Diphyllobothriidae


Published on:
June 7, 2018

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Torres, P. and Yera, H. 2018. Diphyllobothriidae. In: J.B. Rose and B. Jiménez-Cisneros, (eds) Global Water Pathogen Project. http://www.waterpathogens.org (Robertson, L (eds) Part 4  Helminths)   http://www.waterpathogens.org/book/diphyllobothriidae Michigan State University, E. Lansing, MI, UNESCO.
https://doi.org/10.14321/waterpathogens.38

Acknowledgements: K.R.L. Young, Project Design editor; Website Design 
(http://www.agroknow.com)

Last published: June 7, 2018
Authors: 
Patricio Torres (Austral University of Chile)Hélène Yera (Paris Descartes University)

Summary

The Diphyllobothriidae family includes zoonotic species of importance in human and animal health that are grouped into genera Dibothriocephalus, Diphyllobothrium, Adenocephalus, and Spirometra. The species of the first three genera have larval stages (coracidia) that develop in the aquatic environment  and have intermediate host copepods (procercoids) and fish (plerocercoids or sparganums) in their life cycles. In addition, all include a definitive host, where they develop into the adult stage (mammals or birds, the latter in only some species of Dibothriocephalus). In Spirometra spp., the plerocercoid develops in amphibians, snakes, birds and mammals, including humans, and their definitive host corresponds to canids and felids.

Humans become infected by species of so-called broad fish tapeworms (Dibothriocephalus, Diphyllobothrium and Adenocephalus) by eating raw or undercooked fish (freshwater or marine). These parasites can cause severe gastrointestinal symptoms. Spirometra spp. can cause severe disease, depending on the final location of larvae in humans. This infection is acquired by drinking water contaminated with infected copepods, ingesting undercooked meat of amphibians, snakes, birds or mammals or applying flesh of these animals on open wounds.

The aquatic environment becomes contaminated with the feces of the definitive host, containing the parasite eggs: D. latus in mainly humans and dogs, also other domestic and wild mammals; D. dendriticus in fish-eating birds, mainly gulls (Laridae); A. pacificus in marine mammals such as sea lions and eared seals and Diphyllobothrium in whales and fur seals; and canids and felids in the life cycle of Spirometra. The spread of D. dendriticus, A. pacificus, and Diphyllobothrium would be by wild hosts will be greater than the spread by humans.

The spread of parasite eggs derived from human infections by Dibothriocephalus, Diphyllobothrium, and Adenocephalus can partly be controlled by latrine use and treating wastewater. The dissemination of parasites in the family Diphyllobothriidae present in wild or domestic hosts is more difficult to avoid. While wastewater treatment is part of the transmission control options, efficient health education to create awareness of adequate thermal and freezing processes in the preparation of fish dishes will also have an impact. Strict regulations of food prepared with raw fish in restaurants, timely treatment of patients and availability of clean drinking water, in the case of Spirometra, could contribute to an improved control of these re-emerging infections.

1.0 Epidemiology of the Disease and Pathogens

1.1 Global Burden of Disease

1.1.1 Global distribution
1.1.1.1 Dibothriocephalus, Diphyllobothrium, Adenocephalus, and Spirometra

The Diphyllobothriidae family includes the so-called broad tapeworms that are widely distributed all around the world, with some being agents of human infection. They are commonly present in the wildlife among fish, mammal and bird hosts that make an important reservoir (Muller, 2002; Chai et al., 2005). Recently, taxonomy of broad tapeworms included in the family Diphyllobothriidae have been revised, including the followings genera with species of greater importance in health: Dibothriocephalus with D. latus (syn. Diphyllobothrium latum), D. dendriticus (syn. Diphyllobothrium dendriticum), and D. nihonkaiensis (syn. Diphyllobothrium nihonkaiense); Diphyllobothrium with Di. stemmacephalum and Di. balaenopterae (syn. Diplogonoporus balaenopterae); Spirometra and Adenocephalus (Waeschenbach et al., 2017).

In the early 1970´s the worldwide prevalence of human infection was estimated at 9 million cases (WHO, 1979). Recent data indicate that 20 million people are infected worldwide with D. latus (Muller, 2002; Chai et al., 2005; Scholz and Kuchta, 2016). Currently, there are estimated about 270; 1,000; 1,000; and 2,000 human cases of infection by Di. balaenopterae, D. dendriticus, Adenocephalus pacificus, and D. nihonkaiensis, respectively (Scholz and Kuchta, 2016). The zoonosis occurs most commonly in countries where the consumption of raw fish is a frequent practice. Flesh (marinated fillets, carpaccio, ceviche, sushi, etc.) of infected fish with plerocercoid larvae could transmit the infection to humans.

The areas of endemicity are Japan, the Baltic countries, Scandinavia, Western and Eastern Russia, North America (Pacific Northwest) and South America. In Japan, especially along the coast of the Sea of Japan, about 100 cases per year have been reported since the 1970s (Oshima and Wakai, 1983). In the coastal areas of the Okhotsk Sea, human prevalence ranges from 1.0 to 3.3% (Lloyd, 1998).

Human infection by Dibothriocephalus spp. is decreasing in some historical areas of endemicity as the Baltic or Scandinavian countries (Raisanen and Puska, 1984) and in North America (Dick, 2008; Jenkins et al., 2013). In contrary, due to the increasing popularity of dishes utilizing raw or undercooked fish, human infection is emerging or re-emerging in some European countries, particularly in Italian and French speaking areas of the perialpine lakes (Dupouy-Camet and Peduzzi, 2004; Scholz et al., 2009), and in South America (Torres et al., 1989b, 1993, 2004a; Semenas et al., 2001; Sampaio et al., 2005; Knoff et al., 2011; Kuchta et al., 2015a). In Peru, prevalence of infection by A. pacificus is around 2%, but has been as high as 7.5% in some regions (Cabrera, 2003).

Due to the globalization of fish trading and to travels and immigration, sporadic cases occurred in no endemic countries and exotic cases (cases with exotic species) occurred in endemic countries (Yera et al., 2006; Wicht et al., 2007, 2008a, 2008b; Paugam et al., 2009; Knoff et al., 2011; Semenas, 2014; An et al., 2017). In 2010, 214 million people lived outside their country of origin, these movements of people contributing to the displacement of foodborne parasites to different regions of the world, as evidenced by example in Dibothriocephalus spp. in groups of refugees (Robertson et al., 2014). Scholz and Kuchta (2016), gives precise data on the geographical distribution of dibothriocephalosis.

1.1.1.2 Europe

Dibothriocephalus latus has been considered to be the principal species infecting humans in Europe. It is autochthonous in northeastern Europe and subalpine lakes. Microscopic eggs of this species (morphologically diagnosed) were found in coprolites from 4,000 BC in France and Germany (Le Bailly et al., 2005). Human become infected with D. latus by eating raw freshwater fish as perch (Perca fluviatilis), pike (Esox lucius), Arctic char (Salvelinus alpinus), burbot (Lota lota), and other fish containing plerocercoid larvae of parasite. Recently, molecular identification of samples allowed the diagnosis of sporadic exotic clinical cases locally acquired from imported raw fish or overseas acquired in travelers. In this way D. nihonkaiensis cases were reported in France and Switzerland (Yera et al., 2006; Wicht et al., 2008a, 2008b; Paugam et al., 2009), D. dendriticus in Switzerland and Czech Republic (De Marval et al., 2013; Kuchta et al., 2013), A. pacificus (Kuchta et al., 2014; Pastor-Valle et al., 2014), Di. balaenopterae (Clavel et al., 1997; Pastor-Valle et al., 2014) and D. latus cases in Spain (Esteban et al., 2014).

1.1.1.3 North America

Dibothriocephalus latus seems to be native in North America and occurred in indigenous people and dogs (Dick, 2008). Most cases have been reported in the Great Lakes region, central Canada (Manitoba), and Alaska. Human become infected with D. latus by eating raw freshwater as pikeperch or walleye (Sander canadensis and S. vitreus). Dibothriocehalus dendriticus is found commonly throughout arctic and subarctic regions parasitizing piscivorous birds and mammals. Humans have infrequently been infected with this species in north Canada and Alaska (Kuchta et al., 2013). Humans become infected with D. dendriticus by eating raw salmonid including whitefish (Coregonus spp.) (Scholz and Kuchta, 2016). Others species D. dalliae, D. lanceolatum, D. ursi, D. alascensis (syn. Diphyllobothrium alascense) (Rausch and Adams, 2000), and more recently D. nihonkaiensis have been responsible for human infections in the Pacific coast of North America (British Columbia, Canada and North-West, USA) (Wicht et al., 2008b; Fang et al., 2015). Recently, Kuchta et al. (2017) reported plerocercoids of D. nihonkaiensis in the pink salmon, Oncorhynchus gorbuscha, from Resurrection Creek, Alaska.

1.1.1.4 South America

Dibothriocephalus latus was introduced in 20th century in Chile and Argentina where it is endemic in the southern freshwater ecosystems (Neghme et al., 1950; Torres et al., 2004a, 2012; Semenas and Úbeda, 1997; Semenas et al., 2001; Semenas, 2014; Mercado et al., 2010). In Chile, first human autochthonous case was reported in 1950 by Neghme et al. Around 90 cases of dibothriocephalosis attributed to D. latus have been published of patients who have consumed trouts, Oncorhynchus mykiss or Salmo trutta, both hosts of D. latus and D. dendriticus in lakes from southern Chile (Torres et al., 1993, 1998, 2004a; Torres et al., 2001; Mercado et al., 2010: Rosas and Weitzel, 2014). Some of the autochthonous cases have been identified by finding the microscopic egg parasites in feces of patient, other by studying adult stage or their proglottids, that can distinguished morphologically between the 3 species of diphyllobotriids identified in Chile, included D. latus, D. dendriticus and A. pacificus, this last in species of marine fish (Torres et al., 1981b; Andersen et al., 1987; Kuchta et al., 2013; Hernández-Ortz et al., 2015), or by genetic testing (Mercado et al., 2010). Prevalence of infection by D. latus fluctuated between 0.2% and 3.4% in humans living near of some lakes in southern Chile (Torres et al., 1989b, 1998, 2004a). Same authors indicated that domestic dogs are an important reservoir of dibothriocephalosis in the region.

In Argentina, to date there have been about 50 cases of human dibothriocephalosis reported, where the first case was identified in a European patient in 1906, but the first autochthonous case was recorded in 1982 in a fisherman frequent in the Andean-Patagonian lakes that had consumed introduced salmonids (Semenas, 2014).

More recently, based in the review and references cited by Knoff et al. (2011), about 70 cases attributed to D. latus or other unidentified species have been published in Brazil, including autochthonous and non-autochthonous cases (Eduardo et al., 2005a, 2005b; Tavares et al., 2005; Santos and Faro, 2005; Emmel et al., 2006; Capuano et al., 2007; Mezzari and Wiebbeling, 2008; Llaguno et al., 2008; Knoff et al., 2011). The cases in Brazil have been mainly related to the consumption of raw, smoked or insufficiently cooked imported salmonids. However, the presence of Diphyllobothrium plerocercoids in marketed marine fish, such as the pink cusk-eel, Genipterus brasiliensis, has been reported (Knoff et al., 2008). Recently, A. pacificus has been identified in the South American fur seal, Arctophoca australis (syn. Arctocephalus australis), a definitive host in the coast of Brazil (Jacobus et al., 2016).

Adenocephalus pacificus is the native South American diphyllobothriid. Eggs of this species (morphologically diagnosed) were found in coprolites from 2,700 to 2,850 BC in a coastal site in North-Central Peru (Reinhard and Urban, 2003) and at the site of Tiliviche in Northern Chile from 4,110-1,950 BC (Ferreira et al., 1984). Human infections have been reported with about 1,000 (Flores et al., 2002; Kuchta et al., 2015b), 19 (Sagua et al., 2001; Mercado et al., 2010), and 13 (Gallegos and Brousselle, 1991) human cases from Peru, Chile and Ecuador, respectively. Also, Kuchta et al. (2015b) mentioned a case of a student from Peru who was in Argentina and was infected with a tapeworm identified as Lueheella sp. (syn. Spirometra) and after revision of specimen revealed characteristics of A. pacificus. The origin of the infection in that patient it is not clear. The parasite displays a relatively low host specificity (found mainly in otariid mammals, and in accidental hosts such as man, dog and jackal. Human become infected with A. pacificus by eating raw marine fish as cebiche, tiradito, and chinguirito.

A human case identified as Diplogonoporus sp. was recorded in Chile by Wilhelm (1958); however, Figure 1 of the article shows proglottids with lateral and non-ventral and submedian genital pores as with Diphyllobothrium (syn. Diplogonoporus) (Torres et al., 1977).

Figure 1. Adult of Dibothriocephalus latus eliminated by a human patient

1.1.1.5 Asia

Cases of human infection by D. nihonkaiensis corresponds mainly to northeastern Asia (Scholz and Kuchta, 2016; Kuchta et al., 2017) and it would be the main cause of dibothriocephalosis in Japan and in Russia (Yamane et al., 1986; Dick et al., 2001; Cai et al., 2017). In Japan, numerous species of Diphyllobothrium have been reported (Kuchta et al., 2015a). However, molecular identification showed recently that most of these species were D. nihonkaiensis. Arizono et al. (2009b) based on a study of 149 cases, estimated that the mean incidence of infection by D. nihonkaiensis in Kyoto was 0.32 cases/100,000 inhabitants per year between 1988 and 2007, while in 2008, 1 case/100,000 inhabitants was detected.

In Korea, using genetic probes around 77 cases of D. nihonkaiensis have been identified since 1998 (Jeon et al., 2009; Kim et al., 2014; Shin et al., 2014; Song et al., 2014; Choi et al., 2015; Go et al., 2015; Park et al., 2015).

In China, about 11 cases of D. latus were reported between 1927 and 2011, mostly identified by morphological characteristics (Chen et al., 2014). Between 2008 and 2015 about 12 cases of D. nihonkaiensis have been reported (Chen et al., 2014; Zhang et al., 2015; Cai et al., 2017). Two of 4 cases reported recently acquired the infection in Canada and Russia (Cai et al., 2017). Also, a pediatric case of D. latus was confirmed by molecular analysis and attributed to consumption of imported raw fish from endemic areas (An et al., 2017).

In Russia, the parasite was reported as Diphyllobothrium klebanovskii, a synonym of D. nihonkaiensis, before the use of molecular identification (Arizono et al., 2009a). Few human cases, probably due to A. pacificus, have been reported in Sakhalin Island (Far East Russia) Kuchta et al., 2015b). Dibothriocephalus dendriticus has infrequently been reported in human in the region of Siberia, especially from the surroundings of Lake Baikal (Kuchta et al., 2013).

About 200 cases of human infections due to Diplogonoporus tapeworms, a synonymous of Diphyllobothrium, have been recorded in Japan (Yamasaki et al., 2012). Also, have been identified a case in Korea (Chung et al., 1995).

Twenty four human cases of D. stemmacephalum have been reported in Kanagawa Prefecture and the southwestern part of Japan as well as Okinawa Island and a coastal village in Korea (Yamasaki et al., 2016).

1.1.1.6 Australia and New Zealand

Autochthonous human cases of dibothriocephalosis have not been identified in Australia (Bearup, 1957), but one case by D. nihonkaiensis in New Zealand was reported (Yamasaki and Kuramochi, 2009).

1.1.1.7 Africa

In fecal samples of children from Nigeria have been founded eggs morphologically identified as D. latus (Ihesiulor et al., 2013), but more studies with adult parasites and molecular testing are required.

Different species of Dibothriocephalus have been responsible for few human cases from the India, Indonesia, Malaysia, Middle East, Mongolia, Saudi Arabia, and Taiwan (Alkhalife et al., 2006; Rohela et al., 2006; Devi et al., 2007; Lou et al., 2007; Margono et al., 2007; Myadagsuren et al., 2007). Some cases were probably imported from areas of endemicity (Alkhalife et al., 2006).

1.1.1.8 Sparganosis

Despite being a common infection in definitive hosts, cats and dogs, throughout much of the world, human cases of infection by Spirometra plerocercoid larvae or sparganosis occur mostly in East and Southeast Asia. Human sparganosis has cosmopolitan distribution and its endemic area include the East and Southeast Asia (China, Japan, Korea, Thailand, and Vietnam), and the USA, with sporadic cases reported elsewhere in Africa, South America, Europe, and Australia (Rahman et al., 2011; Tappe et al., 2013; Liu et al., 2015; Eberhard et al., 2015; Kuchta et al., 2015a). Imported cases have been described from people after travelling in endemic countries (Botterel and Bourée, 2003).

More than 1,685 cases have been reported worldwide, with more than 80.6% of them from China, 7.9% from Korea, 3.7% from USA and Thailand, 1.5% from Japan and between 0.1-0.5% of the cases from other countries (Argentina, Australia, Czech Republic, Ecuador, France, India, Italy, Paraguay, Sri Lanka, Tanzania, The Republic of Mozambique, Uganda, United Kingdom, and Venezuela) (Liu et al., 2015). In South America cases of human sparganosis also have been reported from Bolivia, Brazil, Colombia, Guiana, and Peru (Oda et al., 2016).

Spirometra erinaceieuropaei is found more frequently in the Old World and S. mansonoides occurs mainly in the New World. Recently, the DNA sequences were obtained for the first time from the Spirometra spp. from Brazil and were reported two species, Spirometra sp. 1 in the ocelot Leopardus pardalis, and Spirometra sp. 2 in the hoary fox Lycalopex vetulus and in the wild tropical rattlesnake (Crotalus durissus) (Almeida et al., 2016). These species differed from Asian isolates of S. erinaceieuropaei (Almeida et al., 2016). Spirometra decipiens is distributed in Brazil, China, and Korea and recently, a morphological study of two mitochondrial cox1 and nad3 genes of the Spirometra species from USA were sequenced concluding that also S. decipiens is present in that country (Jeon et al., 2016). These same authors suggest that S. decipiens was spread from South to North America through their hosts, dogs and cats.

A sister species to S. erinaceieuropaei has been responsible for few human cases in Ethiopia and Soudan (Eberhard et al., 2015). Thirty cases of sparganosis have been reported from Africa (Schmid and Watschinger, 1972). Molecular studies are need to determine if this species is related to species of Spirometra known to commonly infect, as definitive hosts, to large carnivores in East Africa (for example, S. theileri or S. pretoriensis).

Human cases of sparganosis by Sparganum proliferum have been recorded in North and South America and Asia (Noya et al., 1992). Kuchta et al. (2015a) compiled 16 cases reported from Japan, China, USA, Venezuela, Paraguay, and Czech Republic. Also, two cases were reported in Thailand (Anantaphruti et al., 2011). Some cases of infection in cats, dogs and feral hogs have been identified (Drake et al., 2008).

1.1.2 Symptomatology, morbidity, and mortality

Human dibothriocephalosis can cause important morbidity. In addition to be responsible of intestinal disorders and anemia, which can lead to specialized consultations and expensive complementary analyses; the parasitosis can have a substantial psychological impact on patients and their families, because proglottids of parasite are evacuated over a long period of time. The average cost for the management of a single dibothriocephalosis case has been evaluated to € 400 (Desvois et al., 2001). Human infection becomes patent (passage of proglottids or eggs in stools) after approximately 15 to 30 days after ingestion of plerocercoid (Freeman and Jamieson, 1972; von Bonsdorff, 1977; Curtis and Bylund, 1991; Go et al., 2015) and depend of diphyllobothriid and host species and other factors (Section 1.3). The infection is often asymptomatic (Kamiya and Ooi, 1991; Torres, 2013). People are just alerted of the presence of the tapeworm in the intestine because of the expulsion of chains of proglottids with the stools (Torres, 2013). The adult parasite can also be found accidentally when performing a colonoscopy (Lai and Steinhart, 2007; Stanciu et al., 2009). It is found attached to the mucosa in the small intestine (mostly in the ileum and jejunum). It measures from 1 m to 12 m, but could be up to 25 m in D. latus species (Table 1). It could live 10 to 15 years, but an infection lasting about 30 years has been reported (von Bonsdorff, 1977).

Table 1. Geographical distribution and morphological characteristics of adult stages of Dibothriocephalus spp., Adenocephalus pacificus, and Diphyllobothrium balaenopterae

Characteristics

D. latusa

D. nihonkaiensisb

D. dendriticusc

A. pacificusd

Di. balaenopteraee

Geographical distribution

Asia, Europe, North America, and South America (Chile and Argentina; also some human cases in Brazil)

 

China , Japan, South Korea, Far East, Russia up to Kamchatka Peninsula,

North America;

Human sporadic cases in France, Switzerland, and New Zealand.

North Hemisphere: Arctic, subarctic and temperate areas. South America (Southern Chile and Argentina)

South Hemisphere: Australia, Argentina Brazil, Chile, Ecuador, Peru, , Uruguay,

South Africa,

North hemisphere: Canada, Japan, Russia. United States, (Sporadic cases in Spain)

Human cases in Japan. Sporadic cases in Korea and Spain

In marine mammals in South Atlantic Ocean, Arctic and Antarctic, and Pacific Ocean.

Eggs

(range of length/wide by µm)

in different hosts

48 to 76 by 33 to 59.3

(Humans)

54 to 76 by 35 to 58 (Humans)

49 to 70 by 30 to 52 (Humans)

55 to 69 by 38 to 52

(dogs)

55 to 62 by 35 to 41

(gulls)

48.5 to 60 by 36 to 50 (humans)

41 to 67.5 by 35 to 48

(marine mammals)

49 to 60 by 36 to 45

(dogs)

57 to 81 by 34 to 51 (Humans)

Worm size: Length (in m) by wide in mm

as long as 25 length,

10 to 12 by 8 to 20 (Humans)

as long as 6 m by 5 to12

(Humans)

2 by 20

(Humans)

0.1 to 1.4 by 9

(dogs)

0.3 to 1 by 5.6 to 10

(gulls)

as long as 2 by 5 to 10 (Humans)

0.8 length

1.1 by 10 (dogs)

(marine mammals)

as long as 6.4 by 21

(Humans)

2.8 length (dogs)

7.4 by 10

(marine mammals)

Scolex length by wide in mm

Spatulate, ovalate or cordate.

1.5 to 2.7 by 0.6 to 1.0

Spatulate,

1.2 to 2.8 by 0.7 to 1.5

Spatulate or lanceolate,

0.5 to 2.2 by 0.5 to 0.8

Lanceolate, elongate, or oval,

1.8 to 4.0 by 0.6 to 3.5

Subglobular,

1.3 to 2.5 by 1.3

Neck length by wide in mm

4.1 to 30 by 0.6 to 1.4

6 to 16.8 by 1.2 to 1.3

0.6 to 3.2 by 0.3 to 0.8.

0.7 to 1.1 length.

Not present

Proglottids (PR)

Up to 4000

Up to 2200

Up to 968

Up to 269

Up to 2,648

Size of gravid proglotids (PR): Wide by length in mm

10 to 15 by 2 to 5 (Humans)

In experimental infections (1 to 2 months) last PR can be longer than wide.

6.7 to 12 by 1.9 to 2.5 (Humans)

2.4 to 10 by 0.7 to 3.9 (Humans)

Distal PR 1.8 to 3.1 by 2.3 to 8.0. Gravid PR with concave laterals margins. Pointed projections in the union of PR.

4 to 6.6 by 4 to 5.5 with tegmental papilla – like protuberances between margin PR and male gonopore

2.8 to 11.2 by 0.4 to 2.1

Distribution of vitelline follicles (VF) and testes (TE)

In lateral fields without confluence anterior to GA, except in some PR longer than wide. Area usually free of VF and TE between neighboring PR

Similar to D. latum

In lateral fields, often with confluence anterior to GA. VF not overlaps the uterine coils. No area free of VF and TE between neighboring PR

In lateral fields, slightly overlapping ends of loops of gravid uterus. TE not confluent at anterior margin of PR. VF sometimes confluent at anterior margin of PR

In two laterals and central field between 2 uteri, excepting areas around uteri and both GA. Sometimes VF confluent at anterior margin of PR

Gonopores in genital atrium (GA) surrounded by papillae

Yes, Ventral in first half of ventral surface

Yes, Ventral, on the middle at 1/3 or 1/4 anterior of the PR

Yes. In anterior third of PR. In PR longer than wide is posterior to the middle line

No, gonopores separates

Yes, with 2 GA, near anterior margin of PR, surrounded by papillae

Cirrus sac (CS). Length by wide in µm

Round or oval, with microtriches. 375 to 640 by 245 to 390. Horizontal or oblique in sagittal section

Oval, 430 to 480 by 275 to 390. Open obliquely in the genital atrium

Oblique or oval with microtriches. Somewhat oblique in sagittal section.

Subspherical to pyriform, unarmed. 105 to 325 by 89 to 243 in lateral view

Unarmed, pyriform, 262 to 600 by 250 to 368

Seminal vesicle (SV) Length by wide in µm

172 to 357 by 130 to 233. Dorso-caudal to CS. Visible ventrally in whole mounts (WM). Obtuse angle between CS and SV longitudinal axis

Round to elliptical 300 to 420 by 170 to 300. Dorso-caudal to the CS. Visible ventrally in WM. Sharp angle between CS and SV longitudinal axis

Round shape. Dorso-caudal to CS. Generally not ventrally visible in WM

Oval in ventral view 160 to 377 by 111 to 299 to almost elliptical in lateral view 76 to 223 by 106 to 291 in dosoventral length. Angle between CS and longitudinal axis (LA) of PR: 46°-94°. Angle between LA of SV and CS: 20°-143°

Ovoid to spherical. 151 to 230 by 110 to 177 extending slightly posterior from proximal end of CS

Shape of ovary

Variable. Dumbbell – like with 2 rounded or elongate lobes

Kidney shaped, without horns

Variable. Dumbbell shape. With 2 flattened elongated or round lobes

Asimetric bilobed, reticulate extended anteriorly adjacent to proximal uterine loops

Bilobed , reticulate extended anteriorly for both sides of the proximal coils

Number of uterine coils to each side/Uterine pore (UP)

4 to 8 extended up to posterior margin of CS. UP to 260-1240 µm, posterior to GA

6 to 7 no extended to the anterior margin of GA. UP to 150 to 300 µm of GA

2 to 9 extended up to CS. UP to 100 µm of GA

4 to 8 extended up to border of male gonopore. UP: 111 to 299 µm from vaginal aperture

3 to 5 loops extended up to the anterior margin of GA. UP to 100 to 200 µm from GA

Microtriches size (in µm)

Up to 4

8 to 9

7 to 10

Gladiade spinitriches (2) and capilliform filitriches

3.8

References: aAndersen, 1971, 1975; Andersen et al., 1987; von Bonsdorff, 1977; Dick and Poole, 1985; Kamo, 1999; Kuchta et al., 2015b; Rausch and Hilliard, 1970; Scholz et al., 2009; Torres, 2013; Torres et al., 1989a, 1993; Yamane et al., 1989a; Yazaki et al., 1984; bAndo et al., 2001; Arizono et al., 2009b; Choi et al., 2015; Fang et al., 2015; Jeon et al., 2009; Kamo, 1999; Wicht et al, 2008b; Yamane et al., 1986; Yamasaki and Kuramochi, 2009; Yera et al., 2006; cAndersen, 1975; Andersen et al., 1987; Dick and Poole, 1985; Figueroa et al., 1979, 1980; Kuchta et al., 2013; Rausch and Hilliard, 1970; Torres et al., 1981b; dBaer et al., 1967; Cattan et al., 1977; Escalante and Miranda, 1986; Hernández-Ortz et al., 2015; Jacobus et al., 2016; Jimenez et al., 2012; Kuchta et al., 2015b; Maejima et al., 1981; Miranda et al., 1968; Rausch et al., 2010; Torres et al., 1983; Tsuboi et al., 1993; eArizono et al., 2008; Clavel et al., 1997; Kino et al., 2002; Maejima et al., 1990; Nakamura et al.,1993; Rausch, 1964; Suzuki et al., 1988, Suzuki et al., 1993.

When the infection is symptomatic, the clinical signs are limited and polymorphous: common manifestations may include abdominal discomfort, abdominal pain, diarrhea, transit disorders, constipation, weight loss, asthenia, and vertigo. An extensive field investigation in Finnish Karelia showed that the worm carriers had a very significant higher frequency of fatigue, weakness, diarrhea and numbness of extremities (von Bonsdorff, 1977). A sensation of hunger and “craving for salt” was also associated with worm infection. Uncommon manifestations are headache, allergic reactions, pain in the tongue exacerbated with food, intestinal obstruction and a case in which D. latus was found in the gallbladder at cholecystectomy (von Bonsdorff, 1977). Patients can present low B12 levels, eosinophilia and, rarely, anemia (Nyberg et al., 1961; Kamiya and Ooi, 1991; Vuylsteke et al., 2004; Paugam et al., 2009).

Megaloblastic anemia due to vitamin B12 deficiency has been described in case of prolonged or heavy infection to D. latus in malnourished Finnish population after world war II (von Bonsdorff, 1977), but it is only rarely reported in other cestode infection (Golay and Mariaux, 1995). Approximately 80% of the B12 intake is absorbed by the worm. The mechanism is a parasite-mediated dissociation of the vitamin B12-intrinsic factor complex within the gut lumen, making B12 unavailable to the host. Megaloblastic anemia occurs in 2% or less people infected with D. latus; but about 40% of them may have low B12 levels (Nyberg et al., 1961). In Chile, 3 cases of megaloblastic anemia have been reported associated to D. latus infection (Osorio et al., 1974; Cristoffanini et al., 1976; Donoso-Scroppo et al., 1986). This deficiency may produce damage to the nervous system, including peripheral neuropathy or central nervous system degenerative lesions (von Bonsdorff, 1977). Unlike D. latus, infection with A. pacificus is seldom associated with megaloblastic anemia or vitamin B12 deficit and has been reported in only one case (Jimenez et al., 2012). Iron deficiency anemia has rarely been associated with human infection (Medina et al., 2002; Stanciu et al., 2009).

The diagnosis of human infection is based on the identification of proglottids expulsed with the stool or found after colonoscopy and identification of eggs after stool examination. In this last case, it is important consider that eggs of Bothriocephalus (adult parasites of fish) ingested accidentally by humans, are similar from that of Dibothriocephalus, Diphyllobothrium or Adenocephalus, using light microscopy by inexperienced microscopists. Differences between genera can be established by molecular identification (Yera et al., 2013). Analysis of nuclear genes (28S) allowed the identification of Bothriocephalus species. That of mitochondrial genes (cox1, NADH dehydrogenase subunit 3 nd3) allowed differentiating the most common Dibothriocephalus, Diphyllobothrium or Adenocehalus species involved in human infections (Yera et al., 2008). Analysis of nuclear genes (18S) is necessary to identify other species related to Dibothriocephalus or Diphyllobothrium (Waeschenbach et al., 2017). Recently, Lestinova et al. (2016) using light and scanning microscopy established differences between eggs of D. latus, D. nihonkaiensis, D. dendriticus, and A. pacificus. Also, a pyrosequencing method to differentiate among D. dendriticus, Dibothriocephalus ditremus (a not zoonotic parasite that live as adult in aquatic birds), D. latus, D. nihonkaiensis, Di. stemmacephalum, Di. balaenopterae, A. pacificus, Sp. decipiens, and Sp. proliferum has been described based on the mitochondrial cytochrome c oxidase subunit 1(cox1) gene as a molecular marker (Thanchomnang et al., 2016).

Human sparganosis can be responsible of death when the larvae invade vital organs (Phunmanee et al., 2001; Liu et al., 2015). It causes important morbidity. Severe health consequences can occur (i.e. blindness, immobilization, respiratory disorders, and neurological disorders) and lead to partial or total incapacity. The larvae migration into tissues can cause non-specific discomfort, eosinophilia, inflammation or abscess and others serious complications to vital organs. The larvae could live for up to 20 years in humans (Lee et al., 2010; Presti et al., 2015; Liu et al., 2015) and over 5-25 years in infection by Sp. proliferum (Liu et al., 2015).The survival of plerocercoids might be in relation with capacity to modify immune reactions in their hosts (Lee et al., 2010). The larvae have different locations in humans: intestine wall, scrotum, epididymis, ureter, subcutis, muscles, brain, cerebellum, eye, spinal cord, urinary tract, pleural cavity, pericardium, abdominal viscera, and oral cavity (Beaver and Rolon, 1981; Gutierrez, 2000; Iamaroon et al., 2002; Eom and Kim, 2009; Anantaphruti et al., 2011; Liu et al., 2015). Usually plerocercoids are slightly encapsulated in the tissues but insufficiently to prevent migration; in most cases the capsule is not present (Mueller, 1961).

The incubation period of human sparganosis depends on the route of infection: it is usually 6–11 days to some months, but it might be up to several years in oral infection (Lv et al., 2010). The first symptoms correspond to the migration of the larvae in the tissues. A review on human sparganosis in different countries reported 26% of cerebrospinal, 13% of subcutaneous, 4% of ocular, 1% of visceral, and 0.9% of sparganosis, but in 54% of cases the location of the infection was unknown (Liu et al., 2015). In Thailand, the locations of plerocercoids were the following: 46% subcutaneous, 35% ocular, 14% cerebral and 6% visceral (Anantaphruti et al., 2011). In Africa, 16 of 30 cases were subcutaneous (Schmid and Watschinger, 1972).

In cerebrospinal sparganosis, larvae invade and grow in the spinal canal or in the cerebral hemispheres, and, in some cases, extend to the cerebellum. The neurological symptoms, depending of the location of the parasite, are various as fatigue, limb weakness, confusion, headache, seizure, memory loss, coma, fever, paresthesia (as example, numbness or tingling) and hemiparesis (Kuchta et al., 2015a).

Subcutaneous sparganosis manifests as subcutaneous tumour or cyst. The lesion is slow growing and could be itchy, inflamed or painful. Sometimes, it is migratory or the larva emerges outside the skin (Eberhard et al., 2015). A single nodule is often present, whereas many nodules can arise in other larval cestode infections in humans, as example due to Taenia solium (Liu et al., 2015).

Eye is the main invasive location, even if ocular sparganosis is rare. The symptoms depend on the depth of the larva invasion. Extraocular infections (mostly subconjunctival or periorbital) are frequent. At the onset of the disease, patients present palpebral edema or conjunctivitis with eye pain, then kerato-conjunctivitis, ulceration of the cornea. Rarely, the retrobulbar area and the anterior chamber are involved. At this stage, patients had hypopyon, foreign body in the eye which can cause blindness and secondary glaucoma (Ament and Young, 2006). Visceral sparganosis can lead to abdominal pain, intestinal obstruction, peritonitis, pulmonary nodules with or without cavitation or eosinophilic pneumonia (Cheng et al., 2014).

Proliferative sparganosis is caused by Sp. proliferum a rare disease that affect particularly to immunocompromised patients (Schauer et al., 2014) .The patients present tumour-like, subcutaneous nodules that can spread to other parts of the skin, muscles, brain, lung, and abdomen or a bone tumor-like (Nakamura et al., 1990). Most cases are lethal (Kuchta et al., 2015a). The plerocercoid larva invades various organs of the body (including bone and spinal cord). It is pleomorphic as it forms continuous branches and buds to produce many progeny in a single site. The progenies can migrate to other tissues and repeat the processes of invasion, proliferation and dissemination. In an immunocompetent patient with molecular identification Sp. proliferum, the infection present only subcutaneous manifestations without signs of a proliferative course as dissemination and branching or budding (Schauer et al., 2014).

Specific imaging signs are present in cerebral sparganosis (Hong et al., 2013; Song et al., 2007) and subcutaneous sparganosis (Sakurawa et al., 2007) at CT, MRI and ultrasonography and could allow the diagnosis of the infection. Eosinophilia is possible. Serologic tests (detection of antibodies or antigens from peripheral blood or cerebrospinal fluid) are useful for diagnosis, but available in endemic countries. They are sensitive and enough specific, but cross-reactivity is possible in patients presenting cysticercosis, echinococcosis, paragonimosis, clonorchiosis, and schistosomosis (Cui et al., 2011). In consequence, diagnosis is based on the histological identification of the larvae in tissues. PCR seems promising for identification of larvae from archival tissues and for species identification.

1.2 Taxonomic Classification of the Agents

1.2.1 Taxonomy and physical description of the agent
1.2.1.1 General aspects

The Diphyllobothriidae family includes zoonotic parasite species of importance in human and animal health which are grouped principally into genera Dibothriocephalus, Diphyllobothrium, Adenocephalus, and Spirometra. In general, the species of those genera have larval stages which develop in the aquatic environment (coracidia), and have intermediary host copepods (procercoids) and fish (plerocercoids or sparganums) in their life cycles. In addition, these species include a definitive host, where they develop into the adult stage in the small intestine (mammals or birds, the latter, in only some species of Dibothriocephalus). Life cycle in Spirometra spp. is similar, but, plerocercoid develops in amphibians, snakes, birds and mammals, including humans, and their definitive host corresponds to canids and felids. Dibothriocephalus spp. and Spirometra spp. occur in freshwater and terrestrial habitats, while Adenocephalus, including A. pacificus, and Diphyllobothrium, including Di. stemmacephalum and Di. balaenopterae, occur in marine habitats (Waeschenbach et al., 2017).

1.2.1.2 Taxonomic position

The parasites of the family Diphyllobothriidae are included in the phylum Platyhelminthes, subphylum Rhabditophora, class Neoophora, subclass Neodermata, infraclass Cestoda, and order Diphyllobothriidea (Ruggiero et al., 2015). Very recently, a new molecular phylogeny of diphyllobothriideans species has been published (Waeschenbach et al., 2017). It is based on analysis of two nuclear genes (almost completed small subunit ribosomal RNA gene, 18S and partial large small subunit ribosomal RNA gene, 28S) and two mitochondrial ones (partial small subunit ribosomal RNA gene, 16S and cytochrome c oxidase subunit 1 gene, cox1). Diphyllobothriideans species are classified into three different families: Cephalochlamydidae, Solenophoridae and Diphyllobothriidae. In this classification, the family Diphyllobothriidae presents 13 valid genera (Waeschenbach et al., 2017). Of these genera, Dibothriocephalus, Diphyllobothrium, Adenocephalus, and Spirometra are more frequently associated with human disease. The genus Dibothriocephalus was resurrected including freshwater and terrestrial species with D. latus, D. nihonkaiensis and D. dendriticus (ex-Diphyllobothrium latum, D. nihonkaiense or D. dendriticum). Diplogonoporus is determined as a junior synonym of Diphyllobothrium, as suggested by previous molecular data (Yamasaki et al., 2012). Diphyllobothrium accommodate parasites from cetaceans including the type species Di. stemmacephalum and Di. balaenopterae.

Six Dibothriocephalus species (D. latus, D. dendriticus, D. nihonkaiensis, D. alascensis, D. dalliae, and D. ursi), 9 Diphyllobothrium species (Di. stemmacephalum, Di. elegans, Di. cameroni, Di. cordatum, Di. hians, Di. lanceolatum, Di. orcini, Di. scoticum, and Di. balaenopterae), and the only Adenocephalus species (A. pacificus) have been reported in humans (Hernandez-Ortz et al., 2015; Scholz and Kuchta, 2016; Waeschenbach et al., 2017). The species most frequently associated with human infection are D. latus, D. nihonkaiensis, D.dendriticus, A. pacificus, and Di. balaenopterae (Yamasaki et al., 2012; Scholz and Kuchta, 2016).

Faust et al. (1929) established Spirometra as a subgenus of Diphyllobothrium considering characteristics of the eggs and uterine coils. However, Mueller (1937) included Spirometra as an independent genus. Of the 11 species considered by Wardle and McLeod (1968), Kamo (1999) cosiders as valid species to S. erinaceieuropaei (syn. S. mansoni, S. erinacei), S. pretoriensis, S. theileri and S. mansonoides. Of the Spirometra species, the most related to human cases are S. erinaceieuropaei, S. mansonoides, and S. decipiens. Sparganum proliferum corresponds to a larval stage (plerocercoid), while the adult is unknown. Plerocercoid has been considered be an aberrant form of S. erinaceieuropaei or S. mansonoides. Through the use of genetic testing it was concluded that Sp. proliferum is different from S. erinaceieuropaei and molecular data suggest it is a species of Spirometra (Miyadera et al., 2001; Okamoto et al., 2007; Liu et al., 2015; Almeida et al., 2016).

1.2.1.3 Physical description

1.2.1.3.1 Adult stage

Morphological characteristics of Dibothriocephalus spp., A. pacificus, and Di. balaeopterae are indicated in Table 1. Adults can be very long (Figure 1); D. latus can reach 25 m in length by 2 cm in width (von Bonsdorff, 1977). At their anterior end they have a scolex with two longitudinal grooves or bothria (Figure 2), ventral and dorsal ones, which adhere to the mucosa of the small intestine. The scolex presents glands whose ducts open in the tegument (Kuperman and Davydov, 1982). A neck posterior (Figures 2-3) to the scolex of variable length gives rise to a strobila consisting of segments or proglottids (Figure 3), usually wider than they are long. The tegument has finger-like projections or microtriches that increase the surface area 10-fold facilitating an efficient transport of all of the dietary requirements into the parasite (Dalton et al., 2004).

Figure 2. Scolex showing a bothrium (B), and the neck (N) of adult stage of D. latus. (Scanning electron microscopy, SEM)

Figure 3. Scolex (S), neck (N) and immature proglottids (I) of adult of D. latus. Semichon´s acetic carmine stain

The male and female reproductive organs included in each proglottid a set (Dibothriocephalus, Diphyllobothrium, Adenocephalus, Spirometra), or a paired (Di. balaenopterae) of reproductive organs (Bray et al., 1994; Kamo, 1999). Male reproductive organs include testes (Figures 4-5) with their respective efferent ducts that join together in a deferens duct which widens forming an external muscular seminal vesicle (Figure 6) attached to the proximal part of the cirrus sac (Figure 6) including the copula organ (cirrus) that opens in the ventral gonopore (Figure 6). In Spirometra spp. the dorsal portion of the cirrus sac represents the seminal vesicle from other genera. The female organs include an ovary, usually lobed (Figure 5), in the posterior region of the proglotid provided with an oviduct, connected to a structure called the ootype that is surrounded by a Mehlis gland, from which originated the uterus. The ootype communicates with the vagina and vitelline reservoir that stores vitelline cells formed in the vitelline follicles. The vagina opens into a ventral gonopore behind the male gonopore (Figure 6). The uterus has a ventral opening (uterine pore), located behind the female gonopore (Figure 6).

Figure 4: Gravid proglottid of D. latus: genital atrium (G), seminal vesicle (SV), uterus (U) testes and vitelline glands (TV). Semichon´s acetic carmine stain

Figure 5. Gravid proglottid of Dibothriocephalus dendriticus: genital atrium (G), uterus (U) testes and vitelline glands (TV), ovary (O). Semichon´s acetic carmine stain

Figure 6. Longitudinal section of gravid proglottid of D. latus: male (M) and female (F) gonopores and uterine pore (P) in ventral area, seminal vesicle (SV) dorsal and posterior to cirrus sac (C), and uterus (U) with eggs. Hematoxylin and eosin stain

Proglottids of Dibothriocephalus spp. and Diphyllobothrium spp. have a common genital atrium that includes male and female gonopores. In Adenocephalus and Spirometra there is no common genital atrium because female and male gonopores open separately (Bray et al., 1994; Hernández-Ortz et al., 2015). Diphyllobothrium balaenopterae presents 2 genital atria (Bray et al., 1994). Adenocephalus differs from other Diphylobothriidae with the presence of protuberance like- papillae separated by semi-circular transverse grooves on the ventral surface, between the male gonopore and the anterior border of the proglottids (Rausch et al., 2010; Hernández-Ortz et al., 2015). The uterus of Spirometra resembles a spiral with 2 to 7 compressed and anteriorly extended coils. In Dibothriocephalus, Diphyllobothrium, and Adenocephalus the uterus has several loops which may be parallel and extend laterally or rosette-shaped. Spirometra eggs tend to be pointed at one end (Figure 7) (Bray et al., 1994), while in other genera they are rounded (Figure 8).

Figure 7. Egg of Spirometra sp. from feces of domestic cat infected with an adult stage: operculum (O)

Figure 8. Egg of Dibothriocephalus sp. from feces of a human patient infected with an adult stage: operculum (O)

Adults of S. erinaceieuropaei measure about 1 m long by 1 cm wide and S. mansonoides measure up to 1.5 m long by 0.7 cm wide (Mueller, 1974; Bowman et al., 2002). The scolex of S. mansonoides measures up to 0.5 mm long (Mueller, 1974) with 2 bothria of 1 mm in length; the U-shaped uterus terminates in the anterior of the proglottid and presents two sections: anterior heavy outer coils and posterior narrow inner coils joined by a narrow duct (Bowman et al., 2002). In S. mansonoides the uterus form is simple, uniform and always presents two anterior turns as opposed to the uterus of S. erinaceieuropaei which lacks uniformity in the number of loops, besides having irregular arrangements and sizes (Mueller, 1974; Scioscia et al., 2014). A uterus with 5-7 coils on each side and a gravid proglottid with a width of 3.1-4.2 mm by 2.1-3.2 long in S. erinaceieuropaei; and a uterus with 4-4.5 coils and gravid proglottids with a width of 6.8-10.8 mm by 1.1-1.2 mm long in S. decipiens have been reported (Jeon et al., 2015). Both species show differences in the cox1 gene (Jeon et al., 2015). Okino et al. (2017) obtained a clonal population from a single adult specimen designated as a Kawasaki triploid strain whose partial sequence of mitochondrial cytochrome c oxidase subunit 1 gene showed more than 98% similarity with isolates from Australia, China and South Korea. However, gravid proglottids of Kawasaki strain showed 3 to 4 uterine coils, showing similarity with S. decipiens. Although the etiological agents of sparganosis in Asia can be divided into two major groups by mtDNA sequences (Eom et al., 2015; Jeon et al., 2015, 2016), their specific status is highly ambiguous, particularly in parthenogenetic triploid clones and further taxonomic reconsiderations are needed (Okino et al., 2017).

1.2.1.3.2 Eggs and larvae

Eggs of Diphyllobothriidae species are surrounded by a thin proteinaceous vitelline capsule secreted by vitelline cells at the time of fertilization, and a shell of highly resistant protein (sclerotin) formed by a polyphenol/quinone tanning process on the inner surface of the vitelline capsule (Conn and Swiderski, 2008). Eggs have a suture at one end, forming an operculum (Figures 7-9). Internally the eggs have a zygote and vitelline cells. The size of the eggs is variable and depends of various factors, such as the host species, the intensity of infection and the time of exposition (Andersen and Halvorsen, 1978).

Figure 9. Uterus of D. latus with eggs: operculum (O). SEM

Table 1 shows the interval of measurements of eggs corresponding to Dibothriocephalus spp., A. pacificus, and Di. balaenopterae. The size (length, width and length/width ratio) and surface characteristics of the egg shell, using light and scanning electron microscopy have made it possible to distinguish some diphyllobothriideans, such as those that commonly infect humans (Lestinova et al., 2016). Eggs of Spirometra spp. measure about 60-66 µm in length by 35-36 µm wide (Lee et al., 1990; Bowman et al., 2002; Liu et al., 2015).

Coracidium: During their development two syncytial embryonic envelopes appear: a) an outer envelope, surrounding the inner embryonic components and larval stage, and b) an inner envelope, forming a ciliated embryophore (Figure 10), surrounding the hexacanth larva (Figure 10). Both structures formed the coracidium (Conn and Swiderski, 2008) that measures about 50 µm. The hexacanth larva has a tegument (Grammeltvedt, 1973) and different cell types (Swiderski and Mackiewicz, 2004) and 3 pairs of hooks (Figure 10) that differ significantly between D. latus, D. nihonkaiensis, and D. dendriticus (Bylund, 1975; Yamane et al., 1989b).

Figure 10. Coracidium of D. latus: hooks (H) of hexacanth larva and cilia (C). Thionin stain

Procercoid: It develops from the hexacanth larva that hatches in the gut of the first intermediate host (copepod species component of zooplancton) and after being located in its hemocele (Figure 11) (Janicki and Rosen, 1917; Guttowa, 1961; Torres et al., 2004b, 2007). Infective procercoid for the second intermediate host presents calcareous corpuscles, glands with ducts well developed and a posterior invagination (cercomer) containing the embryonic hooks (Figures 12-13) (Janicki and Rosen, 1917; Grammeltvedt, 1973; Torres et al., 2004b). The tegument of D. dendriticus procercoids presents microtriches up to 1.7 µm long (Malmberg, 1971; Grammeltvedt, 1973). Infective procercoids in D. latus measure between 180 and 610 µm long, depending on the host species, exposure time and intensity of infection (Guttowa, 1961; Torres et al., 2004b). In D. dendriticus they measure between 288 and 576 µm (Halvorsen, 1966) and in Spirometra spp. they reach about 260 µm long by 44-100 µm wide (Liu et al., 2015).

Figure 11. Copepod, Tumeodiaptomus diabolicus, infected with procercoids (PR) of D. latus

Figure 12. Procercoids of D. latus extracted from T. diabolicus, showing ducts of glands (D), calcareous corpuscles (C) and cercomer (CE)

Figure 13. Cercomer (CE) of procercoid of D. latus showing some of their hooks

Plerocercoid: these develop in the second intermediate host from infecting procercoids in ingested copepods. The plerocercoids measure 1 mm to several cm in length and present a scolex with 2 bothria (Figure 14). The tegument includes microtriches (Figure 15). Plerocercoids are located in the mesenteries and in different viscera and muscles of fish. The morphological characteristics of the principal species are indicated in Table 2. These larvae present two types of frontal glands (Green and Golden glands) in the parenchyma of scolex and neck (Gustafsson and Vaihela, 1981; Kuperman and Davydov, 1982). The plerocercoids have a muscular layer separating the parenchyma in an internal or medullar region and an external or cortical region (Figure 16), except, in Spirometra spp. where there are groups of muscle fibers unevenly distributed. Spirometra plerocercoids measure between 1 mm to 50 cm long (Mueller, 1966; Bo and Xuejian, 2006; Anantaphruti et al., 2011; Tappe et al., 2013; Lee et al., 2014) and present a scolex with 2 bothria and tegument with microtriches up to 2.4 µm in S. erinaceieuropaei (Yamane, 1968). Sparganum proliferum plerocercoids can reproduce asexually through a process of branching and budding producing abundant progeny that invade various tissues (Anantaphruti et al., 2011; Tappe et al., 2013; Lee et al., 2014). They measure 38 mm long by 3 mm wide (Noya et al., 1992) and their tegument have irregular branches and buds, lacking bilateral symmetry. The parenchyma presents cavities covered by tegument which appear to contain gelatinous material (Noya et al., 1992; Drake et al., 2008).

Figure 14. Plerocercoid of D. dendriticus. SEM

Figure 15. Microtriches in the tegument of D. dendriticus plerocercoid. SEM

Table 2. Morphological characteristics of plerocercoids of Dibothriocephalus spp. and Adenocephalus pacificus

Characteristics

D. latusa

D. nihonkaiensisb

D. dendriticusc

A. pacificusd

Scolex

Normally retracted and not visible. Anterior extremity with transverse dorso-ventral ridge

Evaginated, with dorsal and ventral median grooves like lips

Partially retracted. Interbothridial groove visible. Transverse fissure (frontal pit) at anterior end varies in depth

Extruded with bothrias of 0.5 to 1.4 mm length

Body

Wrinkled. Body widest at the anterior third

Wrinkled with cylindrical external appearance, body widest at the middle.

Wrinkled

Wrinkled. Body widest at the anterior region

Segmentation and primordia

Absent

Absent

Present in large larvae

Absent

Size: length/wide in mm

2 to 50/1 to 1.1

6.2 to 10.7/0.9 to 1.3

9 to 30/1 to 3. Relaxed in water up to 45 or more mm in length

4 to 25/0.5 to 2.4

Microtriches: length in µm

1 to 3

2.5 to 10

3 to 16

3 to 4

Subtegumentary longitudinal muscles (N° of layers)

1

1

1 to 2

NRe

Frontal glands

In the scolex. Extend into first third of body

In the scolex region

In the scolex region

NR

Location

Usually free in musculature or body-cavity; also in pike and salmonids encysted.

Encysted or free in the musculature of anadromous salmonids

Usually encysted in viscera or in body cavity or free in musculature.

Encysted in the surface of stomach, intestine, liver or peritoneum of marine fish

aAndersen et al., 1987; Andersen and Gibson, 1989; von Bonsdorff, 1977; von Bonsdorff and Bylund, 1982; Torres et al., 1983, 1989a; bSuzuki et al., 2010; Yamane et al., 1986; cAndersen et al., 1987; Andersen and Gibson, 1989; Halvorsen, 1970; Kuchta et al., 2013; Kuhlow, 1953; Torres et al., 1981b; Yamane et al., 1989a; dEscalante, 1983; Escalante et al.,1988; Escalante and Miranda, 1986; Tantalean, 1975; Tantalean and Huiza, 1994; eNot Reported

Figure 16. Histological cross section of D. dendriticus plerocercoid: tegument (T), and cortical (CO) and medullar (ME) regions, separated by a muscular (MU) layer. Hematoxylin and eosin stain

1.3 Transmission

1.3.1 Life cycle, routes of transmission, and reservoirs

Adults of D. latus live in the small intestine of humans, domestic dog, domestic cat, foxes and other fish-eating mammals (von Bonsdorff, 1977; Andersen et al., 1987). Parasite eggs, eliminated with the feces of the host, develop a ciliated larva (coracidium) when in contact with fresh water, depending on the temperature. The coracidium will then hatch, influencing this process the wavelength and intensity of light (Grabiec et al., 1963). In the water coracidia can live between 1 to 3 days depending on the temperature and relative depletion of their energy reserves. Following ingestion of the coracidia by copepods, four possible situations may occur: 1) Hatching of hexacanth larva in the gut and the displacement to the hemocele which develop into the infecting procercoid for planktivorous fish, 2) hatched larvae can be damaged by host enzymes leaving only a few surviving to develop in the hemocele, 3) hatched larvae die by action of the enzymes of the host and those who come to hemocele would die by the host response, and 4) the coracidia remain unhatched in the copepod due to unfavorable conditions.

Table 3 shows host copepod species of D. latus. The infective procercoids measure 500-600 µm in length after 15 to 20 days of exposure to laboratory temperature in Cyclops strenuus (Janicki and Rosen, 1917), while in Eudiaptomus transylvanicus (syn. Diaptomus coeruleus) and Eudiaptomus gracilis reach 270-500 µm and 240-450 µm in length, respectively at 12 to 15 days at 18-20°C (Guttowa, 1961). On the other hand, procercoids grow to between 180 to 610 µm in length from 11 days of infection at 20 ± 1°C in Tumeodiaptomus diabolicus (syn. Diaptomus diabolicus) (Torres et al., 2004b). The establishment of infections in humans depends on their eating habits and whether they are associated to the consumption of dishes made with raw fish, cold smoked, treated by salting or undercooked. Of the 29 species of copepods (Table 3), 18 are highly susceptible to D. latus, being 15 (83%) of the order Calanoida and only 3 (17%) of the order Cyclopoida. The susceptibility of some species of copepods depends on their age, physiological compatibility, ecological conditions and geographic origin of host and parasite (Guttowa, 1961; Halvorsen, 1966; Bylund, 1969; Sharp et al., 1990, Marcogliese, 1995).

Table 3. Zooplanktonic copepod hosts of Dibothriocephalus latus in different countries

Country

Copepod hosts

Reference

Australia

Mesocyclops leuckartia,c

Guttowa, 1961; von Bonsdorff, 1977

Australia

Gladioferens brevicornisb,c

Guttowa, 1961; von Bonsdorff, 1977

Australia

Boeckella minutab

Guttowa, 1961; von Bonsdorff, 1977

Australia

Microcyclops varicansa,c

Guttowa, 1961; von Bonsdorff, 1977

Australia, Poland

Eucyclops serrulatusa,c

Guttowa, 1961; von Bonsdorff, 1977

Chile

Tumeodiaptomus diabolicus (= Diaptomus diabolicus)b

Torres et al., 2004b, 2007

Chile

Boeckella gracilisb

Torres et al., 2004b, 2007

Chile

Mesocyclops araucanusa,c

Torres et al., 2004b, 2007

Finland, Poland

Cyclops vicinusa

Guttowa, 1961; von Bonsdorff, 1977

Finland, Poland

Eudiaptomus zachariasib,c

Guttowa, 1961; von Bonsdorff, 1977

Finland, Poland

Cyclops insignisa,c

Guttowa, 1961; von Bonsdorff, 1977

Finland, Germany, Karelia, Norway, Poland, Switzerland

Cyclops strenuus (= C. furcifer)a

Guttowa, 1961; von Bonsdorff, 1977

Germany, Poland

Eudiaptomus transylvanicus(= E. coeruleus)b

Guttowa, 1961; von Bonsdorff, 1977

Germany, Finland, Ireland, Karelia, Norway, Poland, Switzerland

Eudiaptomus gracilisb

Hickey and Harris, 1947; Guttowa, 1961; von Bonsdorff, 1977

Karelia

Thermocyclops oithonoides (= Mesocyclops oithonoides)a,b

Guttowa, 1961; von Bonsdorff, 1977

Karelia, Finland, Germany

Eudiaptomus graciloidesb

Guttowa, 1961; von Bonsdorff, 1977

Norway

Acanthodiaptomus denticornisb

Cyclops lacustrisa,c

Guttowa, 1961; von Bonsdorff, 1977

Russia

Arctodiaptomus ulomskyib

Guttowa, 1961; von Bonsdorff, 1977

USA

Acanthocyclops vernalisa,c

Guttowa, 1961; von Bonsdorff, 1977

USA

Eurytemora affinisb

Guttowa, 1961; von Bonsdorff, 1977

USA

Tropocyclops prasinusa,c

Guttowa, 1961; von Bonsdorff, 1977

USA

Skistodiaptomus oregonensis (=Diaptomus oregonensis)b

Guttowa, 1961; von Bonsdorff, 1977

USA

Skistodiaptomus mississippiensis (= D. mississippiensis)b

Guttowa, 1961; von Bonsdorff, 1977

USA

Leptodiaptomus siciloides (= D. siciloides)b

Guttowa, 1961; von Bonsdorff, 1977

USA

Leptodiaptomus minutus (= D. minutus)b,c

Guttowa, 1961; von Bonsdorff, 1977

USA

Aglaodiaptomus leptopus (= D. piscinae)b

Guttowa, 1961; von Bonsdorff, 1977

USA

Onychodiaptomus sanguineus (= D. sanguineus)b

Guttowa, 1961; von Bonsdorff, 1977

USA

Mastigodiaptomus albuquerquensis (= D. albuquerquensis)b

Guttowa, 1961; von Bonsdorff, 1977

aOrder Cyclopoida; bOrder Calanoida; cSpecies less viable to infection or only accidentally infected

Infections in E. gracilis cause a significant drop in consumption of oxygen in the host after 6-10 days of exposure, when there is greater demand for building materials and secretion of metabolites by the parasite which could induce a physiological imbalance in the host (Klekowski and Guttowa, 1968). In mixed infections by D. latus and Proteocephalus sp. in E. gracilis the procercoids can cause death of the host during organogenesis (Guttowa, 1966).

Infected copepods transmit the infection to planktivorous fish. The procercoids reach the gut and migrate to the wall of the stomach or intestines, mesenteries, viscera and muscles. In fish, procercoid develops into plerocercoid. These larvae are found free (Figures 17-18) or encapsulated in the tissues (Figures 19-20), depending on the host species. Dibothriocephalus latus present about 17 fish host species distributed in 6 orders (Table 4). Infection in native fish from Chile, such as Galaxias spp., Percichthys trucha, Odontesthes mauleanum, and Basilichthys australis have only been observed in 2 lakes with a low intensity of infection in contrast to the high parasitic load of introduced rainbow trout (Torres et al., 1989b, 1998, 2004a, 2012).

Figure 17. Free plerocercoid (P) of Dibothriocephalus sp. in the muscular surface of a rainbow trout, Oncorhynchus mykiss

Figure 18. Histological section of the liver of a rainbow trout with a free plerocercoid (P) of Dibothriocephalus sp. Hematoxylin and eosin stain

Figure 19. Encapsulated plerocercoids (P) of Dibothriocephalus sp. in the stomach of a rainbow trout

Figure 20. Histological section of muscle of a rainbow trout with an encapsulated plerocercoid (P) of Dibothriocephalus sp. Hematoxylin and eosin stain


Table 4. Fish hosts of plerocercoids of Dibothriocephalus spp. and Adenocephalus pacificus

Species: Dibothriocephalus latusa

Area

Hosts

Europe (EU)

 

Gadiformes:

burbot, Lota lota

Perciformes:

perch, Perca fluviatilis

ruffe, Gymnocephalus cernuus;

Sander, Sander lucioperca

EU and Asia

Salmoniformes:

brown trout, Salmo trutta

char, Salvelinus fontinalis

EU and North America

Esociformes:

pike, Esox lucius

EU and North America and South America

Salmoniformes:

rainbow trout, Oncorhynchus mykiss

North America

Perciformes:

yellow perch, Perca flavescens

wallege, Sander vitreus

sauger, Sander canadensis

South

America

 

 

Salmoniformes:

Coho salmon, Oncorhynchus kisutch

Osmeriformes:

puye, Galaxias maculatus

puye, Galaxias platei

Atheriniformes:

cauque, Odontesthes mauleanum

silverside, Basilichthys australis

Perciformes:

creole perch, Percichthys trucha

Species: Dibothriocephalus dendriticusb

Asia

Salmoniformes:

Baikal omul, Coregonus migratorius

EU

Gasterosteiformes:

nine-spined stickleback Pungitius pungitius

Salmoniformes:

common whitefish, Caregonus lavaretus

vendace, Coregonus albula

lake Constance whitefish, Coregonus gutturosus

whitefish, Coregonus pallasii

Valaam whitefish, Coregonus widegreni

Osmeriformes:

grayling, Thymallus thymalus

EU and North America

Gasterosteiformes:

three-spined stickleback, Gasterosteus aculeatus

Salmoniformes:

Arctic char, Salvelinus alpinus

EU and North America and South America

Salmoniformes:

rainbow trout, Oncorhynchus mykiss;

brown trout, Salmo trutta

North America

Salmoniformes:

Atlantic salmon, Salmo salar

lake white fish, Coregonus clupeiformis

Osmeriformes:

rainbow smelt, Osmerus mordax

North America and South America

Salmoniformes:

char, Salvelinus fontinalis

South America

Salmoniformes:

Coho salmon, Oncorhynchus kisutch

Osmeriformes:

puye, Galaxias maculatus

Perciformes:

Percichthys trucha

Atheriniformes:

cauque, Odontesthes mauleanum

silverside, Basilichthys australis

Siluriformes:

tollo, Diplomystes camposensis.

Species: Dibothriocephalus nihonkaiensisc

Asia

Salmoniformes:

masu salmon, Oncorhynchus masou;

pink salmon, Oncorhynchus gorbuscha,

chum salmon, Oncorhynchus keta

North America

Salmoniformes:

Oncorhynchus gorbuscha

Species: Adenocephalus pacificusd

South America

(coast of Peru)

Perciformes:

Peruvian grunt, Anisotremus scapularis.

Corvine drum, Cilus gilberti.

common dolphinfish, Coryphaena hippurus

Peruvian weakfish, Cynoscion analis

kingfish, Menticirrhus ophicephalus

Peruvian banded croaker, Paralonchurus peruanus

Lorna drum, Sciaena deliciosa

Callao drum, Sciaena callaensis

Pacific mackerel, Scomber japonicus

Atlantic Spanish mackerel, Scamberomorus maculatus

Eastern Pacific bonito, Sarda chiliensis

Palm ruff, Seriolella violacea

paloma pompano, Trachinotus paitensis

Jack mackerel, Trachurus murphyi

Peruvian rock seabass, Paralabrax humeralis

Siluriformes:

Colombian shark fish, Ariopsis seemanni.

Peruvian sea catfish, Galeichthys peruvianus.

Ophidiiformes:

pink cusk-eel, Genypterus maculatus.

Gadiformes:

Peruvian hake, Merluccius gayi peruanus

Mugiliformes:

flathead grey mullet, Mugil cephalus

Carcharhiniformes:

sickle fin smooth-hound, Mustelus lunulatus

Pleuronectiformes:

fine flounder, Paralichthys adspersus

South America

(coast of Chile)

Plerocercoids identified as Diphyllobothriidae or Diphyllobothrium sp.e

Perciformes:

corvina drum, Cilus gilberti.

Gadiformes:

South Pacific hake, Merluccius gayi gayi

Southern hake, Merluccius australis;

Southern blue whiting, Micromesistius australis,

Ophidiiformes:

pink cush-ell, Genipterus maculatus.

aAndersen and Gibson, 1989; De Vos and Dick, 1989; Halvorsen and Wissler, 1973; Janicki and Rosen, 1917; Kuchta et al., 2015a; Mercado et al., 2010; Revenga, 1993; Scholz et al., 2009; Thanchomnang et al., 2016; Torres et al., 1989b, 1998, 2004a, 2012

bBylund, 1972; Chubb, 1980; De Vos and Dick, 1989; Gustafsson and Vaihela, 1981; Henricson, 1977; Kuchta et al., 2013; Moravec, 2004; Revenga, 1993; Thanchomnang et al., 2016; Torres, 1990; Torres et al., 1981b, 1989b, 1998, 2004a, 2012; Vik, 1957

cAndo et al., 2001; Fang et al., 2015; Jeon et al., 2009; Kuchta et al., 2017; Suzuki et al., 2010; Yamane et al.,1986; Yanagida et al., 2010

dEscalante, 1983; Escalante and Miranda, 1986; Chero et al., 2014a, 2014b, 2014c; Iannaconne and Alvariño, 2009Kuchta et al., 2015b; Tantalean, 1975; Tantalean and Huiza, 1994

eGarcías et al., 2001; George-Nascimento and Huet, 1984; George- Nascimento et al., 2011; MacKenzie and Longshaw, 1995; Oliva and Ballón, 2002 

Chubb (1980) in their review indicate the following species of fish that were experimentally infected with procercoids: E. lucius, Gymnocephalus cernua, and P. fluviatilis. Similarly, in E. lucius, G. cernua, P. fluviatilis, Rutilus rutilus and O. mykiss plerocercoids infection have been restored after administering orally. Thus, planktivorous fish can transmit the infection to fish-eating fish that act as paratenic hosts, for example, E. lucius, P. fluviatilis and L. lota in the northern hemisphere (von Bonsdorff and Bylund, 1982) or O. mykiss and P. trucha in the lakes of southern Chile (Torres et al., 1989b, 1998, 2012) and Argentina (Revenga, 1993).

Seven of the 10 species of copepods susceptible to infection by D. dendriticus (Table 5) belong to the order Cyclopoida, all described in Europe and North America. In South America the host copepods are unknown. The procercoids of Dibothriocephalus spp. in natural infections of C. strenuus abyssorum reduce the reproductive capacity of females, the feeding and the respiration rate as well as causing a reduction in motility, and impaired escape responses, which results in the individual being more vulnerable to predators (Pasternak et al., 1995).

Table 5. Zooplanktonic copepod hosts of Dibothriocephalus dendriticus in different countries

Area

Copepods

Reference

Canada

Aglaodiaptomus leptopus (= Diaptomus leptopus)b

Wright and Curtis, 2000

Canada

Leptodiaptomus minutus (= Diaptomus minutus)b

Wright and Curtis, 2000

Finland

Germany

Norway

United  Kingdom

 

Cyclops strenuusa

Rahkonen and Valtonen, 1998;

Kuhlow, 1953;

Halvorsen, 1966, Halvorsen, 1967;

Sharp et al., 1990;

Norway

Eudiaptomus gracilisb

Cyclops abyssoruma

Cyclops lacustrisa

Mesocyclops leuckartia

Halvorsen, 1966

Norway

USA

Cyclops scutifera

Halvorsen, 1966;

Johnson, 1975; Meyer and Vik, 1963

USA

Diacyclops bicuspidatus (= Cyclops bicuspidatus)a

 

Meyer and Vik, 1963

USA Acanthocyclops vernalis (= Cyclops vernalis)a Meyer and Vik, 1963

aOrder Cyclopoida; bOrder Calanoida

Fish-eating birds of 9 families (Accipitridae, Alcidae, Vorvidae, Gaviidae, Pandionidae, Pelecanidae, Podicipedidae, Sternidae, and Laridae) distributed in about 50 species represent the final host of D. dendriticus (Kuchta et al., 2013). In South America, Larus dominicanus (Chile and Argentina) and Chroicocephalus maculipennis (Chile) have been recorded with natural and experimental infections (Figueroa et al., 1979, 1980; Torres et al., 1981a, 1981b; Semenas, 2014). Also, infection is present in humans, dogs, cats, red foxes, bears and other mammals (Rausch and Hilliard, 1970; Andersen et al., 1987; Torres et al., 2004a, Wicht et al., 2008a; Kuchta et al., 2013; Catalano et al., 2015). Parasite eggs in the feces of their host spread in freshwater ecosystems developing coracidia in 5 to 6 days at 25°C. Coracidia require light for hatching (Hilliard, 1960; Grabiec et al., 1963). Coracidia infect copepods (Table 5) then develop into procercoid in 14 to 20 days measuring 288 to 576 µm long in C. strenuus (Halvorsen, 1966).

Copepods transmit the infection to planktivorous fish in which the procercoids migrate to the gastrointestinal wall and then the mesenteries, viscera and muscles where they develop into the plerocercoid stage. Table 4 lists 22 species of fish host for D. dendriticus distributed in 6 orders. Thirteen of these species are salmoniforms. The location of D. dendriticus in the muscles is less frequent than in D. latus (Dick and Poole, 1985; Torres et al., 1998, 2004a, 2012; Rozas et al., 2012). The plerocercoids of Dibothriocephalus spp., like other helminths induce granulomatous inflammatory response that encapsulates the parasite in order to isolate and eventually destroy it (Feist and Longshaw, 2008), a situation described by different authors (Gonzalez et al., 1978; Sharp et al., 1989, 1992; Torres et al., 1991, 2002, 2012; Revenga et al., 1995). However this reaction can vary drastically between hosts. Bylund (1972) observed varying degrees of response to D. dendriticus in 3 host species: 1) in Coregonus lavaretus the plerocercoids encyst in the gastrointestinal wall due to a more efficient response that mitigates their migration to other viscera, 2) the response in Salmo trutta is slower allowing penetration of the larva to the gastrointestinal wall and then to different organs where they are encapsulated, and 3) in Coregonus albula they are not encapsulated, remaining free in the abdominal cavity and causing significant damage with their migration. The lack of encapsulation of Dibothriocephalus spp. resulting in greater necrosis, destruction of blood vessels, and hemorrhaging, all of which were observed in some native fish from Lake Panguipulli in Chile (Torres et al., 2012). Experimentally, D. dendriticus plerocercoids in fish held at 14 to 15°C exhibit greater activity and motility from the body cavity to the heart, pericardium and muscles that are found in fish held at 7.5-11°C (Rahkonen and Valtonen, 1998).

The adult stage of D. nihonkaiensis lives in the intestine of humans and the brown bear (Ursus arctos middendorffi) (Yamane et al., 1986; Kamo, 1999; Yera et al., 2006; Arizono et al., 2009a; Yamasaki et al., 2012; Catalano et al., 2015; Choi et al., 2015). Also, Scholz and Kuchta (2016) mentioned as host to Ursus arctos piscator, U. americanus, U. thibetanus, Canis lupus, C. familiaris, Vulpes vulpes, Neovison vison and Sus scrofa. Cyclops strenuus, a freshwater copepod, is an experimental first intermediate host (Eguchi, 1926), but natural copepod hosts are considered to be not known marine species. There are 3 Salmoniformes anadromous species listed as hosts of plerocercoids (Table 4), and which are unlike D. latus that develop in freshwater fish. Dibothriocephalus nihonkaiensis plerocercoids only have been detected in the musculature (Yamane et al., 1986; Suzuki et al., 2010).

Adult stage of A. pacificus develops in the small intestine of marine mammals: Otaria byronia (syn. Otaria flavescens), Arctophoca (syn. Arctocephalus) philippii, A. australis forsteri, A. pusillus pusillus, A. pusillus doriferus, A. gazella, and A. tropicalis in the southern hemisphere, and Callorhinus ursinus, Neophoca cinerea, and Eumetopias jubatus in the northern hemisphere; also develop in humans, Canis familiaris and Canis mesomelas (Kuchta et al., 2015b; Hernández-Ortz et al., 2015; Jacobus et al., 2016). Parasite eggs are eliminated with the feces of the final host completing their development in seawater over 3 days at 22°C and hatching on the fourth day (Escalante et al., 1987). Their first intermediate hosts are unknown but probably correspond to marine copepods. Twenty two species of marine fish included in 7 orders act as intermediate or paratenic hosts, identified in the coast of Peru (Table 4); in Chile the specific identification of plerocercoids of diphyllobothrideans registered in marine fish, included in similar orders, genus or species of the fish host of A. pacificus, have not been verified (Table 4). The lack of records of A. pacificus plerocercoids in the muscles of fish is particularly striking and could be associated with a low level of infection or lack of application of more sensitive detection techniques in this location (Torres and Puga, 2011).

Diphyllobothrium stemmacephalum develops as adult in dolphins and porpoises (Yamasaki et al., 2016). The principal sources of human infection are still unknown (Scholz and Kuchta, 2016).

Diphyllobothrium balaenopterae develops its adult stage in humans and dogs (Rausch, 1964; Yamasaki et al., 2012), but marine mammals are the principal hosts as Mysticeti Balaenoptera acutorostrata, Balaenoptera borealis, Megaptera novaeangliae, Balaenoptera musculus, and Balaenoptera physalus (Kamo, 1999). Kamo et al. (1973) infected a marine planktonic copepod (Oithona nana) an important potential prey for fish larvae, distributed in tropical and subtropical waters of the Atlantic Ocean, in the Mediterranean Sea and the Pacific and Indian Oceans (Cepeda et al., 2012). Some suspected marine fish intermediate hosts are reported, as Engraulis japonica (Clupeiformes / Engraulidae), Sardinops melanostictus (Clupeiformes, Clupeidae), Katsuwonus pelamis (Perciformes, Scombridae) (Yamasaki et al., 2012), as well as Thynnus spp. (Perciformes, Scombridae) and Trachurus japonicus (Perciformes, Carangidae) (Kino et al., 2002). Nevertheless, today the true source of human infection is unknown.

Domestic and wild canids and felids are the final hosts of Spirometra spp.. Adult parasites living in the intestine of Canis lupus familiaris, Cerdocyon thous, Lycalopex fulvipes, L. griseus, L. gymnocercus, L. vetulus, Felis silvestris catus, Leopardus geoffroyi, L. guigna guigna, L. pardalis, L. tigrinus, L. wieddi, Panthera onca, Puma concolor, and P. yagouaroundi in South America (Oda et al., 2016).

Two cases of infection by the adult stage of S. erinacei have been reported in human in Korea (Lee et al., 1984). Also, these authors cited 4 additional cases reported in Japan and 2 cases from China.

The eggs of Spirometra spp. are eliminated with the host feces and then develop coracidia in fresh water. Cyclopoid copepods act as the first intermediate hosts, developing procercoids; and frogs, water snakes, birds, and mammals act as the second intermediate/paratenic hosts being located in different tissues as plerocercoid.

Spirometra erinaceieuropaei adult develops principally in dogs, but is also identified in cats (Mueller, 1974). Copepod species of Mesocyclops, Eucyclops, and Paracyclops act as the first intermediate hosts (Venturini, 1989; Denegri and Reisin, 1993; Bowman et al., 2002). Lee et al. (1990) infected tadpoles of Pelophylax nigromaculatus (syn. Rana nigromaculata) with Mesocyclops leuckarti and Eucyclops serrulatus infected with procercoids. Also, S. mansonoides adult develops in the small intestine of domestic and wild felids (Bowman et al., 2002). Coracidia develop in fresh water and infect copepods of the genus Cyclops, such as C. vernalis and C. viridis (Corkum, 1966; Mueller, 1966). The second intermediate host (amphibians, rodents or water snakes) becomes infected through the ingestion of infected copepods; plerocercoids can also develop in the domestic cat (Bowman et al., 2002).

A revision of Spirometra in China reported 7 species of Rana, and 1 species of the genus Microhyla, Hyla, and Paa, respectively. Also, the authors indicate some reptilian hosts: 6 species of Elaphe, 1 species of Natrix, Naja, Zaocys, Dinodon, Deinagkistrodon, Xenochrophis, and 2 species of Ptyas and Bungrarus, respectively (Liu et al., 2015)

In Malaysia, Rana limnocharis and R. cancrivora, and in Australia Litoria caerulea have been reported as hosts of Spirometra (Liu et al., 2015).

Plerocercoids of S. erinaceieuropaei have been registered in the reptilians Natrix natrix and N. tessellate from Turkey (Yildirimhan et al., 2007).

In USA have been reported infection by plerocercoids in amphibians as Lithobates catesbeianus (syn. Rana catesbeiana) and Lithobates clamitans (syn. Rana clamitans) and in 7 species of reptilians, including 4 species of Nerodia (syn. Natrix), Thamnophis proximus proximus (syn. Thamnophis sauritus proximus), Lampropeltis getula holbrooki and Coluber constrictor flaviventris; in mammals infection was detected in Didelphis virginiana, Procyon lotor and in Urocyon cinereoargenteus (Corkum, 1966).

Oda et al. (2016) review and include new records of Amphibia and Reptilia hosts of Spirometra from South America, including between Amphibia, 3 species of Rhinella, 2 species of Pristimantis, 2 species of Hypsiboas,1 species of Leptodactylus, Dermatonotus, Aplastodiscus, and Scinax, respectively (Oda et al., 2016). Authors include between reptilians, 3 species of Erythrolamprus, Bothrops, Chironius, 2 species of Xenodon and Philodryas, and 1 species of Ameiba, Amphisbaena, Aplastodicus, Drymarchon, Clelia, Leptophis, Lygophis, Mastigodryas, Salvator, Scinax, Simophis, Philodryas, Micrurus, and Crotalus.

In Australia, the existence of two genotypes for S. erinaceieuropaei have been reported, with a genotype in the Litoria caerulea frog different to that found in the dog, fox, cat, tiger snake and python (Zhu et al., 2002).

Humans acquire the sparganosis by ingestion from the first intermediate host in non-drinking water or swimming in unsafe water (Zhou et al., 2005; Eberhard et al., 2015) or, more likely with raw or undercooked second intermediate or paratenic hosts, as frogs or snakes, and crocodiles (Magnino et al., 2009; Liu et al., 2015). The plerocercoid larvae leave their intermediate or paratenic hosts to enter the human body, which is warmer. Infective larvae are released into the digestive tract and penetrate the intestinal wall to develop procercoid to plerocercoid or become established as the latter stage migrating through the intestinal wall, then into the muscle, the viscera or subcutaneous tissue. Also, in East and Southeast Asia, the zoonosis is frequent due to the use of freshwater frogs and snakes-based plasters on eyes or opened wounds in traditional medicine practices, as example, in China, Thailand, and Vietnam (Acha and Szifres, 2003; Wiwanitkit, 2005; Anantaphruti et al., 2011; Liu et al., 2015). Drinking untreated contaminated water is a major infection route in Thailand and in Korea, especially in women and children in the last case (Wiwanitkit, 2005; Anantaphruti et al., 2011; Liu, 2015)

1.3.2 Incubation and pre-patent periods

The incubation period in Dibothriocephalus spp., A. pacificus and Di. balaenopterae infections is variable and depends on the size of the worms, intensity of infection, nutritional status of the host and degree of sensitivity to antigen parasites. Clinical signs usually are not pathognomonic, however, a frequent clinical sign is the periodic elimination of strobila pieces up to 50 cm long with human stools (Kamo et al., 1986; Torres, 2013) and up to 1 m every 1 or 2 months in dogs (Magath, 1929), a situation which occurs after the pre-patent period and when the parasite is mature. The pre-patent period of Dibothriocephalus, Adenocephalus and Spirometra species is variable in different hosts (Table 6). In the majority of species it fluctuates between 6 and 30 days, except in A. pacificus, whose period varies between 20 and 55 days.

Table 6. Pre-patent periods of some Diphyllobothriidae species

Species

Host

Pre-patent Period (Days)

Reference

Adenocephalus pacificus

Dogs

20 to 55

Escalante and Miranda, 1986; Escalante and Chico-Ruiz, 1988

Dibothriocephalus latus

Hamsters

16 to 22

Andersen, 1971

Dibothriocephalus latus

Humans

25 to 30

von Bonsdorff, 1977

Dibothriocephalus latus

Dogs

18 to 27

Torres et al., 1989a

Dibothriocephalus latus

Cats

27 to 29

Torres et al., 1989a

Dibothriocephalus dendriticus

Humans

15

Freeman and Jamieson, 1972; Curtis and Bylund, 1991

Dibothriocephalus dendriticus

Hamsters

7 to 13

Andersen, 1971

Dibothriocephalus dendriticus

Dogs

12 to 18

Torres et al., 1981b, 1989a

Dibothriocephalus dendriticus

Gulls, Larus dominicanus

12

Torres et al., 1981b, 1989a

Dibothriocephalus dendriticus

Gulls, Chroicocephalus maculipennis

13

Torres et al., 1981a

Dibothriocephalus dendriticus

Gulls, L. argentatus

6

Sharp et al., 1990

Dibothriocephalus nihonkaiensis

Hamsters

20

Yamane et al., 1986

Spirometra erinaceieuropaei

Dogs

15 to 30

Venturini, 1989; Denegri and Reisin, 1993

Spirometra erinaceieuropaei

Dogs

16 to 18

Lee et al., 1990

Spirometra erinaceieuropaei

Cats

15

Lee et al., 1990

Spirometra erinaceieuropaei

Cats

8 to 10

Ooi et al., 2000

Spirometra erinaceieuropaei

Cats

12

Lin et al., 2010

Spirometra mansonoides

Cats

12 to 13

Mueller, 1959

1.3.3 Period of communicability

The adult of D. latus have a great capacity for egg production. A patient can eliminate about 122,000 eggs/g feces with a total of 20 to 40 million eggs daily (von Bonsdorff and Bylund, 1982). Between 50,000 and 400,000 eggs/g of feces were eliminated daily over a 4 month period in an experimental human infection with 5 worms (Kamo et al., 1986). Adults of D. latus can live 10 to 15 years in the human intestine, although there have been cases of infections lasting up to 30 years (Bonsdorff, 1977). In Lake Panguipulli in Chile, prevalence in humans (2.8%) and dogs (4.5%) in the Choshuenco locality and 1.8% in the dogs at the Panguipulli locality in part favoring a high prevalence of infection (83% to 93%) in O. mykiss, living in that lake (Torres et al., 2004a, 2012; Torres and Puga, 2011). Control of infection in humans and domestic animals can consequently reduce infection of fish which are food source for the local population and constitute a tourist attraction for sport fishing.

Dibothriocephalus dendriticus lives around 6 months in the gull, Larus argentatus; a humoral response has been demonstrated during infection (Sharp et al., 1990), which would be effective in the removal of the parasite but would not cause a protective immunity against reinfection. However, infection in a human case could remain for 2 years (Wicht et al., 2008a).

Spirometra erinaceieuropaei eliminate about 70,000 eggs/g feces after 15 days of infection (Lee et al., 1990). Ooi et al. (2000) estimated in 428,000 eggs/g of feces per day and a total of 14,416,000 eggs per day in a single worm infection in a cat. Spirometra mansonoides in cats can live for about 3.5 years (Mueller, 1974).

1.4 Population and Individual Control Measures

1.4.1 Treatment options

Dibothriocephalus species are sensitive to niclosamide (2 g in adult or 50 mg/Kg in children older than 2 years-old, on an empty stomach in two doses an hour apart) (Bylund et al., 1977). The tablets are taken with very few water during a hydric diet (Bourée et al., 2012). The drug inhibits the oxidative phosphorylation of the parasite mitochondria leading to the diminution of energetic capacities, an acid lactic accumulation and the death of the worm (Weinbach and Garbus, 1969). The drug inhibits also the defensive mechanisms of the parasite against the host intestinal proteases. Niclosamide is not available in many countries but is the preferred drug because it is not absorbed in the intestine (Flisser and Willingham, 2013). In general, the drug is well tolerated and no toxic effects have been reported in therapeutic doses besides it has no contraindications and can be used during gestation (Flisser and Willingham, 2013). Niclosamide is also efficient on other cestodes as Taenia solium, Taenia saginata and Hymenolepis nana. Niclosamide is effective, but its administration is sometimes difficult to respect; which may be responsible for therapeutic failure (Bourée et al., 2012).

Dibothriocephalus species are sensitive to praziquantel (25 mg/Kg/day in one dose) (Bylund et al., 1977). This dose is usually used to cure D. latus and D. nihonkaiensis infections, lower dose 10 mg/Kg/day are effective against A. pacificus infection (Scholz et al., 2009). The tablets are taken at the end of a meal. The drug is active on the parasite calcium channels, leading to an increase of the cellular membrane permeability and the muscular paralysis of the worm. It could inhibit the absorption of the adenosine by the parasite, which could not synthetize purine de novo. Also, praziquantel is efficient on adult (Taenia spp.) and larva (cysticercus) of cestodes, as trematodes (Schistosoma spp.) and flukes (except, Fasciola hepatica). Praziquantel is well tolerated but should be avoided during pregnancy, breastfeeding and in the first trimester of pregnancy and in children younger than 2 years-old as well as in patients with cirrhosis and hepatic problems (Flisser and Willingham, 2013). Side effects are frequent but transitory and consist in nausea, headaches, dizziness, drowsiness and itch (Chai, 2013). The administration of each drug should be accompanied posteriorly (1-2 h) by a saline purge (Torres, 2013). This prevents the stay of the worm in the intestine avoiding a prolonged action of the enzymes of the host, which contributes to damage and alter it, which can make difficult the search of the scolex in the feces after the treatment, aspect necessary to verify the cure parasitological (Torres, 2013). Moreover, a vitamin B12 supplementation can be initiated in case of depletion. Whatever the treatment, its efficiency has to be controlled one (D. latus) or two months (A. pacificus) after by stool examination, according pre-patent period of parasites (Table 6).

Treatment of sparganosis is surgical. All the nodular masses should be removed to avoid recurrence if the scolex of larvae is let in tissues. If surgery is not possible, praziquantel (120 mg/Kg/day in three doses in adult and 150 mg/Kg/day in three doses in children) is given for 2 days (Liu et al., 2015). In cerebral sparganosis, chemotherapy has been associated to surgery. Moreover, combined dexamethasone and praziquantel allows a better outcome in patients (Hong et al., 2013). Mebendazole has been given after surgical treatment of subcutaneous sparganosis (Tung et al., 2005). In a case of pulmonary sparganosis, two cure of praziquantel (600 mg/day for 5 days) were efficient (Cheng et al., 2014). In proliferative sparganosis, mebendazole and praziquantel (40 mg/Kg/day three times for 2 weeks) did not show efficiency (Moulinier, 1982).The efficiency of the treatment and the absence of recurrence has to be controlled by serology. Antigen or antibody titers should decrease then disappear in case of complete recovery.

Most of the cases of human infection by Sp. proliferum are lethal due to the widespread dissemination and their continuous branching of asexually multiplying plerocercoids. Attempts of surgical removal are usually unsuccessful (Kuchta et al., 2015a).

1.4.2 Hygiene measures

Hygiene measures aim to avoid human contamination. It will be difficult to stop the life cycle of the parasites as species of Diphyllobothriidae are widely present in wildlife.

Treatment of people and domestic animals, principally dogs, known to be infected is a first step in prevention as it would decrease contamination of water.

As people could be asymptomatic wearers, the use of sanitary facilities and sewage treatment plants would reduce water contamination too. The impact of water treatment is limited because excreta from wild and domestic mammals (infected with Dibothriocephalus, Diphyllobothrium, Adenocephalus, or Spirometra spp.) and birds (infected with D. dendriticus) are important source of aquatic environment contamination and are difficult to control.

Eating well-cooked fish prevent infection. Some authors recommend cooking fish at least for ≥ 10 min at 55°C kills the plerocercoid larvae (Dick et al., 2001). European Food safety Authority (EFSA, 2011) for parasites other than trematodes specifies a heat treatment with a core temperature of 60°C or more for 1 min. Reaching such a core temperature depends on the thickness and composition of the product, in fillets that a 3 cm thick should be heated for 10 min (EFSA, 2011). The U.S. Food and Drug Administration (FDA, 2011) guidelines for cooking fish should suffice to inactivate plerocercoids. The guidelines for fish are as follows: cook the fish to an internal temperature of 145ºF (62.7°C) for 15 s; to 155ºF (68.3°C) for comminuted fish, such as fish cakes, and 165ºF (73.8°C) for stuffed fish.

Freezing fish before consumption prevent the infection, too. Some authors considered freezing at least at -10°C during 24 h kills the plerocercoid larvae (Raether and Hanel, 2003). EFSA (2011) recommends specifically for Dibothriocephalus plerocercoids the application of -18°C in all parts of the product for at least 24 h. Also, EFSA considered for different zoonotic parasites other than trematodes, the freezing treatment must lower the temperature in all part of the product to either -20°C or lower for not less than 24 h, or to -35°C or lower for not less than 15 h. FDA (2011) recommend the freezing and storing at an ambient temperature of -20°C or below for 7 days (total time); or freezing at an ambient temperature of -35°C or below until solid and storing at an ambient temperature of -35°C or below for 15 h; or freezing at an ambient temperature of -35°C or below until solid and storing at an ambient temperature of -20°C or below for 24 h. In some countries (e.g. France and Switzerland), the legislation requires that any fish intended to be consumed raw must be frozen at a temperature lower than -20°C, during at least 24 h. These standards have the advantage of also protecting from others zoonosis transmitted by fish, such as anisakidosis. In general, these measures are not done by individual consumers: In practice, fish have been conserved in the usual freezers during one week.

Salting fish reduce plerocercoid larvae infectivity. Fish should be in 12% NaCl (Raether and Hanel, 2003). The time requires killing the larvae, from several days to weeks, depend on the thickness of the fish and the volume of salt. Smoking and pickling are not effective against the plerocercoïds larvae. Information campaigns for the consumers could be useful as they must make them aware with the dangers of raw fish consumption.

The plerocercoids of D. latus survive in the tissues of dead fish for up to 10 days in river water (Feachem et al., 1983). The D. latus plerocercoids survive 1 to 7 days, 72 h, and 10 min at -10°C, 4°C and 50°C, respectively (Peduzzi and Boucher-Rodoni, 2001). Plerocercoids in fish of 9 Kg can be effectively killed at -6°C for 7 days, in fish of 2 Kg in 6 days at -6°C, and in those of 0.7 kg at -6°C in 3 days. At -18°C the larvae die in 4 or 2 days in fish of 2 Kg or 0.5 Kg, respectively (Feachem et al., 1983).

Only in 8 cases the adult stage of Spirometra has been reported in humans (Lee et al. 1984) in contrast to the 1.685 cases of sparganosis (Liu et al., 2015). By this, treatment of infected people will have no impact in preventing transmission as human is an impasse. In contrary, treatment of the farm definitive mammal hosts (cats and dogs) would decrease water contamination (Liu et al. 2015). In the cities of Yunfu and Zhaoqing the prevalence of infection by S. erinaceieuropaei was of 47.2% and 12.6%, respectively in dogs. Infection in the cats was of 64.4% and 35.9% in Yunfu and Zhaoqing, respectively (Hong et al., 2016). Treatment of wild definitive mammal hosts (others canids and felids) is difficult.

The use of drinking water or the filtration of non-drinking water prevents infection; most sparganosis cases reported from Thailand it is by the consumption of drinking contaminated water with infected copepods (Wiwanitkit, 2005; Anantaphruti et al., 2011). Installation of water fountains or areas with open water for public use should be encouraged in regions of endemicity (Nithiuthai et al., 2004).

Eating well-cooked frogs or aquatic snakes prevent the infection (Anantaphruti et al. 2011; Liu et al. 2015). Hong et al (2016) determined through a survey that the consumption of cooking meat with frying and/or porridge and/or soup would be the most probable means of transmission of sparganosis by frogs in Guangzhou, China. These authors found prevalences of 51.2% and 35.1% in wild R. tigrina rugulosa and R. limnocharis, however, sparganosis was not found in farmed frogs. Cooking with 55°C during at least 5 min kills the plerocercoid larvae (Dick, 2008). There is no appropriate data on the effect of freezing on amphibians or reptiles meat except an Australian regulation for the export of fish or their products indicating for crocodiles exposed or suspected of having sparganosis to place, after processing, at -12°C or cooler at the thermal center for a minimum of 5 days (Magnino et al 2009). Public regulation of hunting or sale of wild animals, particularly frogs and snakes, could prevent human infection (Li et al., 2011; Liu et al., 2015).

Avoiding traditional medical practices, using poultices made of frogs or snakes on skin or eye, prevent infection (Anantaphruti et al., 2011; Li et al., 2011; Liu et al., 2015). Information campaigns for people in endemic countries on the dangers of raw frogs or snakes consumption or use as poultices and of non-drinking water would help to the prevention and health care of sparganosis.

2.0 Environmental Occurrence and Persistence

2.1 Detection Methods

Detecting copepods infection by microscopic examination may be performed by directly observing the procercoids in hemocele (Fig. 11), by extraction by mild compression between slide and cover slips (Guttowa, 1963; Torres et al., 2004b, 2007) or dissection, when the wall thickness makes it difficult to observe the larvae (Guttowa, 1963; Sharp et al., 1990).

The infection in copepods has been determined in their aquatic environments (Guttowa, 1963; Johnson, 1975) (Table 7) and its detection depends in part, on the sampling system included seasonal period, collection sites and the development status and sex of the host (Guttowa, 1965; Chubb, 1980). Additionally, morphological descriptions of procercoids have not been evaluated criteria for specific differentiation. The procedure most often used to determine the susceptibility of copepods has been through its experimental infection with coracidia obtained by in vitro incubation of eggs of D. latus, D. dendriticus, D. nihonkaiensis, Di. balaenopterae, and Spirometra spp. (Janicki and Rosen, 1917; Eguchi, 1926; Essex and Magath, 1931; Bearup, 1957; Guttowa, 1961; Halvorsen, 1966; Kamo et al., 1973; Venturini, 1989; Lee et al., 1990; Denegri and Reisin, 1993; Wright and Curtis, 2000; Torres et al., 2007; Lin et al., 2010).

Table 7. Natural infection in copepods by Dibothriocephalus spp.

Country

Parasite

Host

Period

Sector A

Prevalence %

(# of Samples)

Sector B

Prevalence %

(# of Samples)

Reference

Finland,

Lake Karpero

D. latus

Thermocyclops oithonoides

June

23.3%

7/30

20.6%

6/29

Guttowa, 1963

Finland,

Lake Karpero

D. latus

Thermocyclops oithonoides

June

18.5%

5/27

17.6%

3/17

Guttowa, 1963

Finland,

Lake Karpero

D. latus

Thermocyclops oithonoides

July

12.5%

1/8

22.2%

2/9

Guttowa, 1963

Finland,

Lake Karpero

D. latus

Thermocyclops oithonoides

July

12.5%

2/17

14.2%

2/14

Guttowa, 1963

Finland,

Lake Karpero

D. latus

Cyclops strenuus

June

51.7%

43/83

52.3%

11/21

Guttowa, 1963

Finland,

Lake Karpero

D. latus

Cyclops strenuus

June

51.5%

16/31

37.5%

12/32

Guttowa, 1963

Finland,

Lake Karpero

D. latus

Cyclops strenuus

July

33.3%

3/9

27.2%

3/11

Guttowa, 1963

Finland,

Lake Karpero

D. latus

Cyclops strenuus

July

18.0%

2/11

0

0/2

Guttowa, 1963

USA, in waters near Cobb Fish Hatchery, Enfield, Maine

D. dendriticus

Cyclops scutifer

Mid-July

1.7%

1/200

NRa

Johnson, 1975

aNR: Not Reported

The plerocercoids measure 1 mm to several cm long and their detection on the surface of muscles and in the viscera is feasible by naked eye observation (Figs. 17, 19). However those of small size or located in the thickness of muscles or organs must be detected using a stereomicroscope. The presence of plerocercoids in muscles is of primary importance in human transmission of infections. Its detection is performed by visual inspection, slicing, candling, or pressing (EFSA, 2011; FDA, 2011). The pressing technique in plaques has proved more efficiently than candling in the detection of D. latus and D. dendriticus (Torres and Puga, 2011). Digestion is more expensive in terms of reagents than candling and glass compression techniques. Plerocercoids of D. latus and D. dendriticus in O. mykiss are more abundant in the ventral than dorsal musculature (Torres and Puga, 2011).

2.2 Data on Occurrence

2.2.1 Occurrence in the environment

Diphyllobothriideans eggs are sensitive to environmental conditions and to develop must achieve adequate water courses such as rivers or lakes where meet the biotic and abiotic characteristics required for their life cycle. As the temperature rises from 8 to 30°C, the development of D. latus coracidia increases in fresh water; at 8-10°C in 39-42 days, at 15°C in 18-20 days, at 18-20°C in 8-9 days, at 25°C in 7 days, and at 30°C in 6 days. However, hatching rate decreases (Guttowa, 1961) .The eggs remain viable for years at temperatures near 0°C; but they can do not survive at 35°C (Guttowa, 1961). Eggs in water depths greater than 20 m do not hatch and those that found in feces contaminating soil or ice die by desiccation after 3 days in winter (Feachem et al., 1983). Dibothriocephalus latus eggs remain viable in soil for 2 months and in water for 5 months (Khromenkova et al., 1995). The oxygen concentration can influence their development through their aerobic metabolism. The best biotopes are shallow littorals with water temperatures of 15-20°C, with vegetation favoring the development of copepods and fish (von Bonsdorff and Bylund, 1982). The minimum concentration of oxygen in the water for the hatching of coracidium of D. latus is 1.4 mg/L at 24°C. The eggs can survive at low concentrations of oxygen for months and develop coracidia when the optimum concentration is reestablished (von Bonsdorff, 1977).

Coracidia (Fig. 10) can survive for 72 h at 8-10°C, 48-72 h at 15°C, 48 h at 18-20°C, 24-36 h at 25°C, and 12 to 24 h at 30°C, respectively (Guttowa, 1961). The larvae survive 4 days in 0.4 0/00 saline (von Bonsdorff and Bylund, 1982) and could tolerate the brackish water of the coastal areas, estuaries and innermost bays where the salinity is 0.2-0.4 0/00 (Bonsdorff and Bylund, 1982). The capacity to develop coracidia from D. latus eggs from human infection reaches between 74.3% and 95% (Essex and Magath, 1931; Guttowa, 1959). In dog infections that capacity reaches 45% to 52% (Tarassow, 1934; Guttowa, 1959).

Hatching coracidia of D. dendriticus live 12-72 h at room temperature in fresh water (Sharp et al., 1990). At 10°C the eggs develop coracidia in 25 days reaching their peak at 41 days, with full development of the population of eggs at about 2.5 months; at 29 days the eggs start to hatch with a peak at 65 days (Wright and Curtis, 2000). These same authors obtained larvae at 5 days at 20°C with a peak at 9 days with the full development of the population of eggs taking 25 days. Coracidia may develop slowly at 4°C, hatching between 11 and 30 months, favoring the life cycle in those lakes that range from 8 to 14°C during the ice free period in northern North America and Northern Europe as well as in Arctic Lakes with maximum temperatures not exceeding 4°C (Wright and Curtis, 2000). Dibothriocephalus dendriticus eggs die within 6-8 days when kept at 37°C (Kuhlow, 1953). Their eggs are developed at 7°C in 48 days, at 20 °C in 10 to 11 days, and at 23°C in 8 days, respectively (Chubb, 1980).

Dibothriocephalus nihonkaiensis develops experimentally in a freshwater copepod (Eguchi, 1926). Adenocephalus pacificus coracidia hatch in sea water after incubation of the eggs at 22°C for 3 days (Escalante et al., 1987).

Spirometra mansonoides eggs die when exposed to 37°C for 6 to 8 days, but at 4°C they can survive for 5 years, although their viability gradually decreases (Mueller, 1961, 1974). Eggs develops experimentally coracidia that hatch after 7 days at 25°C and infecting procercoids develop between 11 and 12 days at 20-25°C in Paracyclops fimbriatus, reaching a mean of 210 mm long by 120 µm wide (Venturini, 1989; Denegri and Reisin, 1993). The coracidia develop in 10 days at 25 to 27°C (Mueller, 1959) or in 10 to 14 days at 23°C (Mueller, 1974). In Cyclops vernalis, infective procercoids develop between 9 and 14 days at 24 to 27°C (Mueller, 1966).

In areas of the Kama reservoir in Russia where raw sewage is eliminated untreated the infection levels of D. latus in fish are fairly high, especially in areas where chemical residues are also eliminated in water (Artamoshin and Khodakova, 1976). The wastewater effluent from the town of Abakan in Russia contained 0.6-4.1 D. latus eggs/L and effluent from a septic tank of a hospital contained 52.1 eggs /L. The ground around the septic tank contained 1,830 eggs /Kg and in the areas surrounding the fishing sites, 95% of E. lucius was infected (Gerasimov, 1987). Eggs of D. latus were found in 5% of fresh vegetables in the markets of Nasiriyah and Suq-AL-Shuyukh, Iraq (Al-Mozan et al., 2015). Additionally, they were reported as an indicator on the use of untreated sewage in the watering and fertilizing of farm land (Al-Mozan et al., 2015). Also 3.2% of lettuces acquired in supermarkets in the city of Dourados-MS, Brazil were contaminated with eggs of Dibothriocephalus sp. (Correa et al., 2012). Mixed human and farm effluent from three state farms contained 300-1,600 helminth eggs per m3, including Dibothriocephalus, Ascaris, Enterobius, Trichuris, and Taenia (Arkhipova, 1977). The passage of effluent through a treatment plant only halved the egg count (Arkhipova, 1977). Eggs of diphyllobothriidean were found in most of the samples of sludge collected from night-soil disposal plants in Kochi Prefecture, Japan (Takenouchi et al., 1980). In recreational areas in the lower Don region of Russia, most of the sites had either no sanitation or inadequate facilities resulting in heavy contamination of the soil with ova identified as Dibothriocephalus and other helminths where about 43.1% of soil samples contained 17 to 206 ova /Kg (Khromenkova, 1993).

Data on occurrence of eggs of Diphyllobothriidae in the environment do not really reflect the epidemiological situation of the infection. On the opposite, the prevalence of plerocercoids in receptive fish is an indication of the presence of the infection in a given region.

2.2.2 Occurrence in copepods, fish, and others intermediate and paratenic hosts

Most copepod host records correspond to experimental studies in D. latus (Table 3) and D. dendriticus (Table 5). Studies on occurrence of infection by procercoids in copepods under natural conditions are scarce (Table 7). It has been estimated that for the development of D. latus a minimum density of copepods (1,000/m3) is necessary (von Bonsdorff, 1977). In the Don River, Russia, where 8 to 13 human cases of D. latus were recorded annually and 0.3% of predatory fish were infected; the densities of 7 copepod host species ranged from 200 to 2,000/m3 in an area where ​​parasite eggs were released into the river with effluent from households, boats and ships (Khromenkova et al., 1995).

In M. leuckarti in the Luohe City in China a prevalence of 3.5% by S. erinaceieuropaei with an intensity of 3 to 5 procercoids was found (Lin et al., 2010). Nine Mesocyclops species have been recorded as hosts of Spirometra procercoids in China (Liu et al., 2015). In Korea, M. leuckarti and E. serrulatus as hosts of S. erinaceieuropaei have been identified (Lee et al., 1990).

Data on the occurrence, including prevalence, mean intensity and/or mean abundance of infection in fish, for D. latus and D. dendriticus in lakes and rivers of different countries are given for distinct hosts in Table 8. In South America, the highest prevalence and intensity or abundance of infection by D. latus are recorded principally in introduced salmonids such as O. mykiss and S. trutta in Chile and O. mykiss and Salvelinus fontinalis in Argentina. Dibothriocephalus latus and D. dendriticus infection has been recorded in native fish from 1989 in 2 of 21 lakes investigated in southern Chile (Torres et al., 1989b, 1990, 1991, 1992, 1998, 2012). Similar observations have been made on Lake Moreno in Argentina (Table 8). The frequency of finding plerocercoids of D. latus and D. dendriticus ranges from 55.8% to 73.3% and 9.1% to 43% respectively, in the muscles of O. mykiss in Lake Panguipulli, Chile (Torres et al., 2004a, 2012; Torres and Puga, 2011). Examination of 200 (Nicolaud et al., 2005) and 960 (Dupouy-Camet et al., 2015) fillets of P. fluviatilis from Lake Léman showed an overall prevalence of 7% (4-10%) and 0.9% (0.3-1.5%), respectively.

Table 8. Prevalence, mean intensity and/or mean abundance of infection by Dibothriocephalus latus and D. dendriticus in different fish hosts and countries

Area

Parasite

Fish Host

Prevalence Percentage (# of Samples)

Mean

Intensitya

Mean

Abundanceb

Reference

Argentina, Lake Moreno

D. latus

Oncorhynchus mykiss

28.1%

(32/114)

NRc

1.4

Revenga, 1993

Argentina, Lake Moreno

D. latus

Salvelinus fontinalis

9.1%

(1/11)

NR

0.1

Revenga, 1993

Argentina, Lake Moreno

D. latus

Percichthys trucha

18.8%

(6/32)

NR

0.3

Revenga, 1993

Argentina, Lake Moreno

D. dendriticus

O. mykiss

57.9%

(66/114)

NR

7.2

Revenga, 1993

Argentina, Lake Moreno

D. dendriticus

S. fontinalis

27.3%

(3/11)

NR

0.5

Revenga, 1993

Canada, Great Central Lake

D. dendriticus

O. nerka

93.2%

(55/59)

4.0

NR

Ching, 1988

Canada, Lake Igalugaajuruluit

D. dendriticus

S. alpinus

41.8%

(117/280)

73.0

NR

Gallagher and Dick, 2010

Chile, Lake Caburgua

D. latus

O. mykiss

3,7%

(1/27)

3.0

NR

Torres et al., 1991

Chile, Lake Caburgua

D. dendriticus

O. mykiss

44.4%

(12/27)

3.3

NR

Torres et al., 1991

Chile, Lake Calafquén

D. latus

O. mykiss

4,3 %

(1/18)

4.0

NR

Torres et al., 1991

Chile, Lake Calafquén

D. dendriticus

O. mykiss

27.8%

(5/18)

2,2

NR

Torres et al., 1991

Chile, Lake Colico

D. latus

O. mykiss

28.6%

(6/21)

6.8

NR

Torres et al., 1991

Chile, Lake Colico

D. latus

S. trutta

75%

(3/4)

34.0

NR

Torres et al., 1991

Chile, Lake Colico

D. dendriticus

O. mykiss

47.6%

(10/21)

2.6

NR

Torres et al., 1991

Chile, Lake Colico

D. dendriticus

S. trutta

75%

(3/4)

26.7

NR

Torres et al., 1991

Chile, Lake Cucao

D. dendriticus

O. mykiss

5.1%

(4/79)

1.1

NR

Rozas et al., 2012

Chile, Lake Huillinco

D. dendriticus

O. mykiss

3.6%

(5/138)

1.1

NR

Rozas et al., 2012

Chile, Lake Llanquihue

D. dendriticus

O. mykiss

18.8 %

(6/32)

6.2

NR

Torres et al., 1991

Chile, Lake Maihue

D. latus

S. trutta

50.0%

(1/2)

37.0

NR

Torres et al., 1991

Chile, Lake Maihue

D. dendriticus

O. mykiss

3.0%

(1/33)

1.0

NR

Torres et al., 1991

Chile, Lake Maihue

D. dendriticus

S. trutta

50.0%

(1/2)

10.0

NR

Torres et al., 1991

Chile, Lake Natri

D. dendriticus

O. mykiss

4.7%

(5/107)

1.1

NR

Rozas et al., 2012

Chile, Lake Panguipulli

D. latus

O. mykiss

78.0 %

(32/41)

48.4

NR

Torres et al., 1991

Chile, Lake Panguipulli

D. latus

O. mykiss

83.3%

(25/30)

87.1

NR

Torres et al., 2004a

Chile, Lake Panguipulli

D. latus

Basilichthys australis

7.1%

(1/14)

1.0

NR

Torres et al., 2004a

Chile, Lake Panguipulli

D. latus

Odontesthes mauleanum

69.2%

(9/13)

4.6

NR

Torres et al., 2004a

Chile, Lake Panguipulli

D. latus

Percichthys trucha

52.2

(12/23)

3.3

NR

Torres et al., 2004a

Chile, Lake Panguipulli

D. latus

Galaxias maculatus

2.1

(1/47)

1.0

NR

Torres et al., 2004a

Chile, Lake Panguipulli

D. latus

O. mykiss

92%

(71/77)

33.0

30.4

Torres and Puga, 2011

Chile, Lake Panguipulli

D. latus

O. mykiss

93%

(26/28)

35.5

32.9

Torres et al., 2012

Chile, Lake Panguipulli

D. latus

B. australis

10.4%

(5/48)

7.2

0.8

Torres et al., 2012

Chile, Lake Panguipulli

D. latus

O. maulenaum

43.2%

(19/44)

5.8

2.5

Torres et al., 2012

Chile, Lake Panguipulli

D. latus

P. trutta

44.0 %

(11/25)

4.4

1.9

Torres et al., 2012

Chile, Lake Panguipulli

D. dendriticus

O. mykiss

65.9%

(27/41)

7.0

NR

Torres et al., 1991

Chile, Lake Panguipulli

D. dendriticus

O. mykiss

76.7%

(23/30)

5.7

NR

Torres et al., 2004a

Chile, Lake Panguipulli

D. dendriticus

B. australis

14.3

(2/14)

1.0

NR

Torres et al., 2004a

Chile, Lake Panguipulli

D. dendriticus

O. mauleanum

23.1

(3/13)

1.0

NR

Torres et al., 2004a

Chile, Lake Panguipulli

D. dendriticus

O. mykiss

48.1%

(37/77)

5.9

2.8

Torres and Puga, 2011

Chile, Lake Panguipulli

D. dendriticus

O. mykiss

57.1%

(16/28)

5.0

2.9

Torres et al., 2012

Chile, Lake Panguipulli

D. dendriticus

B. australis

4.5 %

(2/44)

NR

0.05

Torres et al., 2012

Chile, Lake Panguipulli

D. dendriticus

O. mauleanum

2.1%

(1/48)

NR

0.04

Torres et al., 2012

Chile, Lake Puyehue

D. latus

O. mykiss

20%

(6/30)

1.2

NR

Torres et al., 1991

Chile, Lake Puyehue

D. dendriticus

O. mykiss

60.0%

(18/30)

3.2

NR

Torres et al., 1991

Chile, Lake Ranco

D. latus

O. mykiss

6.7%

(2/30)

8.0

NR

Torres et al., 1991

Chile, Lake Ranco

D. dendriticus

O. mykiss

16.7%

(5/30)

4.6

NR

Torres et al., 1991

Chile, Lake Riñihue

D. latus

O. mykiss

44.9%

(31/69)

9.2

NR

Torres et al., 1991

Chile, Lake Riñihue

D. latus

O. mykiss

35.7%

(50/140)

5.2

NR

Torres et al., 1998

Chile, Lake Riñihue

D. latus

G. maculatus

0.5%

(1/184)

1.0

NR

Torres et al., 1998

Chile, Lake Riñihue

D. latus

P. trucha

11.1%

(7/63)

1.3

NR

Torres et al., 1998

Chile, Lake Riñihue

D. dendriticus

O. mykiss

53.6%

(37/69)

11.1

NR

Torres et al., 1991

Chile, Lake Riñihue

D. dendriticus

O. mykiss

17.9

(25/140)

2.4

NR

Torres et al., 1998

Chile, Lake Riñihue

D. dendriticus

G. maculatus

0.5%

(1/184)

1.0

NR

Torres et al., 1998

Chile, Lake Riñihue

D. dendriticus

B. australis

0.5%

(1/192)

1.0

NR

Torres et al., 1998

Chile, Lake Riñihue

D. dendriticus

P. trucha

3.2%

(2/63)

2.0

NR

Torres et al., 1998

Chile, Lake San Antonio

D. dendriticus

O. mykiss

66.7%

(4/6)

1.1

NR

Rozas et al., 2012

Chile, Lake Villarrica

D. latus

O. mykiss

6.9%

(2/29)

1.5

NR

Torres et al., 1991

Chile, Lake Villarrica

D. dendriticus

O. mykiss

44.8%

(13/29)

6.1

NR

Torres et al., 1991

Chile, Lake Villarrica

D. dendriticus

S. trutta

33.3%

(1/3)

2.0

NR

Torres et al., 1991

Chile, Lake Rupanco

D. dendriticus

O. mykiss

7.4%

(2/27)

1.5

NR

Torres et al., 1991

Chile, Lake Tarahuín

D. dendriticus

O. mykiss

50.9%

(81/159)

1.1

NR

Rozas et al., 2012

Chile, Lake Todos los Santos

D. dendriticus

O. mykiss

14.3% (2/14)

2.0

NR

Torres et al., 1991

Finland, Lake Inari

D. dendriticus

S. trutta m. lacustris

90%

(109/121)

NR

52.9

Rahkonen and Koski, 1997

Finland, Lake Inari

D. dendriticus

S. trutta m. lacustris

83.0

(73/88)

NR

50.2

Rahkonen and Koski, 1997

France, Lake Léman

D. latus

Perca fluviatilis

29.2%

(7/24)

NR

NR

Dupouy-Camet et al., 2015

France, Lake Léman

D. latus

Esox lucius

100%

(6/6)

NR

NR

Dupouy-Camet et al., 2015

France, Lake Léman

D. latus

Lota lota

28.6%

(2/7)

NR

NR

Dupouy-Camet et al., 2015

France, Lake Léman

D. latus

P. fluviatilis (fillets)

0.9%

(9/960)

NR

NR

Dupouy-Camet et al., 2015

France, Lake Léman

D. latus P. fluviatilis (fillets)

7%

(14/200)

NR

NR

Nicolaud et al., 2005

Germany, Baltic Sea (coast)

D. dendriticus

S. trutta

2.9%

(1/35)

1.0

0.1

Unger and Palm, 2016

Germany, Freshwater streams

D. dendriticus

S. trutta

5.9%

(1/17)

4.0

0.2

Unger and Palm, 2016

Italy, Lake Como

D. latus

P. fluviatilis

25.4%

(108/426)

1.2

0.31

Gustinelli et al., 2016

Italy, Lake Como

D. latus

E. lucius

84.2%

(16/19)

28.3

23.8

Gustinelli et al., 2016

Italy, Lake Como

D. latus

L. lota

3.6%

(2/55)

1.5

0.05

Gustinelli et al., 2016

Italy, Lake Iseo

D. latus

P. fluviatilis

7.6%

(35/428)

1.3

0.10

Gustinelli et al., 2016

Italy, Lake Iseo

D. latus

E. lucius

71.4%

(5/7)

16.4

11.7

Gustinelli et al., 2016

Italy, Lake Iseo

D. latus

L. lota

3.8%

(1/26)

3.0

0.12

Gustinelli et al., 2016

Italy, Lake Lario

D. latus

P. fluviatilis

30.0%

(609/183)

1.3

NR

Wicht et al., 2009

Italy, Lake Maggiore

D. latus

P. fluviatilis

6.6%

(42/635)

1.1

0.07

Gustinelli et al., 2016

Italy, Lake Maggiore

D. latus

E. lucius

100%

(1/1)

1.0

1.0

Gustinelli et al., 2016

Italy, Lake Orta

D. latus

P. fluviatilis

33.3%

(5/15)

NR

NR

Peduzzi and Boucher-Rodoni, 2001

Italy/Switzerland, Lake Maggiore

D. latus

P. fluviatilis

7.8%

(24/309)

NR

NR

Peduzzi and Boucher-Rodoni, 2001

Norway, Lakes Storvatn and Forsanvatn

D. latus

S. trutta

10.2%

(22/214)

2.2

0.2

Kuhn et al., 2016

Norway, Lakes Rekvatn and Fjellfrosvatn

D. latus

S. trutta

45.7%

(52/113)

5.7

2.6

Kuhn et al., 2016

Norway, Lakes Makkvatn, Skilvatn, Sagelvvatn, and Fefjerdevatn

D. latus

S. trutta

45.6%

(122/268)

 

39.4

17.9

Kuhn et al., 2016

Sweden, Lake Bjellojaure

D. dendriticus

Salvelinus alpinus

83.2%

(544/654)

8.8

NR

Henricson, 1977

Switzerland, Lake Bienne

D. latus

P. fluviatilis

3.7%

(3/81)

NR

NR

Golay and Mariaux, 1995

Switzerland, Lake Bienne

D. latus

E. lucius

14.3%

(1/7)

NR

NR

Golay and Mariaux, 1995

Switzerland, Lake Morat

D. latus

P. fluviatilis

5.3%

(1/19)

NR

NR

Golay and Mariaux, 1995

Switzerland, Lake Morat

D. latus

E. lucius

12.5%

(1/8)

NR

NR

Golay and Mariaux, 1995

United Kingdom (Scotland), Loch Awe

D.dendriticus and D. ditremus

O. mykiss

75%

(3/4)

5.0

NR

Dorucu et al., 1995

United Kingdom (Scotland), Loch Rannoch

D.dendriticus and D. ditremus

O. mykiss

42.8%

(3/7)

20.6

NR

Dorucu et al., 1995

United Kingdom (Scotland), Lake of Mentheith

D.dendriticus and D. ditremus

O. mykiss

33.3%

(7/21)

NR

NR

Dorucu et al., 1995

United Kingdom (Scotland), Loch Lomond

D.dendriticus and D. ditremus

S. trutta

25%

(8/2)

2.0

NR

Dorucu et al., 1995

United Kingdom (Scotland), Loch Lomond

D. dendriticus

Coregonus lavaretus

38.5%

(26/10)

NR

NR

Dezfuli et al., 2007

United Kingdom (Scotland), Loch Maragan

D. dendriticus and D. ditremus

S. trutta

45,7%

(16/35)

27.1

NR

Dorucu et al., 1995

United Kingdom (Scotland), Carbeth Reservoir

D. dendriticus and D. ditremus

S. trutta

10%

(2/20)

1.5

NR

Dorucu et al., 1995

United Kingdom (Scotland), Loch Awe

D. dendriticus and D. ditremus

S. trutta

100%

(20/20)

23.8

NR

Dorucu et al., 1995

United Kingdom (Scotland), Dunalastair Reservoir

D. dendriticus and D. ditremus

S. trutta

60.0%

(9/15)

18.7

NR

Dorucu et al., 1995

United Kingdom (Scotland), Cochno Loch

D. dendriticus and D. ditremus

S. trutta

33.3%

(1/3)

NR

NR

Dorucu et al., 1995

United Kingdom (Scotland), Loch Leven

D. dendriticus and D. ditremus

S. trutta

33.3%

(1/3)

4.0

NR

Dorucu et al., 1995

United Kingdom (Scotland), Hill Loch

D. dendriticus and D. ditremus

S. trutta

57.0%

(4/7)

68.2

NR

Dorucu et al., 1995

United Kingdom (Scotland), Loch Rannoch

D. dendriticus and D. ditremus

S. trutta

57.8%

(11/19)

8.0

NR

Dorucu et al., 1995

United Kingdom (Scotland), Bruncrooks

D. dendriticus and D. ditremus

S. trutta

25%

(1/4)

1.0

NR

Dorucu et al., 1995

United Kingdom (Scotland), Talla Reservoir

D. dendriticus and D. ditremus

S. trutta

26.6%

(4/15)

4.2

NR

Dorucu et al., 1995

United Kingdom (Scotland), Loch Venachar

D. dendriticus and D. ditremus

S. trutta

50.0%

(1/2)

2.0

NR

Dorucu et al., 1995

United Kingdom (Scotland), Carron Valley Reservoir

D. dendriticus and D. ditremus

S. trutta

13.6%

(3/22)

2.3

NR

Dorucu et al., 1995

United Kingdom (Scotland), Whiteader Reservoir

D. dendriticus and D. ditremus

S. trutta

15.7%

(3/19)

11.3

NR

Dorucu et al., 1995

United Kingdom (Scotland), Fruid Reservoir

D. dendriticus and D. ditremus

S. trutta

22.2%

(2/9)

1.0

NR

Dorucu et al., 1995

 aMean of parasites by infected fish; bMean of parasites by examined fish; cNot Reported

Table 9 show the prevalence (3.5-23.5%) and mean intensity (1-3.3 plerocercoids/infected fish) of infection with A. pacificus in the coast of Peru. Plerocercoids have not been found in the muscles of different marine fish species.

Table 9. Prevalence and mean intensity of infection by Adenocephalus pacificus in some fish from the coast of Peru

Fish

Prevalence Percentage

(# of Samples)

Mean Intensitya

Reference

Cilus gilberti

23.3

(24/103)

3.3

Chero et al., 2014c

Coryphaena hippurus

11.7

(33/282)

1.1

Escalante et al.,1988

Cynoscion analis

4.7

(2/43)

1.0

Escalante, 1983

Ariopsis seemanni

12.5

(4/32)

1.5

Escalante and Miranda, 1986

Genypterus maculatus

16.7

(1/6)

1.0

Escalante, 1983

Sciaena deliciosa

6.6

(25/381)

NRb

Tantalean, 1975

Sciaena deliciosa

14.3

(16/112)

1.1

Escalante and Miranda, 1986

Sciaena deliciosa

17.1

(6/35)

1.3

Chero et al., 2014a

Sciaena callaensis

23.5

(4/17)

1.8

Escalante, 1983

Trachinotus paitensis

75.0

(12/16)

1.5

Escalante and Miranda, 1986

Trachurus symmetricus murphy

5.7

(4/70)

1.0

Escalante et al., 1988

Paralabrax humeralis

3.5

(369/13)

1.4

Ianaconne and Alvariño, 2009

Merluccius gayi peruanus

6.3

(2/32)

1.0

Escalante, 1983

Merluccius gayi peruanus

4.8

(3/62)

1.0

Chero et al., 2014b

Merluccius gayi peruanus

12.9

(21/163)

1.3

Escalante et al., 1988

Paralichthys adspersus

7.4

2/27

1.0

Escalante and Miranda, 1986

aMean of parasites by infected fish; bNot Reported

In Japan, Oncorhynchus keta, O. masou and O. gorbuscha presented prevalence of 51.1%, 10.8% and 18.5% and mean intensities of 3.9, 3.7, and 1 plerocercoids/infected host with D. nihonkaiensis, respectively (Suzuki et al., 2010). In Hokkaido, juvenile of O. masou do not show any signs of infection before migrating to the sea. However, adults return with prevalence up to 50%, suggesting that these fish could acquire the infection during their migration to the ocean (Awakura et al., 1985; Suzuki et al., 2010; Yanagida et al., 2010).

Fish infection can have economic impact in aquaculture and in the commercialization, export and import of fishes. The presence of parasites in feral or farmed fish would be a high risk for human health, but this risk could be more significant if the plerocercoids were found in the muscle of the fish (Torres et al., 2010, 2012). This last and the higher prevalence and intensity of infection by species of diphyllobothriid of major importance in health are more frequently in feral fish (Torres and Puga, 2011; Torres et al., 2002, Torres et al., 2004a, 2010, 2012). Extensive trade of fresh or chilled fish has led to the description of exotic infections in human living in non-endemic countries (Pastor-Valle et al., 2014) or in endemic countries by non-autochthonous species (Yera et al., 2006; Wicht, 2007, Wicht et al., 2008b; Paugam et al., 2009). It results an impact on the epidemiology of diphyllobothriosis, with a risk of dissemination of the parasitosis (Kuchta et al., 2014).

Dibothriocephalus dendriticus and D. ditremus, both lives as an adult in piscivorous birds and have been identified in farmed O. mykiss in Scotland with prevalence of 1.5% to 11.7% of the hatch trout from 3 raceways (Wootten and Smith, 1979). In a fish farm on the Lake Särkijärvi in Finland, D. dendriticus with prevalence of 2-86% and 1-83% and mean intensity of 1-1.5 and 3.1 plerocercoids in S. trutta m. lacustris and S. trutta m. trutta were identified, respectively (Rahkonen et al.,1996). Dibothriocephalus ditremus was found in visceral organs, musculature and pericardial cavity in stocks of S. salar reared in freshwater cage systems in lakes of Ireland (Rodger, 1991). Prevalence of 4.8% to 6.6% and mean intensity with 1 to 3 plerocercoids of D. dendriticus was reported in viscera of O. mykiss cultured in freshwater cages from 2 farms in the Kola Peninsula, Russia (Karasev et al., 1997). Plerocercoids of D. dendriticus caused an epizootic in 10-18 cm long hatchery-reared O. mykiss in Southern Newfoundland, Canada; causing 7,500 fish died (Khan, 2009). Dibothriocephalus dendriticus was found in viscera of O. mykiss with a prevalence of 6.7% and mean intensity of 1 plerocercoid in a freshwater farm at Lake Tarahuin on Chiloe Island, Chile (Torres et al., 2010). Also, plerocercoids identified as Dibothriocephalus sp. was reported in a trout, O. mykiss, cultivated in the Comau fiord, Chile (Torres et al., 2002).

In China has been reported prevalence of 2.9-91.2% and mean intensities of 1-8.6 plerocercoids/infected hosts in the frogs Rana tigrina, R. limnocharis, R. nigromaculata, R. temporaria chensinensis, R. plancy, R. guentheri, R. cancrivora, Microhyla ornate, Hyla arborea, and Paa yunnanensis (Liu et al., 2015). Also, the authors indicate prevalence of 1.3-100% and mean intensities of 1.8-77.1 larvae/ infected host in the snakes Elaphe plumbea, E. carinata, E. taeniura, E. radiate, E. mandarinus, E. carinata, Natrix tigerini lateralis, Naja naja atra, Zaocys dhumnades, Dinodon rufozonatum, Ptyas korros, P. mucosux, Bungrarus multicinctus, B. albolabris, Deinagkistrodon acutus, and Xenochrophis piscator. In Malaysia is reported R. limnocharis and R. cancrivora with prevalences of 8.8%-21.5%, respectively, and in Australia Litoria caerulea with 4.9% of prevalence (Liu et al., 2015). The occurrence of plerocercoids in lizards and snakes is most likely due the ingestion of infected anurans than by acquisition by infected copepods with procercoids (Oda et al., 2016).

In South America, Leptodactylus latrans (syn. Letodactylus ocellatus) in Uruguay showed 48.6% of prevalence with intensities of up to 22 plerocercoids (Dei-Cas et al., 1976). Tadpoles of Rhinella arenarum (syn. Bufo arenarum) in experimental infections in Argentina presented intensities of up to 14 plerocercoids (Venturini, 1989) and in Pseudopaludicola falcipes and Odontophrynus americanus maximum intensities were of 36 and 7 plerocercoids, respectively (Denegri and Reisin, 1993). In Louisiana, L. catesbeianus and L. clamitans showed prevalence of 1.7% and 4.9%, respectively; also, in 7 species of reptiles, including 4 species of Natrix plus T. proximus proximus, L. getula holbrooki and C. constrictor flaviventris were reported prevalence of 4.8% to 72.7%; in mammals as D. virginiana and P. lotor prevalence was of 60% and 44.8%, respectively; infection was also present in a specimen of U. cinereoargenteus (Corkum, 1966).

Also, sparganosis can have economic impact in high endemic areas related to the induced disabilities.

2.3 Persistence (Survival)

2.3.1 Methods for assessing survival

The viability of eggs of Dibothriocephalus, Diphyllobothrium, Adenocephalus, and Spirometra can be measured by their incubation at optimum temperatures, to obtain the stage of coracidium. The survival of the infective plerocercoids is essentially determined by their capacity of movement; however, their infectivity capacity can be assessed in some cases by experimental infection in laboratory animals. For example, A. pacificus plerocercoids live for 8 to 28 h in ceviche juice, however its infectivity in dogs is limited to 2 h in that liquid (Escalante and Chico-Ruiz, 1988). The criteria recommended by the ESFA (2011) and FDA (2011) European Food Safety Authority or the U.S. Food and Drug Administration for killing plerocercoid stages in fish to prevent human infection are presented in section 1.4.2: Hygiene measures.

3.0 Reducing Environmental Contamination

3.1 Sanitation Technologies

Data on reductions of Dibothriocephalus latus by sanitation management are very scarce and some those given here correspond to von Bonsdorff´s book “Diphyllobothriasis in man” (von Bonsdorff, 1977).

The elimination of eggs from waste waters is essential to avoid the spread of infection to aquatic environments in endemic regions. The eggs are sensitive to environmental conditions and die at temperatures below 5°C in the case of D. latus (von Bonsdorff and Bylund, 1982). The eggs of D. dendriticus also can develop slightly at 4°C (Wright and Curtis, 2000). Eggs isolated from faeces die at 10 min in a solution of 1.8 mg Cl/L but in stools 10 h in 2.5-4.0% solution of chloride of lime is required, although this treatment does not ensure absolute safety (von Bonsdorff, 1977). Also, boiling water is effective to kill D. latus in feces eggs in containers using ratio of 3:1 of water to feces (von Bonsdorff, 1977). Different chemical agents are effective in killing the eggs of D. latus as example alkali, acids, chloroform, ether, alcohol, formaldehyde (von Bonsdorff, 1977, von Bonsdorff and Bylund, 1982).

Dibothriocephalus eggs die in garbage piles where they are exposed to putrefaction and are sensitive to dehydration and will lose their viability in excrements deposited in dry and well aerated latrines (von Bonsdorff, 1977).

Very efficient purification plants are required to eliminate eggs of Dibothriocephalus. Carefully constructed and properly dimensioned purification plants of “three step type” with biological and chemical purification are capable of retaining tapeworm eggs up to 95%. Overloading reduces the purifying effect. Adding filtering through a sand filter or, better, a membrane will give a 100% purifying effect which is certainly obtained by the large plants (von Bonsdorff, 1977). The efficacy of small sewage plants in residential institutions in Russia in decontaminating waste water from helminth eggs, including D. latus, concluded that mechanical processing was 76.4% to 89.4% effective, and that surface filtration beds were fully effective but that the efficacy of drip bio filters was poor (Talapa, 1978).

In Kaliningrad, Russia, the sewage used to irrigate a market garden co-operative following 24 h of mechanical processing, contained on arrival at the irrigation field an average of 8.6 helminth eggs/L, mainly Dibothriocephalus, Ascaris and Trichuris. The soil of areas under overhead irrigation was found to be heavily contaminated but plots where underground irrigation was used were free from ova (Romanenko et al., 1976).

Removing eggs of D. latus in waste water has been made possible by flocculation with aluminum sulfate at 90 to 120 g/m3, at least by 60 min (Doschi, 1972).

From several gums, hydroxypropyl triammonium chloride guar gum (HPTAC-guar) was selected as the most adequate coagulant-flocculant for the class of municipal wastewater obtaining a diminution of 99% of helminth eggs of D. latus, Ascaris lumbricoides, H. nana and Toxocara canis (Zamudio-Perez et al., 2013). Furthermore, the reduction of chemical oxygen demand (COD) and turbidity removals was 46% and 39%, respectively (Zamudio-Perez et al., 2013). These same authors obtained removals of 100% for helminth eggs using the natural gum mixed with Ca (OH)2, and 47% for COD and 30% for turbidity.

If reduction of Dibothriocephalus eggs in water could be efficient in limiting transmission of the parasitosis, reduction of the first intermediate host in water would give the same result on both dibothriocephalosis and sparganosis. Cyclops spp. in the raw water could be completely removed by synergistic effect of chlorine dioxide pre-oxidation followed by coagulation process at chlorine dioxide dosage of 0.9 mg/L. The sort and amount of organic substance in the treated water by chlorine dioxide pre-oxidation were evidently less than that of pre-chlorination and the mutagenicity of drinking water treated by pre-dosing chlorine dioxide was substantially reduced compared with prechlorination (Lin et al., 2007).

It is proven that the quality of the sludge mixed with grass clippings at a ratio of 6:1 volume/volume after having passed a windrow composting process for 8 weeks can be classified as class A biosolids as the levels of remaining fecal coliforms were <3 most probable number per g dry solid and all human parasites were destroyed (Sreesai et al., 2013).

Dibothriocephalus latus eggs may also be rendered innocuous by treatment with chlorine. A 2.5-4% solution of chloride of lime kills the eggs in feces within 10 h (von Bonsdorff, 1977)

Cyclops spp. of zooplankton propagated excessively in eutrophic water body and could not be effectively inactivated by the conventional disinfections process like chlorination due to its stronger resistance to oxidation (Lin et al., 2007). It was found that chlorine dioxide possessed better inactivation effect than chlorine. Cyclops can be completely inactivated after 30 min of contact time by low dosage of chlorine dioxide (1.0 mg/L) (Zhao et al., 2007). The size of Cyclops and its life activity are important influencing factors by oxidants (Zhang et al., 2009).

Up to 56% of the eggs of D. latus were inactivated at 222 nm of water surface radiation dose (up to 5 mJ/cm2), but the eggs of Opisthorchis felineus were only inactivated up to 85% (Lipatov et al., 2016). These differences are attributed to some features of eggs shells and showed the importance of considering the morphological and chemical composition of the eggs that can vary between groups or species of helminth parasites, when the criteria of water treatment is evaluated (Lipatov et al., 2016).

Data on control of dibothriocephalosis have focused on the treatment of wastewater containing parasite egg disseminated by the definitive host through their feces. Nevertheless, simple individual or collective means could be efficient in reducing the number of parasites in water. Elimination of parasites after medical and veterinary treatment should be burned, incinerated or buried in the soil.

Traffic on lakes has also been an efficient way to transmit eggs. Nowadays, the situation is under control as most boats are compelled to keep their black waters on board but this will not apply on small unroofed boats (Khromenkova, 1995). Sanitary treatment of fish for the control of the source of human infection is revised in Part I, Section 1.4.4 Hygiene measures).

It seems that prevalence of human dibothriocephalosis and sparganosis over time depended of human habits: how foods were stored or consumed, if drinking water and latrines were used (Le Bailly et al., 2007; Piers, 2015).

References

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