perch, Cvmatogaster 

aggregata, along the west coast of North 

America. Canadian Journal of Zoology 66:1514- 

1517. 

Kabata, Z. 1979. Parasitic Copepoda of British Fishes. 

The Ray Society, London, 152:1-468. 

. 1988. Copepoda and Branchiura. Pages 3-123 

in L. Margolis and Z. Kabata, eds. Guide to the 

Parasites of Fishes of Canada. Part II. Crustacea. 

Canadian Special Publication of Fisheries and 

Aquatic Sciences No. 101, Department of Fisheries 

and Oceans, Ottawa, Canada. 184 pp. 

Margolis, L., and J. R. Arthur. 1979. Synopsis of 

the parasites of fishes of Canada. Bulletin of the 

Fisheries Research Board of Canada 199:1—269. 

McDonald, T. E., and L. Margolis. 1995. Synopsis 

of the Parasites of Fishes of Canada: Supplement 

(1978-1993). Canadian Special Publication of 

Fisheries and Aquatic Sciences, Department of 

Fisheries and Oceans, Ottawa, Canada 122:1—265. 

Noble, E. R., S. B. Collard, and S. N. Wilkes. 1969. 

A new philichthyid copepod parasitic in the mucous 

canals of surfperches (Embiotocidae). Journal 


Odenweller, D. B. 1975. The life history of the shiner 

surfperch Cymatogaster aggregata Gibbons, in 

Anaheim Bay, California. Pages 107-115 in E. D. 

Lane and C. W. Hill, eds. The Marine Resources 

of Anaheim Bay. State of California, The Resources 

Agency, Department of Fish and Game, 

Fish Bulletin 165. 195 pp. 

Sekerak, A. D. 1970. Parasitic copepods of Sehastodes 

alutus, including Chondracanthus triventricosus 

sp. nov. and Colobomatus kyphosus sp. nov. 

Journal of the Fisheries Research Board of Canada 

27:1943-1960. 

, and H. P. Arai. 1977. Some metazoan parasites 

of rockfishes of the genus Sebastes from the 

northeastern Pacific Ocean. Syesis 10:139-144. 

West, G. A. 1992. Eleven new Colobomatus species 

(Copepoda: Philichthyidae) from marine fishes. 

Systematic Parasitology 23:81 — 133.


67(2), 2()(X) pp. 255-258 

Research Note 

Parasites of the Green Treefrog, Hyla cinerea, from Orange Lake, 

Alachua County, Florida, U.S.A. 

TARA L. CREEL,1'4 GARRY W. FOSTER,2 AND DONALD J. FORRESTER^ 

Department of Pathobiology, College of Veterinary Medicine, University of Florida, Gainesville, Florida 

32611, U.S.A. (e-mails: ' tlc@ufl.edu; 2 FosterG@mail.vetmed.ufl.edu; 3 ForresterD@mail.vetmed.ufl.edu) 

ABSTRACT: Four species of parasites (1 trematode, 2 

nematodes, and 1 protozoan) were identified from 60 

green treefrogs, Hyla cinerea (Schneider), collected 

in north-central Florida, U.S.A. The most prevalent 

parasites were the nematode Cosmocercella haberi 

(Steiner) Baker and Adamson (23%) and the protozoan 

Opalina sp. Purkinje and Valentin (47%). The 

trematode, Clinostomum attemiatum Cort, had a prevalence 

of 2%, and the other nematode, Rhabdias sp. 

Stiles and Hassall, had a prevalence of 5%. Seven 

females and seven males were infected with C. haberi. 

The prevalence and intensity of C. haberi were 

correlated positively with host size (wet weight and 

snout-vent length). There was no statistically significant 

difference between gender and intensity of C. 

haberi infection. Fourteen females and 14 males were 

infected with Opalina sp. The prevalence of Opalina 

sp. was correlated negatively with host size. Both C. 

haberi and Opalina sp. have been reported previously 

from H. cinerea. The green treefrog represents a new 

host record for C. attemiatum and Rhabdias sp. 

KEY WORDS: Hyla cinerea, Hylidae, green treefrog, 

helminths, Trematoda, Clinostomum attemiatum, Nematoda, 

Cosmocercella haberi, Rhabdias sp., Protozoa, 

Opalina sp., prevalence, intensity, Florida, U.S.A. 

The green treefrog, Hyla cinerea (Schneider, 

1799), is a small, bright green, yellow, or greenish-gray 

treefrog with a sharply defined light 

stripe along the upper jaw and side of the body. 

Its North American range extends from Delaware 

south to the Florida Keys, west to Texas, 

and north to Illinois. Hyla cinerea is found predominantly 

near permanent water. In the southern 

parts of its range it breeds from March to 

October (Behler and King, 1997) and is probably 

the most abundant species of treefrog in the 

Gainesville region of north-central Florida (Kilby, 

1945). 


255 

Many parasites have been reported from hylid 

frogs in the United States and Canada. Walton 

(1946) listed primarily nematodes, trematodes, 

and protozoans as being parasitic in H. cinerea. 

Esch and Fernandez (1993) suggested factors 

that may influence parasite populations. Two of 

these included gender and host age or, as may 

be inferred for some animals, host size. To our 

knowledge there is no standard aging technique 

for treefrogs; however, Koller and Gaudin 

(1977) stated that "larger (hence older) frogs" 

usually have a greater species diversity and 

greater intensity of infections than "smaller, 

younger individuals." It was assumed for this 

study that host size is a rough indicator of host 

age. 

We are not aware of any comprehensive 

studies on the parasites of green treefrogs in 

Florida. The purpose of this study was to examine 

the parasites of H. cinerea in north-central 

Florida, to determine the prevalence and 

intensity of parasitic infections, and to determine 

whether relationships exist between gender, 

wet weight, snout-vent length, and parasitic 

infections. 

Sixty green treefrogs were collected from 

Orange Lake (29°27'20"N 082°10'20"W), about 

32 km southeast of Gainesville, Florida, U.S.A. 

Treefrogs were collected from a small stand of 

oak trees at the edge of the lake using the PVC 

pipe technique described by Boughton (1997). 

The PVC pipes were checked twice a week, during 

the day. Thirty treefrogs were collected from 

September to October 1998, and 30 treefrogs 

were collected from January to February 1999. 

All laboratory work was conducted at the Wildlife 

Disease Research Laboratory of the University 

of Florida's College of Veterinary Medicine. 

Treefrogs were killed with tricaine methane sulfonate 

(MS-222) following the methods of Gold- 



Table 1. Prevalence, intensity, abundance, and location of parasites in 60 green treefrogs collected from 

Orange Lake, Alachua County, Florida, U.S.A., 1998-1999. 

Prevalence 

Intensity 

Parasite species Mean Range Abundance Location* 

Trematoda 

Clinostomum attenuatum^ 

Nematoda 

Cosmocercella haberi 

Rhabdias sp.t 

Protozoa 

Opalina sp. 

23 

5 

47 

94 

3 

1-236 

2-5 

0.02 

21.6 

0.15 

SK 

CL,LI,SI,ST 

LU 

CL,LI,SI 

Location in host: CL = cloaca; LI = large intestine; LU = lungs; SI = small intestine; SK = skin; ST = stomach. 

New host record. 

berg et al. (1996) and dissected within 24 hours 

of capture. Gender, wet weight, and snout-vent 

length were recorded for each individual. The 

skin, liver, heart, lungs, esophagus, stomach, 

small intestine, large intestine, cloaca, bladder, 

and kidneys were evaluated for parasites in separate 

Petri dishes under a dissecting microscope. 

Protozoans were fixed in Zn-PVA and stained 

with Giemsa. The trematode was fixed in Roudabush's 

AFA, stained with acetocarmine, and 

mounted in neutral Canada balsam. Nematodes 

were fixed in 70% ethanol containing 10% glycerine 

and mounted in lactophenol for identification. 

Voucher specimens have been deposited 

in the United States National Parasite Collection 

(USNPC), Beltsville, Maryland, U.S.A. The 

prevalence and intensity of parasites were correlated 

with wet weights and snout-vent lengths 

of H. cinerea using Pearson product moment 

correlations. A f-test was used to determine 

whether gender was related to intensity of Cosmocercella 

haberi (Steiner, 1924) Baker and Adamson, 

1977, infections (Minitab, 1998). We did 

not conduct statistical tests on Clinostomum attenuatum 

Cort, 1913, and Rhabdias sp. Stiles 

and Hassall, 1905, because of their low prevalences. 

Terminology used follows Bush et al. 

(1997). 

Thirty-one female and 29 male green treefrogs 

were collected from Orange Lake (mean 

wet weight ±SD = 3.5g±1.5g; mean snoutvent 

length ± SD = 4.2 cm ± 0.6 cm). The 

prevalences, intensities, abundances, and locations 

of parasites are listed in Table 1. Twentytwo 

treefrogs had no parasites, 22 had only 

Opalina sp., 8 had only C. haberi, 4 had both 


C. haberi and Opalina sp., 2 had both C. haberi 

and Rhabdias sp., 1 had both Rhabdias sp. and 

Opalina sp., and 1 had both C. attenuatum and 

Opalina sp. No lesions were associated with the 

parasites. 

One green treefrog was infected with C. attenuatum 

(USNPC No. 88956) encysted under 

the skin on the back. We used 2 features to identify 

the trematode as C. attenuatum rather than 

C. complanatum, which also occurs in amphibians 

(McAllister, 1990): the body is uniform in 

width (rather than wider in the hindbody as in 

C. complanatum), and the testes and ovary are 

postequatorial (rather than medial as in C. complanatum) 

(Cort, 1913; Baer, 1933; Ukoli, 

1966). Yamaguti (1971) indicated that C. attenuatum 

is found in frogs, primarily species of the 

genera Bufo Laurenti, 1768, and Rana Linnaeus, 

1758. The definitive hosts include the great blue 

heron (Ardea herodias Linnaeus, 1758), American 

bittern (Botaurus lentiginosus Rackett, 

1813), green-backed heron (Butorides striatus 

Linnaeus, 1758), and double-crested cormorant 

(Phalacrocorax auritus Lesson, 1831). Hyla cinerea 

is a new host record for C. attenuatum. 

Fourteen green treefrogs were infected with 

C. haberi (USNPC Nos. 88959 and 88960). Cosmocercella 

haberi has been reported previously 

in H. cinerea by Steiner (1924) and Walton 

(1946). A voucher specimen of C. haberi from 

H. cinerea was collected in Arkansas and deposited 

in the USNPC by C. T. McAllister in 

1994 (USNPC No. 84259). This nematode is a 

fairly common parasite of hylids and has been 

identified in other species such as Hyla versicolor 

LeConte, 1825; Hyla arenicolor Cope,

1866; and Hyla wrightorum (Taylor, 1939) 

(Campbell, 1968; Goldberg et al., 1996). Cosrnocercella 

haberi was found in the stomach, 

small intestine, large intestine, and cloaca of H. 

cinerea. Seven females and 7 males were infected 

with the parasite. The prevalence of C. 

haberi was correlated positively with the wet 

weights (r = 0.283, P = 0.029) and snout-vent 

lengths (r = 0.268, P = 0.039) of H. cinerea. 

There was no statistically significant difference 

between gender and intensity of C. haberi infection. 

The intensity of C. haberi infection was 

correlated positively with the wet weights (r = 

0.678, P = 0.008) and snout-vent lengths (r = 

0.760, P = 0.002) of the 14 infected green treefrogs. 

Three green treefrogs were infected with 

Rhabdias sp. (USNPC No. 88958). This nematode 

was found at low intensities in the lungs. 

Rhabdias spp. are considered "cosmopolitan" in 

reptiles and amphibians (Baker, 1978). Other hylids, 

such as Hyla regilla Baird and Girard, 

1852, and Pseudacris crucifer (Wied-Neuwied, 

1838) have been reported having these parasites 

(Koller and Gaudin, 1977; Muzzall and Peebles, 

1991; Yoder and Coggins, 1996). There is no 

record of Rhabdias sp. from H. cinerea. 

Twenty-eight green treefrogs were infected 

with Opalina sp. Purkinje and Valentin, 1835 

(USNPC No. 88957). Opalina obtrigonoidea orbiculata 

has been reported previously in the 

green treefrog by Walton (1946). A voucher 

specimen of Opalina sp. from H. cinerea was 

collected in Arkansas and deposited in the 

USNPC by C. T. McAllister in 1994 (USNPC 

No. 84277). Opalina sp. is a common parasite 

in treefrogs (McAllister, 1991). It has also been 

reported in Hyla avivoca Viosca, 1928; Pseudacris 

clarkii Baird, 1854; and Hyla chrysoscelis 

Cope, 1880 (McAllister, 1991; McAllister et 

al., 1993; Bolek and Coggins, 1998). Opalina 

sp. was found at high intensities in the small 

intestine, large intestine, and cloaca of H. cinerea. 

Fourteen females and 14 males were infected 

with this protozoan. The prevalence of 

Opalina sp. was correlated negatively with the 

wet weights (r = -0.357, P = 0.005) and snoutvent 

lengths (r = -0.387, P = 0.002) of H. cinerea. 

Schorr et al. (1990) studied the population 

changes of Opalina spp. and found population 

declines and even loss of the parasite at 

metamorphosis of some anurans. No change, 

however, was observed in others. They attribut- 


ed the decline or loss of Opalina spp. in some 

hosts to morphological and physiological changes 

in the host at metamorphosis. The negative 

correlation of host wet weights and snout-vent 

lengths with prevalence of Opalina sp. may 

therefore be due to the loss of parasites as the 

treefrogs increase in size or age. The intensity 

of Opalina sp. infection was not determined, because 

of the large numbers of Opalina sp. per 

treefrog. 

We thank the following for their contributions 

to this work: Dr. Kathryn Sieving for assistance 

in the development of this project; Katie L. Heggemeier 

and Jeremy J. Anderson for assistance 

in the field; Dr. Mike Kinsella for aid in identifying 

parasites, especially Clinostomum attenuatum, 

and reviewing the manuscript; Dr. Charles 

H. Courtney for assistance in the statistical analysis 

of the data; and Dr. Marilyn G. Spalding for 

reviewing the manuscript. This research was 

funded in part by the University of Florida College 

of Agriculture and the Department of Wildlife 

Ecology and Conservation. This is Florida 

Agricultural Experiment Station Journal Series 

No. R-07053. 


Baer, J. G. 1933. Note sur un nouveau trematode, 

Clinostomum lophophallum sp. nov., avec quelques 

considerations generales sur la famille des 

Clinostomidae. Revue Suisse de Zoologie 40:317- 

342. 

Baker, M. R. 1978. Morphology and taxonomy of 

Rhabdias spp. (Nematoda: Rhabdiasidae) from 

reptiles and amphibians of southern Ontario. Canadian 


, and M. L. Adamson. 1977. The genus Cosmocercella 

Steiner 1924 (Nematoda: Cosmocercoidea). 

Canadian Journal of Zoology 55:1644- 

1649. 

Behler, J. L., and F. W. King. 1997. National Audubon 

Society Field Guide to North American 

Reptiles and Amphibians. Alfred A. Knopf, New 

York. 743 pp. 

Bolek, M. G., and J. R. Coggins. 1998. Endoparasites 

of Cope's gray treefrog, Hyla chrysoscelis, and 

western chorus frog, Pseudacris t. triseriata, from 

southeastern Wisconsin. Journal of the Helminthological 


Boughton, R. G. 1997. The use of PVC pipe refugia 

as a trapping technique for Hylid treefrogs. M.S. 

Thesis, University of Florida, Gainesville. 96 pp. 





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. 

Cort, W. W. 1913. Notes on the trematode genus Clinostomum. 

Transactions of the American Microscopical 

Society 32:167-182. 

Esch, G. W., and J. C. Fernandez. 1993. A Functional 

Biology of Parasitism. Chapman and Hall, 

New York. 337 pp. 

Goldberg, S. R., C. R. Bursey, E. W. A. Gergus, B. 

K. Sullivan, and Q. A. Truong. 1996. Helminths 

from three treefrogs Hyla arenicolor, Hyla wrightorum, 

and Pseudacris triseriata (Hylidae) from 

Arizona. Journal of Parasitology 82:833-835. 

Kilby, J. D. 1945. A biological analysis of the food 

and feeding habits of two frogs, Hyla cinerea and 

Rana pipiens sphenocephala. Quarterly Journal of 

the Florida Academy of Science 8:71-104. 

Roller, R. L., and A. J. Gaudin. 1977. An analysis 

of helminth infections in Bufo boreus (Amphibia: 

Bufonidae) and Hyla regilla (Amphibia: Hylidae) 

in southern California. Southwestern Naturalist 

21:503-509. 

McAllister, C. T. 1990. Metacercaria of Clinostornum 

complanatum (Rudolphi, 1814) (Trematoda: Digenea) 

in a Texas salamander, Eurycea neotenes 

(Amphibia: Caudata), with comments on C. marginatum 

(Rudolphi, 1819). Journal of the Helminthological 


. 1991. Protozoan, helminth, and arthropod parasites 

of the spotted chorus frog, Pseudacris clarkii 

(Anura: Hylidae), from north-central Texas. 


58:51-56. 

, S. E. Trauth, S. J. Upton, and D. H. Jamieson. 

1993. Endoparasites of the bird-voiced 

treefrog, Hyla avivoca (Anura: Hylidae), from Ar- 



Research Note 

kansas. Journal of the Helminthological Society of 


Minitab, Inc. 1998. Minitab Version 12 for Windows 

95. State College, Pennsylvania. 

Muzzall, P. M., and C. R. Peebles. 1991. Helminths 

of the wood frog, Rana sylvatica, and spring peeper, 

Pseudacris c. crucifer, from southern Michigan. 



Schorr, M. S., R. Altig, and W. J. Diehl. 1990. Populational 

changes of the enteric protozoans Opalina 

spp. and Nyctotherus cordiformis during the 

ontogeny of anuran tadpoles. Journal of Protozoology 

37:479-481. 

Steiner, G. 1924. Some nemas from the alimentary 

tract of the Carolina treefrog (Hyla carolinensis 

pennant) with a discussion of some general problems 

of nematology. Journal of Parasitology 11: 

1-32. 

Ukoli, F. M. A. 1966. On Clinostomum tilapiae n. sp. 

and C. phalacrocoracis Dubois, 1931 from Ghana, 

and a discussion of the systematics of the genus 

Clinostomum Leidy, 1856. Journal of Helminthology 

40:187-214. 

Walton, A. C. 1946. Parasites of the Hylidae (Amphibia—Hylinae) 

V. Anatomical Record 96:592- 

593. 

Yamaguti, S. 1971. Synopsis of the Digenetic Trematodes 

of Vertebrates, Vol. I. Keigaku Publishing 

Co., Tokyo, Japan. 1074 pp. 

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. 

Atypical Specimens of Helminth Parasites (Anoplocephala perfoliata 

and Thelazia lacrymalis} of Horses in Kentucky, U.S.A. 

HEATHER D. BAIR,' EUGENE T. LYONS,U THOMAS W. SwERCZEK,2 AND SHARON C. 

TOLLIVER1 

1 Gluck Equine Research Center and 2 Livestock Disease Diagnostic Center, Department of Veterinary Science, 

University of Kentucky, Lexington, Kentucky 40546-0099, U.S.A. (e-mail: elyonsl@pop.uky.edu) 

ABSTRACT: During a survey of internal parasites in 

horses at necropsy at a diagnostic laboratory in Kentucky, 

U.S.A., in 1998, atypical specimens of 2 species 

were found. Two specimens of the cecal tapeworm, 



Anoplocephla perfoliata, were fused at the midportion 

of each individual. One eyeworm, Thelazia lacrymalis, 

had 3 rather than the normal 2 uteri. 

KEY WORDS: atypical morphology, Cestoda, cecal 

tapeworm, Anoplocephala perfoliata, Nematoda, eyeworm, 

Thelazia lacrymalis, horses, Kentucky, U.S.A.

Several horses, all with unknown antiparasitic 

treatment, were examined at necropsy in Kentucky, 

U.S.A., in 1998 in a prevalence survey 

for various species of internal parasites. Specimens 

of 2 species were atypical. One was the 

cecal tapeworm, Anoplocephala perfoliata 

(Goeze, 1782) Blanchard, 1848. The other was 

the eyeworm, Thelazia lacrymalis (Gurlt, 1831) 

Raillet and Henry, 1910. 

The usual habitat of A. perfoliata in the horse 

is the large intestine, mainly the cecum. In past 

surveys of dead horses in Kentucky, prevalence 

of A. perfoliata was about 50-60% and no differences 

in infection with age of the horse were 

found (Benton and Lyons, 1994). Detrimental 

effects of A. perfoliata are not always evident. 

Some of the problems, mainly at the attachment 

sites of the tapeworms, are ulceration, inflammation, 

edema, and a resulting diphtheritic 

membrane (Proudman and Trees, 1999). Possible 

life-threatening effects attributed to A. perfoliata 

are intussusception, perforation, and hypertrophied 

small intestine (Proudman and 

Trees, 1999). 

Among 265 A. perfoliata found in a 29-yearold 

Thoroughbred gelding in the present study 

were 2 atypical specimens joined together at the 

midportion (Fig. 1). Possibly there had been incomplete 

separation of 2 eggs during embryogenesis. 

Alternatively, in early development, 

there may have been injury to 1 specimen, and 

the other somehow partially invaded the afflicted 

individual. The authors were unable to find any 

reference in the literature to this type of anomaly 

in A. perfoliata. However, several other types of 

abnormalities, including 1-4 extra suckers on 

the scolex and tri- and tetraradiate strobila, have 

been reported for A. perfoliata (Lyons et al., 

1997). 

Thelazia lacrymalis uses muscid flies, e.g., 

Musca autumnalis (deGeer, 1776), as intermediate 

hosts. Negative effects caused by 7". lacrymalis 

are usually limited to conjunctivitis and 

excessive lacrimation (Patton and McCracken, 

1981). In past surveys for T. lacrymalis, about 

40-50% of horses under 5-6 years of age were 

infected; older horses had much lower prevalences 

(Lyons et al., 1986). This eyeworm species 

is associated with several parts of the eyes, 

including the lacrimal glands, lacrimal ducts, 

conjunctival sac, and nictitating membrane 

gland plus ducts. Typically, females in most 

groups of nematodes have a double or bifurcate 


Figure 1. Anomalous Anoplocephala perfoliata: 

2 specimens joined in midportion, 1 smaller (S) 

than the other. Scale bar = 10.0 mm. 

reproductive system consisting of a vulva, vagina, 

and 2 uteri (Fig. 2A) and 2 ovaries. In the 

present study, 1 of 3 female T. lacrymalis recovered 

from the eyes of a yearling male Thoroughbred 

had 3 uteri (Fig. 2B). This aberration 

was observed by chance, because all female T. 

lacrymalis in the survey were examined for the 

presence of embryos with the aid of a compound 

microscope. The specimen with the 3 uteri accidentally 

ruptured at the location shown in the 

accompanying photomicrograph (Fig. 2B), 

which was taken to record the embryos. Later, 

it was realized that the presence of 3 uteri was 

not normal. No references could be found regarding 

such an anomaly in T. lacrymalis. Hyman 

(1951) mentioned that polydelphic female 

nematodes may have more than 2, and as many 

as 10 or 11, ovaries and uteri. This situation occurs 

particularly in the Physalopteridae, which 

are spirurids (Hyman, 1951). Thelazia spp., 

while also spirurids, are in a different family. 

Chandler (1924) found 3 instead of the usual 2 

ovaries and uteri in the ascarid, Ascaris lumbricoides 

(Linnaeus, 1758), and considered this 

highly unusual. 

Causes of anomalies of internal parasites are 



2A 

Figure 2. Photomicrographs of Thelazia lacrymalis. Part of vagina (V) and uterus (U). A. Normal 

individual with 2 uteri. B. Abnormal individual with 3 uteri and embryos visible in each branch. Scale 

bar = 50 u,m. 

difficult to document. It is of interest that Becklund 

(1960) recorded an association of phenothiazine 

given to sheep and morphological 

anomalies of male Haemonchus contortus (Rudolphi, 

1803) Cobb, 1898. 

This investigation was done in connection 

with a project of the Kentucky Agricultural Experiment 

Station and is published with the approval 

of the director as paper No. 99-14-120. 


Becklund, W. R. 1960. Morphological anomalies in 

male Haemonchus contortus (Rudolphi, 1803) 

Cobb, 1898 (Nematoda: Trichostrongylidae) from 

sheep. Proceedings of the Helminthological Society 


Benton, R. E., and E. T. Lyons. 1994. Survey in 

central Kentucky for prevalence of Anoplocephala 

perfoliata in horses at necropsy. Veterinary Parasitology 

55:81-86. 

Chandler, A. C. 1924. A note on Ascaris lumbricoides 


with three uteri and ovaries. Journal of Parasitology 

10:208. 

Hyman, L. H. 1951. The Invertebrates, Volume 3. 

Acanthocephala, Aschelminthes, and Entoprocta. 

McGraw-Hill Book Co., Inc., New York. 572 pp. 

Lichtenfels, J. R. 1975. Helminths of domestic equids. 

Proceedings of the Helminthological Society of 

Washington 42 (special issue): 1-92. 

Lyons, E. T., S. C. Tolliver, J. H. Drudge, T. W. 

Swerczek, and M. W. Crowe. 1986. Eyeworms 

(Thelazia lacrymalis) in one to four-year-old 

Thoroughbreds at necropsy in Kentucky (1984- 

1985). American Journal of Veterinary Research 

47:315-316. 

, , K. J. McDowell, and J. H. Drudge. 

1997. Atypical external characteristics of Anoplocephala 

perfoliata in equids in central Kentucky. 



Patton, S., and M. D. McCracken. 1981. The occurrence 

and effect of Thelazia in horses. Equine 

Practice 3:53-57. 

Proudman, C. J., and A. J. Trees. 1999. Tapeworms 

as a cause of intestinal disease in horses. Parasitology 

Today 15:156-159.



Anniversary Award 

The Helminthological Society of Washington 

FRANK W. DOUVRES 

J. Ralph Lichtenfels, right, presents the 1999 Anniversary Award to Frank W. Douvres 

Mr. President, Members and Guests, Ladies and Gentlemen, as Chair of the Awards Committee, 

I am honored to be able to present, on behalf of the Helminthological Society of Washington, the 

1999 Anniversary Award to an outstanding scientist, the world authority on the in vitro cultivation 

of nematode parasites of livestock and a friend and mentor to many in our society, Dr. Frank W. 

Douvres. 

Frank was born to immigrant parents in the Borough of Harlem in New York City, April 16, 

1927. He grew up there, speaking Greek at home, and received an outstanding education at Benjamin 

Franklin High School, where he graduated in 1943 at the age of 16, ranking third in his 

class, just behind his classmate Daniel Patrick Moynihan. Frank's classmates correctly predicted 

that Moynihan would go into politics, but they were off the mark when they predicted that Frank 

Douvres would become a Russian Commissar. This prediction was based on Frank's outspoken 

support for the Russian war effort against Germany in World War II. 

Frank completed 2 years of premed at Fordham University in December 1944. He transferred to 

the University of Maryland in January 1945, again in premed, but April 12, 1945, just before his 

eighteenth birthday, he enlisted in the navy as a hospital corpsman. On learning that Frank had 

enlisted, Germany immediately surrendered. Later, when Frank completed basic training, Japan 

surrendered! 

Frank was discharged from the Navy in 1947 and returned to the University of Maryland, where 

he switched his major to microbiology and received his B.S. degree in 1948, before reaching the 

age of 20 yr. At the University of Maryland Frank began to meet some really interesting people 

who called themselves parasitologists, so he decided to stay and work on a graduate degree. He 

was interested in ichthyology and completed his Master of Science degree at Maryland in 1951 

after completing a study program that included a survey of the parasites of fish. 

The parasitologist at Maryland was William O. Negherbon, who counted among his students 

Frank Tromba, T. Bonner Stewart, Conrad Yunker, Will Smith, and Les Costello. Professor Negerbon 

was studying rabies and he hired Frank Tromba to collect little brown bats, for which he 

paid $2 each. Douvres helped Tromba collect bats and along the way discovered a new stomach 

261 



worm, Rictularia lucifugus (Douvres, 1956) in the bats. Frank published the description of the new 

species in the Proceedings of the Helminthological Society of Washington. He went on to complete 

a Ph.D. at the University of Maryland on the microanatomy of Rictularia lucifugus under Professor 

Josh Brown in 1958. 

In 1953 Frank married Angelica "Kiki" Vlangas, whom he met in the Greek community of 

Baltimore. Typical of Frank, he told Kiki on their first date that he was going to marry her. After 

working briefly as a cook in a New York diner (and seriously considering staying in the restaurant 

business), Frank followed the example of his fellow graduate students Frank Tromba and Bonner 

Stewart and obtained a job with the U.S. Department of Agriculture. Frank was hired by Benjamin 

Schwartz, Chief of the Zoological Division of the Bureau of Animal Industry, who had obtained 

some new money for work on parasites of cattle. 

His first assignment was at Tifton, Georgia. Frank worked at Tifton with Harry Herlich, Bonner 

Stewart, and Dale Porter from 1953 until 1955 on parasites of cattle. It was at Tifton where Frank 

did his landmark work on "The Morphogenesis of the Parasitic Stages of Ostertagia ostertagi" 

the "Morphogenesis of the Parasitic Stages of Trichostrongylus axei and T. colubriformis," and 

"Keys to the Identification and Differentiation of the Immature Parasitic Stages of Gastrointestinal 

Nematodes of Cattle." These papers are standard references, still in use today. 

After transferring to Beltsville in 1955, Frank teamed up again with his old pal from graduate 

school, Frank Tromba, and with John Lucker on numerous studies on the morphogenesis and development 

of nematode parasites of cattle, sheep, and pigs. 

During the time Frank was a student at Maryland and later at Beltsville, he, like many of us, 

was fortunate to have available the advice and expertise of MayBelle Chitwood. Frank called her 

"coach." 

In 1959, Lou Diamond, who was then working at Beltsville, invited Frank to try some of his 

nematodes in Diamond's media developed for the in vitro culture of protozoa. The success that 

they had with these experiments changed the direction of Frank's research. For the next 25 years, 

Frank Douvres made breakthrough after breakthrough in successfully culturing important nematode 

parasites of large food animals in clear, cell-free media. 

In addition to Lou Diamond, Frank credits Paul Weinstein with mentoring his early in vitro 

cultivation work. Clearly, however, Frank Douvres became the recognized world authority on the 

in vitro cultivation of nematode parasites of animals. He collaborated with Frank Tromba on the 

cultivation and the description of developmental stages of parasites of swine, including Stephanunis 

dentatus and Ascaris suum, and, with John Lucker, Halsey Vegors, Don Thompson, and later Harry 

Herlich, Rob Rew and Lou Gasbarre, on parasites of cattle. 

From the late 1960s until Frank retired in 1985, he was assisted by George Malakatis, a worldclass 

technician with an international reputation of excellence, earned first in the navy with Bob 

Kuntz and Harry Hoogstraal and later at Beltsville with Frank. 

In the early 1980s Frank began a short but extremely productive collaboration with Joe Urban 

that included numerous papers, perhaps the most significant of which were (1) Douvres and Urban. 

1983. Factors contributing to the in vitro development of Ascaris suum from second-stage larvae 

to mature adults. Journal of Parasitology 69:549-558 and (2) Douvres and Urban. 1986. Development 

of Ascaris suum from in vivo—derived third-stage larvae to egg-laying adults in vitro. 

Proceedings of the Helminthological Society of Washington 53:256-262. During this period a distinguished 

visiting scientist from China worked with Frank and Joe and was a coauthor on several 

of their papers. Dr. Xu Shoutai, Chief, Shanghai Laboratory of Animal Schistosomiasis, spent a 

productive 6-month sabbatical with Frank learning his in vitro methods. 

Frank credits Dr. A. O. Foster with strong support and encouragement for the in vitro work. 

Others who worked with Frank and benefited from his expertise included Lou Gasbarre, Ray Fetterer, 

Rob Rew, and Bob Romanowski. Frank asked me to be sure to mention some of the support 

staff who made significant contributions to his research, including Ray Rew, Ken Goodson, and 

Don Thompson. 

Frank retired from the U.S. Department of Agriculture in December of 1985 and became an 

international consultant, traveling to Townsville, Australia, where he instructed Bruce Copeman's 


ANNIVERSARY AWARD 263 

laboratory on the in vitro cultivation of nematodes for several months prior to the 1986 International 

Congress of Parasitology in Brisbane. 

After the Australian trip, Frank settled into retirement and took up the role of grandfather, which 

he now plays for grandsons Christopher and Tim and daughter Nicky. Like everything else in his 

professional life, Frank plays the grandfather role with enthusiasm, a strong personal style, and a 

sense of duty and devotion. 

It was these same values that made Frank Douvres not just an outstanding scientist, but one of 

the most unforgettable personages for his colleagues and friends. At meetings, Frank could be 

counted on for a direct, to-the-point question meant to be provocative. Not everyone understood 

and appreciated this approach, but things were never dull when Frank was around. 

In retirement, Frank has continued to be active in his church and the National Association of 

Retired Federal Employees and was the local NARFE chapter president in 1995, prior to a serious 

illness from which a long recovery is now almost complete. Frank and Kiki have also generously 

supported HelmSoc through the activities of the Brayton H. Ransom Memorial Trust Fund. 

The Anniversary Award of the Helminthological Society of Washington is given either for scientific 

achievement or for service to the Society. Dr. Frank Douvres qualifies in both respects, 

having served in most of the offices of the society and on the editorial board. On behalf of the 

society, it is a great pleasure for me to present the 1999 Anniversary Award to Dr. Frank W. 

Douvres. Congratulations, Frank. 



666th Meeting: Beltsville Agricultural Research 

Center, United States Department of Agriculture, 

Beltsville, Maryland, 13 October 1999. 

President Eric Hoberg presided over the business 

meeting and the scientific session, which 

consisted of 3 presentations: Dr. Benjamin Rosenthal 

provided an overview of the phylogeography 

of deer ticks in eastern North America, 

Dr. John Carroll spoke on black-legged ticks and 

Lyme disease in Maryland, and Dr. Eric Hoberg 

provided a summary of nematode parasites of 

ruminants in the Mackenzie Mountains. New 

members included Santiago Mas-Coma (Spain), 

Eun-Taek Han (Korea), Marie-Claude Durette- 

Desset (France), Pan Cangsang (People's Republic 

of China), Richard Botzler (U.S.A.), Austin 

Maclnnis (U.S.A.), and Scott Monks (Mexico). 

667th Meeting: Sabang Restaurant, Wheaton, 

Maryland, 17 November 1999. The anniversary 

MINUTES 

Six Hundred Sixty-Sixth Through 

Six Hundred Seventieth Meeting 

J. Ralph Lichtenfels 

November 17, 1999 

dinner meeting and program were presided over 

by President Eric Hoberg. The slate of officers 

for 2000 was elected and installed by the membership 

in attendance: Dennis J. Richardson, 

president; Lynn K. Carta, vice president; Pat 

Carney, recording secretary; and Nancy Pacheco, 

corresponding secretary—treasurer. Willis A. 

Reid, Jr., and Janet W. Reid continued in office 

as editors. Dr. Ralph Lichtenfels introduced the 

recipient of the Anniversary Award, Dr. Frank 

Douvres. Dr. Douvres reviewed his research career, 

particularly his pioneering work with the in 

vitro culture of nematodes. Dr. Hoberg's final 

action as president was to turn the meeting over 

to the new president, Dr. Dennis Richardson. Dr. 

Richardson's first action was to adjourn the 

667th meeting of the society and advise the 

membership that the next meeting would be held 

at the National Museum of Natural History, 

Smithsonian Institution, Washington, DC, on 

Wednesday, 19 January 2000, at 1900 h, with 

William Moser serving as the host. 



668th Meeting: National Museum of Natural 

History, Smithsonian Institution, Washington, 

DC, 19 January 2000. President Dennis Richardson 

presided over the business meeting, 

which he summarized for the membership, and 

reminded the membership that the 669th meeting 

of the society would be held at the Johns 

Hopkins Montgomery County Center in Maryland, 

with Dr. Thomas Simpson in charge of 

making the local arrangements. He then introduced 

William Moser, who chaired the scientific 

session, which consisted of 4 papers: the first 

paper, authored by Mr. Dan Holiday and Dr. 

Dennis Richardson and presented by Mr. Holiday, 

dealt with archaeoparasitology on the Chiribaya 

Culture of southern Peru; the second, by 

Dr. Jeff Bates, provided an overview of the molecular 

phylogeny of the Adenophorea; and the 

third, by Dr. Jon Norenburg, reviewed his phylogenetic 

studies of the phylum Nemertea. The 

final speaker was Dr. Duane Hope, who provided 

an overview of the phylogenetic relationship 

between the marine nematode genera Rhabdodemania 

and Pandolaimus. New members included 

Benjamin Rosenthal (U.S.A.) and Alan 

Fedynich (U.S.A.). 

669th Meeting: Johns Hopkins Montgomery 

County Center, 22 March 2000. The business 

meeting was opened by the vice president, Lynn 

Carter, and presided over by President Dennis 

Richardson. President Richardson welcomed 

members and guests to the meeting, and a moment 

of silence was observed in memory of recently 

deceased society members James H. 

Turner, Bryce C. Walton, Richard M. Sayer, 

Francis G. Tromba, Everett L. Schiller, and Marion 

M. Farr. President Richardson then introduced 

Dr. Alan L. Scott, who chaired the scientific 

program, which consisted of 2 presentations. 

Dr. David Sullivan summarized his work 

on the formation and inhibition of heme poly- 


mers in parasites. His presentation was followed 

by Dr. Christopher V. Plowe's discussion of a 

molecular marker for chloroquine-resistant falciparum 

malaria. Following the presentations 

and questions from the members and guests, 

President Richardson thanked the speakers for 

their informative and digestible summaries of 

malaria and schistosomiasis at the molecular 

level, and he also thanked Dr. Tom Simpson, 

who arranged the meeting. Finally, he reminded 

the membership that the last meeting of the season 

would be held at the New Bolton Center, 

University of Pennsylvania, Kennett Square, together 

with the New Jersey Society of Parasitologists, 

on 6 May 2000. New members included 

Al Canaris (U.S.A.), Peter Hotez (U.S.A.), 

Nicole Havas (U.S.A.), John Janovy, Jr. 

(U.S.A.), and Alan Scott (U.S.A.). 

670th Meeting: New Bolton Center, University 

of Pennsylvania, Kennett Square, with the New 

Jersey Society of Parasitologists, 6 May 2000. 

The business meeting was presided over by 

President Richardson. Dr. Jay Farrell presided 

over the scientific meeting, which consisted of 3 

presentations. Dr. Thomas Klei discussed immunity 

to equine strongyle infections. His paper 

was followed by Dr. David Sibley's discussion 

of motility and invasion of Toxoplasma. The final 

presentation was provided by Dr. James B. 

Lok on the Dauer pathway in Caenorhabditis 

elegans as a model for regulation of infective 

larval development in parasitic nematodes. New 

members included Ian Whittington (Australia), 

M. Rocio Ruiz de Ybanez (Spain), Francisco Jimenez-Ruiz 

(U.S.A.), Glen Dappen (U.S.A.), 

Alan Kocan (U.S.A.), Aaron McCormick 

(U.S.A.), Robin LePardo (U.S.A.), Megan Collins 

(U.S.A.), Mike Barger (U.S.A.), Megan 

Ryan (U.S.A.), Tamara Cook (U.S.A.), Kashinath 

Ghosh (U.S.A.), and Richard Clopton 

(U.S.A.).


67(2), 2000 p. 265 

Aguirre-Macedo, L., 85 

Akahane, H., 244 

Almeida, S. C. de, 210 

Amin, O. R., 40, 71 

Bailey, J., 71 

Bair, H. D., 218, 258 

Benz, G. W., 190 

Boczori, K., 230 

Bolek, M. G., 202 

Bolette, D. P., 114 

Botzler, R. G., 135 

Bouamer, S., 169 

Brooks, D. R., 1 

Buck, S., 135 

Bullard, S. A., 190 

Bursey, C. R., 60, 109, 118, 129 

Canaris, A. G., 250 

Champoux, L., 26 

Cheam, H., 118 

Coady, N. R., 32 

Coggins, J. R., 202 

Creel, T. L., 255 

Dailey, M. D., 165 

Daras, M. R., 241 

Diaz-Camacho, S. P., 244 

Doi, T., 224 

Domke, W., 71 

Duclos, L. M., 197 

Durette-Desset, M.-C1., 66 

Eidelman, W. S., 71 

Elsey, R. M., 122 

Emery, M. B., 133 

Forrester, D. J., 124, 255 

Foster, G. W., 124, 255 

Fried, B., 236, 241 

Fujino, T., 236 

Fujisaki, A., 224 



AUTHOR INDEX FOR VOLUME 67 

Fukuda, K., 236 

Galicia-Guerrero, S., 129 

Garcia-Prieto, L., 92, 244 

Goater, T. M., 253 

Goldberg, S. R., 60, 109, 118, 129, 

165 

Hanelt, B., 107 

Hasegawa, H., 224 

Heckmann, R. A., 40 

Hemdal, J., 190 

Hoberg, E. P., 1 

Ichikawa, H., 236 

Janovy, J., Jr., 107 

Jepps, S. F, 253 

Jimenez-Ruiz, F. A., 145 

Joy, J. E., 133 

Kinsella, J. M., 124, 250 

Koga, M., 244 

Kritsky, D. C., 76, 145 

Ladd-Wilson, S., 135 

Lamothe-Argumedo, R., 244 

Leon-Regagnon, V., 92 

Lichtenfels, J. R., 189, 261 

Lyons, E. T., 218, 258 

Machado, P. M., 210 

Marcogliese, D. J., 26 

Martinez-Cruz, J. M., 244 

Mendoza-Franco, E., 76, 85 

Miyata, A., 224 

Moler, P. E., 124 

Morand, S., 169 

Muzzall, P. M., 181 

Nakano, T., 236 

Nguyen, V. H., 40 

Nickol, B. B., 32 

Noda, K., 244 

Olson, K. D., 218 

Osorio-Sarabia, D., 244 

Ouellet, M., 26 

Overstreet, R. M., 190 

Pavanelli, G. C., 210 

Perez-Ponce de Leon, G., 92 

Pfeifer, G., 71 

Pham, N. D., 40 

Pham, V. L., 40 

Razo-Mendivil, U., 92 

Reid, J. W., 189 

Richardson, D. J., 197 

Rodrigue, J., 26 

Rodriguez-Canul, R., 85 

Salgado-Maldonado, G., 129 

Sanchez-Alvarez, A., 92 

Santos, A., Ill, 66 

Scholz, T., 76, 85 

Scott, T. P., 122 

Sey, O., 145 

Shinohara, T., 236 

Simcik, S. R., 122 

Sisbarro, S., 241 

Smales, L. R., 51 

Spraker, T. R., 218 

Swerczek, T. W., 258 

Takemoto, R. M., 210 

Ten-ell, S. P., 124 

Tolliver, S. C., 218, 258 

Vidal-Martinez, V., 85 

Walser, C. M., 109 

Wargin, B., 230 

West, M., 122 

Williams, E. H., Jr., 190 

KEYWORD AND SUBJECT INDEX FOR VOLUME 67 

Abbreviata anomala, 109 

Abbreviata sp., 109 

Abomasal nematodes, 135 

Abundance, 26, 60, 122, 129, 181, 

202, 210, 255 

Acanthocephala, 32, 40, 60, 114, 

122, 124, 133, 181, 210, 250 

Acanthocephalorhynchoid.es cholodkowskyi, 

comb, n., 40 

Acanthocephalus dims, 181 

265 


Acanthopagrus berda, 145 

Acanthopagrus bifasciatus, 145 

Acanthopagrus latus, 145 

Acanthorhodeus fortunensis, 40 

Acari, 124


Agamidae, 109 

Alaska, U.S.A., 218, 250 

Alligator snapping turtle, 122 

Amblyomma dissimile, 124 

Ambystoma andersoni, 92 

Ambystoma dumerilii, 92 

Ambystoma lermaensis, 92 

Ambystoma mexicanum, 92 

Ambystoma tigrinum, 92 

Ameloblastella gen. n., 76 

Ameloblastella chavarriai comb. 

n., 76 

Ameloblastella mamaevi comb, n., 

76 

Ameloblastella platensis comb, n., 

76 

American toad, 202 

American white pelican, 244 

Amphibia, 26, 92, 129, 133, 92, 

165, 202, 224, 255 

Anatomy, 51 

Ancyrocephalinae, 85 

Anguilliformes, 190 

Anomotaenia ericetorum, 250 

Andrias japonicus, 224 

Angiostoma onychodactyla sp. n., 

60 

Angiostoma plethodontis, 133 

Angiostomatidae, 60 

Anisakiasis, 71 

Anoplocephala perfoliata, 258 

Anura, 92, 129, 165, 202, 224, 255 

Aphanoblastella gen. n., 76 

Aphanoblastella mastigatus comb. 

n., 76 

Aphanoblastella robustus comb. 

n., 76 

Aphanoblastella travassosi comb. 

n., 76 

Aphriza virgata, 250 

Aplectana incerta, 129 

Aploparaksis daviesi, 250 

Aploparaksis diagonalis, 250 

Aploparaksis leonovi, 250 

Aquaculture, 181, 190 

Argentina, 76 

Argyrops filamentosus, 145 

Argyrops spinifer, 145 

Arkansas, U.S.A., 122 

Armadillidium nasatum, 32 

Artnadillidium vulgare, 32 

Asian pond loach, 224 

Atheriniformes, 190 

Atypical morphology, 258 

Australia, 51, 109 

Australian water dragon, 109 

Aves, 32, 244, 250 

Aviary, 114 

Batracholandros salamandrae, 133 

Biodiversity, 1 

Biogeography, 85, 92 

Biomphalaria glabrata, 236, 241 

Biosphere, 1 

Black-bellied plover, 250 

Black-tailed prairie dog, 197 

Blarina brevicauda, 32 

Brachycoelium storeriae, 133 

Brachylaima fuscatum, 250 

Brazil, 210 

Brevimulticaecum tenuicolle, 122 

Brevitritospinus subgen. n., 40 

Bristol Bay, Alaska, U.S.A., 250 

British Columbia, Canada, 253 

British West Indies, 190 

Brook trout, 181 

Brown trout, 181 

Bufo americanus americanus, 202 

Bufo marinus, 92, 129 

Bufo marmoreus, 129 

Bufo valliceps, 92 

Bufonidae, 26, 92, 202, 129 

Bullfrog tadpole, 26 

Calidris mauri, 250 

Calidris ptilocnemis, 250 

California, U.S.A., 71, 135, 165 

California treefrog, 165 

Callorhinus ursinus, 218 

Calydiscoides flexuosus, 145 • 

Canada, 26, 253 

Capriniana sp., 181 

Capsalidae, 190 

Carabidae, 107 

Carnivora, 32 

Carolinensis tuffi sp. n., 66 

Case history, 71 

Catadiscus rodriguezi, 92 

Catfish, 76 

Ca the be (fish), 40 

Caudata, 224 

Cavia porcellus, 197 

Cecal tapeworm, 258 

Cenotes, 76 

Centrorhynchus spinosus, 124 

Centrorhynchus sp., 129 

Cephalogonimus americanus, 92 

Cephalouterina leoi, 60 

Cestoda, 109, 118, 124, 181, 197, 

202, 210, 250, 258 

Chaetodon lunula, 190 

Chaetodontidae, 190 

Channidae, 40 

Charadrii, 250 

Chilodonella sp., 181 

Chinchilla lanigera, 197 

Chlaenius prasinus, 107 


Cichla monoculus, 210 

Cichlasoma aureum, 85 

Cichlasoma callolepis, 85 

Cichlasoma friedrichstahli, 85 

Cichlasoma geddesi, 85 

Cichlasoma helleri, 85 

Cichlasoma lentiginosum, 85 

Cichlasoma managuense, 85 

Cichlasoma salvini, 85 

Cichlasoma sp., 85 

Cichlasoma trimaculatum, 85 

Cichlasoma urophthalmus, 85 

Cichlidae, 85, 210 

Ciliophora, 181 

Clinostomum attenuaturn, 255 

Clinostomum sp., 210 

Cloacinidae, 51 

Cobitis biwae, 224 

Cockroach, 32 

Coleoptera, 107 

Colobomatus embiotocae, 253 

Common wallaroo, 51 

Community structure, 202, 210 

Congridae, 190 

Connecticut, U.S.A., 197 

Contracaecum sp., 210 

Copepoda, 181, 224, 253 

Copper-tailed skink, 118 

Cosrnocercella haberi, 255 

Cosmocercoides variabilis, 202 

Cricetidae, 66 

Crustacea, 32, 181, 224, 253 

Cryptoblepharus poeciloplcurus, 

118 

Cryptobranchidae, 224 

Ctenophorus caudicinctus, 109 

Ctenophorus fordi, 109 

Ctenophorus isolepis, 109 

Ctenophorus reticulatus, 109 

Ctenophorus scutulatus, 109 

Cylindrotaenia decidua, 118 

Cymatogaster aggregata 

Cynomys ludovicianus, 197 

Cyprinidae, 40 

Cystacanth, 26, 33, 60, 114, 124, 

129, 133 

Cytopathology, 236 

Dactylogyridae, 76, 85 

Deer mouse, 32 

Delagoa threadfin bream, 145 

Demidueterospinus subgen. n., 40 

Diagnostic Parasitology Course, 39 

Dictymetra nymphaea, 250 

Digenea, 26, 92, 124, 165, 202, 

210, 236, 241, 250, 255 

Diplectanidae, 145 

Diplectanum cazauxi, 145

Diplectanum sillagonum, 145 

Diplectanum spp., 145 

Diplodus noct, 145 

Diplostomatidae, 26 

Diplostomiim (Austrodiplostomum) 

compactum, 210 

Diplostomiim sp., 26, 210 

Domestic mouse, 32, 197, 230, 241 

Domestic spiny mouse, 197 

Diymarchon corais couperi, 124 

Dwarf bearded dragon, 109 

Eastern indigo snake, 124 

Echeneidae, 190 

Echeneis neucratoides, 190 

Echinocotyle tenuis, 250 

Echinopaiyphium recurvatum, 250 

Echinostoma caproni, 241 

Echinostoma trivolvis, 236 

Echinostome metacercariae, 202 

Ecology, 202, 210, 218, 250 

Editors' Acknowledgments, 223 

Egretta alba, 244 

Eleutherodactylus rhodopis, 92 

Embiotocidae, 253 

Etnoia cyanura, 118 

Endohelminths, 1, 26, 32, 40, 51, 

60, 66, 71, 76, 85, 92, 107, 109, 

114, 118, 122, 124, 129, 133, 

135, 145, 165, 169, 181, 197, 

202, 210, 218, 224, 230, 236, 

241, 244, 250, 255, 258 

Epinephelus areolatus, 145 

Epinephelus rnorio, 190 

Epinephelus tauvina, 145 

Estado de Mexico, Mexico, 92 

Eiibothriurn salvelini, 181 

European rabbit, 197 

Eustrongyloides sp., 124 

Experimental infection, 32 

Extraintestinal infection, 32 

Eyefluke, 26 

Eyeworm, 258 

Falcaustra chelydrae, 122 

Falcaustra wardi, 122 

Ferret, 197 

Fibricola sp., 92 

Fishes, 40, 76, 85, 145, 181, 190, 

210, 224, 253 

Florida, U.S.A., 124, 190, 255 

French Polynesia, 118 

Frog, 26, 92, 165, 224, 255 

Gambusia xanthosoma, 190 

Gehyra oceanica, 118 

Genus revision, 40, 165, 169 

Gigantorhynchidae, 114 

Global Taxonomy Initiative, 1 

Glucocorticoid treatment, 230 

Glypthelmins californiensis, 92 

Glypthelmins facial, 92 

Glypthelmins parva, 92 

Glypthelmins quieta, 92 

Glypthelmins sp., 92 

Gnathostorna cf. biniicleatum, 244 

Gnathostoma doloresi, 224 

Gnathostoma sp., 124 

Gnathostomatidae, 124, 224 

Golden hamster, 197 

Gordioidea, 107 

Gordius difficilis, 107 

Gorgoderina attenuata, 92 

Gorgoderina sp., 202 

Grand Cayman Island, 190 

Great egret, 244 

Green treefrog, 255 

Growth, 241 

Guatemala, 85 

Guinea pig, 197 

Gulf of Mexico, 190 

Gyrodactylus sp., 181 

Haematoloechus coloradensis, 92 

Haematoloechus cornplexus, 92 

Haematoloechus illimis, 92 

Haematoloechus longiplexus, 92 

Haematoloechus medioplexus, 92 

Haematoloechus pulcher, 92 

Haematoloechus sp., 92 

Haemonchus contortus, 135 

Halipegus occidualis, 92 

Heligmosomoidea, 66 

Helminthological Society of Washington: 

Anniversary Award, 261 

Articles of Incorporation, 141 

Constitution and By-Laws, 138 

Meeting Schedule, 65, 240 

Membership Application, 143, 

Minutes of Meetings, 263 

Mission and Vision Statements, 

144, 272 

Helminths, 1, 26, 32, 40, 51, 60, 

66, 71, 76, 85, 92, 107, 109, 114, 

118, 122, 124, 129, 133, 135, 

145, 165, 169, 181, 197, 202, 

210, 218, 224, 230, 236, 241, 

244, 250, 255, 258 

Hemiramphidae, 145 

Hemiramphus marginatus, 145 

Hermann's tortoise, 169 

Heteroconger has si, 190 

Heteromyidae, 197 

High-carbohydrate diet, 241 

Hispid pocket mouse, 32 


Hookworms, 218 

Horse, 258 

Host specificity, 85, 190 

Human infection, 71 

Hyla arenicolor, 92 

Hyla cadaverina, 165 

Hyla cinerea, 255 

Hyla eximia, 92 

Hylidae, 92, 165, 255 

Hymenolepis nana, 197 

Hynobiidae, 60 

INDEX 267 

Ichthyobodo sp., 181 

Ichthyophthirus multijiliis, 181 

ICR mice, 241 

India, 145 

Indiana, U.S.A., 190 

Indo-Pacific tree gecko, 118 

Inducible nitric oxide, 230 

iNOS, 230 

Insecta, 114 

Intensity, 32, 60, 109, 118, 122, 

124, 129, 133, 135, 181, 197, 

202, 210, 250, 253, 255 

International Code of Zoological 

Nomenclature (Fourth Edition), 

75 

Intestine, 236 

Inventory, 1 

Isopoda, 32 

Jalisco, Mexico, 92, 129 

Japan, 60, 224 

Japanese clawed salamander, 60 

Japanese giant salamander, 224 

Japanese threadfin bream, 145 

Jird, 197, 236 

Kalicephalus appendiculatus, 124 

Kalicephalus inermis coronellae, 

124 

Kalicephalus rectiphilus, 124 

Kentucky, U.S.A., 258 

King soldierbream, 145 

Kiricephalus coarctatus, 124 

Kowalewskiella cingulifera, 250 

Kreisiella chrysocampa, 109 

Kreisiella lesueurii, 109 

Kuwait, 145 

Lacunovermes sp., 250 

iMinellodiscus furcillatus sp. n., 

145 

Lamellodiscus spp., 145 

Langeronia burseyi sp. n., 165 

Langeronia macrocirra, 165 

iMngeronia parva, 165 

Langeronia provitellaria, 165


Larva, 114, 210, 224, 230, 244 

Lecithodendriidae, 165 

Lepidodactylus lugubris, 118 

Leptodactylus melanonotus, 92 

Lepidotrema kuwaitensis sp. n., 

145 

Lepidotrema longipenis comb, n., 

145 

Life history, 224 

Liga brevis, 250 

Lipotyphyla, 32 

Littoral zone, 250 

Lizard, 109, 118 

Long-tailed chinchilla, 197 

Lophognathus longirostris, 109 

Louisiana, U.S.A., 122 

Loxogenes (Langeronia) macrocirra, 

92 

Lozenge-marked dragon, 109 

Lutjanidae, 190 

Lutjanus campechanus, 190 

Macracanthorhynchus ingens, 124 

Macroclemys temminckii, 122 

Macrocyclops albidus, 224 

Macropodidae, 51 

Macropus rohustus, 51 

Mallee dragon, 109 

Mammalia, 1, 32, 66, 135, 190, 

197, 218, 230, 236, 241, 258 

Manatee, 190 

Marsupalia, 51 

Mastigophora, 181 

Maxvachonia brygooi, 109 

Maxvachonia chabaudi, 118 

Mediorhynchus orientalis, 114 

Mediorhynchus wardi, 114 

Meeting Announcements, 91, 209 

Megalodiscus americanus, 92 

Meriones iinguiculatus, 197, 236 

Mesoccstoides sp., 202 

Mesocoelium brevicaecum, 60 

Mesocoelium rnonas, 92 

Mesocricetus auratus, 197 

Mesocyclops dissimilis, 224 

Metacercaria, 26 

Methylprednisolone, 230 

Metoponorthus pruinosus, 32 

Mexico, 76, 85, 92, 129, 244 

Mexico City, Mexico, 92 

Michigan, U.S.A., 181 

Michoacan, Mexico, 92 

Microtus ochrogaster, 32 

Microtus pennsylvanicus, 32 

Military dragon, 109 

Misgurnus anguillicaudatus, 224 

Mississippi, U.S.A., 190 

Mollusca, 236, 241 

Mongolian gerbil, 197, 236 

Monogenea, 85, 181, 190 

Monogenoidea, 76, 145 

Moorea, 118 

Morocco, 169 

Morphology, 40, 51, 60, 66, 71, 76, 

85, 92, 107, 114, 145, 165, 169, 

224, 244, 258 

Moth skink, 118 

Mourning gecko, 118 

Mouse, 32, 197, 230, 241 

Mudpuppy, 26 

Muscle, 230 

Mus musculus, 32, 197, 230, 241 

Museums for Depositing of Specimens, 

189 

Mustela nivalis, 32 

Mustela putorius favo, 197 

Myxobolus cerebralis, 181 

Myxozoa, 181 

Nebraska, U.S.A., 32, 107 

Necturus maculosus, 26 

Nematoda, 51, 60, 66, 71, 109, 

118, 122, 124, 129, 133, 135, 

169, 181, 202, 210, 218, 224, 

230, 255, 258 

Nernatodeirus odocoilei, 135 

Nematomorpha, 107 

Nemipteridae, 145 

Nemiptenis bipunctatus, 145 

Nemipterus peronii, 145 

Neobenedenia melleni, 190 

Neoechinorhynchus chrysemydis, 

122 

Neoechinorhynchus emydis, 122 

Neoechinorhynchus pseudemydis, 

122 

New book available, 201 

New geographical record, 60, 76, 

85, 92, 107, 109, 118, 122, 124, 

129, 133, 135, 145, 190, 250, 

253, 255 

New host record, 85, 92, 107, 109, 

118, 122, 124, 129, 133, 135, 

145, 190, 197, 250, 255 

New South Wales, Australia, 51 

New taxon, 40, 51, 60, 66, 76, 97, 

145, 165, 169 

Northern fur seal pups, 218 

Northern phalarope, 250 

Northern Territory, Australia, 109 

Norway rat, 197 

Notchedfin threadfm bream, 145 

Numenius phaeopus, 250 

Oaxaca, Mexico, 92, 244 

Obituary Notices: 


Alan F. Bird, 168 

Marion M. Farr, 180 

Michael J. Patrick, 164 

Everett Lyle Schiller, 31 

Obtuse barracuda, 145 

Oceanic gecko, 118 

Ochetosoma elongatum, 124 

Ochetosoma kansense, 124 

Ochetosoma sp., 92 

Ochoterenella digiticauda, 129 

Odocoileus hemionus fuliginatus, 

135 

Ohio, U.S.A., 190 

Oncorhynchus my kiss, 181 

Onychodactylus japonicus, 60 

Oochoristica piankai, 109 

Opalina sp., 255 

Ophicephalus rnaculatus, 40 

Oryctolagus cuniculus, 197 

Osteichthyes, 145, 253 

Oswaldocruzia pipiens, 202 

Otolithes argenteus, 145 

Oxyurid sp., 118 

Oxyuroids, 169 

Pachymedusa dachnicolor, 92 

Pademelon, 51 

Pallisentis, 40 

Pallisentis sensu stricto, new diagnosis, 

40 

Pallisentis tetraodontae, new synonym, 

40 

Pallisentis (Brevitritospinus) vietnamensis 

subgen. et sp. n., 40 

Pallisentis (Pallisentis) pesteri, 

comb, n., 40 

Parana River, Brazil, 210 

Paraphaiyngodon fitzroyi, 109 

Parapharyngodon japonicus, 60 

Pararaosentis gen. n., 40 

Pararaosentis golvani, comb, n., 

40 

Pelecanus erythrorhynchos, 244 

Pennsylvania, U.S.A., 114 

Pentastoma, 124 

Perciformes, 210 

Periplaneta americana, 32 

Perognathus flavescens, 32 

Perognathus hispidus, 32 

Perornyscus leucopus, 32 

Peromyscus maniculatus, 32 

Perornyscus pectoralis, 66 

Persian Gulf, 145 

Petenia splendida, 85 

Pets, 197 

Phalaropus lobatus, 250 

Pharyngodon oceanicus, 118 

Pharyngodonidae, 118, 169

Philichthyidae, 253 

Phodopus sungorus, 197 

Phylogeny, 1 

Physaloptera obtussirna, 124 

Physaloptera sp., 124, 129 

Physocephalus sp., 129 

Pickhandle barracuda, 145 

Pimclodidae, 76 

Pimelodus clarias, 76 

Pisces, 40, 76, 85, 145, 181, 190, 

210, 224, 253 

Plagiorchis morosovi, 251 

Plagiorhynchus cylindraceus, 32 

Plethodon richmondi, 133 

Pluvialis squatarola, 250 

Poeciliidae, 190 

Pogona minor, 109 

Polymorphic magnus, 250 

Popovastrongylus pluteus sp. n., 

51 

Popovastrongylus tasmaniensis sp. 

n., 51 

Popovastrongylus wallabiae, 51 

Prevalence, 26, 32, 60, 109, 118, 

122, 124, 129, 133, 135, 181, 

197, 202, 210, 218, 253, 255 

Proteocephalus macrophallus, 210 

Proteocephalus microscopicus, 210 

Proteocephalus sp., 124, 181 

Proterogynotaenia variabilis, 250 

Protolamellodiscus senilobatus sp. 

n., 145 

Protozoa, 181, 255 

Pseudolamellodiscus sphyraenae, 

145 

Pseudopolystoma dendriticum, 60 

Pseudorhabdosynochus spp., 145 

Pseudoterranova decipiens, 71 

Public aquaria, 190 

Puebla, Mexico, 92 

Puerto Rico, 190 

Pycnoscelis surinarnensis, 114 

Quadrigyridae, 40, 210 

Quadrigyrus machadoi, 210 

Quebec, Canada, 26 

Rainbow trout, 181 

Rana brownorurn, 92 

Rana catesbeiana, 26 

Rana dunni, 92 

Rana forreri, 92 

Rana megapoda, 92 

Rana montezumae, 92 

Rana neovolcanica, 92 

Rana nigromaculata, 224 

Rana rugosa, 224 

Rana vaillanti, 92 

Ransom Trust Fund Report, 249 

Rattus norvegicus, 197 

Ravine salamander, 133 

Red-necked wallaby, 51 

Red Sea seabream, 145 

Reithrodontomys rnegalotis, 32 

Reptilia, 109, 118, 122, 124, 169 

Rhabdias americanus, 202 

Rhabdias fuelleborni, 129 

Rhabdias sp., 255 

Rhamdia guatemalensis, 76 

Ring-tailed dragon, 109 

Robin, 32 

Rock sandpiper, 250 

Rodentia, 32, 66, 197, 230, 236, 

241 

St. Lawrence River, Canada, 26 

St. Paul Island, Alaska, U.S.A., 218 

Salamander, 26, 60, 92, 133, 224 

Salmincola edwardsii, 181 

Salmo trutta, 181 

Salmonidae, 181 

Salvelinus fontinalis, 181 

Sand loach, 224 

Sauria, 109, 118 

Scanning electron microscopy, 

169, 236, 244 

Schistocephalus solidus, 250 

Sciadiclethrutn bravohollisae, 85 

Sciadiclethrum meekii, 85 

Sciadiclethrum mexicanum, 85 

Sciadiclethrum splendidae, 85 

Sciadocephalus megalodiscus, 210 

Sciaenidac, 145 

Seasonal study, 202 

Second-stage larva, 224 

SEM, 169, 236, 244 

Serpinerna trispinosus, 122 

Serranidae, 145, 193 

Shiner perch, 253 

Shorebirds, 250 

Short-tailed shrew, 32 

Siberian hamster, 197 

Sillaginidae, 145 

Sillago siharna, 145 

Siluriformes, 76 

Silver sillago, 145 

Sinaloa, Mexico, 248 

Skrjabinoptera gallardi, 109 

Skrjabinoptera goldrnanae, 109 

Skrjabinoptera sp., 118 

Small-scaled terapon, 145 

Smilisca baudinii, 92 

Snail, 236, 241 

Snake, 124 

Snake-eyed skink, 118 

Snake head mullet, 40 


INDEX 269 

Soldierbream, 145 

Sorex cinereus, 32 

Southern mule deer, 136 

Spain, 169 

Sparidae, 145 

Spauligodon gehyrae, 118 

Spennophilus tridecemlineatus, 32 

Sphyraena chrysotaenia, 145 

Sphyraena jello, 145 

Sphyraena obtusata, 145 

Sphyraenidae, 145 

Spiroxys hanzaki, 224 

Spur-thighed tortoise, 169 

Starling, 32 

Strongyloides sp., 124 

Stump-toed gecko, 118 

Sturnus vulgaris, 32 

Surfbird, 250 

Surinam cockroach, 114 

Survey, 1, 26, 32, 60, 76, 85, 92, 

109, 118, 122, 124, 129, 133, 

135, 145, 197, 201, 218, 250, 

253, 255 

Tadpole, 26 

Tasmania, Australia, 51 

Tasmanian pademelon, 51 

Taxonomic key, 40, 165, 169 

Taxonomy, 1, 40, 51, 60, 66, 76, 

85, 92, 115, 145, 165, 169 

Teladorsagia circumcincta, 135 

Teleostei, 40, 76, 85, 145, 190, 

210, 224, 253 

TEM, 236 

Terapon puta, 145 

Teraponidae, 145 

Terranova caballeroi, 124 

Testudinidae, 169 

Testudo graeca, 169 

Testudo hermanni, 169 

Texas, U.S.A., 66 

Thaparia bourgati sp. n., 169 

Thaparia capensis, 169 

Thaparia carlosfeliui sp. n., 169 

Thaparia contortospicula, 169 

Thaparia domerguei, 169 

Thaparia macrocephala, 169 

Thaparia microcephala, 169 

Thaparia rnacrospiculum, 169 

Thaparia thapari australis, 169 

Thaparia thapari rysavyi, 169 

Thaparia thapari thapari, 169 

Thelazia lacrymalis, 258 

Third-stage larva, 71, 224 

Thirteen-lined ground squirrel, 32 

Thylogale billiardierii, 51 

Toad, 92, 129, 202 

Tortoise, 169


Trachelipus rathkei, 32 

Transmission electron microscopy, 

236 

Trematoda, 60, 124, 165, 202, 241, 

250, 255 

Trichechus manatux, 190 

Trichinella pseudospiralis, 230 

Trichinella spi rails, 230 

Trichocephaloides megalocephala, 

250 

Trichodina sp., 181 

Trichostrongylina, 66, 135 

Trout, 181 

Truttaedacnitis sp., 181 

Tucunare, 210 

Turdus migratorius, 32 

Turtle, 122 

Ultrastructure, 169, 236, 244 

Uncinaria lucasi, 218 

Urodela, 92 

U.S.A., 32, 66, 71, 107, 122, 124, 

133, 135, 165, 181, 190, 197, 

202, 218, 250, 255, 258 

Veracruz, Mexico, 92 

Vietnam, 40 

Vivid metallic ground beetle, 107 

Wallaby, 51 

Wanaristrongyla ctenoti, 109 

Wardium amphitricha, 250 

Wardium squatarolae, 250 

West Virginia, U.S.A., 133 


Western Australia, 109 

Western netted dragon, 109 

Western sandpiper, 250 

Whimbrel, 250 

White-ankled mouse, 66 

Whitefin sharksucker, 190 

Wisconsin, U.S.A., 202 

Wood mouse, 32 

Worm expulsion, 236 

Worm kinetics, 236 

Yellowstrip barracuda, 145 

Yucatan, Mexico, 76 

Zoogeography, 85, 92, 190 

Zoonosis, 197

Name: 

MEMBERSHIP APPLICATION 271 

APPLICATION FOR MEMBERSHIP 

in the 

HELMINTHOLOGICAL SOCIETY OF WASHINGTON 

(Please Type or Print Legibly) 

Mailing Address: 

Present Position and Name of Institution (If you are a student, write "Student" and indicate the 

degree you are working for and the university/department): 

Phone: _ 

E-Mail: 

Highest Degree Earned and the Year Received: 

Fields of Interest: 

FAX: 

If you are established and experienced in your field, would you consent to be a reviewer for 

manuscripts submitted for publication in the Society's journal, Comparative Parasitologyl If so, 

what specific subject area(s) do you feel most qualified to review? 

Signature of Applicant Date 

Signature of Sponsor (a member) 

Mail the completed application along with a check (on a U.S. bank) or money order (in U.S. 

currency) for the first year's dues (US$25 for domestic active members and US$28 for foreign 

active members) to the Corresponding Secretary-Treasurer, Nancy D. Pacheco, 9708 DePaul Drive, 

Bethesda, MD, U.S.A. 20817. 

If you wish to pay by credit card (Visa and MasterCard only), please complete the following: 

Credit Card: Visa® Q MasterCard® H Expiration Date: 

Credit Card Number: O CH d dl I II II I 

Signature of Card Holder: 

Printed Name Exactly as Stated on Card: 

Date 

Amount Charged: US$ 

Helminthological Society of Washington Home Page: http://www.gettysburg.edu/shendrix/helmsoc.html 



THE HELMINTHOLOGICAL SOCIETY OF WASHINGTON 

MISSION AND VISION STATEMENTS 

May 7, 1999 

THE MISSION 

The Helminthological Society of Washington, the prototype scientific organization for parasitological 

research in North America, was founded in 1910 by a devoted group of parasitologists in 

Washington, D.C. Forging a niche in national and international parasitology over the past century, 

the Society focuses on comparative research, emphasizing taxonomy, systematics, ecology, biogeography, 

and faunal survey and inventory within a morphological and molecular foundation. Interdisciplinary 

and crosscutting, comparative parasitology links contemporary biodiversity studies with 

historical approaches to biogeography, ecology, and coevolution within a cohesive framework. 

Through its 5 meetings in the Washington area annually, and via the peer-reviewed Comparative 

Parasitology (continuing the Journal of the Helminthological Society of Washington in its 67th 

Volume), the Society actively supports and builds recognition for modern parasitological research. 

Taxonomic diversity represented in the pages of the Society's journal treats the rich helminth faunas 

in terrestrial and aquatic plants, invertebrates, and vertebrates, as well as parasitic protozoans and 

arthropods. Parasitology, among the most integrative of the biological sciences, provides data critical 

to elucidation of general patterns of global biodiversity. 

THE VISION 

The Helminthological Society of Washington celebrates a century of tradition and excellence 

in global parasitology, solving challenges and responding to opportunities for the future of society 

and the environment. 

Members of the Helminthological Society of Washington contribute to understanding and protecting 

human health, agriculture, and the biosphere through comparative research emphasizing 

taxonomy, systematics, ecology, biogeography, and biodiversity assessment of all parasites. The 

Society projects the exceptional relevance of its programs to broader research and education in 

global biodiversity and conservation biology through the activities of its members and its journal, 

Comparative Parasitology. 


*Edna M. Buhrer 

* Mildred A. Doss 

* Allen Mclntosh 

* Jesse R. Christie 

•Gilbert F. Otto 

* George R. LaRue 

*William W. Cort 

* Gerard • Dikmans 

* Benjamin Schwartz 

*Willard H. Wright 

*Aurel O. Foster 

*Carlton M. Herman 

*May Belle Chitwood 

*Elvio H. Sadun 

E. J. Lawson Soulsby 

David R. Lincicome 

Margaret A. Stirewalt 

•Leo A. Jachowski, Jr. 

* Horace W. Stunkard 

•Kenneth C. Kates 

*Everett E. Wchr 

*George R. LaRue 

* Vladimir S. Ershov 

•Norman R. Stoll 

•"Horace W. Stunkard 

•Justus F. Mueller 

John F. A. Sprent 

Bernard Bezubik 

Hugh M. Gordon 

•W. E. Chambers 

*Nathan A. Cobb 

* Howard Crawley 

*Winthrop D. Foster 

•Maurice C. Hall 

•Albert Hassall 

•Charles W. Stiles 

•Paul Bartsch 

•Henry E. Ewing 

•William W. Cort 

•Gerard Dikmans 

•Jesse R. Christie 

•Gotthold Steiner 

•EmmettW. Price 

•Eloise B. Cram 

•Gerald Thome 

•Allen Mclntosh 

•Edna M. Buhrer 

•Benjamin G. Chitwood 

•Aurel O. Foster 

•Gilbert F. Otto 

•Theodor von Brand 

•May Belle Chitwood 

•Carlton M. Herman 

Lloyd E. Rozeboom 

•Albert L. Taylor 

David R. Lincicome 

Margaret A. .Stirewalt 

•Willard H. Wright 

•Benjamin Schwartz 

•Mildred A. Doss 

* Deceased. 

ANNIVERSARY AWARD RECIPIENTS 

1960 

1961 

1962 

1964 

1965 

1966 

1966 

1967 

1969 

1969 

1970 

1971 

1972 

1973 

1974 

1975 

1975 

1976 

1977 

1978 

1979 

HONORARY MEMBERS 

1959 

1962 

1976 

1977 

1978 

1979 

1980 

1981 

•Philip E. Garrison 

*Joseph Goldberger 

•Henry W. Graybill 

1931 

1931 

1931 

1937 

1945 

1952 

1953 

1956 

1956 

1956 

1956 

1961 

1963 

1963 

1968 

1972 

1972 

1975 

1975 

1975 

1975 

1975 

1976 

1976 

1976 

1976 

1977 

*O. Wilford Olsen 

*Frank D. Enzie 

Lloyd E. Rozeboom 

*Leon Jacobs 



Louis S. Diamond 

•Everett L. Schiller 

Milford N. Lunde 

J. Ralph Lichtenfels 

A. James Haley 

*Francis G. Tromba 

Thomas K. Sawyer 

Ralph P. Eckerlin 

Willis A. Reid, Jr. 

Gerhard A. Schad 

Franklin A. Neva 

Burton Y. Endo 

Sherman S. Hendrix 

Frank W. Douvres 

E. J. Lawson Soulsby 

Roy C. Anderson 

Louis Euzet 

John C. Holmes 

Purnomo 

Naftale Katz 

"Robert Traub 

•Alan F. Bird 

•Maurice C. Hall 

•Albert Hassall 

•George F. Leonard 

•Everett E. Wehr 

•Marion M. Farr 

•John T. Lucker, Jr. 

George W. Luttermoser 

•John S. Andrews 

•Leo A. Jachowski, Jr. 

•Kenneth C. Kates 

•Francis G. Tromba 

A. James Haley 

•Leon Jacobs 

•Paul C. Beaver 

•Raymond M. Cable 

Harry Herlich 

Glenn L. Hoffman 

Robert E. Kuntz 

Raymond V Rebois 

Frank W. Douvres 


Thomas K. Sawyer 

*J. Allen Scott 

Judith H. Shaw 

Milford N. Lunde 

•Everett L. Schiller 


Louis S. Diamond 

Mary Hanson Pritchard 


1990 

1991 

1992 

1993 

1994 

1995 

1996 

1997 

•Charles A. Pfender 

•Brayton H.-Ransom 

•Charles W. Stiles 

1977 

1979 

1979 

1979 

1980 

1981 

1981 

1983 

1984 

1985 

1986 

1986 

1987 

1988 

1988 

1988 

1989 

1989 

1989 

1990 

1990 

1991 

1991 

1991 

1994 

1994

VOLUME 67 

JULY 2000 

CONTENTS 

(Continuedfrom Front Cover) 

NUMBER 2 

RESEARCH NOTES 

CANARIS, A. G., AND J. M. KINSELLA. Helminth Parasites of Six Species of Shorebirds (Charadrii) from 

Bristol Bay, Alaska, U.S.A. . 250 

JEPPS, S. K, AND T. M. GOATER. Colobomatus embiotocae (Copepoda: Philichthyidae) from Shiner Perch, 

Cymatogaster aggregate (Osteichthyes: •Embiotocidae) in Canadian Waters 253 

CREEL, T. L., G. W. FOSTER, D. J. FORRESTER. Parasites of the Green Treefrog, Hyla cinerea, from Orange 

Lake, Alachua County, Florida, U.S.A 255 

BAIR, H. D., E. T. LYONS, T. W. SWERCZEK, AND S. C. TOLLIVER. Atypical Specimens of Helminth Parasites 

(Anoplocephala perfoliata and Thelazia lacrymalis) of Horses in Kentucky, U.S.A 258 

ANNOUNCEMENTS AND NOTICES 

OBITUARY NOTICES -. . 164, 168, 180 

MUSEUMS FOR DEPOSITING OF SPECIMENS 189 

NEW BOOK AVAILABLE 201 

MEETING NOTICES , 209 

EDITORS' ACKNOWLEDGMENTS . : . 223 

HELMINTHOLOGICAL SOCIETY OF WASHINGTON MEETING SCHEDULE „. 240 

REPORT OF THE BRAYTON H. RANSOM MEMORIAL TRUST FUND 249 

PRESENTATION OF THE 1999 ANNIVERSARY AWARD 261 

MINUTES OF MEETINGS OF THE HELMINTHOLOGICAL SOCIETY OF WASHINGTON . 263 

AUTHOR INDEX , 265 

KEY WORD AND SUBJECT INDEX , . . . 265 

MEMBERSHIP APPLICATION 271 

MISSION AND VISION STATEMENT OF THE HELMINTHOLOGICAL SOCIETY OF WASHINGTON _.. 272 

Date of publication, 24 July 2000 

* * * 

PRINTED BY ALLEN PRESS, INC., LAWRENCE, KANSAS 66044, U.S.A.

Comparative Parasitology 67(2) 2000 - Peru State College

Create successful ePaper yourself

Delete template?

Save as template?