Fig. 1. Scanning electron micrographs of acanthocephalan praesomae.
Phylum of Metazoa.
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 (Fig. 4, Fig. 16), which are cylindrical appendages of the praesomal tegument. The tube-shaped metasoma (= trunk) is bounded by a solid body wall, enclosing the pseudocoel, which is mainly filled with male or female sexual organs.
Additional morphological features as well as biological characteristics determine the affiliation to one of the three classes: Archiacanthocephala, Palaeacanthocephala, and Eoacanthocephala.
Class 1: Archiacanthocephala Meyer 1931: species have terrestrial life cycles; mammals (birds) are final hosts, and insects (millipedes) intermediate hosts; in addition, paratenic hosts are often involved; main longitudinal vessels of the lacunar system run dorsally and ventrally; usually eight uninucleate cement glands; few tegumental nuclei; ligament sacs inside the pseudocoel, also in adult worms (Fig. 16). The important 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: in general, aquatic life cycles; fish (and also 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
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Concerning the attachment to the host's intestinal wall, two groups can be distinguished: perforating and 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 (Fig. 4, Fig. 5, Fig. 6, 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. 5). On the other hand, the tipped proboscis hooks seem to use the collagen layers as suitable substrates of anchorage (Fig. 6). In Fig. 6 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. 4). 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.
Such species either continuously or occasionally change their point of attachment, exposing them to the posteriadly directed intestinal peristalsis. In infrapopulations of Echinorhynchus 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 Neoechinorhynchus 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).
Many Acanthocephalans possess a long neck which may comprise a bulbus such as the Pomphorhynchus species (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 Acanthocephalus 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. And 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 Pomphorhynchus 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 Pomphorhynchus laevis, Eocollis arcanus or Acanthocephalus 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 “by 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 pomafostomi, 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).
In non-perforating acanthocephalans the proboscis itself is usually kept in a more or less invaginated condition creating a deep proboscis cavity (Fig. 4) 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. 10 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 (Fig. 7, Fig. 8). 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. 7), and the tegument of the anterior half of the proboscis (Fig. 8) 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 (Fig. 7, Fig. 8). In accordance with their tapered and tipped construction (Fig. 7B) the hooks are in reach of lipid deposits which are not (yet) in contact with the surface of the praesoma. Behind the septum between prae- 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 contract with the praesomal surface during the exposition of a worm to the labelled nutrient, the shoulders too revealed a markable label (Fig. 8), suggesting that enzymes localised at the praesomal surface were involved. Uptake of amino acids as well as monosaccharides also occurs at the surface of the praesoma (Fig. 9), but the metasomal tegument seems to be the major absorptive surface for these substances.
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. Intraintestinal 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 Macracanthorhynchus hirudinaceus, a fibrous whitish nodule with reddened annulation around it can be seen.
The following descriptions of the acanthocephalan tegument are based on reviews by Starling, Miller and Dunagan, and Taraschewski. The tegument of acanthocephalans is a syncytium of up to 2 mm in thickness (Macracanthorhynchus hirudinaceus). It either contains numerous small nuclei (Fig. 13C) 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. 13C). 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. 1). 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. 13C). These components together build up the body wall (Fig. 13C). The tegument shows a typical stratification and a differentiation related to the praesoma-metasoma organisation of the acanthocephalan body.
The tegument of the praesoma is separated from that of the trunk by a septum composed of fibres and adherent amorphic matter (Fig. 12B). 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. 13A) as well as lipid (especially in eo- and palaeacanthocephalans, Fig. 13A, Fig. 15) than the metasomal tegument. Interestingly, a submersion of the tegumental nuclei only occurs in the praesoma. The lemnisci harbouring the nuclei (Fig. 16) do not show a specific stratification like the tegument they branch away from. They too contain lacunar spaces, and are very rich in lipid.
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 antiparasitic enzymes or antibodies with the surface coat. Eoacanthocephalans and palaeacanthocephalans reveal a lipoid, non fuzzy surface coat which may reach a thickness of several microns (Fig. 11) 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 antiparasitic peptides of the host's defense system (Fig. 12B). Unfortunately, the chemical properties of the acanthocephalan surface coat have not been extensively studied to date.
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 stabilising fibres in them (Fig. 15B). The other systematic groups have retained their individual crypts (Fig. 12A, 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.
Irrespective of the systematic affiliation of the worms the hooks (Fig. 13A, B, Fig. 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 (Fig. 13, Fig. 15A, B). The striped layer does not markably differ between eo- and palaeacanthocephalans but in eoacanthocephalans the crypts are not fused but entangle with fingerform protrusions of the tegument's basal membrane inside the hooks (Acanthocephalan Infections/Fig. 7). In both of these subclasses the tipped hooks are capable of discharging lipid substances through their pores (Fig. 13A, B). Mucus-like carbohydrates also contribute to the excreted matter (Fig. 13B) and, rather likely, also enzymes are contained in it which thus far can only be hypothesised. The grease-like surface coat of the hooks my be very voluminous (Fig. 15). Amazingly, however, the hooks are also capable of absorbing nutrients from the host tissue surrounding them (see Food Uptake).
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. 13A, B) as well as smooth endoplasmic reticulum, indicating elevated metabolic activity.
Contrary to the way how acanthocephalans usually are shown in drawings made from dead worms (Fig. 16), in vivo and in situ the acanthocephalan proboscis normally is kept in a semi-invaginated position, especially among species with superficial attachment (Fig. 5). 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.
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 their two-component structure composed of an outer myogenic, contractile belt and a cytoplasmic core enclosed by it (Fig. 13C, Fig. 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. 13C), as do the retractor muscles (proboscis retractor-, Fig. 14B, receptacle-, Fig. 14B and neck retractor musculature), for instance. 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 two-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 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:
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.
Acanthocephalan reproduction as well as the fine structure and genesis of the oocytes and spermatocytes show some unique features.
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 Moniliformis 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 finding a in sexual mate and copulation. In female worms the sexual organs lie within two ligament sacs (Fig. 16) which rupture in palaeacanthocephalans and some eoacanthocephalans. The male sexual organs are located within only one ligament sac (Fig. 16B).
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. 16B). 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 (Acanthocephalan Infections/Fig. 6). The cement locks the female vagina after copulation until the first embryonated eggs are released, and forms typical copulatory caps on the posterior tips in inseminated females. 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. 16B). 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. 16B).
The female reproductive system consists of two major tubes:
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. 16A).
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. 16A). 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 Moniliformis 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 feces. 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. 16A) 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.
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 (Fig. 17). 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 (Fig. 17), 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:
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. Fertilised ovaries (and unfertilised 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.
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, parasitising in green sunfish, mate finding follows a very simple pattern. Individuals of both sexes usually are 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 Behaviour). 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 many open questions on acanthocephalan copulation and fertilisation.
The following steps of fertilisation have been documented: oocysts become fertilised 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 fertilisation membrane around the zygote. The zygote now becomes ovoid and gives rise to a fertilisation gap between its surface and the supporting syncytium of the ovary.
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.
