Hostname: page-component-78c5997874-s2hrs Total loading time: 0 Render date: 2024-11-19T12:16:12.909Z Has data issue: false hasContentIssue false
  • English
  • Français

Occurrence of Schistosoma bovis on Pemba Island, Zanzibar: implications for urogenital schistosomiasis transmission monitoring

Published online by Cambridge University Press:  08 August 2018

Tom Pennance Open the ORCID record for Tom Pennance [Opens in a new window] ,
Shaali M. Ame ,
Armour Khamis Amour ,
Khamis Rashid Suleiman ,
Fiona Allan ,
David Rollinson  and
Bonnie L. Webster Open the ORCID record for Bonnie L. Webster [Opens in a new window]
Tom Pennance
Affiliation:
Natural History Museum, Cromwell Road, London SW75BD, UK London Centre for Neglected Tropical Disease Research, Imperial College London, School of Public Health, Norfolk Pl, Paddington, London W2 1PG, UK Cardiff University, Cardiff CF10 3AT, UK
Shaali M. Ame
Affiliation:
Public Health Laboratory, Chake Chake, Pemba, United Republic of Tanzania
Armour Khamis Amour
Affiliation:
Public Health Laboratory, Chake Chake, Pemba, United Republic of Tanzania
Khamis Rashid Suleiman
Affiliation:
Public Health Laboratory, Chake Chake, Pemba, United Republic of Tanzania
Fiona Allan
Affiliation:
Natural History Museum, Cromwell Road, London SW75BD, UK London Centre for Neglected Tropical Disease Research, Imperial College London, School of Public Health, Norfolk Pl, Paddington, London W2 1PG, UK
David Rollinson
Affiliation:
Natural History Museum, Cromwell Road, London SW75BD, UK London Centre for Neglected Tropical Disease Research, Imperial College London, School of Public Health, Norfolk Pl, Paddington, London W2 1PG, UK
Bonnie L. Webster*
Affiliation:
Natural History Museum, Cromwell Road, London SW75BD, UK London Centre for Neglected Tropical Disease Research, Imperial College London, School of Public Health, Norfolk Pl, Paddington, London W2 1PG, UK
*
Author for correspondence: Bonnie L. Webster, E-mail: [email protected]
Rights & Permissions [Opens in a new window]

Abstract

The causative agent of urogenital schistosomiasis, Schistosoma haematobium, was thought to be the only schistosome species transmitted through Bulinus snails on Unguja and Pemba Island (Zanzibar, United Republic of Tanzania). For insights into the environmental risk of S. haematobium transmission on Pemba Island, malacological surveys collecting Bulinus globosus and B. nasutus, two closely related potential intermediate hosts of S. haematobium were conducted across the island in November 2016. Of 1317 B. globosus/B. nasutus collected, seven B. globosus, identified through sequencing a DNA region of the mitochondrial cytochrome oxidase subunit 1 (cox1), were observed with patent infections assumed to be S. haematobium. However, when the collected cercariae were identified through sequencing a region of the cox1 and the nuclear internal transcribed spacer (ITS1 + 2), schistosomes from five of these B. globosus collected from a single locality were in fact S. bovis. The identified presence of S. bovis raises concerns for animal health on Pemba, and complicates future transmission monitoring of S. haematobium. These results show the pertinence for not only sensitive, but also species-specific markers to be used when identifying cercariae during transmission monitoring, and also provide the first molecular confirmation for B. globosus transmitting S. bovis in East Africa.

Keywords

BulinuscattlePembaSchistosoma bovisSchistosoma haematobiumschistosomiasisschistosomessnailsZanzibar
Type
Special Issue Research Article
Information
Parasitology , Volume 145 , Special Issue 13: 2017 Autumn Symposium of the British Society for Parasitology The multi-disciplinarity of parasitology: Host-parasite evolution in an ever changing world , November 2018 , pp. 1727 - 1731
Check for updates
Creative Commons
Creative Common License - CCCreative Common License - BY
This is an Open Access article, distributed under the terms of the Creative Commons Attribution licence (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted re-use, distribution, and reproduction in any medium, provided the original work is properly cited.
Copyright
Copyright © Cambridge University Press 2018

Introduction

The snail-borne neglected tropical disease (NTD), schistosomiasis, is the most important freshwater parasitic disease of humans associated with poverty, poor sanitation and lack of safe water supplies (Steinmann et al., Reference Steinmann, Keiser, Bos, Tanner and Utzinger2006; Hotez et al., Reference Hotez, Alvarado, Basáñez, Bolliger, Bourne, Boussinesq, Brooker, Brown, Buckle, Budke, Carabin, Coffeng, Fèvre, Fürst, Halasa, Jasrasaria, Johns, Keiser, King, Lozano, Murdoch, O'Hanlon, Pion, Pullan, Ramaiah, Roberts, Shepard, Smith, Stolk, Undurraga, Utzinger, Wang, Murray and Naghavi2014), with an estimated 180–200 million people primarily from low- and middle-income countries being infected (GBD 2016 Disease and Injury Incidence and Prevalence Collaborators, 2017). Ambitious goals to eliminate schistosomiasis have been announced by the WHO as part of its roadmap to overcome the global impact of NTDs by 2020–2025 (WHO, 2012). Whilst mass drug administration, behavioural change through education and snail control are having a major impact on schistosomiasis, further research into schistosome transmission biology together with better tools for transmission monitoring and surveillance are required to help achieve and monitor the success of these ambitious goals (Stothard et al., Reference Stothard, Campbell, Osei-Atweneboana, Durant, Stanton, Biritwum, Rollinson, Ombede and Tchuem-Tchuenté2017). Schistosomiasis is also a disease of animals, with large numbers of domestic livestock affected worldwide but the actual veterinary and economic impact is largely unknown (De Bont and Vercruysse, Reference De Bont and Vercruysse1997, Reference De Bont and Vercruysse1998).

There are 25 recognized species of mammalian schistosomes that cause human and animal infections, which can be split into four Schistosoma species groups (Webster et al., Reference Webster, Southgate and Littlewood2006). The largest group is the Schistosoma haematobium group containing nine species that are all transmitted through Bulinus snails (Brown, Reference Brown1994) with two species, S. haematobium and S. bovis, being responsible for the majority of all human (Hotez and Kamath, Reference Hotez and Kamath2009) and livestock infections (De Bont and Vercruysse, Reference De Bont and Vercruysse1997), respectively. Central to this group is S. haematobium, a major human schistosome species being the most widespread and prevalent across Africa and solely responsible for human urogenital schistosomiasis with often severe pathology (Schwartz, Reference Schwartz1981; Leutscher et al., Reference Leutscher, Ramarokoto, Reimert, Feldmeier, Esterre and Vennervald2000; Bustinduy et al., Reference Bustinduy, King, Scott, Appleton, Sousa-Figueiredo, Betson and Stothard2014; Kjetland et al., Reference Kjetland, Hegertun, Baay, Onsrud, Ndhlovu and Taylor2014; Christinet et al., Reference Christinet, Lazdins-Helds, Stothard and Reinhard-Rupp2016). Schistosoma bovis is a pathogen of domestic livestock and some artiodactylids (Standley et al., Reference Standley, Mugisha, Dobson and Stothard2012), with its distribution commonly overlapping with that of S. haematobium across mainland Africa (Moné et al., Reference Moné, Mouahid and Morand1999), and utilising a wide range of Bulinus (Southgate and Knowles, Reference Southgate and Knowles1975a, Reference Southgate and Knowles1975b; Stothard et al., Reference Stothard, Lockyer, Kabatereine, Tukahebwa, Kazibwe, Rollinson and Fenwick2004). These two species, among others, are also able to hybridize and inter-specific hybridization is now recognized in West Africa with possible detrimental consequences on disease control (Huyse et al., Reference Huyse, Webster, Geldof, Stothard, Diaw, Polman and Rollinson2009; Webster et al., Reference Webster, Diaw, Seye, Webster and Rollinson2013; Léger and Webster, Reference Léger and Webster2017).

Pemba and Unguja Islands (Zanzibar Archipelago, United Republic of Tanzania) have been historically identified as ‘model islands’ for implementing multiple effective infectious disease control and elimination programmes in sub-Saharan Africa (Pennance et al., Reference Pennance, Person, Muhsin, Khamis, Muhsin, Khamis, Mohammed, Kabole, Rollinson and Knopp2016). For schistosomiasis control, Zanzibar also offers an advantage due to the allopatric transmission of S. haematobium through a single snail host, Bulinus globosus, on both Islands (Stothard et al., Reference Stothard, Loxton, Rollinson, Mgeni, Khamis, Ameri, Ramsan and Savioli2000), whereas across most of sub-Saharan Africa, multiple Schistosoma and Bulinus species occur in sympatry (Brown, Reference Brown1994), complicating control interventions and surveillance. Urogenital schistosomiasis was highly endemic on both islands but is now targeted for elimination (Knopp et al., Reference Knopp, Mohammed, Ali, Khamis, Ame, Albonico, Gouvras, Fenwick, Savioli and Colley2012, Reference Knopp, Person, Ame, Mohammed, Ali, Khamis, Rabone, Allan, Gouvras and Blair2013).

As we move towards or reach elimination, there becomes a need for more sensitive methods to monitor the levels of transmission when egg–patent human infections become scarce (Le and Hsieh, Reference Le and Hsieh2017; Stothard et al., Reference Stothard, Campbell, Osei-Atweneboana, Durant, Stanton, Biritwum, Rollinson, Ombede and Tchuem-Tchuenté2017), the risk of infection and also a way to prove transmission interruption when it is finally reached. Xenomonitoring is a nucleic acid-based molecular diagnostic used to monitor the transmission of several vector-borne diseases (Cunningham et al., Reference Cunningham, Lingley, Haines, Ndung'u, Torr and Adams2016; Minetti et al., Reference Minetti, LaCourse, Reimer and Storhard2016; Cook et al., Reference Cook, Pilotte, Minetti, Williams and Reimer2017), including to some extent schistosomiasis where tools are being developed for the xenomonitoring of snails that could support schistosomiasis transmission and elimination monitoring (Hamburger et al., Reference Hamburger, Hoffman, Kariuki, Muchiri, Ouma, Koech, Sturrock and King2004; Allan et al., Reference Allan, Dunn, Emery, Stothard, Johnston, Kane, Khamis, Mohammed and Rollinson2013; Lu et al., Reference Lu, Zhang, Mutuku, Mkoji and Loker2016; Abbasi et al., Reference Abbasi, Webster, King, Rollinson and Hamburger2017). The first stage for snail xenomonitoring for schistosomiasis is the identification of patent schistosome infections within the snails and collecting cercariae shed from them. Here, we report on the molecular identification of these cercariae and the infected snails collected from Pemba Island (Zanzibar) and how the findings complicate the development of robust molecular xenomonitoring protocols for ongoing and future transmission monitoring.

Methods

Malacological surveys and Schistosoma collection

In November 2016, as part of a larger ongoing molecular xenomonitoring study on Pemba, Bulinus snails were collected, by scooping, from human freshwater contact sites in eight shehias (smallest division of administrative regions), examined and individually induced to shed cercariae following previous methods (Allan et al., Reference Allan, Dunn, Emery, Stothard, Johnston, Kane, Khamis, Mohammed and Rollinson2013). An experienced microscopist identified schistosome cercariae, which were individually pipetted in 3.5 µL aliquots onto Whatman FTA cards (Whatman, Part of GE Healthcare, Florham Park, USA) for long-term deoxyribonucleic acid (DNA) storage. After shedding, all infected snails were preserved in 100% ethanol for future morphological and molecular characterization.

Schistosoma and Bulinus identification

DNA from individual cercariae was eluted from the FTA cards (Webster et al., Reference Webster, Rabone, Pennance, Emery, Allan, Gouvras, Knopp, Garba, Hamidou, Mohammed, Ame, Rollinson and Webster2015) and characterized by amplification and sequencing of the mitochondrial cytochrome oxidase subunit 1 (cox1) and partial nuclear internal transcribed spacer (ITS1 + 2) DNA regions (Webster et al., Reference Webster, Emery, Webster, Gouvras, Garba, Diaw, Seye, Tchuente, Simoonga, Mwanga, Lange, Kariuki, Mohammed, Stothard and Rollinson2012).

To determine the species of the infected snails, total genomic DNA was extracted from the whole snail tissue using the DNeasy Blood & Tissue Kit (Qiagen, Manchester, UK), with minor changes to the standard protocol in that quantities of the digest reagents were doubled and digests were incubated for at least 12 h. From each snail, a 623 base pair region of the mitochondrial cox1 gene was amplified and Sanger sequenced using primers BulCox1 and CO2 following previous protocols (Kane et al., Reference Kane, Stothard, Emery and Rollinson2008). The sequence data were manually edited in Sequencher v5.1 (http://genecodes.com) before being compared with reference sequence databases for Bulinus (Kane et al., Reference Kane, Stothard, Emery and Rollinson2008) and Schistosoma (Webster et al., Reference Webster, Emery, Webster, Gouvras, Garba, Diaw, Seye, Tchuente, Simoonga, Mwanga, Lange, Kariuki, Mohammed, Stothard and Rollinson2012, Reference Webster, Diaw, Seye, Webster and Rollinson2013) to confirm species.

Results

In total, 1317 B. globosus and B. nasutus were collected, seven of these snails (Table 1) from Kinyasini (6) and Chambani (1) shehia were shedding schistosome cercariae (Fig. 1). The infected snails were identified as B. globosus with two cox1 haplotypes recognized (GenBank accession numbers: MH014040 and MH014041) which matched those snails previously reported from Pemba (Kane et al., Reference Kane, Stothard, Emery and Rollinson2008). Cercariae collected from these were assumed initially to be the human parasite S. haematobium; however, molecular characterizations of the cercariae from five of these snails, collected from a stream in Kinyasini (Kinya6), were identified as S. bovis (Table 1). Two different S. bovis cox1 haplotypes [Genbank accessions: S.b (i) MH014042 and S.b (ii) MH014043] (Table 1) were identified from these five snails; three snails producing S. bovis cercariae of a single haplotype and two snails producing S. bovis cercariae of both haplotypes suggesting that they had been infected by more than one miracidium.

Fig. 1. Map outlining shehias (smallest division of administrative regions) on Pemba Island, Zanzibar (United Republic of Tanzania) showing the location and images of two freshwater bodies in Kinyasini (Kinya2 and Kinya6) and one in Chambani (Cham10) where Schistosoma haematobium (red) and Schistosoma bovis (green) cercariae were recovered from Bulinus globosus. GPS coordinates for sites (latitude and longitude in decimal degrees): Kinya2 (−5.02033°, 39.73855°); Kinya6 (−5.03560°, 39.73850°); Cham10 (−5.35805°, 39.79182°).

Table 1. Showing the collection sites and genetic profiles of the Bulinus and schistosome cercariae analysed

Two Bulinus globosus cox1 haplotypes [Genbank accessions: (a) MH014040 and (b) MH014041]. Two S. haematobium cercariae cox1 haplotypes, Genbank accessions: S.h (i) MH014046 and S.h (ii) MH01404 and the two S. bovis cox1 haplotypes, Genbank accessions: S.b (i) MH014042 and S.b (ii) MH014043. ITS1 + 2 profiles showed no intra species variation (Genbank accessions: S.h MH014047 and S.b MH014044).

The other two infected snails shed S. haematobium cercariae and were collected from a pond in Chambani (Cham10) and a different stream site in Kinyasini (Kinya2). The S. haematobium cercariae from Kinyasini and Chambani, respectively, were of two different S. haematobium cox1 haplotypes [Genbank accessions: S.h (i) MH014046 and S.h (ii) MH014045] with only single haplotypes produced from each snail. These haplotypes matched those identified as group 2 S. haematobium cox1 haplotypes found only in the Indian Ocean Islands (Webster et al., Reference Webster, Emery, Webster, Gouvras, Garba, Diaw, Seye, Tchuente, Simoonga, Mwanga, Lange, Kariuki, Mohammed, Stothard and Rollinson2012).

ITS1 + 2 profiles showed no intra-species variation (Genbank Accessions: S.h MH014047 and S.b MH014044) and were identified as either S. bovis or S. haematobium by the three inter-specific single nucleotide polymorphisms (Webster et al., Reference Webster, Emery, Webster, Gouvras, Garba, Diaw, Seye, Tchuente, Simoonga, Mwanga, Lange, Kariuki, Mohammed, Stothard and Rollinson2012).

Discussion

The detection of S. bovis on Pemba Island poses a potentially new threat to domestic livestock and wildlife health in Zanzibar (De Bont and Vercruysse, Reference De Bont and Vercruysse1997, Reference De Bont and Vercruysse1998; Standley et al., Reference Standley, Mugisha, Dobson and Stothard2012). The site where S. bovis transmission was identified had grazing cattle (see Fig. 1, Kinya6) in close proximity to the water where the shedding snails were collected; therefore, it is quite likely that ongoing transmission is being maintained. Moreover, the movement of infected cattle could enable the spread of the infection particularly as B. globosus are found throughout most of the island (Stothard et al., Reference Stothard, Mgeni, Alawi, Savioli and Rollinson1997).

The presence of S. bovis complicates the monitoring of S. haematobium transmission since both parasites are shown here to infect the same intermediate snail host and cannot be distinguished from each other easily by microscopy. Therefore, S. bovis-infected B. globosus could be falsely identified as infected with S. haematobium, or vice-versa, complicating urogenital schistosomiasis transmission monitoring. This accentuates the need for routine molecular identification of schistosome infections in snails during malacological surveys (Minetti et al., Reference Minetti, LaCourse, Reimer and Storhard2016), and the development of more species-specific xenomonitoring tools to differentiate S. bovis and S. haematobium transmission (Webster et al., Reference Webster, Rollinson, Stothard and Huyse2010; Abbasi et al., Reference Abbasi, Webster, King, Rollinson and Hamburger2017). The identification of schistosome cercariae shed from snails is often presumed to be of a particular species due to the snail host involved or the locality of the transmission. Our findings strongly emphasize that these assumptions are not accurate and transmission dynamics of different species may change over time and space. The assumed transmission of only S. haematobium by B. globosus on Zanzibar and the non-identification of these S. bovis infections would have led us to believe that the level of S. haematobium transmission is much higher than it actually is, hampering ongoing and future urogenital schistosomiasis transmission monitoring and surveillance.

Schistosoma haematobium and S. bovis hybridization has also been detected in sympatric West African areas (Webster et al., Reference Webster, Diaw, Seye, Webster and Rollinson2013). Zanzibar was considered to be an allopatric area for S. haematobium (Webster et al., Reference Webster, Emery, Webster, Gouvras, Garba, Diaw, Seye, Tchuente, Simoonga, Mwanga, Lange, Kariuki, Mohammed, Stothard and Rollinson2012) but the identification of this sympatry with S. bovis could, in time, lead to inter-species hybridization. The potential consequences of hybridization include increased host associations of hybrids, possible zoonotic transmission and hybrid vigour (Huyse et al., Reference Huyse, Webster, Geldof, Stothard, Diaw, Polman and Rollinson2009; Webster et al., Reference Webster, Diaw, Seye, Webster and Rollinson2013; Léger and Webster, Reference Léger and Webster2017). Investigating the origin of S. bovis being transmitted on Pemba, by genetic comparison with other mainland strains of S. bovis, may help elucidate how this parasite has been imported to Zanzibar. Since the eradication of the tsetse fly, the vector of human and African animal trypanosomiasis, on Unguja Island (Vreysen et al., Reference Vreysen, Saleh, Ali, Abdulla, Zhu, Juma, Dyck, Msangi, Mkonyi and Feldmann2000), there has been an increase of cattle farming (Mdoe, Reference Mdoe2003) facilitated by the import of cattle under strict guidelines of the United Republic of Tanzania's Animal Resources Management Act (1999). Bovine schistosomiasis however is widely ignored/unknown as a veterinary health problem, and therefore is currently not included in these guidelines. This oversight could offer some explanation to how and within what time scale the introduction, or multiple introductions, of S. bovis may have occurred. Additionally, the prevalence and intensity of S. bovis in local cattle and other potential artiodactylid hosts (Standley et al., Reference Standley, Mugisha, Dobson and Stothard2012), such as the Ader's duiker (Cephalophus adersi) endemic to Zanzibar, should be determined to assess the impact on livestock and wildlife health. However, diagnosing S. bovis from the definitive host remains challenging, with the detection of S. bovis eggs in the stool being difficult and the more sensitive method of observing adult worms in the host being only possible post-mortem via dissection. An antigen-based test with promising diagnostic performance has been developed (de la Torre-Escudero et al., Reference de la Torre-Escudero, Manzano-Román, Pérez-Sánchez, Barrera, Siles-Lucas and Oleaga2012), which could offer a sensitive method for judging the epidemiology of S. bovis in Pemba.

Due to the difficulty in classifying species within the Bulinus africanus species complex (Kane et al., Reference Kane, Stothard, Emery and Rollinson2008), previous findings on snail–schistosome compatibilities should be treated with some caution. This molecular confirmation of B. globosus naturally transmitting S. bovis in East Africa gives credibility to a previous observation (Mwambungu, Reference Mwambungu1988), and dispels previous claims of B. globosus being naturally refractory (Christensen et al., Reference Christensen, Mutani and Frandsen1983) or only an intermediate host in West Africa (Diaw and Vassiliades, Reference Diaw and Vassiliades1987; Ndifon et al., Reference Ndifon, Betterton and Rollinson1988). Previous evidence for compatibility of B. nasutus with S. bovis in East Africa is also tainted with contradicting evidence, some showing natural infections (Dowdeswell, Reference Dowdeswell1938; Kinoti, Reference Kinoti1964b) going against failed experimental infections (Southgate and Knowles, Reference Southgate and Knowles1975a, Reference Southgate and Knowles1975b; Southgate et al., Reference Southgate, Rollinson, Ross and Knowles1980) and a lack of naturally infected B. nasutus in other endemic areas (Kinoti, Reference Kinoti1964a; Southgate et al., Reference Southgate, Rollinson, Ross and Knowles1980; Mutani et al., Reference Mutani, Christensen and Frandsen1983). It is likely that S. bovis has a broad intermediate host range in East Africa utilising several Bulinus species, as it has also been identified from B. ugandae (Malek, Reference Malek1969), B. africanus (McClelland, Reference McClelland1955; Teesdale and Nelson, Reference Teesdale and Nelson1958; Kassuku et al., Reference Kassuku, Christensen, Monrad, Nansen and Knudsen1986) and B. forskalii (McClelland, Reference McClelland1955). Therefore, studies to confirm the intermediate snail host vectoral capacity and specificity of S. bovis to B. globosus or indeed other endemic Bulinus species on Pemba, including B. nasutus and B. forskalii, are required to determine the transmission potential and possible spread of this emerging schistosome in Zanzibar.

Acknowledgements

Thanks to Said Mohammed Ali and staff at the Public Health Laboratory – Ivo de Carneri for making the surveys and collections possible, and also to Dr Steffi Knopp at the Swiss Tropical and Public Health Institute in Basel for helping to identify study sites. Thanks also to the Natural History Museums DNA Sequencing Facilities for the sequencing services.

Financial support

The authors would like to thank the London Centre for Neglected Tropical Disease Research (LCNTDR) and University of Georgia Research Foundation, Inc., which was funded by the Bill & Melinda Gates Foundation for the Schistosomiasis Consortium for Operational Research and Evaluation (SCORE) project, for funding TP and BW, respectively, for travel and expenses to undertake the field collections in November 2016. FA is financially supported by the Wellcome Trust (SCAN Project WT104958MA). The authors would also like to acknowledge the Natural History Museum's Departmental Investment Fund for the financial support facilitating the molecular work.

Conflict of interest

None.

Ethical standards

Not applicable.

References

Abbasi, I, Webster, BL, King, CH, Rollinson, D and Hamburger, J (2017) The substructure of three repetitive DNA regions of Schistosoma haematobium group species as a potential marker for species recognition and interbreeding detection. Parasites and Vectors 10, 364.Google Scholar
Allan, FE, Dunn, AM, Emery, AM, Stothard, JR, Johnston, DA, Kane, RA, Khamis, AN, Mohammed, KA and Rollinson, D (2013) Use of sentinel snails for the detection of Schistosoma haematobium transmission on Zanzibar and observations on transmission patterns. Acta Tropica 128, 234–240.Google Scholar
Brown, DS (1994) Freshwater Snails of Africa and Their Medical Importance, 2nd Edn. London, UK: Taylor & Francis.Google Scholar
Bustinduy, A, King, C, Scott, J, Appleton, S, Sousa-Figueiredo, JC, Betson, M and Stothard, JR (2014) HIV and schistosomiasis co-infection in African children. The Lancet Infectious Diseases 14, 640–649.Google Scholar
Christensen, NØ, Mutani, A and Frandsen, F (1983) A review of the biology and transmission ecology of African bovine species of the genus Schistosoma. Zeitschrift Für Parasitenkunde 69, 551–570.Google Scholar
Christinet, V, Lazdins-Helds, JK, Stothard, JR and Reinhard-Rupp, J (2016) Female genital schistosomiasis (FGS): from case reports to a call for concerted action against this neglected gynaecological disease. International Journal for Parasitology 46, 395–404.Google Scholar
Cook, DAN, Pilotte, N, Minetti, C, Williams, SA and Reimer, LJ (2017) A superhydrophobic cone to facilitate the xenomonitoring of filarial parasites, malaria, and trypanosomes using mosquito excreta/feces [version 2; referees: 2 approved]. Gates Open Research 1, 7. doi: 10.12688/gatesopenres.12749.1.Google Scholar
Cunningham, LJ, Lingley, JK, Haines, LR, Ndung'u, JM, Torr, SJ and Adams, ER (2016) Illuminating the prevalence of Trypanosoma brucei s.l. in Glossina using LAMP as a tool for xenomonitoring. PLoS Neglected Tropical Diseases 10, e0004441.Google Scholar
De Bont, J and Vercruysse, J (1997) The epidemiology and control of cattle schistosomiasis. Parasitology Today 13, 255–262.Google Scholar
De Bont, J and Vercruysse, J (1998) Schistosomiasis in cattle. Advances in Parasitology 41, 285–364.Google Scholar
de la Torre-Escudero, E, Manzano-Román, R, Pérez-Sánchez, R, Barrera, I, Siles-Lucas, M and Oleaga, A (2012) Molecular cloning, characterization and diagnostic performance of the Schistosoma bovis 22.6 antigen. Veterinary Parasitology 190, 530–540.Google Scholar
Diaw, OT and Vassiliades, G (1987) Epidémiologie des schistosomoses du bétail au Sénégal. Revue d'Elevage et de Médecine Vétérinaire Des Pays Tropicaux 40, 265–274.Google Scholar
Dowdeswell, RM (1938) Schistosomiasis in the Kavirondo district of Kenya colony. Transactions of the Royal Society of Tropical Medicine and Hygiene 31, 673–688.Google Scholar
GBD 2016 Disease and Injury Incidence and Prevalence Collaborators (2017) Global, regional, and national incidence, prevalence, and years lived with disability for 328 diseases and injuries for 195 countries, 1990–2016: a systematic analysis for the Global Burden of Disease Study 2016. The Lancet 390, 1211–1259.Google Scholar
Hamburger, J, Hoffman, O, Kariuki, HC, Muchiri, EM, Ouma, JH, Koech, DK, Sturrock, RF and King, CH (2004) Large-scale, polymerase chain reaction-based surveillance of Schistosoma haematobium DNA in snails from transmission sites in coastal Kenya: a new tool for studying the dynamics of snail infection. American Journal of Tropical Medicine and Hygiene 71, 765–773.Google Scholar
Hotez, PJ and Kamath, A (2009) Neglected tropical diseases in sub-Saharan Africa: review of their prevalence, distribution, and disease burden. PLoS Neglected Tropical Diseases 3, 1–10.Google Scholar
Hotez, PJ, Alvarado, M, Basáñez, M-G, Bolliger, I, Bourne, R, Boussinesq, M, Brooker, SJ, Brown, AS, Buckle, G, Budke, CM, Carabin, H, Coffeng, LE, Fèvre, EM, Fürst, T, Halasa, YA, Jasrasaria, R, Johns, NE, Keiser, J, King, CH, Lozano, R, Murdoch, ME, O'Hanlon, S, Pion, SD, Pullan, RL, Ramaiah, KD, Roberts, T, Shepard, DS, Smith, JL, Stolk, WA, Undurraga, EA, Utzinger, J, Wang, M, Murray, CJ and Naghavi, M (2014) The Global Burden of Disease Study 2010: interpretation and implications for the neglected tropical diseases. PLoS Neglected Tropical Diseases 8, e2865.Google Scholar
Huyse, T, Webster, BL, Geldof, S, Stothard, JR, Diaw, OT, Polman, K and Rollinson, D (2009) Bidirectional introgressive hybridization between a cattle and human schistosome species. PLoS Pathogens 5, e1000571.Google Scholar
Kane, RA, Stothard, JR, Emery, AM and Rollinson, D (2008) Molecular characterization of freshwater snails in the genus Bulinus: a role for barcodes? Parasites and Vectors 1, 15.Google Scholar
Kassuku, A, Christensen, NO, Monrad, J, Nansen, P and Knudsen, J (1986) Epidemiological studies on Schistosoma bovis in Iringa Region, Tanzania. Acta Tropica 43, 153–163.Google Scholar
Kinoti, G (1964 a) A note on the susceptibility of some gastropod molluscs to Schistosoma bovis and S. mattheei. Annals of Tropical Medicine & Parasitology 58, 270–275.Google Scholar
Kinoti, G (1964 b) Observations on the transmission of Schistosoma haematobium and Schistosoma bovis in the Lake Region of Tanganyika. Bulletin of the World Health Organization 31, 815.Google Scholar
Kjetland, EF, Hegertun, IEA, Baay, MFD, Onsrud, M, Ndhlovu, PD and Taylor, M (2014) Genital schistosomiasis and its unacknowledged role on HIV transmission in the STD intervention studies. International Journal of STD & AIDS 25, 705–715.Google Scholar
Knopp, S, Mohammed, KA, Ali, SM, Khamis, IS, Ame, SM, Albonico, M, Gouvras, A, Fenwick, A, Savioli, L and Colley, DG (2012) Study and implementation of urogenital schistosomiasis elimination in Zanzibar (Unguja and Pemba islands) using an integrated multidisciplinary approach. BMC Public Health 12, 930.Google Scholar
Knopp, S, Person, B, Ame, SM, Mohammed, KA, Ali, SM, Khamis, IS, Rabone, M, Allan, F, Gouvras, A and Blair, L (2013) Elimination of schistosomiasis transmission in Zanzibar: baseline findings before the onset of a randomized intervention trial. PLoS Neglected Tropical Diseases 7, e2474.Google Scholar
Le, L and Hsieh, MH (2017) Diagnosing urogenital schistosomiasis: dealing with diminishing returns. Trends in Parasitology 33, 378–387.Google Scholar
Léger, E and Webster, JP (2017) Hybridizations within the genus Schistosoma: implications for evolution, epidemiology and control. Parasitology 144, 65–80.Google Scholar
Leutscher, P, Ramarokoto, C-E, Reimert, C, Feldmeier, H, Esterre, P and Vennervald, BJ (2000) Community-based study of genital schistosomiasis in men from Madagascar. The Lancet 355, 117–118.Google Scholar
Lu, L, Zhang, S-M, Mutuku, MW, Mkoji, GM and Loker, ES (2016) Relative compatibility of Schistosoma mansoni with Biomphalaria sudanica and B. pfeifferi from Kenya as assessed by PCR amplification of the S. mansoni ND5 gene in conjunction with traditional methods. Parasites and Vectors 9, 166.Google Scholar
Malek, EA (1969) Studies on bovine schistosomiasis in the Sudan. Annals of Tropical Medicine & Parasitology 63, 501–513.Google Scholar
McClelland, WFJ (1955) Two species of Bulinus found naturally infected with a bovine schistosome in Western Kenya. Transactions of the Royal Society of Tropical Medicine and Hygiene 49, 295.Google Scholar
Mdoe, NSY (2003) Livestock and agriculture development in Zanzibar, post-tsetse eradication: a follow-up socioeconomic study. International Atomic Energy Agency confidential report, Vienna. https://doi.org/10.1371/journal.pntd.0002857.s002.Google Scholar
Minetti, C, LaCourse, JE, Reimer, L and Storhard, JR (2016) Focusing nucleic acid-based molecular diagnostics and xenomonitoring approaches for human helminthiases amenable to preventive chemotherapy. Parasitology Open 2, e16.Google Scholar
Moné, H, Mouahid, G and Morand, S (1999) The distribution of Schistosoma bovis Sonsino, 1876 in relation to intermediate host mollusc-parasite relationships. Advances in Parasitology 44, 99–138.Google Scholar
Mutani, A, Christensen, NØ and Frandsen, F (1983) Studies on the relationship between Schistosoma and their intermediate hosts. Parasitology Research 69, 483–487.Google Scholar
Mwambungu, JA (1988) Transmission of Schistosoma bovis in Mkulwe (Mbozi District, Mbeya Region, Southern Highlands of Tanzania). Journal of Helminthology 62, 29–32.Google Scholar
Ndifon, GT, Betterton, C and Rollinson, D (1988) Schistosoma curassoni Brumpt, 1931 and S. bovis (sonsino, 1876) in cattle in northern Nigeria. Journal of Helminthology 62, 33–34.Google Scholar
Pennance, T, Person, B, Muhsin, MA, Khamis, AN, Muhsin, J, Khamis, IS, Mohammed, KA, Kabole, F, Rollinson, D and Knopp, S (2016) Urogenital schistosomiasis transmission on Unguja Island, Zanzibar: characterisation of persistent hot-spots. Parasites and Vectors 9, 646.Google Scholar
Schwartz, DA (1981) Helminths in the induction of cancer II. Schistosoma haematobium and bladder cancer. Tropical and Geographical Medicine 33, 1–7.Google Scholar
Southgate, VR and Knowles, RJ (1975 a) Observations on Schistosoma bovis Sonsino, 1876. Journal of Natural History 9, 273–314.Google Scholar
Southgate, VR and Knowles, RJ (1975 b) The intermediate hosts of Schistosoma bovis in western Kenya. Transactions of the Royal Society of Tropical Medicine and Hygiene 69, 356–357.Google Scholar
Southgate, VR, Rollinson, D, Ross, GC and Knowles, RJ (1980) Observations on an isolate of Schistosoma bovis from Tanzania. Parasitology Research 63, 241–249.Google Scholar
Standley, CJ, Mugisha, L, Dobson, AP and Stothard, JR (2012) Zoonotic schistosomiasis in non-human primates: past, present and future activities at the human-wildlife interface in Africa. Journal of Helminthology 86, 131–140.Google Scholar
Steinmann, P, Keiser, J, Bos, R, Tanner, M and Utzinger, J (2006) Schistosomiasis and water resources development: systematic review, meta-analysis, and estimates of people at risk. The Lancet Infectious Diseases 6, 411–425.Google Scholar
Stothard, JR, Mgeni, AF, Alawi, KS, Savioli, I and Rollinson, D (1997) Observations on shell morphology, enzymes and Random Amplified Polymorphic DNA (RAPD) in Bulinus africanus group snails (Gastropoda: Planorbidae) in Zanzibar. Journal of Molluscan Studies 63, 489–503.Google Scholar
Stothard, JR, Loxton, N, Rollinson, D, Mgeni, AF, Khamis, S, Ameri, H, Ramsan, M and Savioli, L (2000) The transmission status of Bulinus on Zanzibar Island (Unguja), with implications for control of urinary schistosomiasis. Annals of Tropical Medicine and Parasitology 94, 87–94.Google Scholar
Stothard, JR, Lockyer, AE, Kabatereine, NB, Tukahebwa, EM, Kazibwe, F, Rollinson, D and Fenwick, A (2004) Schistosoma bovis in western Uganda. Journal of Helminthology 78, 281–284.Google Scholar
Stothard, JR, Campbell, SJ, Osei-Atweneboana, MY, Durant, T, Stanton, MC, Biritwum, N-K, Rollinson, D, Ombede, DRE and Tchuem-Tchuenté, L-A (2017) Towards interruption of schistosomiasis transmission in sub-Saharan Africa: developing an appropriate environmental surveillance framework to guide and to support `end game’ interventions. Infectious Diseases of Poverty 6, 10.Google Scholar
Teesdale, C and Nelson, GS (1958) Recent work on schistosomes and snails in Kenya. East African Medical Journal 35, 427–436.Google Scholar
Vreysen, MJB, Saleh, KM, Ali, MY, Abdulla, AM, Zhu, ZR, Juma, KG, Dyck, VA, Msangi, AR, Mkonyi, PM and Feldmann, HU (2000) The use of the sterile insect technique (SIT) for the eradication of the tsetse fly Glossina austeni (Diptera: Glossinidae) on the Island of Unguja (Zanzibar). Journal of Economic Entomology 93, 123–135.Google Scholar
Webster, BL, Southgate, VR and Littlewood, DTJ (2006) A revision of the interrelationships of Schistosoma including the recently described Schistosoma guineensis. International Journal for Parasitology 36, 947–955.Google Scholar
Webster, BL, Rollinson, D, Stothard, JR and Huyse, T (2010) Rapid diagnostic multiplex PCR (RD-PCR) to discriminate Schistosoma haematobium and S. bovis. Journal of Helminthology 84, 107–114.Google Scholar
Webster, BL, Emery, AM, Webster, JP, Gouvras, A, Garba, A, Diaw, O, Seye, MM, Tchuente, LAT, Simoonga, C, Mwanga, J, Lange, C, Kariuki, C, Mohammed, KA, Stothard, JR and Rollinson, D (2012) Genetic diversity within Schistosoma haematobium: DNA barcoding reveals two distinct groups. PLoS Neglected Tropical Diseases 6, e1882.Google Scholar
Webster, BL, Diaw, OT, Seye, MM, Webster, JP and Rollinson, D (2013) Introgressive hybridization of Schistosoma haematobium group species in Senegal: species barrier break down between ruminant and human schistosomes. PLoS Neglected Tropical Diseases 7, e2110.Google Scholar
Webster, BL, Rabone, M, Pennance, T, Emery, AM, Allan, F, Gouvras, A, Knopp, S, Garba, A, Hamidou, AA, Mohammed, KA, Ame, SM, Rollinson, D and Webster, JP (2015) Development of novel multiplex microsatellite polymerase chain reactions to enable high-throughput population genetic studies of Schistosoma haematobium. Parasites and Vectors 8, 432.Google Scholar
World Health Organization (2012) Accelerating work to overcome the global impact of neglected tropical diseases - A roadmap for implementation. Available at http://www.who.int/neglected_diseases/NTD_RoadMap_2012_Fullversion.pdf (accessed 25 June 2016).Google Scholar
Figure 0

Fig. 1. Map outlining shehias (smallest division of administrative regions) on Pemba Island, Zanzibar (United Republic of Tanzania) showing the location and images of two freshwater bodies in Kinyasini (Kinya2 and Kinya6) and one in Chambani (Cham10) where Schistosoma haematobium (red) and Schistosoma bovis (green) cercariae were recovered from Bulinus globosus. GPS coordinates for sites (latitude and longitude in decimal degrees): Kinya2 (−5.02033°, 39.73855°); Kinya6 (−5.03560°, 39.73850°); Cham10 (−5.35805°, 39.79182°).

Figure 1

Table 1. Showing the collection sites and genetic profiles of the Bulinus and schistosome cercariae analysed