ISSN: 2455-8400
International Journal of Aquaculture and Fishery Sciences
Research Article       Open Access      Peer-Reviewed

African Diplostomum (sensu Dubois 1961): Minireview on taxonomy and biology

Fred D Chibwana*

Department of Zoology and Wildlife Conservation, University of Dares Salaam P.O. Box 35064, Dar es Salaam, Tanzania
*Corresponding author: Department of Zoology and Wildlife Conservation, University of Dares Salaam P.O. Box 35064, Dar es Salaam, Tanzania, E-mail: fredchibwana@udsm.ac.tz, fredchibwana@yahoo.com
Received: 16 August, 2018 | Accepted: 24 September, 2018 | Published: 25 September, 2018
Keywords: Diplostomum; Tylodelphys; Dolichorchis; Life cyle; Taxonomy; Africa

Cite this as

Chibwana FD (2018) African Diplostomum (sensu Dubois 1961): Minireview on taxonomy and biology. Int J Aquac Fish Sci 4(3): 031-038. DOI: 10.17352/2455-8400.000041

Freshwater fisheries has a significant contribution to development as an important source of human proteins as well as in sport fishing and aquarium. Despite their importance, both wild and aquaculture fish suffer from a problem of parasitism, notably Diplostomum species, causing fish mortalities particularly in fingerlings. Although a considerable effort in understanding Diplostomum species taxonomy, biology and control of fish parasites has been well developed in the northern hemisphere, the knowledge of these aspects in Africa is not clearly known. Therefore the present work reviews the aspects of biology and taxonomy of African Diplostomum. The traditional approach to uncover these aspects would be to complete the life cycles in the laboratory, which would lead to morphological characterisation of all life cycle stages. However, establishing and maintaining life cycles of Diplostomum species is difficult, laborious and expensive. Although, molecular methods have been proven to provide an alternative solution, are not common in Africa due to lack of equipment and expertise. However, improvement of some weaknesses for some studies like providing pictures or diagrams of the Diplostomum species found is recommended. In addition, for laboratories that have the capacity to do molecular analysis, the use of a familiar molecular marker like a barcode region could be a prospective development in future.

Introduction

Freshwater fisheries has a significant contribution to development as an important source of proteins. More than 90% of all freshwater fisheries, i.e. wild capture and aquaculture, occurs in developing countries [1]. Besides providing food and a livelihood for millions of the world’s poorest people, freshwater fisheries contributes to the overall economic income by means of export commodity trade, tourism and recreation [2]. As a consequence, freshwater fisheries has become an important economic activity for both rural and urban populations in Africa and globally. The widening gap between supply and demand for fish products, has further made capture fisheries to be the largest extractive use of wildlife worldwide. Increases in human population, rising incomes and increasing urbanisation coupled with stagnation or decline of supplementary proteins, further exacerbates the situation. As a result most communities globally have responded by venturing into aquaculture to supplement capture fisheries. Despite its progress, African aquaculture needs to address a cascade of challenges including lack of national policies to guide aquaculture development, unfavourable investment policies, the absence of linkages between farmers, lack of research/technology development and extension, and unfavourable investment climates, inadequate quality seed and feed, and above all infectious and parasitic diseases [3,4].

Diplostomum species, especially larval stages namely cercariae and metacercariae are among the main agents of important diseases in fish and aquaculture systems. Diplostomum species are strigeoid digeneans of the family Diplostomidae [5,6]. The family Diplostomidae has four subfamilies: Diplostominae, Crassiphialinae, Alariinae and Codonocephalinae, which are classified on the basis of host specificity and metacercariae types [7]. However, the present review focuses on members of the compound genus Diplostomum [8], that includes three subgenera, often elevated to genera level [9]. They are Diplostomum, Tylodelphys and Dolichorchis. These parasites are ubiquitous in freshwater systems, but the most frequently encountered stages are the metacercariae that reside unencysted in the eyes, cranial and brain cavities of freshwater fishes. Diplostomum can be highly pathogenic to fish and thus threaten the natural and aquaculture practices globally. Fingerlings in particular may experience high rates of mass mortalities when heavily infected with metacercariae, or as a result of massive cercarial penetration [5]. The metacercariae can also impair the fish escape response, diminish fish crypsis and thus may increase their vulnerability to predation [10]. In addition, cercarial penetration can increase the chances of bacterial infection in fish [11] and productivity of farms could be severely compromised.

Many studies on Diplostomum species have been reported in the Northern Hemisphere where their life cycles and general biology have been well investigated. In Africa, on the other hand, a few studies are known and most of them have been restricted to reporting occurrence [12-15]. In general the taxonomy, biology and life cycles of Diplostomum species in Africa remains incomplete. With this regard, the present review focuses on Diplostomum species occurring in the African continent on the aspects of biology, taxonomy and life cycles based on the available literature. I highlighted areas where there is a dearth of information that require further research. This approach is a reflection of my personal view and not an attempt to be exhaustive.

Life cycle and host specificity

Diplostomum species, like other digenean trematodes, have attracted attention of countless studies due to their complex life-cycles, which involve a series of larval stages coupled with sexual and asexual modes of reproduction [16]. In the process they incorporate parasitic stages (sporocyst, metacercaria and adult) and free-living stages (egg, miracidium and cercaria). For a successful completion, diplostomid life cycle requires three hosts, namely, a mollusc intermediate host, a wide variety of fishes and amphibians as second intermediate host and a bird definitive host [6,10].

Sexually matured adults of Diplostomum species are found in the alimentary tract of bird host where they produce eggs that reach the external environment with the host’s faeces. Eggs hatch to release a ciliated free-swimming miracidium that seeks out a molluscan intermediate host. Snail hosts becomes infected after actively being penetrated by the miracidium. Once in the molluscan tissue, the miracidium transforms into a mother sporocyst, which produce asexually embryos that develop into daughter sporocysts generation. Moreover, the second intra-molluscan generations, daughter sporocysts, produce cercariae after another rounds of asexual reproduction [16]. Cercariae are well equipped with features and behaviours geared towards fish-finding, recognition and penetration [6]. After emergence, the cercariae actively infect a wide variety of fishes to become metacercariae. In the host, the metacercaria migrate along specific neural tracts to access sites with relatively minimal immune resistance like visceral organs, optic lobes and the brain so that they remain viable for successful transmission to the next host [16]. Bird hosts normally get infected upon consuming metacercariae in fish hosts [17].

Nonetheless, Diplostomum species life cycles have been well documented in the Palaearctic region, but not fully understood in Africa. Usually, lymnaeid snails (Lymnaea and Radix spp) and planorbid snails (Planorbarius corneus) are the main intermediate hosts [6,18], and in South America the snails of genus Biomphalaria such as B. prona, B. straminea and B. glabrata have been reported as the potential first intermediate [19]. In Africa, on the other hand, the range of snail hosts are far from completely understood, despite several experimental attempts undertaken [13,20]. The only known life cycle is that of Tylodelphys xenopi, in which the freshwater snail Bulinus tropicus is the first intermediate host [21]. Beverley-Burton (1963) also tried to infect Radix natalensis (Lymnaeidae) with Diplostomum (Tylodelphys) mashonense, but could not obtain cercariae. Even so, strigeoid cercariae purported to belong to the genus Diplostomum have been reported from Biomphalaria species from three fish farms in Kibos area within Kisumu, Kenya [22]. Similarly, at Mindu dam in Tanzania, Diplostomum-like furcocercariae had been reported once from the snail Biomphalaria pfeifferi [23,24], but since then B. pfeifferi have not been spotted in the dame. Thus the consistently high prevalence of the metacercariae of T. mashonense in C. gariepinus at Mindu Dam [23,25], brought a suspicion that non-lymnaeid snail species could serve as snail hosts, particularly because lymnaeid snails have not been recorded within or around the dam. As such it was hypothesised that other snails besides lymnaeids and Biomphalaria, could be responsible for the transmission and Bulinus spp were shown to support that hypothesis [25], as shown in figure 1.

In Diplostomum species, host specificity is mostly restricted in the first intermediate host while less specific in the second intermediate host and final hosts. For instance in Europe and North America, metacercariae of Diplostomum species have been recorded from over 150 species of fish from families Percidae, Salmonidae, Coregonidae, Clupeidae, Gobiidae, to name a few [26,27] and a broad range of piscivorous bird species serving as definitive hosts [10]. Similarly in Africa, Diplostomum species have been found in almost every fish family i.e. Characidae, Centrarchidae, Cichlidae, Claridae, Cyprinidae, Hepsetidae, Salmonidae, Schilbeidae to mention but a few [28,29]. However, the catfish Clarias gariepinus (family Claridae) is the most examined fish [29] and references therein. Nonetheless the range of fish hosts in Africa is not well known as the level of Diplostomum studies in Africa is still at an infant stage.

As far as definitive hosts are concerned, in Africa adult Diplostomum have been reported from the Egyptian kite Milvus migrans aegypticus, the Egyptian moorhen Gallinula chloropus chloropus and the giant heron Ardea goliath in Egypt [20,30,31], Pel’s fishing owl Scotopelia peli in Ivory Coast [32], the grey heron Ardea cinerea in Zimbabwe and Tanzania [13,23,33] the African darter, Anhinga rufa rufa in Ghana [34] and the great white egret Ardea alba in Tanzania [23,33]. Generally, studies of Diplostomum species in bird definitive hosts in the African continent are scarce and limited. This is attributed to (i) low sampling efforts in the tropical countries due to inadequate expertise and resources (ii) difficulty in getting study permits to sacrifice some birds as they are either found in the protected areas (national parks and game reserves) or lack of interests in fish parasitology.

Taxonomy of Diplostomum (sensu Dubois, 1970) species in africa

Precise identification of members of the genus Diplostomum is usually difficult because of remarkable morphological similarity within and among species at almost every developmental stage [6]. Also, lack of a well-defined criterion further exacerbates the delineation difficulty within the Diplostomum group [6,35]. Furthermore, the taxonomic problem is aggravated by deformation of the body in the course of fixation, staining and mounting of permanent preparations [6]. In addition, many species have been described on the basis of one or two life cycle stages as a result different stages of the same species have been given different names or different species are known by the same name [6].

The taxonomy of Diplostomum species in Africa in general remains not well understood because the full range of species that may occur in Africa is far from completely known. As a consequence the biology of all reported species such as the range of hosts and life cycle stages (eggs, miracidia, intramolluscan stages, cercariae) is poorly understood. So far in the whole of Africa the adults of only five Diplostomum species have been described; they are D. tregenna [12], D. marahoueense [32], D. magnicaudum [30], D. ghanense [34] and D. ardeae [31]. On the other hand, only four Tylodelphys species adults are known i.e. T. mashonense [13], T. aegypteus [31], T. clavata [36] and T. xenopi [21]. Nevertheless, more than 15 Diplostomum species have been reported as metacercaria in different fish hosts species [29,37,38]. Although some authors tried to name the metacercarial stages to species level, for example D. garrae, D. longicollum, D. montanum and D. tilapiae described by Zhokhov [38], have further exacerbated the taxonomic problem. Moreover the taxonomic status of some Diplostomum species that have been recorded from some fish hosts is confusing. For instance, when the genus Clarias was reviewed by [39], several widespread species i.e Clarias ngamensis, C. melandi and C. capensis of southern Africa, C. mossambicus of central Africa and C. lazera of west and north Africa were synonymized under the name Clarias gariepinus. With this regard, the taxonomic status of Diplostomum (Dolichorchis) tregenna recovered from C. lazera and Tylodelphys mashonense found in C. gariepinus is questionable.

In addition Niewiadomska [9] moved D. marahoueense Baer 1957 to the genus Dolichorchis due to the presence of a genital cone and asymmetrical anterior testes in the adult form. This taxonomic re-evaluation of D. marahoueense implies that other African Diplostomum species with similar features may follow a similar fate. This idea was supported by Zhokhov et al. [40] and Zhokhov [38] who considered Diplostomum tregenna as Dolichorchis tregenna. At the metacercarial stage the fundamental features that enable distinguishing genera or species are the shape of fore- and hindbodies, presence or lack of additional organs of attachment like pseudosuckers, the structure of the holdfast organ, structure of the reserve bladder, the shape and spread of the calcareous bodies [35]. However these features could only be used on T. mashonense, D. tregenna and other known diplostomid metacercariae, but not on the metacercaria of D. marahoueense, which is hitherto unknown. As a result the metacercarial status of D. marahoueense is not understood.

Various literatures regarding Diplostomum species described from Africa reveal a complicated taxonomic situation. In particular, views over the taxonomic position of the named Diplostomum species differ. Dubois [8] considered the genus Diplostomum to be under three subgenera, namely, Diplostomum, Tylodelphys and Dolichorchis (Figure 2). The idea which was supported by Niewiadomska [9], but elevated them to genera. Accordingly, the classification of the three forms was based on the anatomical features of the adults, i.e. the asymmetrical nature of the anterior testis (as in Diplostomum spp.) and the presence of a genital cone (as in Tylodelphys spp.). The genus Dolichorchis is given to materials exhibiting an intermediate position between Diplostomum and Tylodelphys, i.e. asymmetrical anterior testis and the presence of a genital cone. As a result, African material Diplostomum tregenna, D. marahoueense and D. mashonense have been assigned within the genus Dolichorchis [9], although they originally belonged to the genus Diplostomum. However, phylogenetic analysis of Diplostomum mashonense by Chibwana et al. [41] suggested a re-allocation to Tylodelphys as previously viewed by Sudarikov [42].

Furthermore, records of Tylodelphys species in Africa are generally scarce. The only descriptions of adults Tylodelphys species are those of immature T. clavata from the intestine of a jackal, buzzard Buteo rufofuscus in the Democratic Republic of Congo (DRC) [36] and T. xenopi from an experimental host, the African darter Anhinga melanogaster in South Africa [21] and T. mashonense from Ardea cinerea and A. alba [13,33]. Other Tylodelphys species have only been described at the metacercarial stages. For instance Tylodelphylus (Diplostomulum) victorianus was described by Vercammen-Grandjean [43] from the pericardial cavity of Xenopus laevis victorianus in Nyakabere River in DRC. T. grandis was described from C. gariepinus by Zhokov et al. [40] from Lake Tana in Ethiopia. Four other Tylodelphys species, namely, T. claviformis (from Barbus, LabeoBarbus, Garra), T. mutica (from Barbus), T. fusiliformis (from Oreochromis niloticus) and T. clariae (from C. gariepinus) were also described by Zhokov [44] from Lake Tana in Ethiopia. However, there are several metacercariae of Tylodelphys species from numerous fishes in Africa (Table 1), which have not been described. In addition, the adults of these metacercariae and their life cycles are hitherto unknown, and their classifications only end at the genus level.

Techniques and Methods

In order to have a reliable research output, researchers need a good criterion to filter from diversely available techniques and methods. For instance Chappell [45], reported that methods for identification of Diplostomum species depend upon: (i) site of infection in a host; (ii) metacercarial morphology; (iii) infected host species; (iv) chaetotaxy of cercariae (not reliable unless supported by evidence obtained from experimental infections); (v) experimental establishment of the life cycle and recovery of the adult worm. Of the five methods above only three i.e. site of infection, infected host species and metacercarial morphology are the most common in African studies. In other words most studies have been dealing with occurrences. Diplostomum metacercariae have been reported in the cranial cavity [13,14,46,47], eyes [29,48] and abdominal cavities [21]. Although other Diplostomum species recorded by some studies occur in unusual sites in fish host or unusual host, they are neither accompanied with morphometrics nor figures. For example metacercariae and adults of Diplostomum commutatum were isolated from the stomachs and intestines of fishes Pseudotolithus elongatus and Cynoglossus senegalensis, respectively [49]. Generally most of studies on Diplostomum species in Africa are not accompanied with the known taxonomic methods i.e. morphometrics and illustrations. With this regard it is difficult for other researchers to support or refute conclusions.

According to Niewiadomska [7], a precise identification of Diplostomum species requires the a priori knowledge of all the features in the developmental cycle i.e. adult, cercaria and metacercaria. However, a correct description of cercariae and metacercariae relies on traditional and modern methods of morphological research like numerical taxonomy, as well as verification of the correctly identified adult forms obtained experimentally. So far, in Africa, identification of Diplostomum species across the life cycle has not been done due to incomplete information on the cercarial stage [13,20]. In addition, maintaining life cycles using in vivo systems is difficult, laborious and expensive and in vitro cultivation is almost impossible [6]. All these considerations indicate that both the identification of the larval stages and uncovering the diversity of Diplostomum species in Africa is challenging. Furthermore, experimental approaches cannot be applied to a large-scale screening of natural infections in intermediate hosts (fish or snails) due to the impossibility of linking each larval stage with its corresponding adult stage [25]. Although, the application of molecular techniques on Diplostomum has advanced significantly elsewhere [9,50-54], only a few researches in Africa have ventured into these techniques.

As polymerase chain reaction (PCR)-based methods for molecular analysis have started to advance in Africa, and a variety of molecular markers have been applied in different studies. Some of the gene regions that have been used to assess genetic diversity and variability among species and linking of life cycle stages are partial 18S rDNA sequences [23,55], 28S recombinant DNA [47], ITS rDNA (ITS1-5.8S-ITS2 [25,41.48] and the DNA barcode region of cytochrome c oxidase I (COI) [25,56]. T. mashonense is the most analysed diplostomid species in Africa, and almost every molecular marker mentioned above has been tested on it (see above citations). However, studies based on molecular methods on African Diplostomum are few in number, and the majority of them have used different markers, thus making it difficult to compare the findings taxonomically. For instance, it is not clearly understood if the T. mashonense studied by Chibwana et al. [25,41], is similar to that dealt by Moema et al. [47]. In other words, although DNA sequences of those workers’ materials have been deposited in the public nucleotide databases (like GenBank), they cannot be aligned together for identification purposes. As a result, more and more of the so-called Diplostomum species are discovered spatially and temporally, but their sequences cannot be used to harmonise their identities. Increased synonyms in Diplostomum species as shown by Niewiadomska [6] could be the ultimate outcome.

Future prospects

Occurrence of Diplostomum species in their hosts across the life cycle, i.e. in snails, fish or birds, is one of the first steps that need to be performed for the subsequent analyses. However, a correct identification of hosts is one of the most challenging endeavour in Africa, which requires an involvement of experts in specific fields viz. ornithologist, malacologist and fish biologist. Understanding the biology of Diplostomum species and their hosts would help fisheries managers and extension workers to deal with the threat of the disease that might be posed by these parasites. As a consequence, it will raise the economic returns of fish farms as correct identification of species is critical in making correct decisions for disease control. Such information would also be invaluable for planning, implementation and assessment of control strategies for these parasites in fish farms. An experimental establishment of the life cycle of diplostomid species is needed in order to study the biology of the partially studied developmental stages like eggs, miracidia, intramolluscan stages, cercariae and metacercariae. In other words, studies on infectivity, pathology and migration through the fish host of the parasite would only be possible if a life cycle is maintained in the laboratory. Since Diplostomun species can easily be cultivated in the laboratory [6], identification and naming of species should be based on adult stages.

An alternative to completing the life cycle experimentally, which is tedious and laborious, would be DNA sequencing. Unfortunately, in Africa, DNA sequencing has rarely been employed not only in the assessment of Diplostomum, but also other animal groups. In cases where DNA sequencing was applied, there has been little coordination (the sequences of various studies cannot be compared because they targeted exclusive DNA regions) of the four genes (i.e. 18S, 28S, ITS and mtDNA) frequently used with great success in these trematode studies. Consequently, the many sequences available on public databases such as GenBank could not be assembled and analysed together because they do not represent homologous gene regions. To alleviate this difficulty, in future studies, sequencing of all four genes would be an ideal solution in a quest to identify more species. Alternatively, DNA barcoding efforts for trematode species across Africa would be recommented.

DNA barcoding, the use of single locus cytochrome c oxidase subunit I (COI) of mitochondrial DNA, has already been popular in revealing illegal imports of bushmeats (Eaton et al., 2009), labelling requirements violations in marketed fish (Wong and Hanner, 2008; Lowenstein et al., 2009) and accurate identification of potentially toxic tuna (Lowenstein et al., 2010). The uniqueness of the locus COI across species is purported to allow rapid and accurate identification of almost all animal species (Hebert et al., 2003). Although this technique has been widely adopted by other biological fields, the parasitological community has barely used it. In parasitology and Diplostomum in particular, DNA barcoding has been able to reveal diversity and specificity of metacercariae in hosts [52], disentangle cryptic species [53] and linking life cycle developmental stages [25]. As already shown by Besansky et al. (2003), DNA barcoding is potentially a great tool to improve the rate of discovery of parasites’ species diversity and life cycles. The author of this review, therefore recommends the increased use of this genetic region in African diplostomids in order to fill the taxonomic gaps prevailing in Diplostomum species emanating from various morphological challenges. It will also enable quick identification of diplostomid species irrespective of their developmental stage once their sequences are deposited in the public nucleotide databases.

Conclusion

Since the first Diplostomum species in Africa was described in the intestines of Egyptian kite Milvus migrans aegypticus in Sudan by Nazmi (1932), many other Diplostomum have been found in other countries and hosts albeit most studies have been conducted in the sub-Saharan Africa. However, a majority of studies have been reporting their occurrences in fish either in natural waters or fish farms. The influence of most Diplostomum in the fish populations is not well understood. Although, interest on Diplostomum studies has increased, most aspects like their biology, epidemiology, taxonomy and immunology are well known. The traditional approach to uncover these aspects would be to complete the life cycles in the laboratory, which would lead to morphological characterisation of all life cycle stages. However, establishing and maintaining the life cycles of these Diplostomum species using in vivo systems is difficult, laborious and expensive. Although, molecular methods have been proven to provide an alternative solution, are uncommon in African studies due to lack of equipment, resources and expertise. However, improvement of some weaknesses for some studies like providing pictures or diagrams of the Diplostomum species found. For laboratories or researchers that have the capacity to do molecular analysis, the use of a molecular marker that is commonly used like barcode region CO1 and ITS could be a prospective development in future.

  1. Tidwell JH, Allan GL (2001) Fish as food: aquaculture’s contribution. EMBO reports 2: 958–963. Link: https://tinyurl.com/ycnjcyw8
  2. Béné C, Macfayden G, Allison EH (2007) Increasing the contribution of small-scale fisheries to poverty alleviation and food security. Fisheries Bethesda 125. Link: https://tinyurl.com/y7g22qz4
  3. Hecht T, Endemann F (1998) The impact of parasites, infections and diseases on the development of aquaculture in sub-Saharan Africa. Journal of Applied Ichthyology14: 213–221. Link: https://tinyurl.com/yatleg2f
  4. Brummett RE, Williams MJ (2000) The evolution of aquaculture in African rural and economic development. Ecological Economics 33: 193–203. Link: https://tinyurl.com/yaggojj3
  5. Chappell L, Hardie LJ Secombes CJ, (1994) Diplostomiasis: the disease and host-parasite interactions. In A. W. Pike & J. W. Lewis, eds. Parasitic Diseases of Fish. Samara Publishing, Dyfed, Wales 59–86. Link: https://tinyurl.com/yaozxyg6
  6. Niewiadomska K (1996) The genus Diplostomum-taxonomy, morphology and biology. Acta Parasitologica 41: 55–66. Link: https://tinyurl.com/yaozxyg6
  7. Niewiadomska K, (2002) Family Diplostomidae Poirier, 1886. In D. Gibson, A. Jones, & R. Bray, eds. Keys to the Trematoda. CABI Publishing and The Natural History Museum, Wallingford 167–196. Link: https://tinyurl.com/yc2wskuf
  8. Dubois G (1961) Le genre Diplostomum von Nordmann, 1832 (Trematoda: Strigeida). Bulletin de la Société Neuchâteloise des Sciences Naturelles 84: 113–124. Link: https://tinyurl.com/y73hgnzn
  9. Laskowski Z, Niewiadomska K (2002) Systematic relationships among six species of Diplostomum Nordmann, 1832 (Digenea) based on morphological and molecular data. Acta Parasitologica 47: 20–28. Link: https://tinyurl.com/y9tlb8wt
  10. Seppala O, Karvonen A, Valtonen ET (2006) Susceptibility of eye fluke-infected fish to predation by bird hosts. Parasitology 132: 575–579. Link: https://tinyurl.com/y85zj5pr
  11. Pylkkö P (2006) Evidence of enhanced bacterial invasion during Diplostomum spathaceum infection in European grayling, Thymallus thymallus (L.). Journal of fish diseases 29: 79–86. Link: https://tinyurl.com/ycq9ep3z
  12. Nazmi M (1932) LIX.—Diplostomum tregenna, sp. n., a new Trematode parasite of the Egyptian Kite. Journal of Natural History 9: 567–573. Link: https://tinyurl.com/yajhtxn9
  13. Beverley-Burton M (1963) A new strigeid Diplostomum (T) mashonense n.sp (Trematoda: Diplostomatidae) from the grey heron, Ardea cinerea L., in Southern Rhodesia with an experimental demonstration of part of the life cycle. Revue Zoologica Botanica Africa, LXVIII 291–306. Link: https://tinyurl.com/yboeetmp
  14. Musiba MJ, Nkwengulila G (2006) Occurrence of Metacercariae of Diplostomum and Tylodelphys species (Diplostomidae) in Clarias species (Clariidae) From Lake Victoria. Tanzania Journal of Science 32: 89–98. Link: https://tinyurl.com/y77o7yaj
  15. Ibrahim A (2016)  Natural occurrence of Diplostomum spp. in farm-raised African catfish (Clarias gariepinus) from Oyo state, Nigeria. International Journal of Veterinary Science and Medicine 4: 41–45. Link: https://tinyurl.com/yajxhd6a
  16. Galaktionov KV, Dobrovolskij AA (2003) The biology and evolution of Trematodes B. Fried & Æ. T. K. Graczyk, eds., Springer Science+Business Media Dordrecht.
  17. Barber I (2003) The role of parasites in fi sh – bird interactions: a behavioural ecological perspective. In I. G. COWX, ed. Interactions Between Fish and Birds. Blackwell Science Ltd. 221–243.
  18. Faltýnková A (2005) Larval trematodes (Digenea) in molluscs from small water bodies near Šeské Budšjovice, Czech Republic. Acta Parasitologica 50: 49–55. Link: https://tinyurl.com/yadpm26t
  19. Pinto HA, Melo AL (2013) Biomphalaria straminea and Biomphalaria glabrata (Mollusca: Planorbidae) as New Intermediate Hosts of the Fish Eyefluke Austrodiplostomum compactum (Trematoda: Diplostomidae) in Brazil. Journal of Parasitology 99: 729–733.  Link: https://tinyurl.com/y8j6h76c
  20. Khalil L (1963) On Diplostomulum tregenna, the diplostomulum stage of Diplostomum tregenna Nazmi Gohar, 1932 with an experimental demonstration of part of the life cycle. Journal of Helminthology 37: 199–206. Link: https://tinyurl.com/y95areax
  21. King PH, Van As JG (1997) Description of the adult and larval stages of Tylodelphys xenopi (Trematoda: Diplostomidae) from Southern Africa. The Journal of Parasitology 83: 287–295. Link: https://tinyurl.com/y922kj4j
  22. Ndeda V (2013) Occurrence and effect of Diplostomum parasites in cultured Oreochromis niloticus (L.) and distribution in vector snails within Kisumu City, Western Kenya. Ecohydrology and Hydrobiology 13: 253–260. Link: https://tinyurl.com/y9azu2x6
  23. Chibwana FD, Nkwengulila G (2010) Variation in the morphometrics of diplostomid metacercariae ( Digenea : Trematoda ) infecting the catfish , Clarias gariepinus in Tanzania. Journal of helminthology 84: 61–70. Link: https://tinyurl.com/y7rc3bey
  24. Nkwengulila G, Kigadye E (2005) Occurrence of digenean larvae in freshwater snails in the Ruvu basin, Tanzania. Tanzania Journal of Science. Link: https://tinyurl.com/y9nygggq
  25. Chibwana FD (2015) Completion of the life cycle of Tylodelphys mashonense (Sudarikov, 1971) (Digenea: Diplostomidae) with DNA barcodes and rDNA sequences. Parasitology Research 114: 3675–3682. Link: https://tinyurl.com/ycvtblap
  26. Sweeting R (1974) Investigations into natural and experimental infections of freshwater fish by the common eye-fluke Diplostomum spathaceum Rud. Parasitology 69: 291–300. Link: https://tinyurl.com/ybkth2sx
  27. Hglund J, Thulin J (1992) Identification of Diplostomum spp . in the retina of perch Perca fluviatilis and the lens of roach Rutilus rutilus from the Baltic Sea - an experimental study. Systematic Parasitology 21: 1–19. Link: https://tinyurl.com/ybgsuyf6
  28. Van As JG, Basson L (1984) Checklist of freshwater fish parasites from southern africa. South African Journal of Wildlife Research 14: 49–61. Link: https://tinyurl.com/y8npvluy
  29. Grobbelaar A (2014) Ecology of diplostomid (Trematoda: Digenea) infection in freshwater fish in southern Africa. African Zoology 49: 222–232. Link: https://tinyurl.com/yb82xey7
  30. El-Naffar M (1979) Parasites of the Egyptian moorhens. I: Diplostomum magnicaudum sp. nov. with part of its life cycle. Journal of the Egyptian Society of Parasitology 9: 349–358. Link: https://tinyurl.com/ybtyz7zn
  31. El-Naffar M, Khalifa R, Sakla A (1980) Parasitofauna of the Egyptian aquatic birds. II. Trematode parasites of the giant heron (Ardea goliath) in Assiut governorate. Journal of the Egyptian Society of Parasitology 10: 107–116. Link: https://tinyurl.com/ybzqq5h4
  32. Baer J (1957) Trematodes et Cestodes recoltes en Cote d’Ivoire, avec remarques sur lafamille des Dicrocoeliidae Odhner et sur les parasites des Damans. Revue Suisse de Zoologie 64: 547–575. Link: https://tinyurl.com/y967fjb8
  33. Chibwana FD.  Nkwengulila G (2016) Spread of Tylodelphys mashonense (digenea: diplostomidae) by grey heron Ardea cinerea and great white Ardea alba in Lake Victoria, Tanzania. Tanzania Journal of Science 42: 150–159. Link: https://tinyurl.com/yaxnlcub
  34. Ukoli F (1968) Three new trematode parasites of the African darter, Anhinga rufa rufa (Lacepéde and Daudin, 1802) in Ghana. Journal of Helminthology 42: 179–192. Link: https://tinyurl.com/ybo7rolp
  35. Niewiadomska K (1970) An analysis of criteria for generic differentiation within the order Strigeidida (La Rue, 1926). Acta Parasitologica Polonica 18: 277–289. Link: https://tinyurl.com/y859le6c
  36. Dubois G, Fain A (1956) Contribution a l’étude des Strigeida du Congo Beige. I. Bulletin de la Societé neuchâteloise des Sciences naturelles 79: 17–38. Link: https://tinyurl.com/ydhokfg2
  37. Migiro KE (2012) Diplostomum Parasites Affecting Oreochromis niloticus in Chepkoilel Fish farm and Two Dams in Eldoret-Kenya 3: 3–9. https://tinyurl.com/yb2bf5vx
  38. Zhokhov A (2014) Metacercariae of new trematode species of the genus Diplostomum (Trematoda, Diplostomidae) from fishes of Lake Tana, Ethiopia. Inland Water Biology. Link: https://tinyurl.com/yb4njqea
  39. Teugels GG (1986) A Systematic revision of the African Species of the genus Clarias (Pisces: Clariidae). Link: https://tinyurl.com/yb26gyof
  40. Zhokhov AE, Morozova DA, Tessema A (2010) Trematode metacercariae from the cranial cavity of African catfish Clarias gariepinus (Burchell, 1822) from Lake Tana, Ethiopia. Inland Water Biology 3: 160–164. Link: https://tinyurl.com/y8owsas9
  41. Chibwana FD  (2013) Infec tion , Geneti cs and Evol ution A first insight into the barcodes for African diplostomids ( Digenea : Diplostomidae ): rain parasites in Clarias gariepinus ( Siluriformes : Clariidae ). Infection, Genetics and Evolution 17: 62–70. Link: https://tinyurl.com/yaanpq49
  42. Sudarikov VE (1971)  Part II. Order Strigeidida (La Rue, 1926) Sudarikov, 1959: Suborder Strigeata La Rue, 1926: Part V. Metacercariae and mesocercariae. In K. I. Skrjabin, ed. Trematodes of animals and man, Vol. XXIV. Moscow, Russia: Academy of Sciences of the USSR 69–308.
  43. Vercammen-Grandjean PH (1960) Les trématodes du lac Kivu sud (Vermes). Annales du Musee Royal de l’Afrique Centrale, Tervuren. Nouvelle Serie in 4, Sciences Zoologiques 5: 171. Link: https://tinyurl.com/ybr3zf4t
  44. Zhokhov A (2012) Metacercariae of trematodes (Plathelminthes: Trematoda) of Garra dembecha (Actinopterygii: Cyprinidae) from Lake Tana , Ethiopia. Zoosystematica Rossica 21: 193–203. Link: https://tinyurl.com/y8n3ozzd
  45. Chappell LH (1995) The biology of diplostomatid eyeflukes of fishes. Journal of Helminthology 69: 97 -101. Link: https://tinyurl.com/y8aq62e9
  46. Mashego SNm, Saayman JE (1989) Digenetic trematodes and cestodes of Clarias gariepinus (Burchell, 1822) in Lebowa, South Africa, with taxonomic notes. South African Journal of Wildlife Research 17–20. Link: https://tinyurl.com/ya3z6qko
  47. Moema E, King P (2013) Descriptions of diplostomid metacercariae (Digenea: Diplostomidae) from freshwater fishes in the Tshwane area. Onderstepoort Journal  80. Link: https://tinyurl.com/y8g2qp5p
  48. Ndeda VM (2013) Genetic Relatedness of Diplostomum Species ( Digenea : Diplostomidae ) Infesting Nile Tilapia ( Oreochromis Niloticus L.) in Western Kenya. Open Journal of Applied Sciences  201: 441–448. Link: https://tinyurl.com/yb3o8w4p
  49. Abraham J, Akpan PA, Okon O (1995) Occurrence Of Parasites In Pseudotolithus elongatus and Cynoglossus seneglensis in Cross River Estuary, Nigeria. Global Journal of Pure and Applied Sciences 11: 45–49. Link: https://tinyurl.com/yazypg8t
  50. Laskowski Z (1996) Species identification of Diplostomum pseudospathaceum Niewiadomska, 1948 and D. paracaudum (Iles, 1959) metacercariea using DNA polymorphism amplified by. Acta Parasitologica. Link: https://tinyurl.com/yd9ftr95
  51. Galazzo DE, Dayanandan S, Marcogliese DJ, McLaughlin JD (2002) Molecular systematics of some North American species of Diplostomum ( Digenea ) based on rDNA- sequence data and comparisons with European congeners. Canadian Journal of Zoology 80: 2207–2217. Link: https://tinyurl.com/ybmz2nyu
  52. Locke S,  McLaughlin J (2010) Diversity and specificity in< i> Diplostomum</i> spp. metacercariae in freshwater fishes revealed by cytochrome< i> c</i> oxidase I and internal transcribed spacer. International journal for. Link: https://tinyurl.com/y7wwvu8v
  53. Pérez-del-Olmo A, Georgieva S (2014)  Molecular and morphological evidence for three species of Diplostomum (Digenea: Diplostomidae), parasites of fishes and fish-eating birds in Spain. Parasites 7: 502. Link: https://tinyurl.com/y92wlged
  54. S Georgieva M,  Soldánová A, Pérez-del-Olmo DR, Dangel JS, Bernd Sures AK (2013) Molecular prospecting for European Diplostomum(Digenea: Diplostomidae) reveals cryptic diversity. International Journal for Parasitology 43: 57–72. Link: https://tinyurl.com/y7kslgfw
  55. Mwita C, Nkwengulila G (2010) Phylogenetic relationships of the metazoan parasites of the clariid fishes of Lake Victoria inferred from partial 18S rDNA sequences. Tanzania Journal of Science 36: 47–58. Link: https://tinyurl.com/y9cghoo5
  56. Otachi EO (2015) Morphometric and molecular analyses of Tylodelphys sp. metacercariae (Digenea: Diplostomidae) from the vitreous humour of four fish species from Lake Naivasha, Kenya. Journal of Helminthology 89: 404–414. Link: https://tinyurl.com/ya6ejd94
© 2018 Chibwana FD. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.