Acanthocephala

Horst Taraschewski

doi:10.1007/978-3-662-43978-4_15

Acanthocephala

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Horst Taraschewski

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First Online: 23 December 2016

DOI: https://doi.org/10.1007/978-3-662-43978-4_15

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Name

Greek: acantha = thorn, cephale = head.

Classification

Phylum or lower group of Metazoa.

General Information

Adult members of the Acanthocephala are highly specialized heterosexual, intestinal parasites that take up nutrition parenterally since they have no intestine. Vertebrates are used as final (definitive) hosts, arthropods as intermediate hosts (Table 1). The body consists of two major parts, the praesoma and the metasoma. The praesoma comprises the proboscis, armed with a set of specific hooks (Fig. 1, Attachment), a more or less pronounced neck, the proboscis receptacle, and the two lemnisci (Figs. 2 and 3), which are cylindrical appendages of the praesomal tegument. The tube-shaped metasoma (= trunk) is bounded by a solid body wall, enclosing the pseudocoel, which in addition to liquid is mainly filled with male or female sexual organs.

Acanthocephala, Table 1

Some important species of the Acanthocephala

Class/species	Size (adults, mm; egg, E, μm)	Final host	Intermediate host	Paratenic host	Geographic distribution
Archiacanthocephala
Moniliformis moniliformis	m 30–45 f 140–270 E 90–125 × 50–62	Rattus spp., other rodents, occasionally humans, monkeys etc.	Cockroaches	–	Worldwide
Macracantho-rhynchus hirudinaceus	m 50–90 f 200–650 E 90–100 × 50–56	Pigs, occasionally humans, etc.	Beetles (larvae)	–	Worldwide
M. ingens	m 130–150 f 180–300 E 96–106 × 51–54	Raccoons, other mammals	Beetles	Amphibia, reptilia	North America
Prosthenorchis elegans	m 20–40 f 30–55 E 60–65 × 41–43 (78–81 × 49–53)	Monkeys, other mammals	Cockroaches, beetles	–	South America, domestic cycle worldwide
Palaeacanthocephala
Acanthocephalus anguillae	m 5–7 f 10–35 E 100–125 × 12–14	Chub, barbel	Asellus aquaticus	Small cyprinid fish^a	Europe
A. ranae	m 5–12 f 20–60 E 110–130 × 13–16	Amphibia	Asellus aquaticus	–	Holarctic
Echinorhynchus truttae	m 8–11 f 15–20 E 100–110 × 23–26	Salmonid fish	Gammarus spp.	–	Europe
Corynosoma semerme	m 3–5 f 3–5 E 79–100 × 16–29	Seals, birds, occasionally dogs, etc.	Pontoporeia affinis (Amphipoda)	Various marine fish	Holarctic
Pomphorhynchus laevis	m 6–16 f 10–30 E 110–121 × 12–19	Chub, barbel, trout	Gammaridae	Small Cyprinidae and other fish^a	Palaearctic
Filicollis anatis	m 6–8 f 10–25 E 75–84 × 27–31	Ducks, other water birds	Asellus aquaticus	–	Palaearctic
Polymorphus minutus	m 2–3 f 6–10 E 100 × 11–12	Ducks, other water birds	Gammarus spp.	–	Holarctic
Eoacanthocephala
Neoechinorhynchus cylindratus	m 4.5–8.5 f 10–15 E 51–61 × 17–28	Predatory fish (bass, etc.)	Cypria globula (Ostracoda)	Small fish (bluegills, etc.)^a	North America
N. emydis	m 6–15 f 10–22 E 20–25 × 20–22	Turtles	Cypria globula (Ostracoda)	Water snails	North America
N. rutili	m 2–6 f 5–10 E 26–27 × 14–17	Salmonid and other fish	Ostracoda	–	Holarctic
Paratenuisentis ambiguus	m 2.5–8 f 8–14 E 62–72 × 26–31	Eels	Gammarus tigrinus	–	North America, Europe (introduced)

m male, f female, E egg

^aThis host category is not yet sufficiently investigated. Thus it remains doubtful whether these species are true paratenic hosts

Acanthocephala, Fig. 1
SEMs of acanthocephalan praesomae

Acanthocephala, Fig. 2
Life cycle of common acanthocephalan species. A *Macracanthorhynchus hirudinaceus* ; B *Polymorphus minutus* . 1 The adults live in the intestine of their final hosts, being attached by their hooked proboscis. The penetration of the intestinal wall leads to inflamed protrusions (IP) appearing along the outer side. 2 After copulation the adult females excrete eggs for several months (patent period). These eggs are passed fully embryonated (i.e., they contain the hooked acanthor larva) with the faeces of the host. 3–6 Intermediate hosts (*Gammarus* spp. or beetle larvae) become infected by ingesting infective eggs. Inside the intestine the acanthor is released from the egg (4), enters the body cavity, and is transformed into an acanthella larva (5). The latter grows up within 60–95 days (in *M. hirudinaceus*) and is described as an infective larva ( Cystacanth). Infection of the final hosts occurs when they swallow infected intermediate hosts. The young worms reach sexual maturity within 60–90 days in *M. hirudinaceus* (after 20 days in *Polymorphus minutus*) and start egg production (= end of prepatent period). AC, acanthor; BH, body hooks; IP, inflamed protrusion of IW; IW, intestinal wall; PH, proboscis hooks; RA, released acanthor

Acanthocephala, Fig. 3
Life cycle of two common acanthocephalan species parasitizing fish. A *Neoechinorhynchus rutili* ; B *Acanthocephalus anguillae* . 1 Adults are attached to the intestinal wall of their final hosts, trout (A) or chub and other fish (B). 2 Fully embryonated eggs are passed with host’s faeces. 3–6 Intermediate hosts (A ostracod crustaceans, BAsellus aquaticus) are infected by uptake of eggs. Inside their intestine the acanthor larva (4) is released from its eggshell, enters the body cavity and becomes transformed into the acanthella larva (5). This stage differentiates to the infective larva without encystation in about 30–60 days (6) depending on outer conditions. Final hosts are infected by swallowing intermediate hosts. In *A. anguillae* a paratenic host may also become involved. When bleaks and some other fish ingest intermediate hosts (*Asellus aquaticus*), the infective larva enters the fish viscera, but there is no further development, but quick degeneration. *Neoechinorhynchus rutili* and *A. anguillae* reach sexual maturity in about 20–30 or 40–60 days, respectively (prepatent period). Adults live only for about 2–3 months (patent period). AC, acanthor; IP, inflamed protrusion of IW; IW, intestinal wall; LM, lemniscus; PH, proboscis hooks

Additional morphological features as well as biological characteristics determine the affiliation to one of the classes.

System

Class 1: Archiacanthocephala Meyer 1931: species have terrestrial life cycles; mammals or birds are final hosts, and insects (or millipedes) intermediate hosts; in addition, paratenic hosts are often involved; main longitudinal vessels of the lacunar system run dorsally and ventrally; usually there are eight uninucleate cement glands; few tegumental nuclei; ligament sacs inside the pseudocoel, also in adult worms (Fig. 3). The well known orders are:

Order: Apororhynchida
Order: Gigantorhynchida
Order: Moniliformida
- Family: Moniliformidae
  
  Genus: Moniliformis
Order: Oligacanthorhynchida
- Family: Oligacanthorhynchidae
  
  Genus: Macracanthorhynchus
  
  Genus: Prosthenorchis

Class 2: Palaeacanthocephala Meyer 1931: mostly aquatic life cycles; fish (waterbirds, seals) are final hosts, crustaceans intermediate hosts; main vessels of the lacunar system run laterally; two to eight multinucleate cement glands; numerous tegumental nuclei; ligament sacs ruptured in adult worms.

Order: Echinorhynchida
- Family: Echinorhynchidae
  
  Genus: Acanthocephalus
  
  Genus: Echinorhynchus
- Family: Pomphorhynchidae
  
  Genus: Pomphorhynchus
Order: Polymorphida
- Family: Centrorhynchidae
- Family: Plagiorhynchidae
- Family: Polymorphidae
  
  Genus: Corynosoma
  
  Genus: Filicollis
  
  Genus: Polymorphus

Class 3: Eoacanthocephala Van Cleave, 1936: aquatic life cycles; fish (also reptiles, amphibians) are final hosts, and small crustaceans (mostly Ostracoda) intermediate hosts; main vessels of the lacunar system run dorsally and ventrally, only a single, giant uninucleate cement gland; tegument with giant nuclei; ligament sacs generally persistent in adults.

Order: Gyracanthocephala
Order: Neoechinorhynchida
- Family: Neoechinorhynchidae
  
  Genus: Neoechinorhynchus
- Family: Tenuisentidae
  
  Genus: Paratenuisentis

Recently, a fourth class has been erected, the Polyacanthocephala. The few known members of this group have been little studied. Fishes and crocodiles were found to be parasitized.

Important Species

Table 1.

Life Cycles

Figures 4 and 5.

Acanthocephala, Fig. 5
Light micrograph of a longitudinally sectioned (paraffin section) anterior body of *Acanthocephalus lucii* attached to the intestinal wall of a river perch (*Perca fluviatilis*). Only the intestinal mucosa has been ruptured. A thin layer of collagen fibres (*arrow*) interiadly lining this epithelium was not perforated. Also note the proboscis cavity at the anterior tip of the worm showing the normal condition of the proboscis of a non-perforating species. ×20

Attachment

Concerning the attachment to the host’s intestinal wall, two groups can be distinguished: perforating and non-perforating acanthocephalans.

Non-perforating Acanthocephalans

Generally, acanthocephalans that have a short neck do not penetrate deeply into the host’s intestinal wall with their praesoma, but display some mode of shallow attachment, i.e., they do not create lesions reaching as deep as into the muscular layers of the intestinal wall (Figs. 2, 6, and 7, Acanthocephalan Infections, Fig. 8). Accordingly, often even the posterior half of the proboscis does not become surrounded by host tissue (Fig. 6). Layers of connective tissue within the hosts’ intestinal wall often appear to function as penetration obstacles. This might be the stratum compactum in salmonids retaining Echinorhynchus truttae in superficial positions or a collagen layer interiadly lining the intestinal mucosa of perch (Perca fluviatilis) affecting the mode of attachment of Acanthocephalus lucii (Fig. 6). On the other hand, the tipped proboscis hooks seem to use the collagen layers as suitable substrates of anchorage (Fig. 7). In Fig. 7 a necrotic tissue with a slight infiltration of granulocytes and haemorrhagic involvement, typical of the attachment site of Acanthocephalus lucii and other non-perforating species, is shown (Fig. 2). When non-perforating species were experimentally inoculated into small specimens of fish not comprising penetration obstacles in their gut wall, three non-perforating species did not try or succeed in perforating either. And accordingly such species usually cannot be found in toto in extraintestinal locations like perforating species. Paratenuisentis ambiguus and P. lucii, both non-perforators, do not possess colagenolytic proteinases useful in chemical support of penetration activity. So paratenic hosts do not occur in the life cycles of non-perforating acanthocephalans, but postcyclic transmission of intraintestinal worms like Neoechinorhynchus rutili in sticklebacks to predatory brown trout seems to be very common.

Acanthocephala, Fig. 6
TEM of a transversally sectioned proboscis hook of *Acanthocephalus lucii* surrounded by necrotic and inflamed intestinal tissue of *Perca fluviatilis*. The hook: note the perforations in its striped layer (SL); the connective tissue of the hook does not reach into the tip sectioned here. The hook has punctually perforated the layer of subepithelial connective tissue (SC) that can be seen in Fig. 6 at a lower magnification (*arrow*). But the proboscis *in toto* did not perforate this layer (LA: lamina of fine amorphous material lining the mucosal epithelium). The proboscis (not seen) is either in front or behind the plane of the section. ×2,000

Acanthocephala, Fig. 7
(a–c) Micrographs showing the proboscis tip and hooks of various acanthocephalans. (a) Longitudinal semithin section of *Paratenuisentis ambiguus* that had been exposed *in vitro* to [³H]-glyceroltrioleate for 5 min. The proboscis is fully everted, which is not the normal condition *in vivo*. Note the intense label of the “apical organ” (AO). The lipid that has been taken up by the hooks and the surrounding tegument (PT) seems to be transported along the outer membrane and the basal membrane (BM) of the tegument; CH, connective tissue of a hook; PR, proboscis retractor muscle. ×3,500. (b) SEM of the tipped hooks at the anterior proboscis of *Acanthocephalus anguillae*. ×1,600. (c) Longitudinal semithin section of a proboscis hook of *A. anguillae* that has been exposed to [³H]-labeled lipids. Note the labelled tegument of the hook underneath its striped layer. The proboscis tegument (PT) has already absorbed huge quantities of the lipid. ×3,000. CH, connective tissue of the hook

Such species either continuously or occasionally change their point of attachment, exposing them to the posteriadly directed intestinal peristalsis. In infrapopulations of E. truttae in brown trout all specimens have arrived at the posterior end of the small intestine by the time the worms have matured. As has been shown for N. cylindratus, a species that is potentially perforating, infrapopulations with high worm densities lead to enhancement of change of the point of attachment and consequently to greater posterior shift. An interesting feature can be observed in other neoechinorhynchids. Although they occupy superficial positions, they do not seem to migrate or become shifted after an initial period of establishment, due to a firm capsule of collagen fibres enclosing their small, roundish proboscis. A negative point of the proboscis remains in the intestinal wall after deattaching a worm using forceps ( Acanthocephalan Infections, Fig. 5A). Not unlikely, this massive collagen formation is provoked by the excretion of proline ( Amino Acids) or other substances by the praesoma.

A typical non-perforating species is the archiacanthocephalan Moniliformis moniliformis displaying a deep proboscis cavity and shallow attachment ( Acanthocephalan Infections, Fig. 8).

Perforating Acanthocephalans

Many Acanthocephalans possess a long neck which may comprise a bulbus such as the Pomphorhynchus spp. (Palaeacanthocephala) with an inflated neck region (praesoma) (Fig. 1). In the eoacanthocephalan Eocollis arcanus it is the anterior part of the metasoma which forms a bulb. In both cases the bulbus functions as a dowel enabling the worm to occupy a permanent point of attachment at a specific site. The deep and quick perforation of the intestinal wall may be supported by a proteolytic enzyme as shown for Pomphorhynchus laevis which excretes a trypsin-like proteinase into the culture medium. It has a collagenolytic activity and the molecular mass differs slightly among infectious larvae and adult worms removed from fish. The long-necked species A. anguillae does not display such abilities in lysing collagen and accordingly the collagenic stratum compactum of salmonid fishes retains most worms in rather superficial connections with the intestinal wall. In experimental infections in adult rainbow trout the worms take about 60 days to perforate the stratum compactum, in juveniles of the same salmonid host it occurs around 20–30 d.p.i. In the long run, only those worms which succeed in penetrating seem to survive for several months in this host, while the others probably do not have the potential to withstand the intestinal drift. As shown for P. laevis, typical perforating species do not change their site of attachment and thus do not become backwards shifted. In natural populations of fish hosts, species like P. laevis, Eocollis arcanus, or A. anguillae are not only found in positions with a praesoma deeply inbedded inside the intestinal wall, but also partly lying in toto inside the peritoneal cavity or viscera, especially in small specimens or species. Obviously, in such hosts the intestinal wall or the collagen layers within it are not strong enough to withstand the worms’ penetrating activity. In juveniles of goldfish experimentally infected with A. anguillae, the first worms started projecting into the peritoneal cavity with parts of their praesoma up from about 10 d.p.i., worms of about 20–30 d.p.i. were mostly found in various positions like lying with parts of their bodies inside one intestinal loop and projecting into another with the proboscis or anterior body ( Acanthocephalan Infections, Fig. 5). In contrast, at 50 d.p.i. all worms recovered had taken intraperitoneal positions and most of them were already degenerating. Due to this quick death of the worms in extraintestinal positions, one may conclude that they did not leave the intestine “on purpose” but slipped into the peritoneal cavity in toto by lack of penetration obstacles or other features. Thus, the small fishes that became infected in these experiments should not be called paratenic hosts. However, true paratenic hosts exist in the life cycles of certain perforating acanthocephalans. Oncicola pomatostomi, for instance, is parasitic in the intestine of Felidae and Canidae in Southeast Asia and Australia while it has been found under the skin of 19 species of passerine birds where it probably has a certain longevity making the birds true paratenic hosts.

Often Macracanthorhynchus hirudinaceus occupies extraintestinal positions in humans. The migration of this perforating acanthocephalan through the gut wall is very painful. Such infected humans with an intraperitoneally located worm might be named accidental hosts since they do not play a role in the transmission of the acanthocephalan.

Among perforating species a proboscis cavity is formed mainly during the early phase in the final host when the worm has not yet penetrated, later on, the cavity’s depth and frequency of invagination are progressively reduced ( Acanthocephalan Infections, Fig. 5).

Food Uptake

In non-perforating acanthocephalans the proboscis itself is usually kept in a more or less invaginated condition creating a deep proboscis cavity (Fig. 2a) which obviously functions as a funnel collecting remnants of cells and nutrients leaking into the lesion that has been created by the worm. In eo- and palaeacanthocephalans, especially lipid substances such as triacylglycerols are highly abundant as storage lipids in the intestinal wall of fish, ducks, or seals serving as final host. As shown in Fig. 8 lipid matter as well as, for instance, peptides deriving from the granules of eosinophilic granulocytes contribute to the efflux from the necrotic tissue. However, autoradiographic studies by Taraschewski and Mackenstedt with two species of eoacanthocephalans and four palaeacanthocephalans (two non-perforating and two perforating species) show that predominantly lipid substances are absorbed at the worms’ praesoma (Figs. 9 and 10). The “apical organ” of eoacanthocephalans, a structure not yet well understood at the tip of the proboscis, i.e., at the bottom of the proboscis cavity (Fig. 9), and the tegument of the anterior half of the proboscis (Fig. 10) play the most active role in lipid uptake. Interestingly, the proboscis hooks, too, can be considered organs well adapted to the task of lipid uptake (Figs. 9 and 10). In accordance with their tapered and tipped construction (Fig. 9b) the hooks are in reach of lipid deposits which are not (yet) in contact with the surface of the praesoma. Behind the septum between praesomal and metasomal tegument the uptake of a triacylglycerol as well as of vitamin A was very low in in vitro trials. However, if “shoulders” of the metasoma were in contact with the praesomal surface during the exposition of a worm to the labelled nutrient, the shoulders too revealed a markable label (Fig. 10), suggesting that enzymes localized at the praesomal surface were involved. Uptake of amino acids as well as monosaccharides also occurs at the surface of the praesoma (Fig. 11), but the metasomal tegument seems to be the major absorptive surface for these substances.

Acanthocephala, Fig. 8
Longitudinally cut semithin section of the anterior body of an *Acanthocephalus lucii* that had been *in vitro* exposed to [³H]-vitamin A for 15 min. Note the intense label of the anterior half of the proboscis tegument which lines the proboscis cavity normally formed. Behind the septum between the praesomal and the metasomal tegument (*black arrows*) almost no absorbance of the substance offered has taken place. However, it appears that the accidentally formed “shoulder” of the metasoma that obviously was in contact with the praesoma during the exposure has attained some label in its tegument (*white arrows*), suggesting that this part of the metasoma took advantage of enzymatic activity prevailing at the praesomal surface. ×60

Acanthocephala, Fig. 9
Autoradiographically treated longitudinal section of the anterior body of a female *Echinorhynchus truttae* that was exposed to [³H]-lysine for 8 min. Note the less intense label of the presomal tegument (PT) compared to the metasomal tegument. ×100. SE, septum between the presomal and the metasomal tegument (compare Fig. 13b); LA, tegumental lacunar system; LE, lemniscus; N, nucleus

Acanthocephala, Fig. 10
Semithin section of the proboscis of a specimen of *Neoechinorhynchus rutili* attached to the gut wall of a naturally infected juvenile rainbow trout. Note the conspicuous quantities of lipid (LI) accumulating at the worm’s proboscis. They seem to derive from the surrounding necrotic tissue. Also, fused granules from degranulated eosinophilic granulocytes (*arrow*) are abundant, fusing with the lipid drops. In such a methylene-blue-stained section the lipid attains a greenish-golden colour while the fused granules are a deep blue. Thus both substances can be distinguished well. ×100. AO, apical organ

Acanthocephala, Fig. 11
Transmission electron micrograph of a section through the distal part of the praesomal tegument of *Acanthocephalus anguillae* (in a rainbow trout, 90 d.p.i.). Note the thick lipoid surface coat (SC) between the tegument and the necrotic tissue (NT) at the point of attachment, also the fused crypts of the outer membrane (FC) with supporting microfibres in it and the underlying feltwork layer (FL). A section with little osmiophilic content of the fused crypts was chosen in order to show the microfibres in it. ×25,000

Concerning the uptake of nutrients by adult perforating species, the mechanisms do not basically deviate from those described for non-perforating species. Since the whole praesoma is deeply embedded in the gut wall, a funnel for substances leaking into intestinal lumen is not very large. Intraintestinally attached worms that have a bulbus can be easily recognized at the gut’s exterior side ( Acanthocephalan Infections, Fig. 6) but also in species without a bulbus, like M. hirudinaceus, a fibrous whitish nodule with reddened annulation around it can be seen.

Integument

The tegument of acanthocephalans is a syncytium of up to 2 mm in thickness (M. hirudinaceus). It either contains numerous small nuclei (Fig. 12c) or specific numbers of giant nuclei in eoacanthocephalans (Table 1). The nuclei of the tegument of the metasoma (trunk) are not immersed below the tegument (Fig. 12c). In the praesoma (proboscis and neck), however, the nuclei are harboured by the lemnisci; sack-shaped outgrowths of the praesomal tegument projecting into the body cavity (Fig. 3). The tegument is supported by underlying fibres of connective tissue, partly identified as collagen, of equal thickness in all parts of the body, and by cords of circular (only in the metasoma) muscles and longitudinal muscles (in both parts of the body; Fig. 12c). These components together build up the body wall (Fig. 12c). The tegument shows a typical stratification and a differentiation related to the praesoma-metasoma organization of the acanthocephalan body.

Metasoma

The syncytial tegument of the Acanthocephala is delimited by a plasma membrane carrying a filamentous surface coat (Fig. 13a) which has a similar appearance in all systematic groups of these worms regarding the surface of the metasoma (Fig. 13, Table 1). This glycocalyx also covers the pores (openings of the outer membrane’s crypts) of the tegument. It may reach a thickness of up to 1 μm or more and obviously proteoglycanes are present in it. Infective larvae inside the intermediate host’s hemocoel carry the most conspicuous surface coat. The plasma membrane itself forms densely set crypts projecting into the outer part of the tegument. Their greater density in the metasoma compared to the praesoma might have to do with the heavy competition pressure between the trunk surface being located inside the gut lumen, and the host’s intestinal mucosa for the absorbance of nutrients. The crypts have been calculated to increase the worm’s “outer” surface 20- to 80-fold. The crypts have slender necks with electron-dense annulations underneath their outer openings (seen as pores by SEM) and they form branches directly underneath the pores or further inside the tegument. The crypts are considered extracytoplasmic digestive organelles which under the influence of surface hydrolytic enzymes maximize the opportunities of food absorption by these gutless worms. So it might be a point of debate whether the membrane limiting the lumen of the crypts really is an “outer” surface (Fig. 13).
Interiadly, the outer membrane is supported by an electron-dense layer of about 5 μm in thickness. Due to perforations of this “ cuticle” by the crypts, this layer obviously having stabilizing functions is seen as a striped layer in TEM micrographs (Fig. 13). Usually the longitudinal extension of the membrane-crypts considerably exceeds the diameter of the striped layer, forming a spongy belt rich in pinocytotic activity (Fig. 13). This layer is considered a “ vesicular layer” by a few authors.
The feltwork layer adjoining underneath again seems to contribute to the skeletal task of the tegument but its diameter is about five times larger than that of the striped layer ( Acanthella, Fig. 1, showing this feature in the praesoma). It is characterized by fibres displaying no particular order, and normally it contains large amounts of glycogen. Metasomal spines, present in many palae- and eoacanthocephalans (Table 1), have been described as outgrowths of the feltwork layer and are thus invested by a thin cover of a somewhat condensed striped layer. These trunk spines, often ornamenting a wider part of the ventral surface than of the dorsal side, are thought to act as additional holdfast organs.
The more proximal radial layer (Fig. 12c) occupies more than 70 % of the tegumental diameter. This layer is considered to be the main metabolic centre of the acanthocephalan body. During the (allometric) growth of the worms it is mainly the increase in diameter of the radial layer which leads to a thicker tegument. This layer is characterized by radially arranged fibres connected to the basal membrane. Electron-dense matter accumulates around the spokes (Fig. 12c). This layer harbours the tegumental nuclei (Fig. 12c) and the major canals of the lacunar system (Fig. 14c). The lumen of this system is poor in organelles and electron-dense matter (Fig. 14c) and in autoradiographic trials it takes up and/or retains less nutrients than the surrounding cytoplasm. Also its glycogen content is low in contrast to the true radial layer, which usually stores plenty of this carbohydrate. Apparently contractions of the subtegumental musculature (Fig. 12c) act as the motive force for fluid flow inside the lacunas. However, it remains unclear whether these caverns fulfil functions of a circulatory system.
The basal membrane, interiadly bounding the tegumental syncytium, shows a typical labyrinthine structure. Along its distal surface it is supported by amorphic matter, whereas a thin lamina of a fine matrix lines the proximal side of the membrane (Fig. 12c).

Acanthocephala, Fig. 13
(a–c) TEMs of sections through the proboscis tegument and hooks of the eoacanthocephalan *Paratenuisentis ambiguus*. (a) The curved hook which is retracted in this micrograph is sectioned twice, at its connection with the subtegumental connective tissue (CH, connective tissue of the hook) and at its tipped outer portion. Note the lipoid substance (LI) being excreted through the pores in the hook, which (or a similar substance) is also abundant in the surrounding tegument; SC, connective tissue of the presomal tegument; ER, rough endoplasmic reticulum. ×10,000. (b) Cross section through a retracted hook and neighbouring proboscis tegument. The section has been treated according to the electron microscopical PAS-staining method by Thiéry. Note the mucus-like carbohydrates inside (CM, crypt of the hook’s outer membrane) and outside the hook (*arrow*). ×56,000. (c) TEM of the basal part (radial layer) of the metasomal tegument of an adult *Acanthocephalus anguillae*. Note the radially arranged fibres (RF) with fibrous, electron-dense material (FM) attached to them, the labyrinthine basal membrane (LB) and the cords of subtegumental musculature (CM, circular muscle; LM, longitudinal muscle) consisting of an outer myogenic portion (MP) and an inner non-contractile portion (CP); BL, basal lamina; LD, lipid drop; N, nucleus; NU, nucleolus; SC, subtegumental connective tissue. ×50,000

Acanthocephala, Fig. 14
(a, b) Micrographs of acanthocephalan muscles of *Paratenuisentis ambiguus* showing their bi-component construction comprising an outer myogenic belt (MP) and an enclosed cytoplasmic portion (CP). (a) Longitudinally ultrathin-sectioned subtegumental muscle of an infectious larva. The section has been treated according to the electron microscopical PAS method of Thiéry in a mode to visualize glycogen. Note the intense Thiéry label in the core of the muscle; LD, lipid drop; SC, subtegumental connective tissue; SE, septum. ×260. (b) Transversally semithin sectioned body cavity of an adult worm that was exposed to ³H-glucose and then autoradiographically treated. The intense label in the cytoplasmic portion of the receptacle retractor muscles (RR) seems to be due to glucose-metabolites (probably mainly glycogen) incorporated in these muscles; CK, knobs on the muscle’s surface also showing the bi-portion structure; E, egg; LS, ligament strand; OB, ovarian ball; SL, subtegumental longitudinal muscle. ×300. (c, d) Semithin sections of the praesoma (and partly the metasoma: c) of adult acanthocephalans. (c) Longitudinal section showing the cytoplasmic “finger” (CP) of the (inner) receptacle wall projecting into the proboscis. Also note the lacunar system (LS) inside the metasomal tegument (MT); SE, septum between praesomal and metasomal tegument. ×30. (d) Transverse section through the receptacle wall musculature (IW, inner wall; OW, outer wall) of *Acanthocephalus lucii*; note the tubular structure of the proboscis retractor musculature with its cytoplasmic cores (CP) of low density. ×150.

Praesoma

The tegument of the praesoma is separated from that of the trunk by a septum composed of fibres and adherent amorphic matter (Fig. 13b). So even the lacunar cavities are part of two different systems, which might make sense considering the assumed involvement of hydrostatic pressures in the protrusion, invagination, and retraction of the proboscis or the entire praesoma. Within the praesoma mainly the neck possesses lacunar cavities. The praesomal tegument reveals major differences compared to that of the metasoma and these features become more prominent towards the anterior part of the proboscis. Generally, the praesomal tegument contains more amorphous, electron-dense matter, more mitochondria, and rough and smooth endoplasmic reticulum (Fig. 12a) as well as lipid (especially in eo- and palaeacanthocephalans, Figs. 12a and 15) than the metasomal tegument. Interestingly, a submersion of the tegumental nuclei only occurs in the praesoma. The lemnisci harbouring the nuclei (Fig. 3) do not show a specific stratification like the tegument they branch away from. They too contain lacunar spaces, and are very rich in lipid.

Acanthocephala, Fig. 15
(a–c) (a) Schematic drawing of an eoacanthocephalan proboscis hook and surrounding tegument. Note the tegumental cover of the hook’s connective tissue (CH), and the crypts of the outer membrane (CM) entangling with “crypts” of the tegument’s basal membrane externally lining the connective tissue of the hook. These crypts perforate a layer of amorphic matter (AM) covering the connective tissue of the hook’s tegument. LC, lipoid coat on the hooks and on the tegument; PM, finger-shaped protuberance of the subtegumental musculature; SC, subtegumental connective tissue. (b) Schematic drawing of a palaeacanthocephalan proboscis hook and surrounding tegument. Note the tegumental cover of the hook’s connective tissue (CH), the lipoid coat (LC) on the tegument and the hook, the fused crypts of the outer membrane (FC) supported by fibres, and the fused crypts of the basal labyrinth (FB) which possibly are continuous with the latter fused crypts. PM, finger-shaped protuberance of the subtegumental musculature; SC, subtegumental connective tissue. (c) Schematic drawing of an archiacanthocephalan proboscis hook and surrounding tegument. Note that the connective tissue of the hook (CH) has no tegumental cover and is invested only by a lipoid coat (LC) which is discharged by the tegument into the pouch surrounding the hook. The proboscis tegument carries a fuzzy glycocalyx (GL), and the crypts of the outer membrane (CM) are not fused. PM, finger-shaped protuberance of the tegumental musculature; SC, subtegumental connective tissue

The surface coat of the praesomal tegument reveals systematics-related specificities (Fig. 15) and shows interesting links with the host–parasite interactions ( Acanthocephalan Infections). The fine structure and obviously also the chemical composition of the surface coat vary among the classes. Regarding archicacanthocephalans the optical impression of the praesomal glycocalyx resembles that of the metasoma, although it is more coarsely structured and more osmiophilic than the latter. Shedding of the surface coat frequently or often occurs and seems to follow a complexation of host’s anti-parasitic enzymes or antibodies with the surface coat ( Acanthocephalan Infections, Fig. 5A). Eoacanthocephalans and palaeacanthocephalans reveal a lipoid, non-fuzzy surface coat which may reach a thickness of several microns (Fig. 16) and shows a matrix which suggests a liquid or semiliquid condition. In addition to lipid, mucus-like carbohydrates are also present in it. Often osmiophilic films, perhaps representing a “glycocalyx,” can be seen in it, and it is rather likely that these films are shed into the voluminous coat once the outer membrane has become loaded with anti-parasitic peptides of the host’s defense system (Fig. 13b). Unfortunately, the chemical properties of the acanthocephalan surface coat have not been extensively studied to date.

Acanthocephala, Fig. 16
(a) DR of a female acanthocephalan (*Paratenuisentis ambiguus*, Eoacanthocephala) with emphasis on the sexual organs (most muscles omitted). It has been reduced in size (length) compared with the male worm. The ventral ligament sac leading into the uterine bell is specific to eoacanthocephalans. Note the lack of genital ganglia (GA) in the female worm. For the inscriptions of the non-sexual organs see (b). Eggs and floating ovaries inside the ligament sacs are not shown (Fig. 18a). (b) DR of a male acanthocephalan (*Paratenuisentis ambiguus*, Eoacanthocephala) with emphasis on the sexual organs. Most muscles are omitted. The single polynucleate cement gland and the presence of a cement reservoir and a seminal vesicle are specific to eoacanthocephalans. AS, apical sensory organ; BC, bursa copulatrix (evaginated); CG, cerebral ganglion; DL, “dorsal” ligament sac; ES, egg-sorting apparatus; G, genital opening; GS, Saefftigen’s pouch; HO, testes; LE, lemnisk; M, muscle; N, giant nucleus; PR, proboscis with hooks (evaginated); SB, seminal vesicle; SP, sphincter; TE, tegument; UG, uterine bell; UT, uterus; VL, “ventral” ligament sac; ZD, cement gland; ZR, cement reservoir

The (pores of the) praesomal crypts are less densely set and the striped layer measures half or less in diameter than the trunk surface. In Palaeacanthocephala the single crypts are fused underneath the striped layer, forming large caverns with stabilizing fibres in them (Fig. 15b). The other systematic groups have retained their individual crypts (Fig. 15a, c). Generally, the strata of the tegument as described from the trunk cannot be well distinguished: due to the abundance of fibres they often all together appear like a feltwork layer. The metasomal labyrinthine structure of the basal membrane is considerably reduced and instead its coating with amorphous material is more pronounced.

Proboscis Hooks

Irrespective of the systematic affiliation of the worms the hooks (Figs. 9b, 12a, b, and 15) possess a central cone of connective tissue which partly has been demonstrated to contain collagen and/or chitin. This major part of the hook arises from the subtegumental connective tissue. In its proximal part it encircles a finger-like projection of the subtegumental longitudinal musculature (Fig. 15). But this musculature tie is not present in all hooks of all species, implying that not all hooks can be individually retracted.

In eo- and palaeacanthocephalans the fibrous core of the hooks carries a condensed tegumental cover making these holdfast organs pointed (Figs. 12 and 15a, b). The striped layer does not markably differ between eo- and palaeacanthocephalans but in eoacanthocephalans the crypts are not fused but entangle with finger-form protrusions of the tegument’s basal membrane inside the hooks (Fig. 15a). In both of these subclasses the tipped hooks are capable of discharging lipid substances through their pores (Fig. 12a). Mucus-like carbohydrates also contribute to the excreted matter (Fig. 12b) and, rather likely, enzymes are also contained in it which thus far can only be hypothesized. The grease-like surface coat of the hooks may be very voluminous (Fig. 15). Amazingly, however, the hooks are also capable of absorbing nutrients from the host tissue surrounding them.

In archiacanthocephalans the hooks do not bear a tegumental vestment and the naked cone of the connective tissue, thus piercing the tegument, is less pointed than the hooks of the two other systematic groups (Fig. 15c). Obviously, the hooks attain their slippery surface cover by dipping into a pit encircling them; this annular cleft filled with a highly osmiophilic lipoid paste deriving from the surrounding tegument (Fig. 15c).

Uniformly in all systematic groups of the Acanthocephala, the close tegumetal surrounding of the hooks is rich in lipid droplets, mitochondria, and rough (Fig. 12a) as well as smooth endoplasmic reticulum, indicating elevated metabolic activity.

Proboscis Cavity

Contrary to the way in which acanthocephalans are usually shown in drawings made from dead worms (Fig. 3), in vivo and in situ the acanthocephalan proboscis is normally kept in a semi-invaginated position, especially among species with superficial attachment (Fig. 2). Thus a proboscis exhibiting a more or less deep anterior cavity resembling a mouth opening should be part of our idea of these gutless worms. Among eo- and palaeacanthocephalans inside the proboscis cavity the tegumental surface including the hooks appears as a labyrinth with remnants of host cells and tissue between its curves and with grease occupying all external niches of the labyrinth at its bottom plane.

In Archiacanthocephala the proboscis cavity is not filled with grease in its inner part and the labyrinth is lined by the fuzzy glycocalyx described under praesoma.

Musculature

Relatively few investigations have dealt with the fine structure of the acanthocephalan musculature. Some muscles, such as the receptacle retractor muscles, appear obliquely striated, e.g., fibres are connected to Z-line-like structures.

The basic feature of the acanthocephalan musculature is its 2-component structure composed of an outer myogenic, contractile belt and a cytoplasmic core enclosed by it (Figs. 12c and 14). The interior part seems to have a function in energy storage.

Usually glycogen is very abundant in it (Fig. 14a) and in autoradiographic experiments with labelled glucose, the glucose, or more likely metabolites of it like glycogen, accumulates in the cytoplasmic core (Fig. 14b). The cords of subtegumental musculature follow this bi-component composition (Fig. 12c), as do the retractor muscles (Fig. 14b–d). In addition the receptacle retractor muscles also carry small knobs on their surface which have non-contractile cores (Fig. 14b). In all these muscles the central non-contractile portion may contain plenty of organelles, mainly mitochondria, or may be rather electron-lucent, suggesting a higher fluidity than the latter cytoplasm. Inside the proboscis retractor musculature a low viscosity core should enable a quick directional shift of the enclosed cytoplasm when the proboscis cavity is formed or discontinued. An interesting differentiation is shown by the proboscis receptacle musculature enclosing and thus forming the hollow into which the proboscis can be retracted. In palaeacanthocephalans it consists of a double wall which has almost no non-contractile portion (Fig. 14d). In eoacanthocephalans it is considered single-walled but the “receptacle protrusor musculature” exteriadly surrounding the receptacle without being firmly connected to it probably represents the outer wall of the receptacle. It reveals the described 2-portion structure. The inner wall basically consists of a firm, contractile wall but on its dorsal inner side a conspicuous sack-shaped cytoplasmic outgrowth with a very narrow contractile outer cover projects into the posterior half of the proboscis (Fig. 14c). The cytoplasmic finger seems to function as the major glycogen deposit of the praesoma. In archiacanthocephalans the inner wall consists of plane myogenic tissue whereas the outer wall is formed by spirally arranged single muscle cords with non-contractile cores. Due to this spiral arrangement the retraction and protrusion of the praesoma (not only the proboscis can be invaginated) are performed in a torsion-like, screwing fashion.

Excretory System

Excretory products of most acanthocephalans seem to be released exclusively through the body wall, but it is not known whether this takes place through the whole tegument or through special regions. In addition, oligacanthorhynchids and probably other archiacanthocephalans have protonephridia. Their efferent canals either lead into the vas deferens (male) or into the uterine bell (female). Two types of protonephridia are known:

Dendric type: numerous flame cells drain into branched canals which lead into a central canal;
Saccular type: the flame cells drain into an encapsulated bowl and a subsequent central canal.

Excretory products of acanthocephalans seem to be similar to other helminths, containing lactate, succinate, etc. Ethanol, however, has been described as the main excretory product of Moniliformis moniliformis . There is still controversy over whether acanthocephalans are osmoconformers or not, but most species seem to have little osmoregulatory ability.

Reproduction

Acanthocephalan reproduction as well as the fine structure and genesis of the oocytes and spermatocytes show some unique features.

Reproductive Organs

Acanthocephalans are dioecious. Male worms are usually smaller than females, and in addition sexual dimorphism may affect other features such as trunk spination. Only males have a pair of genital ganglia (and a bursal ganglion, so far only described for M. moniliformis), whereas both sexes have a cerebral ganglion. Sensory papillae of the genital region are confined to males. And indeed only males seem to be active in finding a sexual mate and copulation. In female worms the sexual organs lie within two ligament sacs (Fig. 3a) which rupture in palaeacanthocephalans and some eoacanthocephalans. The male sexual organs are located within only one ligament sac (Fig. 3b).

The male gonads and accessory organs are enclosed by the dorsal ligament sac (the ventral sac does not persist in males), and further posteriadly, by the muscular genital sheath (Fig. 3b). The organs are attached to the ligament strand which keeps them in position. Males normally have two testes, but monorchidism is rather frequent. A seminal vesicle may be present (Eoacanthocephala). The vasa efferentia fuse to form a vas deferens, which fuses with one or several ducts of the cement gland(s) to form a genital canal. Cement glands are significant accessory organs (1–8 in number), and eoacanthocephalans have a separate cement reservoir. The cement locks the female vagina after copulation until the first embryonated eggs are released, and forms typical copulatory caps on the posterior tips of inseminated females. But dominant males may also use this secretion to prevent inferior male competitors from fertilizing females of the respective infrapopulation. If protonephridia are present, the genital canal is joined by the (ciliated) excretory canal. The genital (or urogenital) canal leads into the bursa copulatrix (Fig. 3b). The muscular terminal part of the genital canal inside the bursa is considered a penis. Additional accessory organs are the Saefftigen’s pouch and a few glandular structures associated with the bursa that are not yet well known. The fluid-filled muscular Saefftigen’s pouch is connected with the lacunar system of the bursa tegument. By its contraction it regulates the hydrostatic pressure of the bursa and thus its protrusion or invagination (Fig. 3b).

The female reproductive system consists of two major tubes:

The ligament sacs (or the pseudocoel if the sacs are ruptured) that contain the floating ovaries (ovarian balls).
An efferent duct system including a complex egg-sorting apparatus which is unique among helminths.

The two ligament sacs are interconnected at their anterior end. Posteriad, one sac leads into the uterine bell while the other is connected to a lateral opening of the subsequent apparatus (Fig. 3a).

The muscular efferent duct consists of the uterine bell, the egg-sorting apparatus, the uterus, and the vagina which is enclosed by one or two genital sphincters (Fig. 3a). Eggs from the dorsal (Archiacanthocephala) or ventral (Eoacanthocephala) ligament sac are “sucked” into the funnel-shaped bell which leads into a narrow duct. The subsequent egg-sorting apparatus of M. moniliformis consists of two lateral pockets, two dorsal median cells, two anterior ventral median cells, two posterior ventral median cells, and two lappet cells. It ensures that normally only embryonated eggs are found in the host’s faeces. By a complex interaction between the muscular activity of the bell wall and the cells and pockets of the apparatus, only embryonated eggs are allowed to enter the uterus, while immature ones are forced back into the ventral (Archiacanthocephala) or dorsal (Eoacanthocephala) ligament sac. The egg-sorting mechanism is not fully understood. The uterus is surrounded by layers of muscles and fibrous material. The vagina is a narrow duct which connects the uterus with the gonopore (Fig. 3a) and was found to carry glandular appendages in some species. The gonopore of a few species is surrounded by genital spines, and after insemination is generally blocked by a copulatory cap imposed on it by the male until eggs are released.

Gametogenesis

Acanthocephalan reproduction as well as the fine structure and genesis of the oocytes and spermatocytes show some unique features. Acanthocephalan spermatozoa are filiform (Fig. 17) and consist of a nucleocytoplasmic spermatozoan body rich in glycogen, and a flagellum. They measure 20–80 μm in length depending on the species, and obviously do not posses mitochondria or acrosomes. They contain a longitudinal chromatin strand (which is not membrane bound), two lateral rows of “dense inclusions” of unknown function and a centriole which gives rise to the flagellum. The axoneme of the free flagellum consists of microtubules which in most species are arranged in a (9 × 2) + 2 pattern, but also either one or three central tubuli have been found even within one species. The microtubules may show typical dynein arms, but the pattern is not consistent (Fig. 17). Among the phases of spermatogenesis the spermiogenesis is best described. It is characterized by several events:

The centriole of the flagellum migrates from the posterior to the anterior region of the spermatid while the spermatids are still connected in clusters (Fig. 17b) by cytophoral stalks. The flagellum then extends slightly posteriorly (while the nucleus becomes elongated) and finally it extends greatly anteriorly (Fig. 17c). Thus, as a result of this extension the spermatozoan body becomes reversed in relation to the free flagellum.
The nuclear membranes disintegrate to form the nucleocytoplasmic body (Fig. 17a, c). Only a remnant of the nuclear envelope remains.
The mitochondria disappear from the spermatozoan body.
The spermatozoan body detaches from the spermatid’s residual body containing the mitochondria.

Acanthocephala, Fig. 17
(a–c) Acanthocephalan spermatozoan and spermatid morphology. (a) DR of an acanthocephalan spermatozoan. *1, 2* Transverse sections through the spermatozoan body (×2) and through the flagellum (×3). (b) Transmission electron micrograph of a cluster of spermatids of *Echinorhynchus truttae* in the process of nuclear and flagellar elongation, i.e., spermiogenesis. Note the numerous mitochondria (MI) inside the spermatids. ×6,840. (c) TEM of a transverse section through spermatids (ST) and early spermatozoans (SB) of *E. truttae*. Note the apparent lack of mitochondria in the spermatozoan body and the rupturing nuclear envelopes (RN) in some spermatozoans. ×39,900. CL, centriole; CR, chromatin; DI, electron-dense inclusions; FE, flagellum extending from the anterior part of a spermatid posteriad; FL, flagellum; GB, Golgi body; MI, mitochondrion; MT, microtubules with dynein arms; N, nucleus; NE, nuclear envelope; NR, remnant of nuclear envelope; RN, rupturing nuclear envelope; SR, spermatozoan body; ST, spermatid; TF, terminal flagellum

Several species of acanthocephalans have been found to be precocious, e.g., mature spermatozoa have been found in male larvae.

Mature oocytes are spherical cells that lie below the surface of the free-floating ovaries (ovarian balls) and show typical electron-dense inclusions (Fig. 18a, c). The floating ovaries derive from the ovarian primordium of some larval stage. Immature floating ovaries have a thick surface coat and lack microvilli-like structures of their outer membrane. Mature ones consist of two syncytia, i.e., the central oogonial syncytium and the peripheral supporting syncytium. Furthermore they contain developing oocytes which seem to derive from the oogonial syncytium. The superficial supporting syncytium reveals microvilli-like outgrowths of its surface which absorb nutrients from the body cavity, as can be demonstrated by autoradiographic experiments. Fertilized ovaries (and unfertilized mature ovaries of a few species) apparently lose their microvilli (Fig. 18c). The actual process of oogenesis from oogonia to mature oocytes is not yet well known.

Fertilization

Acanthocephalan females may become inseminated subsequently several times. But little is known about how the worms attract each other – if they do so – prior to copulation and insemination. According to observations by Richardson et al. in the palaeacanthocephalan Leptorhynchoides thecatus, parasitizing in green sunfish, mate finding follows a very simple pattern. Individuals of both sexes are usually positioned inside the pyloric ceca in a mode such that their posterior ends extend into the intestinal lumen within the small area from which the ceca orginate. So emigration to find a mate is unnecessary (see also Behavior). There is still a lack of evidence about the function of the copulatory cap which locks the vagina of inseminated females during the prepatent period. Some philosophical debate has been held about the applicability of the “selfish gene theory” preventing males with inferior genes from reproduction. There are still open questions on acanthocephalan copulation and fertilization.

The following steps of fertilization have been documented: oocysts become fertilized whilst lying underneath the surface (syncytium) of an ovary. The flagellum of the sperm attaches to the surface of the ovary (Fig. 18b), which leads to an inflation of the flagellar apex. The subsequent penetration of the spermatozoan through the supporting syncytium into the oocyte (Fig. 18c) apparently initiates meiosis and the formation of polar bodies. The electron-dense inclusions of the mature oocyte move to the periphery and initiate the formation of a fertilization membrane around the zygote. The zygote now becomes ovoid and gives rise to a fertilization gap between its surface and the supporting syncytium of the ovary.

Postzygotic Development

The first step of postzygotic development is the formation of the first larva, the acanthor.

Females of all acanthocephalans release fully embryonated eggs. Prepatent periods of worms from homoiothermic hosts last between 22 days (Polymorphus minutus) and 70 days (Macracanthorhynchus hirudinaceus); in poikilothermic hosts development depends on the temperature. These same two species may remain patent for a maximum of only 25 days (P. minutus) or for up to 10 months (M. hirudinaceus).

After being taken up by the intermediate host the acanthor changes its morphology and becomes an Acanthella – the stage between the acanthor and the larva that is infective to the final (or paratenic) host.

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Authors and Affiliations

Horst Taraschewski
- 1

1.Zoologisches Institut, TH KarlsruheKarlsruheGermany

How to cite

Cite this entry as:: Taraschewski H. (2016) Acanthocephala. In: Mehlhorn H. (eds) Encyclopedia of Parasitology. Springer, Berlin, Heidelberg