Schistosomiasis is a snail-borne neglected tropical disease caused by parasitic blood flukes of the genus Schistosoma. Human urogenital schistosomiasis is targeted for elimination on the Zanzibar Archipelago, United Republic of Tanzania, with multiple interventions being implemented to curtail transmission of the parasite to humans on the islands since 2012. Environmental surveillance for schistosomiasis transmission by collecting intermediate host snails, checking snails for Schistosoma infection, and preserving collected snails and Schistosoma parasites offers the possibility for molecular analyses to investigate the evolutionary/genetic relationships of both snails and parasites. Schistosome transmission on Zanzibar was believed to involve a single schistosome species (Schistosoma haematobium) transmitted via a single intermediate host species (Bulinus globosus). However, our findings demonstrate the locally established presence of S. bovis, responsible for bovine intestinal schistosomiasis, and an extended intermediate host compatibility of S. haematobium with the snail B. nasutus on Pemba. Increased parasite diversity and intermediate host species compatibility may increase the transmission of Schistosoma species on Zanzibar and stretch resources for public health interventions with the need for Schistosoma species specific surveillance.

Bulinus spp. distribution inferred from Stothard et al. [23] and Pennance et al. [29,30]. Schistosoma haematobium infection distribution interpreted from Knopp et al. [18]. Digital shape files for Unguja and Pemba administrative regions were obtained from DIVA-GIS (https://www.diva-gis.org).

https://doi.org/10.1371/journal.pntd.0010585.g001

Ethics statement

Ethical approval for the collection and analyses of the snail and schistosome samples collected during the ZEST project were obtained from the Zanzibar Medical Research Ethics Committee in Zanzibar, United Republic of Tanzania (ZAMREC, reference no. ZAMREC 0003/Sept/011), the "Ethikkommission beiber Basel" (EKBB) in Basel, Switzerland (reference no. 236/11) and the Institutional Review Board of the University of Georgia in Athens, Georgia, United States of America (project no. 2012-10138-0) [18]. The ZEST study is registered with the International Standard Randomized Controlled Trial Number register (ISRCTN48837681). Additional sampling and analyses of snails in Unguja and Pemba were conducted in agreement with the Neglected Diseases Program of the Zanzibar Ministry of Health and the Public Health Laboratory-Ivo de Carneri respectively. All other snail and cercariae samples used were accessioned in the Schistosomiasis Collection at the Natural History Museum [39].

Sampling of Bulinus spp. on Pemba and Unguja islands

Following oral approval to conduct surveys by local Shehas, community leaders that locally govern each area (Shehia), human-freshwater contact sites were located from previous reports or with the help of local residents. Coordinates were taken at 112 freshwater sites (109 on Pemba and three on Unguja) across 20 shehias (Fig 1) using a Garmin GPSMAP 62sc device (Garmin, Kansas City, USA) and each water body was surveyed for the presence of intermediate host snails. Between 1 and 5 surveys were conducted at each human-freshwater contact site across Pemba and Unguja in October 2016, October 2017, February, July and November 2018 and January 2019 (Table 1).

At each site, snails were identified using shell morphology to their genera, with any non-Bulinus africanus group snails (namely B. forskalii group snails) being returned to the collection site. Snails were collected by hand predominantly from submerged vegetation and tree roots around the water’s edge that were in close proximity to access points to the water. Each site was surveyed for 15 minutes by three collectors, starting from access points and then searching as much of the accessible perimeter of the waterbody during this time. Snails morphologically identified as either B. globosus or B. nasutus (species differentiation on morphology alone not possible) were placed in collection pots and transported back to either the Public Health Laboratory-Ivo de Carneri (Chake Chake, Pemba) or the Neglected Diseases Program laboratory (Zanzibar Town, Unguja) where they were counted and housed in plastic trays with bottled water and covered by a glass lid overnight to acclimatise. The following morning (before 08:00), snails were rinsed with bottled water (to mitigate carry over of any cercariae between snails) and examined for cercarial shedding by placing individuals in wells of 12-well ELISA plates filled to approximately two thirds with bottled water and placed under indirect sunlight. Each well was checked using a dissection microscope after two hours and again eight hours after first sunlight to capture schistosomes with different shedding patterns [40]. An experienced microscopist distinguished furcocercous schistosome cercariae from other species using descriptions of Schistosoma spp. under a dissecting microscope [41]; a subset of any shed cercariae were individually captured and pipetted in 3.5 μl onto Whatman FTA cards (Whatman, Part of GE Healthcare, Florham Park, USA) for molecular characterisation. All snails were preserved in 100% ethanol for subsequent molecular characterisation as previously described (see [29]).

A targeted malacological survey was conducted in late January 2019 to collect B. nasutus snails at one site in Kangagani on Pemba previously identified as inhabited by B. nasutus (see [26]) (Fig 1). These snails were maintained in laboratory aquaria (dimensions 45x30x30cm, 40.5L, filled to approximately two thirds full with water from the collection site and equipped with an air pump for continuous aeration) at a maximum density of 100 Bulinus individuals per aquarium. Snails were re-checked for shedding of schistosome cercariae three weeks later in an effort to capture any infections that may have been pre-patent during the initial screen. Aquarium water was replaced twice a week using water from the site of collection (Kangagani). Water was only used in aquaria after storage for at least 48 hours in transparent 15L water containers, allowing for any sediment to settle and eliminate the risk of introducing live schistosome eggs, miracidia or cercariae into the aquaria. Snails were fed on dried lettuce when all lettuce in the tank had been eaten. Dead snails were removed from the aquaria daily.

Archived samples included in the analysis

Bulinus samples, from Unguja and Pemba, accessioned within the Schistosomiasis Collection at the Natural History Museum (SCAN) [39] were included in the study providing material from areas not covered in the malacological surveys described above. Samples were only included if associated geographical information was available. Six snails were included, five from Unguja (Jendele, Miwani, Kinyasini, Mtopepo, Chaani) and one from Pemba (Wingwi) as presented in Table 1. Two of these snails from Miwani and Kinyasini on Unguja were recorded as patent with S. haematobium when they were collected, as this was later confirmed by cercarial molecular analysis as described below. The remaining four Bulinus snails from the SCAN collection were negative for patent Schistosoma infections (S1 Table). One of the infected B. globosus identified in the SCAN repository (MCF389B0F0286, S1 Table) could not be associated with its emerging S. haematobium cercariae as it was preserved together with two other B. globosus also infected with S. haematobium from the same site.

Bulinus spp. molecular characterisation

Since a significant degree of morphological overlap exists between B. globosus and B. nasutus, a molecular marker was used to unequivocally identify a subset of the Bulinus snails collected (n = 504), and those taken from archived specimens (n = 6). The shell was removed by crushing and the use of sterile forceps from the preserved sample and gDNA from whole snail tissue was extracted using either the Qiagen BioSprint 96 DNA Blood Kit following manufacturer’s instructions (Qiagen, Manchester, UK) or the Qiagen DNeasy Blood & Tissue Kit modified protocol (Qiagen, Manchester, UK) using double volumes of the lysis buffers [29]. Since not all snails could be identified using molecular characterisation due to cost and time constraints, a minimum of three non-patent snails per site per malacological survey (except for those collected on Pemba during October 2016 and from Chambani and Uwandani in 2017/2018) were randomly selected for identification. Molecular characterisation was performed for all snails with patent Schistosoma infections and snails retrieved from SCAN.

DNA was extracted from a total of 510 Bulinus spp. snails, from Unguja (11 snails collected from 8 sites) and Pemba (499 snails collected from 63 sites). A 623 bp partial region of mitochondrial cox1 DNA was amplified and Sanger sequenced following previously described methods (see [29]). Sanger sequence data were edited, manually trimmed to 463–621 bp and aligned in Sequencher v5.4.6 (GeneCodes Corp., Michigan, USA) before being collapsed into cox1 haplotype groups. Species identification were confirmed by alignment and phylogenetic analysis (see below) of Bulinus cox1 haplotypes, as presented in S2 Table, with reference data for B. globosus and B. nasutus [42].

Schistosoma spp. cercariae molecular characterisation

From each infected snail either two or six Schistosoma cercariae were processed individually for molecular identification. Six cercariae were individually processed from each infected snail collected between 2016 and 2019 (n = 20 snails) and two cercariae for snails collected as part of the ZEST study (n = 69 snails) [16–18] and made available via SCAN (S3 Table). Following elution of parasite DNA from Whatman FTA cards [43], Schistosoma species identification was confirmed by mito-nuclear genetic profiling targeting the partial mitochondrial cox1 region (956 bp) and the complete nuclear ITS (1–5.8S-2) rDNA region (967 bp) from each individual cercariae as described in [27,28]. Both mitochondrial and nuclear DNA were analysed for species identification and to identify any hybridisation [44]. The cox1 data were also used for genetic diversity and phylogenetic analyses. The sequence data were manually edited and trimmed to 750 bp for cox1, and 880bp for ITS, using Sequencher v5.4.6 (GeneCodes Corp., Michingan, USA). The cox1 species identity was confirmed by comparison to nucleotide sequences using NCBI-BLAST [45] and the ITS species ID was confirmed by comparison to reference data as described [28,29]. Species identification of each cercariae was confirmed by concordance between the cox1 and ITS genetic profiles. Cercariae of identical cox1 sequences were collapsed into cox1 haplotype groups for further phylogenetic analysis (S4 Table). ITS data were not used for phylogenetic analysis since no intra-species diversity was observed.

Phylogenetic cox1 analysis of Bulinus spp. and Schistosoma spp

The Bulinus haplotype data were imported into Geneious v11.1.4 [46] for phylogenetic analysis together with reference data for B. nasutus and B. globosus collected previously from East Africa (Zanzibar, Tanzania, Kenya, Uganda, Mafia Island; [42] and an outgroup of Biomphalaria glabrata available from GenBank (Accession: NC005439) [47]. Haplotype alignments were performed using ClustalW v2.1 [48] executed in PAUP* [49] and then an appropriate evolutionary nucleotide substitution model (HKY + I + G; -lnl 2020.2144, AIC 4052.4287) was selected in MrModelTest v2.4 [50] using the Akaike Information Criterion. Bayesian inference was performed using MrBayes v3.2.7a [51]. The burn-in was set at 3.5 million generations for consistency after confirming that the average standard deviation of split frequencies (ASDOSF) reported from MrBayes output was at least <0.01 by this point. Clades were considered to have high nodal support if Bayesian inference posterior probability was ≥0.95; tree nodes with <0.95 were collapsed in SumTrees v4.4.0 [52].

Schistosoma cercarial haplotype phylogenetic analyses were performed as above with S. curassoni (AY157210; [53]) as the outgroup. Analyses also included published S. haematobium haplotypes from Zanzibar (GU257334 –GU257360; [28]), and S. bovis from Pemba (MH014042 & MH014043; [29] and OK484569 [54]), mainland Tanzania (AY157212; [53]) and Cameroon (MH647141; [55]), with the alignment trimmed to 750 bp to maintain uniform ends. A Templeton, Crandall and Sing’s (TCS) haplotype network analysis was also conducted using PopART [56,57] using the same sequence alignment.

Statistical analysis

Within the molecular analyses S. haematobium cercariae were assigned to either the Group 1 or Group 2 cox1 haplotype group [27], and Chi-squared tests were performed in R v.4.0.0 [58] to investigate any differences in the abundance of the two groups in relation to their snail host species and geographical distribution.

Spatial distribution of Bulinus and Schistosoma species

Bulinus and Schistosoma spp. distribution data were visualised using QGIS v3.0.1 Girona (http://qgis.osgeo.org) and mapped for each site. Digital shape files for Unguja and Pemba administrative regions were obtained from DIVA-GIS (https://www.diva-gis.org).

Patent schistosome infections of Bulinus

Over the six malacological surveys conducted on Pemba between 2016 and 2019, and the one survey conducted in Unguja, a total of 11,116 Bulinus spp. were collected from 65 of the 112 sites. From the subset of the snails that were identified by molecular analysis from each of these sites (n = 504), and those identified from the SCAN repository (n = 6), the majority were B. globosus (n = 433 snails), the remainder were B. nasutus (n = 77 snails). The other 10,606 Bulinus collected remain only morphologically identified as being within the Bulinus africanus species group (Bulinus genus), since morphological differentiation between B. globosus and B. nasutus is not possible. Of the 11,116 Bulinus, 0.2% (n = 20) shed Schistosoma spp. cercariae. These were collected from eight sites: four sites in Kinyasini (n = 16 snails), two in Kizimbani (n = 2 snails), one in Chambani (n = 1 snail) and one in Kangagani (n = 1 snail) as presented in Table 1. The infected snail from Kangagani was collected during the targeted malacological B. nasutus survey, in which 198 individual Bulinus spp. specimens were collected and at the time of collection were not shedding. Although having no observed patent schistosome infections during the first round of shedding, a single B. nasutus was found shedding Schistosoma cercariae 21 days later. No follow up shedding was attempted on the other snails collected in Pemba as they were preserved within 48 hours of collection.

For the malacological surveys conducted at three sites in two shehias (Mtende and Kizimbani Dimbani) on Unguja, only six B. nasutus (Table 1) were collected in total, of which none shed Schistosoma cercariae within 24 hours of their collection. No further shedding attempts were made on these snails which were preserved after the first round of shedding.

Bulinus spp. genetic diversity and distribution

From the subset of Bulinus spp. successfully sequenced, 433 out of 510 from 3 sites across Unguja and 49 sites across Pemba were B. globosus (S1 Table). The remaining 77 snails were identified as B. nasutus collected from 15 sites in Pemba and 5 in Unguja (S1 Table). Bulinus nasutus was molecularly identified only from freshwater bodies near the east coast of Pemba and the southern districts up to the central west areas of Unguja (Fig 2). Where >1 snail was molecularly identified per site, the snails were identified as either B. globosus or B. nasutus coming from the same site, except for from one waterbody on Pemba (Puj11, Fig 2B), where both species co-occurred in the same seasonal pond (S1 Table). Based on the subset of molecularly identified Bulinus, no other mixed B. globosus and B. nasutus populations were observed, however this would have to be fully confirmed with further genetic analysis of a large set of snails from each water body.

All 11 B. globosus and 9 out of 10 B. nasutus cox1 haplotypes were unique to either Unguja or Pemba, with one exception being B. nasutus haplotype 3, as shown in S2 Table, that was detected on both islands. A single clade of B. nasutus specimens from both Unguja and Pemba was observed, whereas the B. globosus isolates from each island fell into two distinct clades (Fig 3). All B. globosus from Pemba fell into one clade containing two sister groups, with those previously identified from Pemba [42]. The three haplotypes from Unguja fell into a another clade containing two sister groups of B. globosus from Eastern Kenya and those previously identified from Unguja (Fig 3) [42].

Schistosoma spp. cercariae identification

From the subset of cercariae identified from 89 of the infected Bulinus collected from Zanzibar between 2016 and 2019 (n = 20) and identified during the ZEST study (n = 69), 82 were shedding S. haematobium with 18 different haplotypes and seven were shedding S. bovis with two haplotypes (S3 and S4 Tables). Both Group 1 and 2 haplotypes of S. haematobium representing mainland African and Indian Ocean islands respectively were identified [27,28] (Figs 4 and S1). Including Schistosoma coinfections, of which there were seven determined by multiple cox1 haplotypes presented in S5 Table, the proportion of snails shedding S. haematobium Group 1 cercariae (n = 41) was similar to those shedding Group 2 cercariae (n = 45). However, the majority of Group 1 S. haematobium infections occurred in Unguja (n = 35), with significantly fewer (n = 6) from Pemba (χ² = 10.0, df = 1, P< 0.01). In contrast, Group 2 S. haematobium cercariae were distributed evenly across the islands (Unguja n = 23 and Pemba n = 22). Most (n = 13 of 18) S. haematobium cox1 haplotypes were unique to either Pemba or Unguja, but five were present across both islands.

Seven snails as presented in S5 Table were confirmed as shedding Schistosoma cercariae with multiple cox1 haplotypes of either S. haematobium (n = 6) or S. bovis (n = 1), indicating they had been infected by multiple miracidia. Furthermore, five of the six S. haematobium infected snails simultaneously shed both Group 1 and Group 2 S. haematobium haplotypes, whilst only one snail was identified shedding two haplotypes of Group 2.

Comparison of mitochondrial cox1 and nuclear ITS profiles from S. haematobium cercariae showed no evidence of hybridisation between S. haematobium and S. bovis. No intraspecies variation in the ITS profiles were observed, with 100% match to reference data [27].

Bulinus observed shedding Schistosoma haematobium and S. bovis

Of the 20 infected snails collected in Pemba listed in S6 Table, 12 were identified as B. globosus infected with S. haematobium and seven as B. globosus infected with S. bovis. The remaining infected Bulinus collected during the targeted survey in Kangagani was identified as B. nasutus (Haplotype 3 in S2 Table), matching that previously reported on Pemba (GenBank Accession: AM921812, see [42]). The two cercariae identified from this snail were S. haematobium of a single cox1 haplotype (Sh3). This S. haematobium haplotype has been previously identified as ‘Group 1’ (GenBank Accession: GU257343, see [28]) representing those from mainland Africa and Zanzibar (Figs 4 and S1 Fig). The infected B. globosus from Unguja from the SCAN repository was shedding ‘Group 2’ cercariae as presented in S6 Table.

Distribution and diversity of Bulinus globosus and B. nasutus

Co-occurrence of B. globosus and B. nasutus was observed at just one site on Pemba (Puj11), generally supporting previous observations that species distribution in freshwater bodies is dictated by species specific ecological factors (such as water conductivity) determined by the geological zones of Zanzibar (see [61,62]). However, it is noteworthy that during the current study it was only feasible to identify a proportion (n = 510 of 11,116) of the B. globosus and B. nasutus accurately through partial cox1 sequencing. Therefore, it is possible that other sites containing both B. globosus and B. nasutus may be identified on the Zanzibar Archipelago in future species identification. The development of a cheap, easily interpreted, rapid diagnostic assay to distinguish between B. globosus and B. nasutus, such as is available for the differentiation of S. haematobium and S. bovis [63], would provide a much needed solution for identifying large numbers of field collected specimens.

The 11 B. globosus cox1 haplotypes identified from the snails collected across Zanzibar fell into two distinct clades representing Unguja and Pemba taxa [24]. The B. globosus from Unguja were more closely related to those previously identified from East Kenya [42], whilst those from Pemba form an independent group distinct from the other East African isolates, suggesting independent origins. This agrees with our current understanding of Zanzibar’s geological formation, whereby Pemba island separated from mainland Africa earlier, during at least the early Pliocene compared to Unguja island during the Pleistocene [61,64]. In contrast, there was no phylogenetic distinction between the B. nasutus from Unguja, Pemba and mainland East Africa, with samples from each region all forming a single clade of multiple haplotypes. As recorded previously, B. globosus is more genetically diverse than B. nasutus (see [65]). Identical haplotypes of B. nasutus were also present on Unguja and Pemba. At this stage of investigation, we postulate that since B. nasutus infected with S. haematobium observed here in Pemba matched haplotypes of B. nasutus in the south of Unguja (see below), it might serve as an intermediate host of urogenital schistosomiasis across the Archipelago, as is the case in Kenya [31,66] and Tanzania [32–37].

A ‘new’ intermediate host of Schistosoma haematobium on Pemba: Bulinus nasutus

The finding of B. nasutus in one locality naturally infected with S. haematobium on Pemba contradicts previous results suggesting that this snail species was not involved with the transmission of urogenital schistosomiasis on Zanzibar [22–24]. Indeed, pre-patent Schistosoma spp. infections previously observed in B. nasutus from Pemba might have been S. haematobium (see [26]) but infections did not appear to result in cercarial production [25].

Schistosoma haematobium cercariae shed from the intermediate snail hosts, analysed here, fall into the two known S. haematobium haplotype groups (1 and 2) previously identified from miracidia collected from infected humans [27,28]. The cox1 haplotypes of cercariae shed from B. nasutus were identified as Group 1, a group predominantly associated with African mainland S. haematobium populations. Possibly only Group 1 S. haematobium is compatible with Pemba B. nasutus, since this same snail species acts as an intermediate host in East Africa [31–37], however more samples would be needed to test this hypothesis. Schistosoma strain specific interactions/compatibilities with intermediate host snails based on differential immune responses, have been well studied and established in strains of S. mansoni and B. glabrata (as summarised in [67]). However, relatively little is understood regarding co-evolution of host/parasite compatibility between S. haematobium group strains and Bulinus species, save some studies investigating geographical isolates (see [68–72]). Such strain dependency or local adaptation reflects the patchy compatibility of Bulinus snail hosts of Schistosoma generally [73], discussed in a previous study in Tanzania, where experimental infections of B. nasutus using a local strain of S. haematobium that usually infects B. globosus were unsuccessful [74]. As demonstrated again here, B. globosus remains the primary host of S. haematobium on Zanzibar, but endemic B. nasutus may also play a minor role in transmission involving specific S. haematobium strains, complicating future monitoring. This hypothesis also provides some explanation to how ‘intermittent’ and/or ‘unstable’ transmission was historically maintained in areas of Zanzibar (such as south Unguja) where only B. nasutus is present currently [7,9,10,75,76].

Study limitations and future work

Several limitations are apparent in the current study. The time and costs required to generate cox1 sequence data limited the number of snail intermediate hosts that could be identified by this means. Development and testing of a rapid diagnostic assay, or alternative, to provide rapid high throughput identification of snails would greatly support future studies. Additionally, few Bulinus specimens were available for analysis from Unguja, so further malacological surveys with molecular sub-sampling here would be beneficial to confirm snail species distributions across the island. Also, since the majority of infected Bulinus spp. collected during the ZEST studies were not accessioned with their associated Schistosoma spp., complete inferences on snail-Schistosoma relationships were not possible. Finally, although cercariae identification was used here to identify Schistosoma species, it would have been of interest to identify a greater number of cercariae per snail infection, as this may have significantly increased the number of coinfections observed from these Bulinus spp. and allow for the potentially immune modulated interactions of Group 1 and Group 2 S. haematobium coinfections, observed in five snails here, to be explored further. In a future study, it would also be of interest to use PCR based methods to identify pre-patent S. haematobium and S. bovis infections in snails to further assess the total number of snails that have been exposed to schistosomes but are not currently contributing to transmission [77].

Implications for future monitoring of schistosomiasis on Zanzibar

The findings discussed here provide implications for future control and elimination efforts of urogenital schistosomiasis on Zanzibar [18,19]. First, it is suggested here that the Ministry of Health, Social Welfare, Elderly, Gender and Children Zanzibar should conduct periodic surveys in areas associated with B. nasutus distribution on Unguja and Pemba. These surveys should include both malacological collections and combined human urine collections with questionnaires (including questions on freshwater usage locally and elsewhere in Zanzibar), followed by targeted treatment of infected individuals and focal snail control, to reduce any ongoing transmission. Second, a monitoring system to check the identity of schistosome cercariae shed from Bulinus spp. snails from Zanzibar to differentiate bovine and human schistosomiasis would enable accurate mapping of both schistosome species, appropriate targeting of urogenital schistosomiasis control interventions, and also monitor any potential hybridization events between S. haematobium and S. bovis that may be occurring in cattle or humans [60].

Nearly a century has passed since the first reports of widespread human urogenital schistosomiasis on Zanzibar, during which time there have been great achievements towards urogenital schistosomiasis elimination. As the battle to eliminate schistosomiasis from the Zanzibar Archipelago continues, our findings emphasize the need to carefully plan future surveillance strategies of transmission on the islands, taking into consideration the presence of bovine Schistosoma species and the capacity for an expanded intermediate host, and therefore geographical, range of S. haematobium.

S1 Table. Associated Bulinus spp. specimen data.

Detailed information on Bulinus spp. specimens used in current study, including species identification determined through molecular analysis (cox1), collection site name including latitude and longitude, schistosome species patency and cox1 haplotype.

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S2 Table. Bulinus spp. cox1 haplotypes observed from Pemba and Unguja, and associated Schistosoma spp. infecting each snail haplotype.

^a Schistosoma haplotype group indicates whether cercariae were identified as mainland Africa (1) or Indian Ocean Island (2) haplotypes (as described in Webster et al. [27]). ^b It was not possible to associate this snail haplotype with its S. haematobium cercariae cox1 haplotype(s) as it was preserved with two other S. haematobium infected B. globosus.

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S3 Table. Associated Schistosoma spp. specimen data.

Detailed information on Schistosoma spp. specimens used in current study, including species identification determined through molecular analysis (cox1 and ITS1-5.8S-ITS2), collection site name including latitude and longitude and Schistosoma cox1 haplotype.

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S4 Table. Schistosoma cox1 haplotypes identified from cercariae from Unguja and Pemba.

^a Schistosoma haplotype group indicates whether cercariae were identified as mainland Africa (1) or Indian Ocean Island (2) cox1 haplotypes (as described in Webster et al. [27]). ^b Island; U = Unguja, P = Pemba.

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S5 Table. Trematode coinfections of Bulinus spp. from Unguja and Pemba.

^a Schistosoma haplotype group indicates whether cercariae were identified as mainland Africa (1) or Indian Ocean Island (2) cox1 haplotypes (as described in Webster et al. [27]). ^b Coinfection indicates the Schistosoma cox1 haplotype group (as described in [27]) infection profile of each Bulinus spp. Sh1 = Schistosoma haematobium mainland African haplotype group 1, Sh2 = S. haematobium Indian Ocean haplotype group 2, Sb1 = S. bovis haplotype 1, Sb2 = S. bovis haplotype 2.

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S6 Table. Bulinus spp. infected with Schistosoma spp. identified from Unguja and Pemba.

^a Schistosoma haplotype group indicates whether cercariae were identified as mainland Africa (1) or Indian Ocean Island (2) haplotypes (as described in Webster et al. [27]).

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S1 Fig. Bayesian inference of the partial mitochondrial cox1 haplotype dataset of Schistosoma haematobium and S. bovis collected from Unguja and Pemba.

Phylogenetic tree constructed using Bayesian inference in MrBayes v3.2.7a [51] under the HKY + I model (-lnL = 1809.8890, AIC 3629.7781, ASDOSF < 0.01 at 1,791,000 generations). Branches with <0.95 posterior probability are collapsed. The branch length scale bar indicates the number of substitutions per site. Text in red indicates Schistosoma haplotypes generated in the current study.

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