Acanthocephalan infections of fish
Don't need the entire report?
Generate a print friendly version containing only the sections you need.Generate report
PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Acanthocephalan infections of fish
Pathogen/sTop of page
OverviewTop of page
Acanthocephalans are endoparasitic worms comprising approximately 1100 species (Golvan, 1994), nearly one-half of which are found as adults in the intestine of fishes. Juvenile worms of many other species occur in the viscera, especially the mesentery and liver, of fishes that act as paratenic hosts. Although these parasites are not usually regarded as having significant economic importance, epizootics are known from hatcheries (Bullock, 1963) and they have been linked to local extinction of natural populations (Schmidt et al., 1974). There is little knowledge of the more subtle effects the parasite has on host populations, but mortality might be significant when hosts are stressed (Jilek, 1979; Connors and Nickol, 1991).
Acanthocephalans occur globally in most fishes, but epizootics and significant economic losses as a result of infection are unusual. Acanthocephalans require vertebrate animals for definitive hosts and arthropods for intermediate hosts. Isopods, amphipods and ostracods are the usual intermediate hosts for aquatic species. Infection occurs when a definitive host consumes the infective cystacanth stage contained in an arthropod or in a paratenic host. There is an increasing awareness that, in some cases, postcyclic transfer of adult worms from fish to fish can occur as a result of predation. Worms are typically recruited into fish populations during the spring, with maturation, egg production and transmission to intermediate hosts in the summer and early autumn. Adult worms usually live about one season.
Acanthocephalans attach to the intestine of definitive hosts by means of a spiny proboscis. Mucosal tissue is damaged at the attachment site, resulting in fibroplasia, which may extend through the submucosa and into the muscularis. Occasionally perforation of the gut wall occurs. The mucosal epithelium is frequently compressed or eroded along the trunk of the worm, and the tips of the villi may be absent. Destruction of intestinal villi and necrotic and degenerative changes in mucosal epithelium almost certainly reduce the absorptive efficiency of the fish intestine.
Some acanthocephalans are known to induce antibody formation in piscine species. Haemorrhage at the attachment site and copious amounts of mucus, from goblet-cell hyperplasia, in parasitized fishes provide an opportunity for antibody-worm interaction, but it is not known if protective immunity develops.
Acanthocephalans lack an alimentary tract and hence uptake of nutrients, derived both from leakage of host tissues and from dietary contents in the intestinal lumen of the host, is through the tegument. Hydrolytic enzyme activity at the tegumental surface probably assists in obtaining nutrients and in rapid penetration by the worm. Some acanthocephalans penetrate deeply and induce formation of a nodule, which extends into the coelom of the host. Such nodules are extensively vascularized. Increased leakage of proteins from the blood into the nodules ensures a steady supply of nutrients for the parasites.
Carbohydrate, in the form of glycogen, is the main substrate for energy, and glycolysis occurs in the presence or absence of oxygen. Lactate and succinate are the main end products, but large amounts of ethanol are excreted by at least one species.
All media in which significant growth has been achieved contain undefined components. Further research for a defined medium is necessary before appreciable progress can be made in determining nutritional requirements, especially for amino acids and lipids.
The consequences of acanthocephalan-induced reductions in energy efficiency and altered metabolism of hosts are likely to be focused more sharply with increasing emphasis on aquaculture. Suitable methods of in vitro cultivation are necessary to elucidate responses of worms to chemotherapeutic agents. Known effects detrimental to commercially cultured fishes are likely to stimulate research directed at testing for chemotherapeutic compounds. When efficacious drugs are found, their toxicity to fishes, aquatic invertebrates and human beings must be assessed. Recent appreciation of the propensity of acanthocephalans to accumulate heavy metals and other elements is likely to stimulate research to evaluate their effectiveness as bioindicators of heavy-metal pollution. The broad spectrum of metals that can be accumulated, even if present in very low concentrations, might make acanthocephalans pre-eminent among organisms used as sentinels in aquaculture systems.
[Derived from: Woo, PTK, ed., 2006. Fish diseases and disorders, Volume 1: Protozoan and Metazoan infections. (2nd edition) Wallingford, UK: CAB International]
Host AnimalsTop of page
Hosts/Species AffectedTop of page
Fishes of most systematic groups are parasitized. Although these parasites are rare in agnathans and elasmobranchs, a few species are known to occur in lampreys (Hoffman, 1999), rays (Brooks and Amato, 1992; Sanmartin et al., 2000; Knoff et al., 2001) and sharks (Yamaguti, 1961). Sturgeons, primitive ray-fin fishes (Chondrostei), have one acanthocephalan species specific to them (Choudhury and Dick, 2001) and are reported to host another 14 species incidentally (Choudhury and Dick, 1998). Bowfins and gars, intermediate ray-fin fishes, harbour another few species (Golvan and de Buron, 1988; Hoffman, 1999). Most of the acanthocephalans of piscine hosts are found in the bony fishes (Teleostei). More species of Acanthocephala are found in the cypriniform families Cyprinidae and Catostomidae than in any other piscine family.
DistributionTop of page
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.Last updated: 10 Jan 2020
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
PathologyTop of page
After ingestion by the definitive host, cystacanths are activated and attach to the wall of the small intestine. Many species apparently remain at the activation site and develop to maturity. Others migrate and localize in specific regions of the intestine (Crompton 1975; Kennedy, 1985). Several factors, including physiological differences along the alimentary tract (Richardson and Nickol, 1999), maximization of sexual congress (Richardson and Nickol, 2000) and ageing (Taraschewski, 2000), have been implicated in site specificity. In most cases, however, it is unknown whether differences in sites occupied result from differences in sites of activation, differential mortality or emigration by the parasites.
Fig. 4. Scanning electron micrograph of an egg from Leptorhynchoides thecatus showing unwrapping of embryonic membranes. Unwrapped membranes entangle aquatic vegetation and anchor egg in habitat of intermediate host feeding. Scale bar = 0.01 mm. (Unpublished micrograph courtesy of M.A. Barger.)
Course of infection
Shortly after infection, the proboscis is surrounded by necrotic tissue, which becomes haemorrhagic and inflamed after a few days. During the second week after infection by species that penetrate deeply (e.g. in fish, Acanthocephalus anguillae, P. bulbocolli, P. laevis), inflamed tissue around the anterior portion of the worm is dominated by monocytes and macrophages maturing into epithelioid cells, and an outer belt of connective tissue appears (Taraschewski, 2000). This chronic stage often results in a fibrous nodule visible on the outer surface of the intestine. Species that do not penetrate deeply (e.g. in fish, Acanthocephalus lucii, E. truttae, N. cylindratus) move about in the intestine and change their point of attachment before the connective-tissue response results in nodules (Adel-Meguid et al., 1995).
Copulation in the definitive host may occur within 24 h of infection (Muzzall and Rabalais, 1975). In laboratory infections of green sunfish (Lepomis cyanellus), copulation within a group of L. thecatus continued at least 12 weeks after infection (Richardson et al., 1997). For most species, egg production starts between 4 and 8 weeks after infection, and it continues for approximately 2 months. The number of eggs produced daily by each female acanthocephalan is unknown for most species, but it appears to be related to the size of the worm. At peak production, a female Macracanthorhynchus hirudinaceus (large worms found in pigs) may produce about 260,000 eggs per day (Kates, 1944), a female Moniliformis moniliformis (intermediate-sized worms in rats) about 4800 (Reyda and Nickol, 2001) and a female Polymorphus minutus (small worms found in waterfowl) about 1700 (Crompton and Whitfield, 1968).
Male worms have shorter lifespans than do females; death of males and subsequent loss from the host may begin shortly after copulation. Although females of some species, especially those parasitic in mammals, may live longer than a year, most in poikilothermic hosts live about one season. There may be more than one generation each year.
Clinical signs and gross pathology
Acanthocephalans in moribund or dead animals are frequently assumed to indicate deleterious effects; worms have been observed extending from the rectum (Schmidt et al., 1974; Fig. 5) or protruding through the trunk (Taraschewski, 2000) of infected fishes. There are, however, numerous instances of exceedingly heavy infections in animals that do not show any obvious disease.
Fig. 5. Photograph of Acanthocephalus dirus attached to the prolapsed rectum of a mottled sculpin (Cottus bairdii). Photograph = actual size. (From Schmidt et al., 1974).
Fig. 6. External view of intestine from a quillback (Carpiodes cyprinus) showing a parasite-induced nodule (P) and expansion of the intestine compared with the normal gut (NG) from a fish infected with Neoechinorhynchus carpiodi. Scale bar = 5 mm. (From Szalai and Dick, 1987.)
Bullock (1963) described trout of several species infected with Acanthocephalus dirus as having underdeveloped musculature, heads that were disproportionately large and a slightly concave dorsolateral surface. He interpreted these as signs of malnutrition.
Fibrotic nodules induced on the intestinal surface (Fig. 6) by deeply penetrating worms are the most frequent signs of infection. At times the viscera may be discoloured or the infected intestine enlarged and inflamed. Pyloric caeca of green sunfish (L. cyanellus) infected with L. thecatus are nearly twice the diameter of those in uninfected fish of the same size (de Buron and Nickol, 1994; Fig. 7). Acanthocephalans occasionally perforate the intestinal wall and protrude into the coelom or attach their proboscides to another organ.
Acanthocephalans embed their spiny proboscis into the mucosal epithelium. Attachment is frequently between villi. At the site of attachment, cells are destroyed and fibroblasts, lymphocytes and macrophages are mobilized below the lamina propria (Dezfuli et al., 1990), where chronic fibrinous inflammation, resulting in an increased amount of connective tissue, causes a thickening (Bullock, 1963). In some species, fibroplasia extends to layers of themuscularis mucosa (de Buron and Nickol, 1994).
Fig. 7. Transverse section through the region of the pyloric caeca of a green sunfish (Lepomis cyanellus) infected with Leptorhynchoides thecatus, showing unparasitized caecum (top) and enlarged parasitized caecum (bottom). Note the compressed and eroded villi along the trunk of the parasite (a) compared with normal villi in the unparasitized caecum. Scale bar = 0.1 mm. (Unpublished micrograph, courtesy of I. de Buron.)
Worms of the deeply penetrating species often possess very long necks and bulbous proboscides, which anchor them deep within the gut wall of their piscine hosts. Dezfuli et al. (2002a) described the cellular structure of a fibrotic tunnel that forms around the neck and proboscis bulb of P. laevis. The tunnel terminates in a capsule, covered by serosa and mesentery, which protrudes several millimetres into the coelomic cavity (Fig. 8). Sometimes these capsules persist as conspicuous fibrinous nodules on the external surface of the alimentary canal, or the proboscis perforates the capsule to emerge free in the coelom or to penetrate the liver or another visceral organ. Similar findings were reported for P. bulbocolli in rainbow darters (Etheostoma caeruleum) (McDonough and Gleason, 1981) and two species of catostomids (Chaicharn and Bullock, 1967). A. anguillae is also known to perforate the intestine of its host and attach to the liver. In goldfish (C. auratus), large portions of this organ are replaced by proliferative tissue, often with patches of pancreas, surrounding the embedded proboscis (Taraschewski, 1989a).
Fig. 8. Section of chub (Leuciscus cephalus) intestine showing penetration of Pomphorhynchus laevis. Note hyperplasia of the lamina propria (arrows) around the neck (cou) and bulbous proboscis (P) of the worm, forming a nodule in the coelom. An anterior portion of the trunk (T) of the worm shows in the intestinal lumen. Scale bar = 1 mm. (Unpublished micrograph, courtesy of I. de Buron.)
Most studies of acanthocephalan-induced lesions reveal areas along the trunk of the worm where the mucosal surface is compressed or desquamated (Fig. 7). In these cases, mucosal folds and tips of villi may be absent and the paramucosal lumen contains large amounts of mucoid material originating from goblet-cell hyperplasia. In green sunfish infected with L. thecatus, there is a significantly greater number of goblet cells in parasitized pyloric caeca than in unparasitized caeca in the same fish (de Buron and Nickol, 1994). This suggests a parasite-induced response that might lessen damage from the erosive nature of the worms. Taraschewski (2000) concluded that mucins from goblet cells also contribute to the expulsion of acanthocephalans from immunized hosts.
Chronic fibrosis, destruction of intestinal villi and necrotic and degenerative changes in mucosal epithelium adversely affect motility and the absorptive efficiency of the fish intestine. This might affect the general health and growth of the host. According to Bristol et al. (1984), there is a negative correlation between the number of acanthocephalans and the amount of body lipid in trout, and Buchmann (1986) demonstrated a negative correlation between the number of Echinorhynchus gadi present and energy stores in Baltic cod (Gadus morhua). Trout infected with P. laevis have lower muscle protein than uninfected fish (Wanstall et al., 1982). Seasonal factors, such as host spawning or reduced winter feeding, exacerbate the protein depletion. Wanstall et al. (1982) suggested that P. laevis induced mobilization of endogenous protein for energy in preference to endogenous carbohydrates or lipids. This phenomenon is not uncommon in fishes under stress. Further, the acanthocephalan Plagiorhynchus cylindraceus, considered of little pathological consequence, has a significant detrimental effect on the flow of food energy through the host and alters its basal metabolism (Connors and Nickol, 1991).
There are few field studies that relate acanthocephalan infections to the condition factor or survivorship of fishes. Probably this is because less fit animals are sampled less frequently as a result of more efficient predation on sick and weak hosts. Nevertheless, Bakker and Mundwiler (1999) and Sasal et al. (2001) have shown acanthocephalans to have an adverse effect on the fitness of their fish hosts.
Mechanism of disease
Several biologists have suggested that acanthocephalans secrete toxic substances that paralyse or kill their hosts or that promote other patent pathological changes, such as emaciation, discoloured viscera and prolapse of the rectum (Holloway, 1966; Fig. 5). No such substance has been isolated, however, and Schmidt et al. (1974) attributed such damage to localized toxaemia and chronic fibrinous inflammation, which result from laceration of cells at the site of attachment.
Action of the spined proboscis results in destruction of mucosal cells and penetration of the intestinal wall, with an accompanying loss of absorptive capability, impaired gut motility and sometimes perforation of the intestinal wall. Apparently this damage is exacerbated by biochemical reactions. Miller and Dunagan (1971) described a pore-like opening and groove on acanthocephalan hooks, and they postulated delivery of a secretion via the proboscis hooks. Taraschewski (1989b) demonstrated proteolytic enzymes incorporated within osmiophilic material associated with the proboscis of Paratenuisentis ambiguus in eels (Anguilla anguilla) and postulated that they are discharged from the worms through pores in the proboscis hooks. Such enzymes with trypsin-like activity secreted by P. laevis in chub (L. cephalus) degrade collagen (Polzer and Taraschewski, 1994) and thus are capable of degrading one or more of the major components of the host's gastrointestinal tissue.
In addition to aggravating damage done to cells and tissues by the proboscis, biochemical reactions might also increase loss of gut motility caused by parasite-induced fibrosis in the intestinal wall. Dezfuli et al. (2002b) found that, when infected with P. laevis, brown trout (S. trutta) have altered distribution and concentrations of calcitonin gene-related peptide, beta-endorphin, metenkephalin, neuropeptide Y, substance P, vasoactive intestinal peptide and bombesin-, cholecistokinin-8- leu-enkephalin-, and serotonin-(5-hydroxytryptamine (5-HT))-like immunoreactive cells. Most of these neuromodulators are known to control gut motility and digestive and absorptive processes. Changes in these functions probably occur in infected fishes.
Host Immune Response
The range of cellular and humoral immune mechanisms described in recent years for teleosts makes it clear that the piscine immune system is relatively well developed (Buchmann et al., 2001). Although there are few studies to demonstrate effective immunological protection against acanthocephalans, there is evidence to suggest that some degree of immunity exists.
Goblet-cell hyperplasia occurs widely in acanthocephalan infections. The covering created by copious secretion of mucus and the presumed presence of antibodies within probably reduce the number of parasites that succeed in establishing (Thomas, 2002).
Apart from a proliferation of goblet cells, increased numbers of eosinophils, neutrophils and monocytes at the attachment site characterize histopathological effects of acanthocephalans. Mobilization of leucocytes occurs regardless of whether the fish species is suitable for development of the parasite, and Hamers et al. (1992) found interspecific differences in the response of leucocytes in fishes parasitized by P. ambiguus. In eels (A. anguilla), a suitable definitive host, the response was much less intense than in carp (Cyprinus carpio) or rainbow trout (Oncorhynchus mykiss), both unsuitable hosts that expel the acanthocephalans within a few days. The leucocytes damage acanthocephalan tegument extensively in carp; hence Hamers et al. (1992) concluded that cellular defence is a factor in determining host specificity for P. ambiguus.
Plasma cells ('type B cell') occur in the inflammatory tissue around the proboscis of P. laevis in rainbow trout, and they are probably responsible for the humoral response produced by the fish (Wanstall et al., 1986). Even though immunoglobulins are relatively slow to develop in fish following an infection, and precipitins are rare (Taraschewski, 2000), antibodies precipitating to acanthocephalan antigens have been reported from sera (Harris, 1970; Szalai et al., 1988) and intestinal mucus (Harris, 1972) of infected fishes. Sera from chub (L. cephalus) held at 10°C and with no history of exposure to P. laevis do not have anti-P. laevis precipitins, but precipitins were detected within 160 days after infection. Parenteral injection of P. laevis antigen also induces a similar response by 150 days after injection (Harris, 1972). Anti-P. laevis precipitins are not found in four other piscine species in which P. laevis occurs but, unlike in L. cephalus, does not reach maturity. This led Harris (1972) to speculate that the antigenic substance is produced only by mature worms.
Despite a humoral response, chub (L. cephalus) have large numbers of P. laevis in a variety of stages of development. Harris (1972) found no evidence of expulsion of worms, even from fish with mature worms.
Concurrent infections with more than one species of acanthocephalan and/or with other helminths of other phyla are frequent. P. bulbocolli and A. dirus occur concurrently in rainbow darters (E. caeruleum), where they occupy sites in close proximity to one another. There appears to be no synergistic effect as both species cause damage as if they were in single-species infections (McDonough and Gleason, 1981). In contrast, numbers of the cestode Proteocephalus exiguus and Neoechinorhynchus sp. show an inverse relation in ciscos (Coregonus artedi), although the two helminth species occupy different intestinal sites. Cross (1934) believed that non-specific immunity limits either P. exiguus or the species of Neoechinorhynchus.
Very little is known about the nature of precipitins in serum of acanthocephalan-infected fishes, and seldom has specific immunoglobulin been demonstrated. Szalai et al. (1988), however, confirmed that anti- Neoechinorhynchus carpiodi precipitins in serum from infected quillback (Carpiodes cyprinus) were not complement-reactive protein or the alpha migrating factor. Partial characterization of chub (L. cephalus) antibody to P. laevis indicated that it is an immunoglobulin M (IgM) type (Harris, 1972). Consequently, precipitating antibody occurs in at least one acanthocephalan infection but its role in limiting infection and the degree of species specificity are not known.
Deep penetration by the proboscis results in fibrinous nodules, which may project several millimetres into the coelom (Fig. 6). Szalai and Dick (1987) reported extensive vascularization of such nodules in quillback (C. cyprinus) induced by N. carpiodi and demonstrated increased leakage of proteins from blood in the region of the nodules. They suggested that the lesion might ensure a limited, but steady, supply of nutrients for the parasites.
P. laevis is another species that induces nodule formation (Fig. 8). Polzer and Taraschewski (1994) found that the worm releases an aminopeptidase and a trypsin-like collagenase into the culture medium. They concluded that acanthocephalan aminopeptidases have a nutritional role but that the collagenase facilitates rapid and deep penetration by the worm into the intestinal tissues. This suggests that proteolytic enzymes degrade peptides at the body surface. In some species, the histolytic action permits rapid and deep penetration, leading to formation of nutrient-supplying nodules.
DiagnosisTop of page
At least some species of fish respond to acanthocephalan infections (natural and in the laboratory) by production of specific antibodies (Harris, 1972). However, there is insufficient information for immunodiagnosis. Acanthocephalans can be diagnosed from eggs passed out in the faeces of the host, but this is not convenient and there is little necessity for the technique. Most infections are detected during post-mortem examination.
EpidemiologyTop of page
A review of seasonal occurrence of acanthocephalan species in piscine hosts (Chubb, 1982) reveals many differences among species and geographical localities. Kennedy (1972) found that feeding response was an important factor in controlling the intensity of Pomphorhynchus laevis in dace (Leuciscus leuciscus) and that water temperature influenced this response. Unless negated by another factor, such as temperature-dependent rejection (Kennedy, 1972), prevalence and intensity should be greatest during times when intermediate hosts with infective larvae constitute an appropriate portion of the diet of definitive hosts. When this occurs throughout the year, there may be no seasonal periodicity in the occurrence of acanthocephalans. For example, P. laevis shows no seasonal periodicity in prevalence, intensity or development and maturation in chub (Leuciscus cephalus) in the Danube River basin of the Czech and Slovak republics where the amphipod intermediate host is available to fish throughout the year (Moravec and Scholz, 1991).
Absence of seasonal periodicity does not imply that rates of recruitment and mortality are constant throughout the year. High water temperatures (under laboratory conditions) reduce the success with which P. laevis establishes in goldfish (Carassius auratus); hence Kennedy (1972) suggested that increased feeding by fish in the summer offset a lower rate of parasite establishment. Thus, seasonal differences in rates of recruitment and mortality (from failure to establish in fish) could occur even if parasite occurrence shows no periodicity.
There are instances in which appropriate invertebrate intermediate hosts are consumed by definitive hosts throughout the year, but there are definite seasonal cycles in infections. Awachie (1965) demonstrated that, even though infective larvae of Echinorhynchus truttae occur in Gammarus pulex and the amphipods are an important part of the food of brown trout (Salmo trutta) all year, seasonal differences in densities of infective larvae lead to fluctuating occurrence in fish. Variations in habits of hosts during the year, for instance, changes in diet, also cause seasonal differences in prevalence and intensity of infections (O'Neill and Whelan, 2002).
Kennedy (1970) suggested that, when maturation of helminths from freshwater fishes shows seasonal cycles, peak egg production almost always occurs late in spring or early in summer. This generalization applies to many, but not all, species of Acanthocephala. Annual cycles in fishes frequently begin with recruitment in spring, and most transmission between fish and invertebrate hosts occurs during summer and early autumn. Development in hosts infected late in the season is retarded by low temperatures (Olson and Nickol, 1995) until spring.
Some acanthocephalans show adaptations directly related to seasonal events in the life histories of fishes. Peak egg production by Fessisentis friedi apparently occurs as pickerel (Esox americanus) move into shallow water to spawn (Muzzall, 1978). Maximum intensity and egg production of Pomphorhynchus bulbocolli in white suckers (Catostomus commersoni) have also been related to migration and spawning (Muzzall, 1980). In Lake Michigan, Echinorhynchus salmonis reaches peak sexual maturity in chinook salmon (Oncorhynchus tshawytscha) during spawning (Amin, 1978). These seasonal patterns result in peak egg production by the acanthocephalans when fishes are in shallow waters, where appropriate invertebrate intermediate hosts are most abundant.
A common type of cycle consists of spring recruitment by fishes after the environment has warmed and fish and invertebrate hosts have become active, summer transmission between fishes and invertebrates, slow development in invertebrates during the winter and completion of larval development with rising temperatures in the spring.
Deviations from the spring 'recruitment cycle' are often related to deviations in fish-invertebrate relationships and environmental disturbances. Prevalence of E. salmonis in yellow perch (Perca flavescens) in Lake Ontario rises in the autumn, after which it declines until the parasite is absent from fish during the summer (Tedla and Fernando, 1970). Watson and Dick (1979) also found that E. salmonis was most common in white-fish (Coregonus clupeaformis) from Southern Indian Lake, Manitoba, in late autumn when the amphipod intermediate host was a prominent whitefish food item.
In instances where invertebrates are rare or absent during winter, the cycle may be altered. This results in recruitment to the definitive host population in the autumn. Maturation then is slow, with egg production occurring in the spring. Eure (1976) found this for Neoechinorhynchus cylindratus in a heated reservoir in South Carolina. Recruitment into largemouth bass (Micropterus salmoides) occurs in the autumn, while ostracod numbers decrease. By November, ostracods are scarce and most acanthocephalans are in bass. Egg production is delayed until spring, when ostracods are again abundant.
Environmental disturbances might not affect host relationships of all parasites similarly. Contrary to Eure's (1976) findings, Boxrucker (1979) detected no seasonal difference in prevalence or intensity of P. bulbocolli in a thermally heated area in Lake Monona, Wisconsin. Instead, he found that, in an unheated area, this species displayed the familiar pattern of increasing prevalence and intensity during the spring followed by declining numbers in autumn. Elevated winter temperatures may have resulted in more rapid parasite development in intermediate hosts, and in fish feeding more extensively on them, than during winter in unheated environments.
ReferencesTop of page
Aloo PA; Dezfuli BS, 1997. Occurrence of cystacanths of Polyacanthorhynchus kenyensis larvae (Acanthocephala) in four teleostean fishes from a tropical lake, Lake Naivasha, Kenya. Folia Parasitologica, 44(3):233-238.
Amin OM, 1987. Key to the families and subfamilies of Acanthocephala, with the erection of a new class (Polyacanthocephala) and a new order (Polyacanthorhynchida). Journal of Parasitology, 73:1216-1219.
Amin OM; Heckmann RA; Inchausty V; Vasquez R, 1996. Immature Polyacanthorhynchus rhopalorhynchus (Acanthocephala: Polyacanthorhynchidae) in venton, Haplias malabricus (Pisces) from Moca Vie River, Bolivia, with notes on its apical organ and histopathology. Journal of the Helminthological Society of Washington, 63(1):115-119.
Awachie JBE, 1965. The ecology of Echinorhynchus truttae Schrank, 1788 (Acanthocephala) in a trout stream in north Wales. Parasitology, 55:747-762.
Aznar FJ; Bush AO; Raga JA, 2002. Reduction and variability of trunk spines in the acanthocephalan Corynosoma cetaceum: the role of physical constraints on attachment. Invertebrate Biology, 12:104-114.
Barger MA; Nickol BB, 1998. Structure of Leptorhynchoides thecatus and Pomphorhynchus bulbocolli (Acanthocephala) eggs in habitat partitioning and transmission. Journal of Parasitology, 84(3):534-537.
Barger MA; Nickol BB, 1999. Effects of coinfection with Pomphorhynchus bulbocolli on development of Leptorhynchoides thecatus (Acanthocephala) in amphipods (Hyalella azteca). Journal of Parasitology, 85(1):60-63.
Beermann I; Arai HP; Costerton JW, 1974. The ultrastructure of the lemnisci and body wall of Octospinifer macilentus (Acanthocephala). Canadian Journal of Zoology, 52:533-555.
Brand Tvon, 1939. The glycogen distribution in the body of Acanthocephala. Journal of Parasitology, 25:22S.
Brooks DR; Amato JFR, 1992. Cestode parasites in Potamotrygon motoro (Natterer) (Chondrichthyes: Potamotrygonidae) from southwestern Brazil, including Rhinebothroides mclennanae n. sp. (Tetraphyllidea: Phyllobothriidae), and a revised host-parasite checklist for helminths inhabiting neotropical freshwater stingrays. Journal of Parasitology, 78(3):393-398.
Buchmann K, 1986. On the infection of Baltic cod (Gadus morhua L.) by the acanthocephalan Echinorhynchus gadi (Zoega) Müller. Scandinavian Journal of Veterinary Science, No. 38:308-314.
Buchmann K; Lindenstrom T; Bresciani J, 2001. Defense mechanisms against parasites in fish and the prospect for vaccines. Acta Parasitologica, 46:71-81.
Bullock WL, 1963. Intestinal histology of some salmonid fishes with particular reference to the histopathology of acanthocephalan infections. Journal of Morphology, 112:23-44.
Buron Ide; Nickol BB, 1994. Histopathological effects of the acanthocephalan Leptorhynchoides thecatus in the ceca of the green sunfish, Lepomis cyanellus. Transactions of the American Microscopical Society, 113(2):161-168.
Byram JE; Fisher FM, 1974. The absorptive surface of Moniliformis dubius (Acanthocephala) II. Functional aspects. Tissue and Cell, 6:21-42.
Chaicharn A; Bullock WL, 1967. The histopathology of acanthocephalan infections in suckers with observations on the intestinal histology of two species of catostomid fishes. Acta Zoologica, 48:19-42.
Choudhury A; Dick TA, 1998. Patterns and determinants of helminth communities in the Acipenseridae (Actinopterygii: Chondrostei), with special reference to the lake sturgeon, Acipenser fulvescens. Canadian Journal of Zoology, 76(2):330-349.
Connors VA; Nickol BB, 1991. Effects of Plagiorhynchus cylindraceus (Acanthocephala) on the energy metabolism of adult starlings, Sturnus vulgaris. Parasitology, 103:395-402.
Crompton DWT, 1975. Relationships between Acanthocephala and their hosts. In: Jennings DH, Lee DL, eds. Symposia of the Society for Experimental Biology, 29. Cambridge, UK: Cambridge University Press, 467-504.
Crompton DWT, 1991. Nutritional interactions between hosts and parasites. In: Toft CA, Aeschlimann A, Bolis L, eds. Parasite-Host Associations. Oxford, UK: Oxford Science Publications, 228-257.
Crompton DWT; Keymer A; Singhvi A; Nesheim MC, 1983. Rat dietary fructose and the intestinal distribution and growth of Moniliformis (Acanthocephala). Parasitology, 86:57-81.
Crompton DWT; Lassiere OL, 1987. Acanthocephala. In: Taylor AER, Baker RR, eds. In Vitro Methods for Parasite Cultivation. London, UK: Academic Press, 394-406.
Crompton DWT; Lockwood APM, 1968. Studies on the absorption and metbolism of D-(u-14C) glucose by Polymorphus minutus (Acanthocephala) in vitro. Journal of Experimental Biology, 48, 411-425.
Crompton DWT; Ward PFV, 1967. Production of ethanol and succinate by Moniliformis dubius (Acanthocephala). Nature, 215:964-965.
Crompton DWT; Whitfield PJ, 1968. The course of infection and egg production of Polymorphus minutus (Acanthocephala) in domestic ducks. Parasitology, 58:231-246.
Cross SX, 1934. A probable case of non-specific immunity between two parasites of ciscoes of the Trout Lake region of northern Wisconsin. Journal of Parasitology, 20:244-245.
DeGiusti DL, 1949. The life cycle of Leptorhynchoides thecatus (Linton), an acanthocephalan of fish. Journal of Parasitology, 35:437-460.
Dezfuli BS; Giari L; Simoni E; Bosi G; Manera M, 2002. Histopathology, immunohistochemistry and ultrastructure of the intestine of Leuciscus cephalus (L.) naturally infected with Pomphorhynchus laevis (Acanthocephala). Journal of Fish Diseases, 25(1):7-14.
Dezfuli BS; Grandi G; Franzoi P; Rossi R, 1990. Histopathology in Atherina boyeri (Pisces: Atherinidae) resulting from infection by Telosentis exiguus (Acanthocephala). Parassitologia, 32:283-291.
Dezfuli BS; Pironi F; Giari L; Domeneghini C; Bosi G, 2002. Effect of Pomphorhynchus laevis (Acanthocephala) on putative neuromodulators in the intestine of naturally infected Salmo trutta. Diseases of Aquatic Organisms, 51(1):27-35.
Dunagan TT; Bozzola JJ, 1992. Morphology of the apical organ in Macracanthorhynchus hirudinaceus (Acanthocephala). Journal of Parasitology, 78:899-903.
Edmonds SJ, 1965. Some experiments on the nutrition of Moniliformis dubius Meyer (Acanthocephala). Parasitology, 55:337-344.
Filipponi C; Taraschewski H; Weber N, 1994. Metabolism of long-chain fatty acids, alchols and alkylglycerols in the fish parasite Paratenuisentis ambiguus (Acanthocephala). Lipids, 29:583-588.
Fried B; Stableford LT, 1991. Cultivation of helminths in chick embryos. Advances in Parasitology, 30:107-165.
Garcia-Varela M; Perez-Ponce de Leon G; de la Torre P; Cummings MP; Sarma SSS; Laclette JP, 2000. Phylogenetic relationships of Acanthocephala based on analysis of 18 S ribosomal RNA gene sequences. Journal of Molecular Evolution, 50:532-540.
Golvan YJ, 1994. Nomenclature of the Acanthocephala. Research and Reviews in Parasitology, 54, 135-205.
Golvan YJ; de Buron I, 1986. Les hôtes des acanthocephales. II - Les hôtes définitifs. 1. Poissons. Annales de Parasitologie, 63:394-375.
Hamers R; Lehmann J; Stürenberg FJ; Taraschewski H, 1992. In vitro study of the migratory and adherent responses of fish leucocytes to the eel-pathogenic acanthocephalan Paratenuisentis ambiguus (van Cleave, 1921) Bullock et Samuel, 1975 (Eoacanthocephala: Tenuisentidae). Fish & Shellfish Immunology, 2(1):43-51.
Hamers R; Taraschewski H; Lehmann J; Mock D, 1991. In vitro study on the impact of fish sera on the survival and fine structure of the eel-pathogenic acanthocephalan Paratenuisentis ambiguus.. Parasitology Research, 77(8):703-708.
Harris JE, 1970. Precipitin production by chub (Leuciscus cephalus) to an intestinal helminth. Journal of Parasitology, 56:1035.
Herlyn H; Piskurek O; Schmitz J; Ehlers U; Zischler H, 2003. The syndermatan phylogeny and the evolution of acanthocephalan endoparasitism as inferred from 18S rDNA sequences. Molecular Phylogenetics and Evolution, 26:155-164.
Hoffman GL, 1999. Parasites of North American Freshwater Fishes. 2nd edn. Ithaca and New York, USA: Comstock Publishing Associates, Cornell University Press, 539 pp.
Holloway HL Jr, 1966. Prosthorhynchus formosum (Van Cleave, 1918) in songbirds, with notes on acanthocephalans as potential parasites of poultry in Virginia. Virginia Journal of Science, New Series, 17:149-154.
Jilek R, 1979. Histopathology due to the presence of Gracilisentis gracilisentis in Dorosoma cepedianum (Le Sueur). Journal of Fish Biology, 14:593-595.
Kates KC, 1944. Some observations on experimental infections of pigs with the thorn-headed worm Macracanthorhynchus hirudinaceus. American Journal of Veterinary Research, 5:166-172.
Kennedy CR, 1970. The population biology of helminths of British freshwater fish. Symposia of the British Society for Parasitology, 9:145-159.
Kennedy CR, 1972. The effects of temperature and other factors upon the establishment and survival of Pomphorhynchus laevis (Acanthocephala) in goldfish, Carassius auratus. Parasitology, 65(2):283-294.
Kennedy CR, 1993. Introductions, spread and colonization of new localities by fish helminth and crustacean parasites in the British Isles: a perspective and appraisal. Journal of Fish Biology, 43(2):287-301.
Král'ová-Hromadová I; Tietz DF; Shinn AP; Spakulová M, 2003. ITS rDNA sequences of Pomphorhynchus laevis (Zoega in Müller, 1776) and P. lucyi Williams & Rogers, 1984 (Acanthocephala: Palaeacanthocephala). Systematic Parasitology, 56(2):141-145.
Laurie JS, 1959. Aerobic metabolism of Moniliformis dubius (Acanthocephala). Experimental Parasitology, 8:188-197.
Malta JCde O; Gomes ALS; Andrade SMSde; Varella AMB, 2001. Massive infestation by Neoechinorhynchus buttnerae Golvan, 1956 (Eoacanthocephala: Neochinorhynchidae) in young "tambaquis" Colossoma macropomum (Cuvier, 1818) cultured in the Central Amazon. Acta Amazonica, 31(1):133-143.
McAlister RO; Fisher FM, 1972. The biosynthesis of trehalose in Moniliformis dubius (Acanthocephala). Journal of Parasitology, 58:51-62.
McDonough JM; Gleason LN, 1981. Histopathology in the rainbow darter, Etheostoma caeruleum, resulting from infections with the acanthocephalans, Pomphorhynchus bulbocolli and Acanthocephalus dirus. Journal of Parasitology, 67(3):403-409.
Meyer A, 1933. Acanthocephala. In: Dr. Bronn’s Klassen und Ordnungen des Tier-reichs, Vol. 4. Leipzig,Germany: Akademische Verlagsgesellschaft, 333-582.
Miller DM; Dunagan TT, 1971. Studies on the rostellar hooks of Macracanthorhynchus hirudinaceus (Acanthocephala) from swine. Transactions of the American Microscopical Society, 90:329-335.
Miller DM; Dunagan TT, 1985. Functional morphology. In: Crompton DWT, Nickol BB, eds. Biology of the Acanthocephala. Cambridge, UK: Cambridge University Press , 73-123.
Moravec F; Scholz T, 1991. Observations on the biology of Pomphorhynchus laevis (Zoega in Müller, 1776) (Acanthocephala) in the Rokytná River, Czech and Slovak Federative Republic. Helminthologia, 28(1):23-29.
Morris SC; Crompton DWT, 1982. The origins and evolution of the Acanthocephala. Biological Reviews of the Cambridge Philosophical Society, 57:85-115.
Muzzall PM, 1978. The host-parasite relationship and seasonal occurrence of Fessisentis friedi (Acanthocephala: Fessisentidae) in the isopod (Caecidotea communis). Proceedings of the Helminthological Society of Washington, 45(1):77-82.
Muzzall PM; Rabalais FC, 1975. Studies on Acanthocephalus jacksoni Bullock, 1962 (Acanthocephala: Echinorhynchidae). I. Seasonal periodicity and new host records. Proceedings of the Helminthological Society of Washington, 42:31-34.
Nickol BB, 1972. Two species of Acanthocephala from Australian fishes with description of Arhythmacanthus paraplagusiarum sp. n. Journal of Parasitology, 58:778-780.
Nickol BB, 2003. Is postcyclic transmission underestimated as an epizootiological factor for acanthocephalans? Helminthologia, 40:93-95.
Noble ER, 1973. Parasites and fishes in a deep-sea environment. Advances in Marine Biology, 11:121-195.
Olson PD; Nickol BB, 1995. Effect of low temperature on the development of Leptorhynchoides thecatus (Acanthocephala) in Lepomis cyanellus (Centrarchidae). Journal of the Helminthological Society of Washington, 62(1):44-47.
Parshad VR; Crompton DWT; Nesheim MC, 1980. The growth of Moniliformis (Acanthocephala) in rats fed on various monosaccharides and disaccharides. Proceedings of the Royal Society of London, B209:299-315.
Read CP; Rothman AH, 1958. The carbohydrate requirement of Moniliformis (Acanthocephala). Experimental Parasitology, 7:191-197.
Reyda FB; Nickol BB, 2001. A comparison of biological performances among a laboratory-isolated population and two wild populations of Moniliformis moniliformis. Journal of Parasitology, 87:330-338.
Richardson DJ; Nickol BB, 1999. Physiological attributes of the pyloric caeca and anterior intestine of green sunfish (Lepomis cyanellus) potentially influencing microhabitat specificity of Leptorhynchoides thecatus (Acanthocephala). Comparative Biochemistry and Physiology. A, Molecular Integrative Physiology, 122(4):375-384.
Richardson DJ; Nickol BB, 2000. Experimental investigation of physiological factors that may influence microhabitat specificity exhibited by Leptorhynchoides thecatus (Acanthocephala) in green sunfish (Lepomis cyanellus). Journal of Parasitology, 86(4):685-690.
Schmidt GD, 1985. Development and life cycles. In: Crompton DWT, Nickol BB, eds. Biology of the Acanthocephala. Cambridge, UK: Cambridge University Press, 273-305.
Starling JA, 1985. Feeding, nutrition and metabolish. In: Crompton DWT, Nickol BB, eds. Biology of the Acanthocephala. Cambridge, UK: Cambridge University Press, 125-212.
Starling JA; Fisher FM, 1978. Carbohydrate transport in Moniliformis dubius (Acanthocephala). II. Post-absorptive phosphorylation of glucose and the role of trehalose in the accumulation of endogenous glucose reserves. Journal of Comparative Physiology, 126:223-231.
Stranack FR; Woodhouse MA; Griffin RL, 1966. Preliminary observations on the ultrastructure of the body wall of Pomphorhynchus laevis (Acanthocephala). Journal of Helminthology, 40:395-402.
Sures B, 2001. The use of fish parasites as bioindicators of heavy metals in aquatic ecosystems: a review. Aquatic Ecology, 35:245-255.
Sures B; Siddall R, 2003. Pomphorhynchus laevis (Palaeacanthocephala) in the intestine of chub (Leuciscus cephalus) as an indicator of metal pollution. International Journal for Parasitology, 33(1):65-70.
Sures B; Steiner W; Rydlo M; Taraschewski H, 1999. Concentrations of 17 elements in the zebra mussel (Dreissena polymorpha), in different tissues of perch (Perca fluviatilis), and in perch intestinal parasites (Acanthocephalus lucii) from the subalpine lake Mondsee, Austria. Environmental Toxicology and Chemistry, 18(11):2574-2579.
Sures B; Taraschewski H; Jackwerth E, 1994. Lead content of Paratenuisentis ambiguus (Acanthocephala), Anguillicola crassus (Nematodes) and their host Anguilla anguilla. Diseases of Aquatic Organisms, 19(2):105-107.
Szalai AJ; Danell GV; Dick TA, 1988. Intestinal leakage and precipitating antibodies in the serum of quillback, Carpiodes cyprinus (Lesueur), infected with Neoechinorhynchus carpiodi Dechtiar, 1968 (Acanthocephala: Neoechinorhynchidae). Journal of Parasitology, 74(3):415-420.
Szalai AJ; Dick TA, 1987. Intestinal pathology and site specificity of the acanthocephalan Neoechinorhynchus carpiodi Dechtiar, 1968, in quillback, Carpiodes cyprinus (Lesueur). Journal of Parasitology, 73(3):467-475.
Taraschewski H, 1989. Acanthocephalus anguillae in intra- and extraintestinal positions in experimentally infected juveniles of goldfish and carp and in sticklebacks. Journal of Parasitology, 75(1):108-118.
Taraschewski H, 1989. Host-parasite interface of Paratenuisentis ambiguus (Eoacanthocephala) in naturally infected eel and in laboratory-infected sticklebacks and juvenile carp and rainbow trout. Journal of Parasitology, 75(6):911-919.
Taraschewski H, 2000. Host-parasite interactions in Acanthocephala: a morphological approach. Advances in Parasitology, 46:1-179.
Tedla S; Fernando CH, 1970. Some remarks on the ecology of Echinorhynchus salmonis (Muller, 1784). Canadian Journal of Zoology, 48:317-321.
Thomas JD, 2003. The ecology of fish parasites with particular reference to helminth parasites and their salmonid fish hosts in Welsh rivers: a review of some of the central questions. Advances in Parasitology, 52:1-154.
Uglem GL; Read CP, 1973. Moniliformis dubius: uptake of leucine and alanine by adults. Experimental Parasitology, 34:148-153.
Valtonen ET; Koskivaara M, 1994. Relationships between the parasites of some wild and cultured fishes in two lakes and a fish farm in central Finland. International Journal for Parasitology, 24(1):109-118.
Van Cleave HJ, 1920. Acanthocephala. Report of the Canadian Arctic Expedition 1913-18 9, 1E-11E.
Van Cleave HJ, 1941. Relationships of the Acanthocephala. American Naturalist, 75:31-47.
Van Cleave HJ, 1947. Analysis of distinctions between the acanthocephalan genera Filicollis and Polymorphus, with description of a new species of Polymorphus. Transactions of the American Microscopical Society, 66:302-313.
Wanstall PW; Robotham PWJ; Thomas JS, 1982. Changes in the energy reserves of two species of freshwater fish during infection by Pomphorhynchus laevis. Parasitology, 85:xxvii.
Wanstall ST; Robotham PWJ; Thomas JS, 1986. Pathological changes induced by Pomphorhynchus laevis Muller (Acanthocephala) in the gut of rainbow trout, Salmo gairdneri Richardson. Zeitschrift für Parasitenkunde, 72(1):105-114.
Ward HL, 1940. Studies on the life history of Neoechinorhynchus cylindratus (Van Cleave, 1913) (Acanthocephala). Transactions of the American Microscopical Society, 59:327-347.
Wayland MT; Sommerville C; Gibson DI, 1999. Echinorhynchus brayi n. sp. (Acanthocephala: Echinorhynchidae) from Pachycara crassiceps (Roule) (Zoarcidae), a deep-sea fish. Systematic Parasitology, 43(2):93-101.
Yamaguti S, 1961. Systema Helminthum, Vol. 5. Acanthocephala. New York, UK: Interscience Publishers.
Young BW; Lewis PD Jr, 1977. Growth of an acanthocephalan on the chick chorioallantois. Proceedings of the Montana Academy of Sciences, 37:88.
Zdzitowiecki K, 2001. New data on the occurrence of fish endoparasitic worms off Adelie Land, Antarctica. Polish Polar Research, 22:159-165.
Zrzavy J, 2001. The interrelationships of metazoan parasites: a review of phylum- and higher-level hypotheses from recent morphological and molecular phylogenetic analyses. Folia Parasitologica, 48(2):81-103.
Aloo P A, Dezfuli B S, 1997. Occurrence of cystacanths of Polyacanthorhynchus kenyensis larvae (Acanthocephala) in four teleostean fishes from a tropical lake, Lake Naivasha, Kenya. Folia Parasitologica. 44 (3), 233-238.
Amin O M, 1978. Effect of host spawning on Echinorhynchus salmonis Muller, 1784 (Acanthocephala: Echinorhynchidae) maturation and localization. Journal of Fish Diseases. 1 (2), 195-197. DOI:10.1111/j.1365-2761.1978.tb00021.x
Amin O M, Heckmann R A, Inchausty V, Vasquez R, 1996. Immature Polyacanthorhynchus rhopalorhynchus (Acanthocephala: Polyacanthorhynchidae) in venton, Haplias malabricus (Pisces) from Moca Vie River, Bolivia, with notes on its apical organ and histopathology. Journal of the Helminthological Society of Washington. 63 (1), 115-119.
Boxrucker J C, 1979. Effects of a thermal effluent on the incidence and abundance of the gill and intestinal metazoan parasites of the black bullhead. Parasitology. 78 (2), 195-206. DOI:10.1017/S0031182000049246
CABI, Undated. CABI Compendium: Status as determined by CABI editor. Wallingford, UK: CABI
CROSS S X, 1934. Research notes.-A probable case of non-specific immunity between two parasites of ciscoes of the Trout lake region of Northern Wisconsin. Journal of Parasitology. 20 (4), 244-245. DOI:10.2307/3272467
Eure H, 1976. Seasonal abundance of Neoechinorhynchus cylindratus taken from large-mouth bass (Micropterus salmoides) in a heated reservoir. Parasitology. 73 (3), 355-370. DOI:10.1017/S003118200004703X
Knoff M, Clemente S C de S, Pinto R M, Gomes D C, 2001. Digenea and acanthocephala of elasmobranch fishes from the southern coast of Brazil. Memórias do Instituto Oswaldo Cruz. 96 (8), 1095-1101. DOI:10.1590/S0074-02762001000800012
Moravec F, Scholz T, 1991. Observations on the biology of Pomphorhynchus laevis (Zoega in Müller, 1776) (Acanthocephala) in the Rokytná River, Czech and Slovak Federative Republic. Helminthologia. 28 (1), 23-29.
Sanmartín M L, Alvarez M F, Peris D, Iglesias R, Leiro J, 2000. Parasite community study of the undulate ray Raja undulata in the Ría of Muros (Galicia, northwest Spain). Aquaculture. 184 (3/4), 189-201. DOI:10.1016/S0044-8486(99)00332-4
Sures B, Steiner W, Rydlo M, Taraschewski H, 1999. Concentrations of 17 elements in the zebra mussel (Dreissena polymorpha), in different tissues of perch (Perca fluviatilis), and in perch intestinal parasites (Acanthocephalus lucii) from the subalpine lake Mondsee, Austria. Environmental Toxicology and Chemistry. 18 (11), 2574-2579. DOI:10.1897/1551-5028(1999)018<2574:COEITZ>2.3.CO;2
Van Cleave HJ, 1920. Acanthocephala. In: Report of the Canadian Arctic Expedition 1913-18 9, 1E-11E,
Watson R A, Dick T A, 1979. Metazoan parasites of whitefish Coregonus clupeaformis (Mitchill) and cisco C. artedii Lesueur from Southern Indian Lake, Manitoba. Journal of Fish Biology. 15 (5), 579-587. DOI:10.1111/j.1095-8649.1979.tb03648.x
Zdzitowiecki K, 2001. New data on the occurrence of fish endoparasitic worms off Adelie Land, Antarctica. In: Polish Polar Research, 22 159-165.
Distribution MapsTop of page
Select a dataset
CABI Summary Records
Unsupported Web Browser:
One or more of the features that are needed to show you the maps functionality are not available in the web browser that you are using.
Please consider upgrading your browser to the latest version or installing a new browser.
More information about modern web browsers can be found at http://browsehappy.com/