Molecular Footprint of Parasite Co-introduction with Nile Tilapia in the Congo Basin

doi:10.21203/rs.3.rs-995291/v1

Download PDF

Research Article

Molecular Footprint of Parasite Co-introduction with Nile Tilapia in the Congo Basin

https://doi.org/10.21203/rs.3.rs-995291/v1

This work is licensed under a CC BY 4.0 License

Version 1

posted

You are reading this latest preprint version

Nile tilapia, one of the most popular aquaculture species worldwide, has been introduced into the Congo Basin several times. In previous morphological studies, we showed that some of the monogenean gill parasites were co-introduced with Nile tilapia and some spilled over to native Congolese cichlids. In this study, we investigated the co-introduced monogeneans of Nile tilapia genetically from three major parts of the Congo Basin; Upper, Middle and Lower Congo. We generated sequences of Congolese native and introduced monogeneans from native and introduced tilapias and evaluate their position in a phylogeny. Additionally, we generated sequences of the same species of monogeneans co-introduced with Nile tilapia in Madagascar and of a native population of Nile tilapia from Burundi. Our results confirm the co-introductions in the Congo. We found that co-introduced parasites are less genetically diverse than native ones, and that there was no geographical pattern between introduced populations. Furthermore, our COI haplotype networks suggest multiple introduction events of Nile tilapia into the Congo Basin. Additionally, we tested the barcoding gap and the performance of mitochondrial COI and nuclear ribosomal ITS-1, 28S and 18S markers. We found a significant intra/interspecific barcoding gap of 15% for COI, but none for the other markers. Our molecular results reveal that Cichlidogyrus halli, C. papernastrema, C. tiberianus, C. cirratus and C. zambezensis are in need of taxonomic revision.

Marine and Freshwater Ecology

Tilapia

Monogenea

Co-introduction

Phylogeny

Barcoding

Marker Performance

mutations

Human-mediated species translocations are ubiquitous and form one of the major challenges to global biodiversity today (Pimentel 2001 et al., Tollefson 2019). As a result of invasive species, the abundance of native plants, insects and other animals has fallen by an estimated 20% since 1900 (Tollefson 2019). Of these translocated species, tilapias are among the most widely introduced aquaculture species and are now found in over 140 countries (Deines et al. 2016). Cultured tilapia comprises species of African cichlids mainly from Oreochromis Günther 1889, Coptodon Gervais 1848, Tilapia Smith 1840 and Sarotherodon Rüppel 1852 and is estimated to produce over 5 million tonnes yearly globally (Deines et al. 2016, FAO 2017). Of these, Nile tilapia, Oreochromis niloticus (Linneaus, 1758), is the most popular and makes up over 75% of the cultured tilapias in Sub-Saharan Africa (FAO 2017). Although Sub-Saharan Africa houses many native tilapias, others were introduced anyway for aquaculture purposes, such as Nile tilapia, which started being cultured after the Second World War in the Democratic Republic of the Congo (Micha 2013). Production spread rapidly throughout the country until 1960 after which production almost halted (Micha 2013). In 1964, the first feral Nile tilapia population was found in the North-East of the basin (Thys van den Audenaerde 1964). From 1996 fish production increased steadily again to modern levels (Toguyeni 2004).

Negative effects of Nile tilapia introductions are reported worldwide, including competition with and subsequent decline of native fish species (Canonico et al. 2005, Shipton et al. 2008, Šimková et al. 2019), hybridization with other tilapias (Trewavas 1983, Nyingi and Agnèse 2007, Firmat et al. 2013, Deines et al. 2014), stimulation of phytoplankton growth (Starling et al. 2002), predation on eggs of aquatic organisms (Alcaraz et al. 2015) and co-introduction of parasites (Soler-Jimenez et al. 2017, Jiménez-García et al. 2001, Šimková et al. 2019, Zhang et al. 2019, Jorissen et al. 2020). One of the dangers of co-introducing parasites is that these parasites might switch hosts and successfully infect native species (‘spillover’, Goedknegt et al. 2016), further increasing the extinction risk of native species. For example, co-introduced blood flukes in Spain spilled over from invasive turtles to native European pond turtles. The native pond turtles experienced increased mortality rates and loss of mobility in tail and limbs because of these newly acquired blood flukes (Iglesias et al. 2015).

Our previous study identified parasite co-introductions with Nile tilapia and spillover to native cichlids in the Congo Basin (Jorissen et al. 2020). The respective parasites belong to Cichlidogyrus Paperna 1960 and Scutogyrus Pariselle and Euzet 1995. They are monogenean flatworms (Dactylogyridea) that mainly infect the gills of African mainland cichlids (Pariselle and Euzet 2009), where they feed on mucus, skin and possibly blood of the hosts (Gonçalves et al. 2009). Currently, 130 valid species of Cichlidogyrus and 7 of Scutogyrus have been described (WoRMS 2021) and DNA sequences of 31 and 3, respectively, are available on Genbank (NCBI 2021). These sequences are generally limited to partial fragments of 28S or 18S+ITS1 (internal transcribed spacer 1 region) rDNA. The genes coding for 18S and 28S are the most conservative markers currently available (Vanhove et al. 2013) and are used for phylogenetic reconstructions (Pouyaud et al. 2006; Wu et al. 2007; Mendlová et al. 2012; Mendlová and Šimková 2014). However, there is a lack of molecular markers for flatworms (Littlewood 2008, Moszczynska et al. 2009, Vanhove et al. 2013) and a lack of sequenced species. For example, only for five species within Cichlidogyrus and Scutogyrus partial or complete sequences of the more variable mitochondrial cytochrome c oxidase subunit I (COI) gene are available (Genbank accession numbers: KT037411.1, MG288510.1, MG970255.1, JQ038226.1, MG970257.1). The first goal of this study is to aid in filling this knowledge gap by adding sequences of Congolese native and introduced species and to evaluate the position of these previously unsequenced species in phylogenetic analyses. Additionally, we will discuss marker performance for mitochondrial COI, ITS-1 rDNA, 18S rDNA and 28S rDNA.

Secondly, because of this molecular knowledge gap, these monogeneans are predominantly diagnosed based on the morphology of their sclerotized hard parts from the reproductive organs and the attachment organ (haptor). However, out of the five morphospecies of Cichlidogyrus and one of Scutogyrus that were co-introduced with Nile tilapia into the Congo Basin (Jorissen et al. 2020), we suspect that three are in need of taxonomic revision: Cichlidogyrus halli (Price and Kirk 1967), C. cirratus Paperna 1964 and C. tilapiae Paperna 1960. A subspecies of C. halli has been described in the past (Paperna 1979), but was later revoked (Pariselle and Euzet 2009). Additionally, we discovered a morphotype of C. halli native to Bangweulu-Mweru, in the Upper Congo Basin, infecting the native Oreochromis mweruensis Trewavas 1983 (Jorissen et al. 2018a). However, genetic information on this morphotype is missing and would be helpful to decide on its taxonomic status. Pouyaud et al. (2006) found three molecular variants of C. halli based on 18S and 28S rDNA, but the low molecular variation did not indicate the need of taxonomical revision at the time. The morphological (and molecular) variation within Cichlidogyrus cirratus could also be underestimated as this species was recently reported in new areas and on new hosts (Zhang et al. 2019, Jorissen et al. 2020 and unpublished data). Finally, Pouyaud et al. (2006) suggested that C. tilapiae might constitute a species complex of morphologically closely resembling taxa. They found larger intraspecific distances based on 18S and 28S rDNA within C. tilapiae than between some specimens of C. tilapiae and C. cubitus Dossou 1982. Our second goal is therefore to evaluate the species status of these three species through molecular data.

Thirdly, genotyping the co-introduced monogeneans of Nile tilapia could offer a more precise picture of the invasive history of the fish. Indeed, parasites can, due to their faster generation time compared to their host, shed more light on the evolutionary and geographical history of their hosts, as predicted by the magnifying glass concept (Nieberding and Olivieri 2007). Moreover, genotyping can reveal biological phenomena like hybridization, as was the case for catfish parasites, which in turn implied historical contact between the catfish host species that currently live in allopatry (Barson et al. 2010). Also the lack of genetic variation can be informative on transmission routes (Hayward et al., 2003). Gyrodactylus anguillae Ergens, 1960 collected from eel populations from three different continents share identical ribosomal DNA sequences, as the result from recent live eel trade (Hayward et al., 2003).

In this study, we focus on the Congo Basin because it is the largest African basin where Nile tilapia was introduced and because the country has a historical tradition of tilapia culture (Welcomme 1988, Micha 2013). Additionally, molecular data from monogenean parasites within the Congo Basin, apart from Lake Tanganyika, is largely lacking. Our expeditions took place in Upper Congo (Bangweulu-Mweru) in the southeast of the basin, Middle Congo around Kisangani (DRC) and Lower Congo downstream of Boma and the tributaries of the Congo around Mbanza-Ngungu (DRC). The boundaries between these three parts are Pool Malebo around Kinshasha and Boyoma Falls upstream from Kisangani (Fig. 1; Alter et al. 2015). We include parasite populations from introduced Nile tilapia from Madagascar because Nile tilapia is well-established there, and mostly the same monogenean species have been co-introduced there as in the Congo Basin (Šimková et al. 2019). Lastly, we sample a native population of Nile tilapia from a pool next to Lake Tanganyika (Burundi) because it is geographically the closest native population of Nile tilapia to the DRC. According to the concept of isolation by distance (IBD), the genetic similarity between populations should decrease with increasing geographic distance (Avise 1994, Poulin and Krasnov 2010). However, introduction events can blur this signal and can also lead to lower genetic diversity in introduced populations compared to their source populations. This can lead to potential founder effects (Mayr 1942, Avise 1994). Therefore, we expect a low diversity in the introduced parasite populations from Congo and Madagascar, compared to the native parasite population from Burundi. Also, if Nile tilapia in Congo and Madagascar originate from a common source of introduction, they will share parasite haplotypes and no signal of IBD will be found.

Data collection

A total of 146 specimens of parasites belonging to 16 described and one undescribed species, of which 15 of Cichlidogyrus and two of Scutogyrus were collected from 7 host species (Table 1). Samples were collected during three field expeditions to the Democratic Republic of the Congo: Kisangani in May-June 2014, Bangweulu-Mweru in August-September 2014 (Jorissen et al. 2018a), Lower Congo in June 2015 (Jorissen et al. 2018b); one to Burundi in September 2013 (Rahmouni et al. 2017a); and one to Madagascar in April 2016 (Šimková et al. 2019). Samples from Lake Kariba in Zimbabwe were caught in September 2016. Fish were collected in the wild, from aquaculture stations or bought at local fish markets and killed with an overdose of MS222. Specimens and sample localities are listed in Addendum 1 and shown in Fig. 1. The gill arches of the right side were dissected in the field and stored in pure ethanol. The left side of the fish was left intact for ichthyological research. Parasites were isolated in the lab using an entomological needle and a Wild M5 stereomicroscope (Heerbrugg, Switzerland). Parasites were cut in half with a scalpel; the anterior body part was fixed in Hoyer’s medium and sealed on a slide with glyceel (Bates 1997) for morphological identification. The pictures of C. cf. halli ‘Burundi’ were taken with a Zeiss Axio Imager Z1 microscope at a magnification of 100 x (oil immersion, 10 x ocular) under differential interference contrast, with an AxioCamMR3 camera and AxioVision v.4.2.8 software. The posterior body part of the parasite was placed in an Eppendorf tube filled with 180µl of T1 buffer, Nucleospin Kit, Macherey Nagel, for DNA extraction and stored at -21°C if extraction was not carried out immediately (see Addendum 1 for the collection numbers of fish hosts, parasite vouchers and GenBank accession numbers of the parasite DNA sequences generated in this study). Parasite slides and fish hosts were both stored in the collections of the Royal Museum for Central Africa, Tervuren, Belgium (RMCA). Fish from the Madagascar expedition and Upper Congo (Bangweulu-Mweru) were stored under collection 2016-15-P. Fish from Middle and Lower Congo were stored under collections 2014-XX-P and 2015-30-P, respectively. Parasite slides are found under RMCA_VERMES_XXX (see Addendum 1).

Molecular analyses

DNA was extracted using the Nucleospin kit (Macherey-Nagel). In the final step 60µl of elution buffer was added instead of 100µl. DNA extracts were stored at -18°C. We used the primers listed in Table 2 for the amplification of fragments of the mitochondrial cytochrome c oxidase subunit 1 (COI, 314bp), complete ITS-1+partial18S (702bp) and partial 28S (653bp). For COI, if initial amplification was unsuccessful we tried again with nested primers, resulting in the 314bp mentioned above. Protocols of DNA amplification are included in Table 3. PCR products were run on 1.5% agarose gels stained with GelRed (Biotum Inc.). Successful amplifications were purified with EXOSAP-IT (Thermo Fisher) in a 5/2 (product/EXOSAP) ratio and incubated at 37 °C for 4 minutes followed by 80 °C for 1 minute. A total of 3.2 µM forward primer was added to the purified PCR products in a 7/5 (purified PCR product/3.2 µM primer) ratio and sent to Macrogen Europe under the EZ-seq service for single direction Sanger sequencing.

Phylogenetic analyses

A total of 38 species were included to build the phylogeny from ribosomal markers, of which 12 we present the first sequences. All sequences are submitted to Genebank and accession numbers are available upon acceptance. Our tree was rooted on Cichlidogyrus pouyaudi Pariselle and Euzet, 1994 as previous phylogenetic research found that it is the most basal taxon of the group (Pouyaud et al. 2006; Wu et al. 2007; Mendlová et al. 2010; Mendlová and Šimková 2014). All sequence chromatograms were visually inspected for sequencing errors and blasted individually on the NCBI website (http://www.ncbi.nlm.nih.gov). The resulting sequences were aligned with MUSCLE (Edgar 2004a, 2004b) under default settings, edited in MEGA 7.0.18 (Kumar et al. 2016) and cleaned-up with Gblocks 0.91b under default parameters (Castresana 2000, Talavera & Castresana 2007). The 28S and 18S+ITS-1 sequences were concatenated using SequenceMatrix (Vaidya et al. 2011). To test whether a concatenation was possible we performed a partition-homogeneity test in PAUPUP 1.0.3.1 (Swofford 2003). From some specimens either 28S or 18S+ITS-1 were successfully amplified and these sequences were aligned with the concatenated sequences to include the maximum number of species and specimens. A total of 30 reference sequences of congeners were obtained from GenBank (see Table 4). JModelTest2 was used to determine the most appropriate model of nucleotide evolution (Darriba et al. 2012) with default parameters, employing the Akaike Information Criterium (AIC). The GTR+G+I model was found as the optimal model for the concatenated dataset with gamma being 0.65 and the proportion of invariant sites 0.29. For the COI dataset TIM1+G+I was the optimal model, but we used GTR+G+I as it was the most similar model being available in MrBayes and RAxMLHPC2. Gamma was 0.2 and the proportion of invariant sites 0.2. Maximum Likelihood (ML) analyses were carried out with RAxMLHPC2 (Stamatakis 2014) via the CIPRES webserver with the GTR+G+I model and 1000 iterations. Branch support was estimated by bootstrapping with 1000 replicates. The Bayesian inference of phylogeny (BI) was performed in MrBayes 3.2.7 (Ronquist & Huelsenbeck 2003) with 4.000.000 generations sampled every 1000 generations and four chains (three cold and one hot). One fourth of the topologies were discarded as burn-in as the standard deviation of split frequencies was below 0.01 for both the concatenated dataset and for the COI dataset. Figtree 1.4.4. (Rambaut 2018) and TreeGraph 2 (Stöver and Muller 2010) were used to visualize and edit the trees. Additionally, we identified phylospecies and evolutionary species with a coalescent tree-based Poisson tree process model and its Bayesian implementation (bPTP, Zhang et al. 2013) on the web server (http://species.h-its.org/ptp/) under default parameters and 0.2 burnin. Trees will be available on TreeBASE upon acceptance.

Haplotype networks and distance matrices

Distance matrices were calculated in PAUPUP 1.0.3.1 (Swofford 2003) using the GTR+G+I model. A median-joining haplotype network (Bandelt et al. 1999) was constructed with PopART 1.7 (Leigh & Bryant 2015) with ε = 0 separately for C. sclerosus Paperna and Thurston, 1969, C. thurstonae and C. tilapiae based on 9, 20 and 17 COI sequences, respectively. For C. halli the genetic distance between specimens was too large for a median-joining haplotype network, see results. COI-alignments were translated to amino acid sequences with MEGA 7.0.18 for information on (non-)synonymous mutations and to check for nuclear mitochondrial DNA copies (numts). To estimate a barcoding gap and species partitioning we used ASAP: Assemble Species by Automatic Partitioning (Puillandre et al. 2020) calculated with the K2P-distance model (Kimura 1980). Nucleotide diversity and polymorphic sites were calculated with DnaSP V.6.12.03 (Rozas et al. 2017). Distance matrices will be available on TreeBASE upon acceptance.

The sequencing of the partial COI-gene resulted in 67 sequences of 314 bp of 13 species after trimming and clean-up in Gblocks; none showed stop codons. We opted for the nested fragment to include the most specimens in the analyses. The partition homogeneity test allowed the concatenation of 18S, ITS-1 and 28S fragments with a p-value of 0.95 (anything above 0.05 was sufficient for concatenation). Our concatenated sequences consisted of 1362 bp divided in fragments of 429, 275 and 658 bp of 18S, ITS-1 and 28S rDNA, respectively. We generated 31 sequences of 10 morphospecies for 18S+ITS-1 rDNA and 85 sequences of 17 morphospecies for 28S rDNA. The ITS-1 fragment was the most variable (60.2% maximum variance, addendum 2) followed by COI (44.3%, addendum 3), 28S (11.9%, addendum 4) and 18S (4.8%, addendum 5-7). For the COI-alignment the topology of both the ML and Bayesian trees was not well supported, especially the deeper branches lacked support. We, therefore, used the COI dataset to compare haplotypes through median-joining haplotype networks (Fig. 2a-c). The COI dataset included 15 morphospecies and was divided in 18 species by ASAP and between 22–28 by bPTP. The concatenated dataset included 37 morphospecies and was divided in 34–38 species by ASAP and between 22–47 by bPTP. The species divisions in the COI dataset between ASAP and bPTP corresponded well and were largely well supported with bPTP. However, for the concatenated ribosomal dataset with bPTP the species divisions were largely unsupported.

Co-introduced parasites of Nile tilapia

Four of the co-introduced species of Cichlidogyrus, C. halli, C. thurstonae, C. tilapiae, and C. sclerosus showed instraspecific variation in COI (Addendum 3). The variation within the halli group was so high (0.6–30.1%, 104 polymorphic sites, Pi=0.1463) that it did not allow a median-joining haplotype network. Within C. halli, we observed nine groups, each separated by more than ten to 30 mutations. The locality Bumaki in the Upper Congo has the most haplotypes of C. halli and these cluster in three groups. Furthermore, the native C. cf. halli ‘Burundi’ and C. halli morphotype 2 sensu Jorissen et al. 2018a showed the highest distance to the introduced specimens (0.2–3.2% 18S, 3.2–10.1% ITS-1, 1.1–1.9% 28S, 24.7–36% COI), with large distances within these native representatives for COI also, but not for the rDNA fragments (0.2% 18S, 2.4% ITS-1, 1% 28S, 23.1–29.5% COI). The COI sequence of C. cf. halli ‘Burundi’ has three non-synonymous mutations compared to introduced C. halli and the two specimens of C. halli morphotype 2 have two and four non-synonymous mutations respectively. One non-synonymous mutation was shared between these three native specimens and two non-synonymous mutations between the two specimens of C. halli morphotype 2. Within the introduced specimens of C. halli in Congo, three of Upper Congo and one from Middle Congo shared the same non-synonymous mutation. Other co-introduced species, C. sclerosus (1–2.7% COI, 16 polymorphic sites, Pi=0.01776), C. tilapiae (0.3–3.4% COI, 10 polymorphic sites, Pi=0.01409) and C. thurstonae Ergens 1981 (0.3–3.4% COI, 13 polymorphic sites, Pi=0.01706) show more modest variation (Addendum 3, 6).

For C. sclerosus, each locality was characterized by a unique parasite haplotype, with the highest variation within Upper Congo. Only one locality in Upper Congo had one non-synonymous mutation. The haplotype from the Middle Congo was more closely related to the one from Madagascar than to the haplotypes from Upper Congo (Fig. 2a). For C. thurstonae on the other hand, the highest diversity was found in Lower Congo and haplotypes are shared by parasites from Madagascar and Upper Congo (Fig. 2b). Specimens from Upper and Middle Congo cluster furthest apart, each separated by at least five respectively from the nearest C. thurstonae specimen from Lower Congo, of which three were non-synonymous. Thus, the three regions of the Congo Basin do not share any C. thurstonae haplotypes. This in contrast to C. tilapiae, where apart from a few unique sequences, haplotypes are shared between the Lower and Middle Congo and between Lower Congo and Madagascar (Fig. 2c). The native parasite specimens from Burundi were clearly distinct from the introduced specimens in Congo and Madagascar. However, the amino acid sequence of these Burundese specimens is identical to all other specimens of C. tilapiae, except for one specimen from Middle Congo, which has two non-synonymous mutations.

Phylogenetic relationships among native and introduced tilapia parasites and evaluation of species status

A Bayesian phylogram constructed with 28S and 18S+ITS-1 sequences of Cichlidogyrus/Scutogyrus representatives of native and introduced tilapia parasites and sequences from GenBank is shown in Fig. 3. The topology of BI and ML trees were very similar (ML tree not shown). Minor differences were on the level of poorly supported clades (bootstrap support <70), or unresolved intraspecific relationships. The basal topology is unclear from our analyses as we observe six monophyletic groups, between which the relations are unresolved. We named each of these groups after the oldest described species within it, following Pouyaud et al. (2006).

The “halli” group has a reference sequence of C. halli from Senegal (Table 4) at the base followed by a polytomy of 18 haplotypes of co-introduced C. halli from Nile tilapia in the Congo basin and Madagascar (0–2.9% 18S, 0.4–7.2%ITS-1, 0–1.4% 28S 1.6–21.9% COI, see Addenda 2–7). More derived of this group are three sequences of native specimens of Nile tilapia from Burundi, Mweru tilapia from Upper Congo and a hybrid Oreochromis host (Vanhove et al. 2018). Cichlidogyrus halli forms its own clade and the closest relative was not revealed by phylogenetic analysis.

The “papernastrema” group houses apart from Cichlidogyrus papernastrema Price, Peebles and Bamford 1969 also Cichlidogyrus zambezensis Douëllou 1993. Our two specimens of C. papernastrema (intraspecific distance of 1.8% 28S, 29.9% COI) do not cluster together. Cichlidogyrus zambezensis, on the contrary, is monophyletic and the distance (0.6% 28S) between specimens on different host species is larger than between two specimens of the same host species (identical 28S), although they all are from the same geographic region. Furthermore, both ASAP and bPTP split C. zambezensis in two species.

The “tiberianus” group has Cichlidogyrus cubitus at the basal position followed directly by C. tiberianus Paperna 1960 (reference sequence from Senegal). The group then splits up in three lineages, one of which includes a polytomy of 23 haplotypes of co-introduced C. thurstonae from Nile tilapia, the reference of C. thurstonae from Madagascar and C. ergensi Dossou 1982. Distances within this lineage are 0–1.7% 18S, 0–4.4% ITS-1, 0–0.6% 28S, 0.3–4.8% COI (Addenda 2–7). However, the support for this group is very low (53 posterior probability and <50 bootstrap value). Directly related to C. thurstonae and C. ergensi are Cichlidogyrus dossoui Douëllou 1993 and C. tiberianus from Upper Congo. Our specimens of C. tiberianus from Upper Congo do not cluster with the reference of C. tiberianus from Senegal. The distance of C. tiberianus between Upper Congo and Senegal is 0.2–0.5% 18S, 8.5–9% ITS-1, 1.4% 28S.

The second lineage in the “tiberianus” group has Cichlidogyrus aegypticus Ergens 1981 at the base followed by C. arthracanthus Paperna 1960 (Senegal) and two specimens of C. sp. 2 from Congolese Redbreast tilapia, which are morphologically similar to C. arthracanthus (Jorissen et al. 2020). Pairwise distances amount to 1–1.4% for 28S between C. sp. 2 and C. arthracanthus and 0.5% within C. sp. 2. Within this group we have a well-supported monophyletic group which includes Cichlidogyrus agnesi Pariselle and Euzet 1995 and C. gallus Pariselle and Euzet 1995 as sister species and the C. bilongi Pariselle and Euzet 1994 and C. flexicolpos Pariselle and Euzet 1995 as sister species (only supported by PP), respectively. The last lineage includes Cichlidogyrus douellouae Pariselle, Bilong Bilong, Euzet 2003.

In the “cirratus” group, we find Cichlidogyrus njinei Pariselle, Bilong Bilong and Euzet 2003 at the base followed by C. acerbus Dossou 1982 and C. cirratus (Senegal) and lastly two specimens of C. cirratus from native Mweru tilapia from Bangweulu-Mweru. Distances within C. cirratus are 0.5% 18S, 9.8% ITS-1, 0–2.8% 28S with the 28S sequences from Mweru tilapia being identical. Furthermore, bPTP suggests that our specimens of C. cirratus belong to a different species than the reference.

In the “tilapiae” group we find all sequences of C. tilapiae from both native and introduced hosts (n = 24, with p-distances ranging from 0–1.7% 18S, 0.7–2.7% ITS-1, 0–1% 28S, 0.3–3.4% COI), with no apparent structure other than that the reference sequence from Senegal is the earliest diverging (Table 4).

The “longicirrus” group is split up in three clades and contains species that infect Hemichromis spp. (Dossou and Birgi 1982, Pariselle and Euzet 2004, Jorissen et al. 2018a) One clade is well-supported and contains a representative of Cichlidogyrus falcifer Dossou & Birgi 1984 from Lower Congo, which clusters with the reference from Senegal and the recently discovered C. polyenso Jorissen, Pariselle and Vanhove 2018 from Lower Congo which clusters with C. longicirrus Paperna 1965.

Another poorly supported clade contains three smaller well-supported groups. The first contains our specimens of C. sclerosus (identical 28S, 3% COI), with their reference sequence (distance of 1.4–1.6% 28S) and a reference of C. amphoratus Pariselle and Euzet 1995. The recently discovered C. consobrini Jorissen, Pariselle and Euzet 2018 clusters with C. irenae Gillardin, Vanhove, Pariselle, Snoeks, Huyse and Volckaert 2011 and C. casuarinus Pariselle, Muterezi Bukinga and Vanhove 2015. These three species have not been considered closely related to each other based before (Pariselle et al. 2015, Jorissen et al. 2018a). Furthermore, both C. irenae and C. casuarinus were found in Lake Tanganyika but on different host species, Gnathochromis pfefferi (Boulenger, 1898) and representatives of Bathybates Boulenger 1898, respectively (Gillardin et al. 2012, Pariselle et al. 2015, Kmentová et al. 2016), whilst C. consobrini is native to the Upper Congo and found on Sargochromis mellandi and Orthochromis katumbii Schedel, Vreven, Manda, Abwe, Manda, Schliewen, 2018 (Jorissen et al. 2018a). Furthermore, we have a clade that consists of representatives of Scutogyrus.

Finally, we have a well-supported clade with two specimens of C. berradae Pariselle & Euzet 2003 (0.2% 28S), C. quaestio Douëllou 1993, C. digitatus Dossou 1982 and C. yanni Pariselle and Euzet 1995. The 28S sequence of C. yanni was identical to C. berradae and was left out of the tree. Jorissen et al. (2018a) predicted all these species were closely related to each other based on the morphology of the sclerites.

In this study, we aim to compare haplotypes of monogenean parasites from introduced populations of Nile tilapia from three regions of the Congo Basin and one from Madagascar. In addition, we generated haplotypes of the parasites from a native population of Nile tilapia in Burundi. We expect the introduced parasite populations from Congo and Madagascar to be low in genetic variation. Additionally, if they share many of the same haplotypes they might have a common introduction origin. Our second goal is to evaluate the species status of co-introduced monogeneans of Nile tilapia through four molecular markers. Lastly, this study aims to add sequences of Congolese native and introduced species (Table 1, addendum 1) and evaluate the position of these previously unsequenced species in phylogenetic analyses.

Co-introduced parasites of Nile tilapia

From the COI haplotype networks it can be inferred that the variation within countries is sometimes higher than between countries. For example, C. thurstonae specimens from Upper Congo and Madagascar are identical but they are different from specimens collected in the Lower and Middle Congo (Fig. 2b). This lack of isolation by distance (IBD) typically reflects recent introduction events, which blurs geographic signals (Hayward et al. 2001). For example, Gyrodactylus anguillae Ergens, 1960 had identical rDNA sequences (ITS-1, 5.8S, ITS-2) in North America, Europe and Australia as a result of live eel trade that started forty to fifty years ago (Hayward et al. 2001). Similarly, Gyrodactylus cichlidarum Paperna, 1968 was co-introduced into Mexico with Nile tilapia after fish introductions started in the 1940s (García-Vásquez et al. 2017). The ITS-1 sequences of G. cichlidarum specimens that spilled over to Mexican poeciliids were almost completely identical to G. cichlidarum from Nile tilapia in Ghana (García-Vásquez et al. 2017). Our results, therefore, strongly point to an identical or geographically similar source of introduction of Nile tilapia in the Upper Congo and Madagascar, and a different source (or sources) for the other Congolese regions.

The high haplotype diversity of C. sclerosus within the Upper Congo (Fig. 2a) can partly be explained by the sampling bias (relatively more specimens were sequenced from this locality), but it also strongly suggests that multiple introductions have taken place in this area, from different geographic source populations. Indeed, the variation is higher compared to that of the population of C. sclerosus on native Nile tilapia in Burundi. Aquaculture of Nile tilapia in the DRC started in the Lubumbashi area, Upper Congo, in the late 1940s (Micha 2013). However, there has also been a period of very low aquaculture activity until 1996 (Toguyeni 2004) and it is not known whether Nile tilapia or its parasites from before 1996 still persist in the basin. In any case, our results refute our initial hypothesis that introduced parasite populations suffer bottlenecks. Similar scenarios have been described for other biological invasions, where introduced populations could maintain a high diversity because of multiple introductions from different source populations (Kolbe et al., 2004; Genton et al., 2005), sometimes followed by intraspecific hybridization (Rosenthal et al., 2008).

Finally, the C. tilapiae haplotypes shared by the Lower and Middle Congo suggest that natural gene flow is possible at this scale, or it could point to a shared introduction source (Fig. 2c).

Marker performance and barcoding gap

Based on our histograms (Fig. 4), we find a significant (P<0.05) barcoding gap for COI at 15% (Fig. 4a) but none for 28S, 18S or ITS-1 (Fig. 4b–d). Visually there is a second gap between 7–11% for COI (Fig. 4a) but this was not found significant by ASAP. It should be noted that our fragment of the COI gene covers just less than a quarter of the total COI gene and constitutes the most variable part (unpublished data). Additionally, the COI dataset itself was the smallest of the four markers because hardly any references were available on Genbank and the amplification success of COI was lower than that of the rDNA markers. Whether 15% is representative for Cichlidogyrus/Scutogyrus should be investigated in the future by including more species.

As for the other markers we can find some clues in the literature to a possible barcoding gap. Within Cichlidogyrus, Rahmouni et al. (2021) found intraspecific variation of ITS-1 in C. nshomboi Muterezi Bukinga, Vanhove, Van Steenberge and Pariselle 2012, up to 1.1% and interspecific variation starting at 3.5%. The 18S sequences of C. nshomboi and Cichlidogyrus casuarinus Pariselle, Muterezi Bukinga, Van Steenberge, Vanhove 2015, were identical and the 28S sequences differed 0.13% between the species. COI intraspecific variation within C. nshomboi amounted to 12.2%. This distance for COI roughly corresponds with what we find in our dataset, but the observed distances for the three rDNA fragments are higher in our study. Representatives of Trinigyrus (Dactylogyridae: Monogenea), which infect siluriforms, have interspecific variation <6% for COI and around 1% for 28S (Franceschini et al. 2020). In Dactylogyrus (Dactylogyridae: Monogenea), which infect European cyprinids, the cut-off value was set at 1.4% for a fragment consisting of partial 18S, complete ITS-1 and partial 5.8S (Šimková et al. 2004). Rahmouni et al. (2017b) found 1% for 28S; 0.4% for 18S and 4.3% for ITS-1 of Dactylogyrus parasitizing North-African congeneric cyprinids. From all these we learn that the barcoding gap for 28S may be around 1% and for 18S below 1%. For ITS-1, this is likely higher than 1% but probably lower than the 15%, which we found for COI of the included species of Cichlidogyrus.

Furthermore, 28S sequences can be identical over large geographic distances (C. thurstonae, C. sclerosus and C. falcifer) between introduced and native populations (C. tilapiae) and even between species (C. berradae and C. yanni; Scutogyrus gravivaginus (Paperna & Thurston 1969), S. bailloni Pariselle & Euzet 1995 and S. longicornis (Paperna & Thurston, 1969)) (Addendum 4). However, whether 28S can be identical between species of Cichlidogyrus/Scutogyrus is uncertain as the obtained references from Genbank can be morphologically misidentified. New sequences of C. berradae, C. yanni, S. gravivaginus, S. longicornis and S. bailloni are needed to verify this. Conversely, rDNA fragments can appear conserved in other closely related monogeneans. Kmentová et al. 2016a found identical 28S and 18S+ITS-1 sequences of Cichlidogyrus casuarinus from Lake Tanganyika from different hosts (Bathybates and Hemibates Regan 1920), although intraspecific morphological variation was observed. The COI fragments, on the other hand, were highly differentiated, with distances between haplotypes reaching 4.7% (Kmentová et al. 2016a). For Kapentagyrus tanganicanus Kmentová, Gelnar and Vanhove 2018 infecting Lake Tanganyika sardines, morphological intraspecific variation was found based on host species (phenotypic adaptation for attachment), COI fragment and possibly seasonality, but this was not reflected in the 28S or 18S+ITS-1 sequences (Kmentová et al. 2018). Benovics et al. (2017) observed identical 18S+ITS-1 sequences between Dactylogyrus vastator Nybelin 1924, from the Po River, Italy and Šuica River, Bosnia and Herzegovina. Marchiori et al. (2015) found identical 18S sequences for different, but closely related species of Ligophorus infecting Mugilidae in Brazil (Dactylogyridae: Monogenea), whilst 28S differed 0.2% and ITS-1 0.4%. In conclusion, rDNA sequences might be conserved in conspecific monogeneans over large geographic distances and between conspecifics from different host species also, but it is unlikely for different species to have identical rDNA sequences. Therefore, there is potential for the rDNA fragments to be used for species delineation in Cichlidogyrus/Scutogyrus, as shown for Gyrodactylus spp. (Matejusová et al. 2001, Mendoza-Palmero et al. 2019) and Dactylogyrus spp. (Benovics et al. 2018). However, at this time, we need to include other methods such as bPTP to interpret results from the rDNA fragments about species status. What the rDNA fragments are good at right now are constructing higher-level phylogenies and the COI-fragment is suited for population level studies and species delineation.

Phylogenetic relationships among native and introduced tilapia parasites and evaluation of species status

Previous phylogenetic studies showed that Scutogyrus forms a monophyletic group within Cichlidogyrus, rendering Cichlidogyrus paraphyletic (Pouyaud et al. 2006; Wu et al. 2007; Mendlová et al. 2012; Mendlová and Šimková 2014). Representatives of Cichlidogyrus formed several well-supported monophyletic groups, but the relation between some of these groups were unresolved (Pouyaud et al. 2006; Wu et al. 2007; Mendlová et al. 2012; Mendlová and Šimková 2014). We focus on and add species from the Congo Basin, which were until now sparsely represented in phylogenetic studies of Cichlidogyrus. Our results largely correspond with the previously published analyses on Cichlidogyrus/Scutogyrus (Pouyaud et al. 2006; Wu et al. 2007; Mendlová et al. 2012; Mendlová and Šimková 2014).

Cichlidogyrus halli forms a species complex (see Jorissen et al. 2018a) supported by molecular data in the present study. ASAP suggests the “halli” group to consist of at least three species; firstly, the native specimen from Burundi, secondly the native specimens from Mweru tilapia from Upper Congo together with the specimen from the O. niloticus x mweruensis hybrid from Upper Congo, and thirdly all introduced specimens of C. halli. The genetic distances within the “halli” group are indeed large (2.1% for 28S; 3.2% for 18S; 10.1% for ITS-1 and 36% for COI). This variation is higher than all other intra/interspecific boundaries stated above. The bPTP method is inconclusive for the rDNA markers, where C. halli is divided in six species, including morphotype 2 and C. cf. halli ‘Burundi’, but the support for these divisions is very low. The divisions in COI are better supported and correspond with ASAP.

Morphotype 2 of C. halli (sensu Jorissen et al. 2018a) from Upper Congo, as drawn and discussed by Jorissen et al. (2018a), corresponds in locality and host species with C. halli ex O. mweruensis on the tree (Fig. 3). Therefore, we suggest that morphotype 2 should be elevated to species level. Similarly, the specimens of C. cf. halli ‘Burundi’ from Lake Tanganyika, Burundi belong to a third species within C. halli, as suggested by ASAP. From the same Burundese population we found a specimen of C. cf. halli ‘Burundi’ with elongated and thickened hooklets pair I compared to C. halli (Fig. 5). However, we strongly feel that the species delineation within the “halli” group should be based on morphology and genetics together. Therefore, new morphological material is needed to resolve this. For species within Cichlidogyrus/Scutogyrus, the genital sclerites are important for species identification (see diagnosis in Pariselle & Euzet 2009). Therefore, for our study, we decided to only keep the body part with the genital sclerites and use the body part with the haptor for genetic analysis (Jorissen et al. 2018a). However, recent work on Kapentagyrus and Cichlidogyrus shows that closely related species might first diverge in haptor morphology before genital sclerites (Messu Mandeng et al. 2015, Kmentová et al. 2016a). This implicates the evolution of these parasites is related strongly to microhabitat (attachment site) and host species (Messu Mandeng et al. 2015, Gobbin et al. 2020). We conclude that morphological features of the haptor are important in this complex for species delimitation.

Cichlidogyrus zambezensis and Cichlidogyrus papernastrema together form a clade. However, both species belong to different groups within the genus based on the morphology of the haptoral hooklets. Cichlidogyrus zambezensis has seven pairs of small hooklets (group A sensu Vignon et al. 2011), whilst in C. papernastrema the first pair is thick and elongated (group B sensu Vignon et al. 2011). Pariselle and Euzet (2003) suggested a division of species of Cichlidogyrus in three groups based on the morphology of haptoral hooklets (uncinuli in the source). Additionally, Vignon et al. (2011) found a high congruence between these morphological groups and the molecular phylogeny, meaning that hooklet morphology is phylogenetically constrained. However, the well-supported clade including C. papernastrema and C. zambezensis has a representative of group A and group B. This suggests this group division might not be supported by phylogenies. Pariselle and Euzet (2003) and Vignon et al. (2011) included a subset of species in their analyses. Vignon et al. (2011) even omitted C. arthracanthus from their analysis because it did not fit any of the three groupings. In conclusion, this could mean that firstly, haptor morphology is not as phylogenetically constrained as previously thought (see the ‘halli’ group above) and secondly, that the division in three groupings - whilst useful for morphological identification, see Pariselle and Euzet 2009 – does not cover the morphological evolution of the haptor within Cichlidogyrus fully.

Both C. papernastrema and C. zambezensis have a copulatory tube with a bulbous thickening in the middle and this could be a synapomorphy for the “papernastrema” group instead of characters of haptor morphology. Other species with a bulbous thickening of the copulatory tube and thus possibly belonging to this group are Cichlidogyrus halinus Paperna 1969, Cichlidogyrus sanjeani Pariselle and Euzet 1997, Cichlidogyrus philander Douëllou 1993, Cichlidogyrus bulbophallus Geraerts and Muterezi Bukinga 2020, Cichlidogyrus pseudozambezensis Geraerts and Muterezi Bukinga 2020 and Cichlidogyrus ranula Geraerts and Muterezi Bukinga 2020. Within these candidate species are several representatives from Haplochromine cichlids and others from southern Africa. Cichlidogyrus zambezensis is monophyletic, but we observe variation between specimens of different host species. Douëllou (1993) reports intraspecific morphological variation between different hosts in C. zambezensis. Cichlidogyrus zambezensis is known from four cichlid hosts, belonging to three cichlid lineages (Douëllou 1993, Vanhove et al. 2013, Jorissen et al. 2018a). Additionally, the bPTP analysis of COI splits our samples of C. zambezensis and the reference as different species. Therefore, C. zambezensis is in need of further study and might consist of multiple species. The monophyly of C. papernastrema is not supported. Even more, the genetic distance between the two specimens of C. papernastrema is larger than between this species and C. zambezensis and above 1% for 28S and 15% for COI. Therefore, it is likely that both specimens belong to different species. Jorissen et al. (2018) redescribed C. papernastrema and noted large variation in thickness of the copulatory tube between specimens. It would be worthwhile to check whether this variation is a good diagnostic character to delineate species in tandem with genetic distances.

In the “tiberianus” group, C. tiberianus from Bangweulu-Mweru does not cluster with the reference sequence from Senegal (Mendlová et al. 2012). This species infects representatives of Coptodon ranging from Senegal to Zimbabwe (Douëllou 1993, Pariselle and Euzet 1995, 1996, 2009, Mendlová et al. 2012, Jorissen et al. 2018a). This is a native range of over 7000 km, including different ichthyographic provinces and bassins and it is, therefore, plausible that C. tiberianus might consist of multiple species within this range. Additionally, the genetic distances between C. tiberianus of Senegal and Upper Congo is above 1% for 28S which also point to multiple species; however the distance in 18S is well below 1%. Fannes et al. (2017) used SEM to investigate the sclerotized parts of C. dossoui and C. tiberianus from Upper Congo because both species are morphologically quite similar and share hosts. The COI genetic distances are smaller between C. tiberianus and C. dossoui from Upper Congo than within C. tiberianus. Therefore, it is not surprising that C. dossoui from Bangweulu-Mweru appears as the sister species to C. tiberianus from Bangweulu-Mweru. Cichlidogyrus tiberianus requires a species status re-evaluation backed by genetic data from across its native range and different host species.

Furthermore, in the “tiberianus” group, C. ergensi is situated within C. thurstonae, but with low support (53 posterior probability and not supported in the ML analysis), thus we do not make inferences to this result. All species in the “tiberianus” group belong to group C based on the morphology of the haptoral hooklets (Pariselle and Euzet 2003, Vignon et al. 2011), except for C. arthracanthus and C. sp. 2, which fall outside of the classification in three main groups. Here again, the division by Pariselle and Euzet (2003) is not completely supported.

Cichlidogyrus cirratus was found to be monophyletic (100 posterior probability and 90 bootstrap support value, Fig. 3). However, the branch lengths within C. cirratus are much longer than for example between the different species of Scutogyrus. The genetic distance between C. cirratus from Bangweulu-Mweru and Senegal is 0.5% 18S, 9.8% ITS-1 and 2.8% for 28S (Addenda 2, 4, 5). This indicates that our samples might represent two separate species. It is also debated whether C. cirratus and C. mbirizei Muterezi Bukinga, Vanhove, Van Steenberge, Pariselle, 2012, are conspecific (Zhang et al. 2019). Scanning electron microscopy revealed that the distinguishing characters between C. cirratus and C. mbirizei (Muterezi Bukinga et al. 2012) on specimens of C. cirratus from China (introduced) could be transformed by turning the specimens over (Zhang et al. 2019). In conclusion, we deem it likely that C. cirratus and possibly C. mbirizei consist of multiple species and that this should be investigated further genetically. Subsequently, an evaluation of the morphological characters within C. cirratus and C. mbirizei is needed.

In Cichlidogyrus tilapiae the reference sequence from Senegal appears basal to all other specimens of native and introduced hosts in the Congo Basin (Fig. 3). We do not find any evidence to contest the species status of C. tilapiae as opposed to Pouyaud et al. 2006 who suggested it is a species complex based on ribosomal DNA.

Our results strongly point to an identical or geographically similar source of introduction of Nile tilapia in Congo and Madagascar, as both regions share identical COI parasite haplotypes. The high haplotype diversity of C. sclerosus within the Upper Congo strongly suggests that multiple introductions have taken place in this area, from different geographic source populations. This refutes our initial hypothesis that introduced parasite populations would suffer genetic bottlenecks. Also, the strong differentiation between parasites from the Upper Congo compared to those from the Middle and Lower Congo suggests different sources of introduction for the latter two regions. Finally, shared parasite haplotypes between the Lower and Middle Congo suggest that natural gene flow is possible at this scale, or it could point to a shared source of introduction.

Considering the genetic markers, we find a barcoding gap at 15% variation for COI, but not for the other markers. This value is quite high compared to other dactylogyrid monogeneans, but it aligns with the only other available study within Cichlidogyrus, which suggests a barcoding gap above 12% (Rahmouni et al. 2021). However, we want to stress that sequences of more species are needed to have a more complete overview of the phylogeny of the group and to estimate a barcoding gap more precisely.

Based on our study, we suggest the need of taxonomic re-evalution for C. halli, C. papernastrema, C. zambezensis, C. tiberianus and C. cirratus as they all potentially represent at least two species. Additionally, within C. halli we find that closely related species can first diverge in haptor morphology and later differentiate in the genital sclerites. Lastly, the grouping of C. papernastrema with C. zambezensis, C. arthracanthus and C. sp. 2 within the “tiberianus” group shows that the division of the genus by Pariselle et al. 2003 based on haptor configuration does not explain the variation within the group fully and that this division is not always phylogenetically supported. We utter the need for a revision of morphological features corresponding with the larger clades in the phylogenetic tree of Cichlidogyrus/Scutogyrus.

Acknowledgements

We thank B Vanschoenwinkel (VUBrussels), I Schön (Royal Belgian Institute for Natural Sciences) and J-L Justine (Musée National d’histoire Naturelle) for independently reviewing this manuscript before submission. We thank S Koblmüller (University of Graz), E Abwe, BK Manda, C Mukweze Mulelenu, M Kasongo Ilunga Kayaba and C Kalombo Kabalika (Université de Lubumbashi), M Collet and P N’Lemvo (Institut Congolais pour la Conservation de la Nature), D Kufulu-ne-Kongo (Ecole Muilu Kiawanga), L Matondo Mbela (Université Kongo), P Nguizani Bimbundi, B Boki Fukiakanda, P Ntiama Nsiku (Institut Supérieur Pédagogique de Mbanza-Ngungu), P Nzialu Mahinga (Institut National pour l’Etude et la Recherche Agronomiques – Mvuazi/Institut Supérieur d’études agronomiques – Mvuazi), M Katumbi Chapwe, E Řehulková, N Kmentová (Masaryk University), the Schreyen-Brichard family (Fishes of Burundi), and JR Rasoloariniaina, J Rasamy Razanabolana and S Rakotomamonjy (Université d’Antananarivo) for administrative, field and lab support, making this study possible. This research was supported by the Belgian Federal Science Policy Office (BRAIN-be Pioneer Project (BR/132/PI/TILAPIA), research grant 1513419N of the Research Foundation – Flanders (FWO-Vlaanderen), the University Development Cooperation of the Flemish Interuniversity Council (VLIR-UOS) South Initiative ZRDC2014MP084, the OCA type II project S1_RDC_TILAPIA and the Mbisa Congo project (2013-2018), the latter two being framework agreement projects of the RMCA with the Belgian Development Cooperation, and the Czech Science Foundation project no. P505/12/G112 - European Centre of Ichtyoparasitology (ECIP). MWPJ was supported by a BOF Reserve Fellowship from Hasselt University. MPMV is receiving support from the special research fund (BOF20TT06). MPMV received travel grant K220314N from the Research Foundation – Flanders (FWO-Vlaanderen), support from the Belgian Directorate-General for Development Cooperation and Humanitarian Aid (CEBioS programme at the Royal Belgian Institute of Natural Sciences), and is financed by the Special Research Fund of Hasselt University (BOF20TT06). The research leading to results presented in this publication was carried out with infrastructure funded by EMBRC Belgium – FWO project GOH3817N. Fieldwork was carried out under permission 06/AR.ED./15 from the General Directorate for Fishery Resources and Fisheries, Ministry of Fisheries Resources and Fisheries of Madagascar, and mission statements 863/2014 (Faculté des Sciences Agronomiques, Université de Lubumbashi), C/075/2015/I.S.P./MBNGU/AUT.AC and AC/076/2015/I.S.P./MBNGU/AUT.AC.

Statements and declarations

The authors have no competing interests to declare that are relevant to the content of this article.

The datasets generated during and/or analysed during the current study are available from the corresponding author on reasonable request. All input data is available through public repositories (see material and methods).

Alcaraz, G., Lόpez-Portela, X., Robles-Mendoza, C. (2015). Response of a native endangered axolotl, Ambystoma mexicanum (Amphibia), to exotic fish predator. Hydrobiologia, 753(1), 73-80.
Avise, J.C. (1994). Molecular markers, natural history and evolution. Chapman and Hall.
Bandelt, H., Forster, P., Röhl, A. (1999). Median-joining networks for inferring intraspecific phylogenies. Molecular Biology and Evolution, 16(1), 37–48.
Barson, M., Přikrylová, I., Vanhove, M.P.M., Huyse, T. (2010). Parasite hybridization in African Macrogyrodactylus spp. (Monogenea, Platyhelminthes) signals historical host distribution. Parasitology, 137, 1585–95.
Bates, J. (1997). The sealing compound: Glyceel. Journal of Nematology, 29(4), 565-566.
Benovics, M., Kičinjaová, M.L., Šimková, A. (2017). The phylogenetic position of the enigmatic Balkan Aulopyge huegelii (Teleostei: Cyprinidae) from the perspective of host-specific Dactylogyrus parasites (Monogenea), with a description of Dactylogyrus omenti n. sp.. Parasites & Vectors, 10, e547.
Benovics, M., Desdevises, Y., Vukić, J., Šanda, R., Šimková, A. (2018). The phylogenetic relationships and species richness of host-specific Dactylogyrus parasites shaped by the biogeography of Balkan cyprinids. Scientific Reports, 8, 13006.
Canonico, G.C., Arthington, A., McCrary, J.K., Thieme, M.L. (2005). The effects of introduced tilapias on native biodiversity. Aquatic Conservation: Marine and Freshwater Ecosystems, 15, 463–483.
Castresana, J. (2000). Selection of conserved blocks from multiple alignments for their use in phylogenetic analysis. Molecular Biology and Evolution, 17(4), 540–552.
Darriba, D., Taboada, G.L., Doallo, R., & Posada, D. (2012). jModelTest 2: more models, new heuristics and parallel computing. Nature Methods, 9, 777(2012).
Deines, A.M., Bbole, I., Katongo, C., Feder, J.L., Lodge, D.M. (2014). Hybridization of native Oreochromis species (Cichlidae) and the introduced Nile tilapia (O. niloticus) in the Kafue River, Zambia. African Journal of Aquatic Science, 39(1), 23-34.
Deines, A.M., Wittmann, M.E., Deines, J.M., Lodge, D.M. (2016). Tradeoffs among ecosystem services associated with global tilapia introductions. Reviews in Fisheries Science and Aquaculture, 24(2), 178–191.
Dossou, C., Birgi, E. (1984). Monogènes parasites d’Hemichromis fasciatus Peters, 1857 (Teleostei: Cichlidae). Annales des Sciences naturelles, Zoologie, Paris, 6, 101–109.
Douëllou, L. (1993). Monogeneans of the genus Cichlidogyrus Paperna, 1960 (Dactylogyridae: Ancyrocephalinae) from cichlid fishes of Lake Kariba (Zimbabwe). with descriptions of five new species. Systematic Parasitology, 50, 159–186.
Edgar, R.C. (2004a). MUSCLE: Multiple Sequence Alignment with high accuracy and high throughput. Nucleic Acids Research, 32(5), 1792-97.
Edgar, R.C. (2004b). MUSCLE: a multiple sequence alignment method with reduced time and space complexity. BMC Bioinformatics, 5(1), 113.
FAO (2017). Regional review on status and trends in aquaculture development in sub-Saharan Africa -2015. FAO fisheries and aquaculture circular no 1135/2. FAO Rome.
Fannes, W., Vanhove, M.P.M., Huyse, T. (2017). Redescription of Cichlidogyrus tiberianus Paperna, 1960 and C. dossoui Douëllou, 1993 (Monogenea: Ancyrocephalidae), with special reference to the male copulatory organ. Systematic Parasitology, 94, 133–144.
Firmat, C., Alibert, P., Losseau, M., Baroiller, J.F., Schliewen, U.K. (2013). Successive invasion-mediated interspecific hybridizations and population structure in the endangered cichlid Oreochromis mossambicus. PLoS ONE, 8(5), e63880.
Franceschini, L., Acosta, A.A., Zago, A.C., Müller, M.I., da Silva, R.J. (2020). Trinigyrus spp. (Monogenea: Dactylogyridae) from Brazilian catfishes: new species, molecular data and new morphological contributions to the genus. Journal of Helminthology, 94, e126.
García-Vásquez, A., Razo-Mendivil, U., Rubio-Godoy, M. (2017). Triple trouble? Invasive poeciliid fishes carry the pathogen Gyrodactylus cichlidarum in the Mexican highlands. Veterinary Parasitology, 235, 37–40.
Genton, B.J., Skykoff, J.A., Giraud, T. 2005. High genetic diversity in French invasive populations of common ragweed, Ambrosia artemisiifolia, as a result of multiple sources of introduction. Molecular Ecology. 14, 4275-4285.
Gillardin, C., Vanhove, M.P.M., Pariselle, A., Huyse, T., Volckaert, F.A.M. (2012). Ancyrochephalidae (Monogenea) of Lake Tanganyika: II: description of the first Cichlidogyrus spp.parasites from Tropheini fish hosts (Teleostei, Cichlidae). Parasitology Research, 110, 305–313.
Goedknegt, M.A., Feis, M.E., Wegner, K.M., Luttikhuizen, P.C., Buschbaum, C., Camphuysen, K.C.J., Van der Meer, J., Thieltges, D.W. (2016). Parasites and marine invasions: ecological and evolutionary perspectives. Journal of Sea Research, 113, 11–27.
Gonçalves, E.L.T., Jerônimo, G.T., Martins, M.L. (2009). On the importance of monogenean helminthes in Brazilian cultured Nile tilapia. Neotropical Helminthology, 3, 53−56.
Hassouna, N., Michot, B., Bachellerie, J.P. (1984). The complete nucleotide sequence of mouse 28S rRNA gene. Implications for the process of size increase of the large subunit rRNA in higher eukaryotes. Nucleic Acid Research, 12(8), 3563–83.
Hayward, C.J., M. Iwashita, K. Ogawa and I. Ernst, 2001. Global spread of the eel parasite Gyrodactylus anguillae (Monogenea). Biological Invasions, 3, 417-424.
Huyse, T., Vanhove, M.P.M., Mombaerts, M., Volckaert, F.A.M., Verreycken, H. (2015). Parasite introduction with an invasive goby in Belgium: double trouble? Parasitology Research, 114, 2789–2793.
Iglesias, R., García-Estévez, J., Ayres, C., Acuňa, A., Cordero-Rivera, A. (2015). Frist reported outbreak of severe spirorchiidiasis in Emys orbicularis, probably resulting from a parasite spillover event. Diseases of Aquatic Organisms, 113, 75–80.
Jiménez-García, M.I., Vidal-Martínez, V.M., Lόpez-Jiménez, S. (2001). Monogeneans in Introduced and Native Cichlids in México: Evidence for Transfer. Journal of Parasitology, 87(4), 907–909.
Jorissen, M.W.P., Pariselle, A., Huyse, T., Vreven, E.J., Snoeks, J., Volckaert, F.A.M., Chocha Manda, A., Kapepula Kasembele, G., Artois, T., & Vanhove, M.P.M. (2018a). Diversity and host specificity of monogenean gill parasites (Platyhelminthes) of cichlid fishes in the Bangweulu-Mweru ecoregion. Journal of Helminthology, 92(4), 417–437.
Jorissen, M.W.P., Pariselle, A., Huyse, T., Vreven, E.J., Snoeks, J., Decru, E., Kusters, T., Wamuini Lunkayilakia, S., Muterezi Bukinga, F., Artois, T. & Vanhove, M.P.M. (2018b).Six new dactylogyrid species (Platyhelminthes, Monogenea) from the gills of cichlids (Teleostei, Cichliformes) from the Lower Congo Basin. Parasite, 25, 64(2018).
Jorissen, M.W.P., Huyse, T., Pariselle, A., Wamuini Lunkayilakia, S., Muterezi Bukinga, F., Chocha Manda, A., Kapepula Kasembele, G., Vreven, E.J., Snoeks, J., Decru, E., Artois, T., Vanhove, M.P.M. (2020). Historical museum collections help detect parasite species jumps after tilapia introductions in the Congo Basin. Biological Invasions, 22(9), 2825–2844.
Kimura, M. (1980). A simple method for estimating evolutionary rate of base substitutions through comparative studies of nucleotide sequences. Journal of Molecular Evolution, 16, 111–120.
Kmentová, N., Gelnar, M., Mendlová, M., Van Steenberge, M., Koblmüller, S., Vanhove, M.P.M. (2016a). Reduced host-specificity in a parasite infecting non-littoral Lake Tanganyika cichlids evidenced by intraspecific morphological and genetic diversity. Scientific Reports, 6, 39605(2016).
Kmentová, N., Gelnar, M., Koblmüller, S., Vanhove, M.P.M. (2016b). First insights into the diversity of gill monogeneans of ‘Gnatochromis’ and Limnochromis (Teleostei, Cichlidae) in Burundi: do the parasites mirror host ecology and phylogenetic history? PeerJ, 4, e1629.
Kolbe, J.J., Glor, R.E., Schettino, L.R.G. et al. 2004. Genetic variation increases during biological invasion by a Cuban lizard. Nature. 431, 177-181.
Kumar, S., Stecher, G., Tamura, K. (2016). MEGA7: Molecular Evolutionary Genetics Analysis version 7.0 for bigger datasets. Molecular Biology and Evolution, 33, 1870–1874.
Leigh, J.W., Bryant, D. (2015). PopART: Full-feature software for haplotype network construction. Methods in Ecology and Evolution, 6(9), 1110–1116.
Littlewood, D.T.J., Rohde, K., Clough, K.A. (1997). Parasite speciation within or between host species? Phylogenetic evidence from site-specific polystome monogeneans. International Journal for Parasitology, 27, 1289–1297.
Littlewood, D.T.J. (2008). Platyhelminth systematics and the emergence of new characters. Parasite, 15, 333–341.
Lockyer, A.E., Olson, P.D., Østergaard, P., Rollinson, D., Johnston, D.A., Attwood, S.W., Southgate, V.R., Horak, P., Snyder, S.D., Le, T.H., Agatsuma, T., McManus, D.P., Carmichael, A.C., Naem, S., Littlewood, D.T.J. (2003). The phylogeny of the Schistosomatidae based on three genes with emphasis on the interrelationships of Schistosoma Weinland, 1858. Parasitology, 126, 203–224.
Marchiori, N.C., Pariselle, A., Pereira, J. Jr., Agnèse, J-F., Durand, J-D., Vanhove, M.P.M. (2015). A comparative study of Ligophorus uruguayense and L. saladensis (Monogenea: Ancyrocephalidae) from Mugil liza (Teleostei: Mugilidae) in southern Brazil. Folia Parasitologica. https://doi.org/10.14411/fp.2015.024
Matejusová, I., Gelnar, M., McBeath, A.J.A., Collins, C.M., Cunningham, C.O. (2001). Molecular markers for gyrodactylids (Gyrodactylidae: Monogenea) from five fish families (Teleostei). International Journal for Parasitology, 31(7), 738–745.
Mayr, E. (1942). Systematics and The Origin of Species. Columbia University Press.
Mendlová, M., Pariselle, A., Vyskočilová, M. & Šimková, A. (2010). Molecular phylogeny of monogeneans parasitizing African freshwater Cichlidae inferred from LSU rDNA sequences. Parasitology Research, 107, 1405–1413.
Mendlová, M., Desdevises, Y., Civáňová, K., Pariselle, A., Šimková, A. (2012). Monogeneans of West African Cichlid fish: Evolution and Cophylogenetic Interactions. PLoS one, 7(5), e37268.
Mendlová, M. & Šimková, A. (2014). Evolution of host specificity in monogeneans parasitizing African cichlid fish. Parasites & Vectors, 7, 69(2014).
Mendoza-Palmero, C.A., Blasco-Costa, I., de Leon, G.P.P. (2019). Morphological and molecular characterisation of a new species of Gyrodactylus von Nordmann, 1832 (Monogenoidae: Gyrodactylidae) of cichlid fishes (Perciformes) from Mexico. Parasitology International, 70, 102–111.
Messu Mandeng, F.D., Bilong Bilong, C.F., Pariselle, A., Vanhove, M.P.M., Bitja Nyom, A.R., Agnèse, F.A. (2015). A phylogeny of Cichlidogyrus spp. (Monogenea, Dactylogyridea) clarifies a host-switch between fish families and reveals an adaptive component to attachment organ morphology of this parasites genus. Parasites & Vectors, 8, 582.
Micha, J.-C. (2013). Fish farming in the Congo Basin: past, present and future. Conference: Nutrition and Food Production in the Congo Basin, Brussels, Belgium, 2013, National Committee for Biological Sciences, 147-171.
Moszczynska, A., Locke, S.A., McLaughlin, J.D., Marcogliese, D.J., Crease, T.J. (2009). Development of primers for the mitochondrial cytochrome c oxidase I gene in digenetic trematodes (Platyhelminthes) illustrates the challenge of barcoding parasitic helminths. Molecular Ecology Resources, 9, 75–82.
Muterezi Bukinga, F., Vanhove, M.P.M., Van Steenberge, M., Pariselle, A. (2012). Ancyrocephalidae (Monogenea) of Lake Tanganyika: III: Cichlidogyrus infecting the world’s biggest cichlid and the non-endemic tribes Haplochromini, Oreochromini and Tylochromini (Teleostei, Cichlidae). Parasitology Research, 111, 2049–2061.
NCBI (2021). Nucleotide query for Cichlidogyrus and Scutogyrus. National Center for Biotechnology Information. https://www.ncbi.nlm.nih.gov/. Accessed 1 March 2021.
Nieberding, C.M., Olivieri, I. (2007). Parasites: proxies for host genealogy and ecology? TRENDS in Ecology and Evolution, 22(3), 156–165.
Nyingi, D.W., Agnèse, J.-F. (2007). Recent introgressive hybridization revealed by exclusive mtDNA transfer from Oreochromis leucostictus (Trewavas, 1933) to Oreochromis niloticus (Linnaeus, 1758) in Lake Baringo, Kenya. Journal of Fish Biology, 70(Supplement A), 148–154.
Paperna, I. (1979). Monogenea of inland water fish in Africa. Annales du Musée royal d’Afrique centrale, sér-in8°, 226, 1–131.
Pariselle, A., Euzet, L. (1995). Gill parasites of the genus Cichlidogyrus Paperna, 1960 (Monogenea, Ancyrocephalidae) from Tilapia guineensis (Bleeker, 1862), with descriptions of six new species. Systematic Parasitology, 30, 187–198.
Pariselle, A., Euzet, L. (1996). (Monogenea, Ancyrocephalidae): gill parasites from West African Cichlidae of the subgenus Coptodon Regan, 1920 (Pisces), with descriptions of six new species. Systematic Parasitology, 34, 109–124.
Pariselle, A., Euzet, L. (2003). Four new species of Cichlidogyrus (Monogenea: Ancyrocephalidae), gill parasites of Tilapia cabrae (Teleostei: Cichlidae), with discussion on relative length of haptoral sclerites. Folia Parasitologica, 50(3), 195–201.
Pariselle, A., Euzet, L. (2004). Two new species of Cichlidogyrus Paperna, 1960 (Monogenea, Ancyrocephalidae) gill parasites on Hemichromis fasciatus (Pisces, Cichlidae) in Africa, with remarks on geographical distribution. Parasite, 11, 359–364.
Pariselle, A., Euzet, L. (2009). Systematic revision of dactylogyridean parasites (Monogenea) from cichlid fishes in Africa, the Levant and Madagascar. Zoosystema, 31(4), 849–898.
Pariselle, A., Muterezi Bukinga, F., Van Steenberge, M., Vanhove, M.P.M. (2015). Ancyrocephalidae of Lake Tanganyika: IV: Cichlidogyrus casuarinus parasitizing species of Bathybatini (Teleostei, Cichlidae): reduced host-specificity in the deepwater realm? Hydrobiologia, 748, 99–119.
Pimentel, D., McNair, S., Janecka, J., Wightman, J., Simmonds, C., O’Connell, C., Wong, E., Russel, L., Zern, J., Aquino, T., Tsomondo, T. (2001). Agriculture, Ecosystems and Environment, 84, 1–20.
Poulin, R., Krasnov, B.R. (2010). The Biogeography of Host-Parasite Interactions. Oxford University Press Inc.
Pouyaud, L., Desmarais, E., Deveny, M. & Pariselle, A.(2006). Phylogenetic relationships among monogenean gill parasites (Dactylogyridae, Ancyrocephalidae) infesting tilapiine hosts (Cichlidae): systematic and evolutionary implications. Molecular Phylogenetics and Evolution, 38, 241–249.
Prenter, J., MacNeil, C., Dick, J.T.A., Dunn, A.M. (2004). Roles of parasites in animal invasions. TRENDS in Ecology and Evolution, 19(7), 385–390.
Puillandre, N., Brouillet, S., Achaz, G. (2020). ASAP: assemble species by automatic partitioning. Molecular Ecology Resources, 21(2), 609–620.
Rahmouni C., Vanhove M.P.M. & Šimková A. (2017a) Underexplored diversity of gill monogeneans in cichlids from Lake Tanganyika: eight new species of Cichlidogyrus Paperna, 1960 (Monogenea: Dactylogyridae) from the northern basin of the lake, with remarks on the vagina and the heel of the male copulatory organ. Parasites & Vectors, 10, 591
Rahmouni, I., Řehulková, E., Pariselle, A., Berrada Rkhami, O., Šimková, A. (2017b). Four new species of Dactylogyrus Diesing, 1850 (Monogenea: Dactylogyridae) parasitizing the gills of Northern Moroccan Luciobarbus Heckel (Cyrpindae): morphological and molecular characterisation. Systematic Parasitology, 94, 575–591.
Rahmouni, I., Vanhove, M.P.M., Šimková, A., Van Steenberge, M. (2021). Conservative divergent evolution in a gill monogenean parasitizing distant cichlid lineages of Lake Tanganyika: Cichlidogyrus nshomboi (Monogenea: Dactylogyridae) from representatives of Boulengerochromini and Perissodini. Evolutionary Biology (in press).
Rambaut, A. (2018). Figtree v1.4.4. Institute of Evolutionary Biology, University of Edinburgh, Edinburgh.
Reyda, F.B., Wells, S.M., Ermolenko, A.V., Ziętara, M.S., Lumme, J.I. (2020). Global parasite trafficking: Asian Gyrodactylus (Monogenea) arrived to the U.S.A. via invasive fish Misgurnus anguillicaudatus as a threat to amphibians. Biological Invasions, 22, 391–402.
Ronquist, F., Huelsenbeck, J.P. (2003). MRBAYES 3: Bayesian phylogenetic inference under mixed models. Bioinformatics, 19(12), 1572–1574.
Rosenthal, D.M., Ramakrishnan, A.P., Cruzan, M.B. 2008. Evidence for multiple sources of invasion and intraspecific hybridization in Brachypodium sylvaticum (Hudson) Beauv. In North America. Molecular Ecology. 17, 4657-4669.
Rozas, J., Ferrer-Mata, A., Sánchez-DelBarrio, J.C., Guirao-Rico, S., Librado, P., Ramos-Onsins, S.E., Sánchez-Gracia, A. (2017). DnaSP 6: DNA Sequence Polymorphism Analysis of Large Datasets. Molecular Biology and Evolution. https://doi.org/10.1093/molbev/msx248
Shipton, T., Tweddle, D., Watts, M. (2008). Species Risk Assessment: Introduction of the Nile Tilapia (Oreochromis niloticus) in the Eastern Cape. Eastern Cape Development Corporation report.
Šimková, A., Plaisance, L., Matĕjusová, I., Morand, S., Verneau, O. (2003). Phylogenetic relationships of the Dactylogyridae Bychowsky, 1933 (Monogenea: Dactylogyridea): the need for the systematic revision of the Ancyrocephalinae Bychowsky, 1937. Systematic Parasitology, 54(1), 1–11.
Šimková, A., Morand, S., Jobet, E., Gelnar, M., Verneau, O. (2004). Molecular phylogeny of congeneric monogenean parasites (Dactylogyrus): a case of intrahost speciation. Evolution, 58(5), 1001–1018.
Šimková, A., Řehulková, E., Rasoloariniaina, J.R., Jorissen, M.W.P., Scholz, T., Faltýnková, A., Mašová, Š., Vanhove, M.P.M. (2019). Transmission of parasites from introduced tilapias: a new threat to endemic Malagasy ichthyofauna. Biological Invasions, 21(3), 803–819.
Sinnappah, N.D., Lim, L.H.S., Rohde, K., Tinsley, R., Combes, C., Verneau, O. (2001). A paedomorphic parasite associated with a neotenic amphibian host: phylogenetic evidence suggests a revised systematic position for Sphyranuridae within anuran and turtle Polystomatoineans. Molecular Phylogenetics and Evolution, 18(2), 189–201.
Soler-Jimenez, L.C., Paredes-Trujillo, A.I., Vidal-Martinez, V.M. (2017). Helminth parasites of finfish commercial aquaculture in Latin America. Journal of Helminthology, 91(2), 110–136.
Stamatakis, A. (2014). RAxML version 8 : a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics, 9, 1312–1313.
Starling, F., Lazzaro, X., Cavalcanti, C., Moreira, R. (2002). Contribution of omnivorous tilapia to eutrophication of a shallow tropical reservoir: evidence from a fish kill. Freshwater Biology, 47, 2443-2452.
Stöver, B.C., Müller, K.F. (2010). TreeGraph 2: Combining and visualizing evidence from different phylogenetic analyses. BMC Bioinformatics, 11, 7 (2010).
Swofford, D.L. (2003). PAUP*. Phylogenetic Analysis Using Parsimony (*and other methods). Version 4. Sinauer Associates, Sunderland, Massachusetts.
Talavera, G., Castresana, J. (2007). Improvement of phylogenies after removing divergent and ambiguously aligned blocks from protein sequence alignments. Systematic Biology, 56(4), 564–577.
Teugels, G.G., Thys van den Audenaerde, D.F.E. (2003). Cichlidae. In Paugy, D., Lévêque, C., Teugels, G.G. (Eds.). The fresh and brackish water fishes of West Africa volume 2 (pp. 714–780). Paris, France and Tervuren, Belgium.
Thys Van Den Audenaerde, D.F.E. (1964). Révision systématique des espèces congolaises du genre Tilapia (Pisces, Cichlidae). Annales du Musée d’Afrique Central, Sciences Zoologiques, 124, 1–155.
Toguyeni, A. (2004). Tilapia production and its global impacts in Central African countries. 6th International Symposium on Tilapia in Aquaculture. Manila, Philippines.
Tollefson, J. (2019). Humans are driving one million species to extinction. Nature, 569(7755), 171.
Trewavas, E. (1983). Tilapiine fishes of the genera Sarotherodon, Oreochromis and Danakilia. British Natural History Museum.
Trewavas, E., Teugels, G.G. (1991). Oreochromis in: Daget, J., Gosse, J.-P., Teugels, G.G., Thys van den Audenaerde, D.F.E. (eds.). Checklist of the freshwater fishes of Africa (CLOFFA). Brussels, Belgium, Tervuren, Belgium and Paris, France.
Vaidya, G., Lohman, D.J., Meier, R. (2011). SequenceMatrix: concatenation software for the fast assembly of multi-gene datasets with character set and codon information. Cladistics, 27(2), 171–180.
Vanhove, M.P.M., Van Steenberge, M., Dessein, S., Volckaert, F.A.M., Snoeks, J., Huyse, T., Pariselle, A. (2013). Biogeographical implications of Zambezian Cichlidogyrus species (Platyhelminthes: Monogenea: Ancyrocephalidae) parasitizing Congolian cichlids. Zootaxa, 3608(5), 398–400.
Vanhove, M.P.M., Pariselle, A., Van Steenberge, M., Raeymaekers, J., Hablützel, P.I., Gillardin, C., Hellemans, B., Breman, F.C., Koblmüller, S., Sturmbauer, C., Snoeks, J., Volckaert, F.A.M., Huyse, T. (2015). Hidden biodiversity in an ancient lake: phylogenetic congruence between Lake Tanganyika tropheine cichlids and their monogenean flatworm parasites. Scientific Reports, 5, 13669.
Vanhove, M.P.M., Briscoe, A.G., Jorissen, M.P.M., Littlewood, D.T.J., Huyse, T. (2018). The first next-generation sequencing approach to the mitochondrial phylogeny of African monogenean parasites (Platyhelminthes: Gyrodactylidae and Dactylogyridae). BMC Genomics, 19, 520.
Vignon, M., Pariselle, A., Vanhove, M.P.M. (2011). Modularity in attachment organs of African Cichlidogyrus (Platyhelminthes: Monogenea: Ancyrocephalidae) reflects phylogeny rather than host specificity or geographic distribution. Biological Journal of the Linnean Society, 102, 694–706.
WoRMS (2021). Cichlidogyrus Paperna, 1960. Taxonomic list. World Register of Marine Species. http://www.marinespecies.org/aphia.php?p=taxdetails&id=517932. Accessed 1 March 2021.
Wu, X.Y., Zhu, X.Q., Xie, M.Q. & Li, A.X. (2007). The evaluation for generic-level monophyly of Ancyrocephalinae (Monogenea, Dactylogyridae) using ribosomal DNA sequence data. Molecular Phylogenetics and Evolution, 44, 530–544.
Zhang, J., Kapli, P., Pavlidis, P., Stamatakis, A. (2013). A general species delimitation method with applications to phylogenetic placements. Bioinformatics, 2013(22), 2869–2876.
Zhang, S., Zhi, T., Xu, X., Zheng, Y., Bilong Bilong, C.F., Pariselle, A., Yang, T. (2019). Monogenean fauna of alien tilapias (Cichlidae) in South China. Parasite, 26, 4.

Table 1. Overview of the sampling design with parasite species listed per host species and per region. Last column depicts “the number of sequenced specimens: 18S+ITS-1/28S/COI sequences”.

Host species	Generic name	Region	Introduced/native	Parasite species	#
Coptodon rendalli (Boulenger, 1897)	Redbreast tilapia	Lower Congo	introduced	C. berradae	4:1/4/0
		Upper Congo	native	C. dossoui, C. quaestio, C. tiberianus	1:1/1/1 2:0/2/0 2:1/2/0
		Madagascar	introduced	C. sp.2	2:0/2/0
Hemichromis stellifer Loiselle, 1979	/	Lower Congo	native	C. falcifer, C. polyenso	1:1/1/1 1:0/1/0
Oreochromis mweruensis Trewavas, 1983	Mweru tilapia	Upper Congo	native	C. cirratus, C. halli morphotype 2, C. papernastrema, C. tilapiae, S. gravivaginus	2:1/2/0 2:0/2/0 1:0/1/0 1:1/1/0 1:0/1/0
Oreochromis niloticus (Linnaeus, 1758)	Nile tilapia	Lake Tanganyika, Burundi	native	C. cf. halli ‘Burundi’, C. tilapiae	1:1/1/1 6:0/3/3
		Lower Congo	introduced	C. halli, C. sclerosus, C. thurstonae, C. tilapiae	14:6/9/6 3:0/1/3 34:5/17/16 22:5/13/10
		Middle Congo	introduced	C. halli, C. sclerosus, C. thurstonae, C. tilapiae	1:0/0/1 1:0/1/1 1:0/1/1 3:0/0/3
		Upper Congo	introduced	C. halli, C. sclerosus, C. thurstonae	8:0/0/8 6:0/0/6 1:0/0/1
		Madagascar	introduced	C. halli, C. sclerosus, C. thurstonae, C. tilapiae, S. longicornis	5:2/3/1 1:0/0/1 4:0/2/2 2:1/1/2 1:0/1/0
Sargochromis mellandi (Boulenger, 1905)	Snaileater	Upper Congo	native	C. consobrini, C. zambezensis	2:1/2/0 1:1/1/0
Serranochromis macrocephalus (Boulenger, 1899)	Purpleface largemouth	Middle Zambezi, Zimbabwe	native	C. zambezensis	3:1/3/0
Tilapia sparrmanii Smith, 1840	Banded tilapia	Upper Congo	native	C. dossoui, C. papernastrema	1:0/0/1 4:0/3/1

Table 2. List of primers used to amplify the gene fragments used in this study.

Gene	Primer	Direction	Sequence (5’-3’)	Reference
COI	ASmit1	F	TTTTTTGGGCATCCTGAGGTTTAT	Littlewood et al. 1997
COI	Cox1_Schisto_3	R	TAATGCATMGGAAAAAAACA	Lockyer et al. 2003
COI	ASmit2 (nested)	R	TAAAGAAAGAACATAATGAAAATG	Littlewood et al. 1997
18S+partialITS-1	S1	F	ATTCCGATAACGAACGAGACT	Sinappah et al. 2001
18S+partialITS-1	IR8	R	GCTAGCTGCGTTCTTCATCG	Šimková et al. 2003
28S	C1	F	ACCCGCTGAATTTAAGCAT	Hassouna et al. 1984
28S	D2	R	TGGTCCGTGTTTCAAGAC	Hassouna et al. 1984

Table 3. PCR conditions and master mix for the different molecular markers used in the phylogenetic analyses. Taq is Platinum Taq DNA polymerase by Invitrogen (PlatTaq).

Primer set

PCR condition

Mastermix

ASmit1 – Schisto3 (COI)

94 °C: 3 min

94 °C: 30 s

44 °C: 30 s 39 x

72 °C: 60 s

72 °C: 7 min

1X PCR Buffer

1.5 mM MgCl₂

0.2 mM dNTP mix

0.4 µM F-primer

0.6 µM R-primer

14.8 µl ddH₂O

1.2 µl Template

3 Units/reaction Taq

ASmit1 – ASmit2 (COI, nested PCR)

94 °C: 3 min

94 °C: 30 s

50 °C: 30 s 39 x

72 °C: 60 s

72 °C: 7 min

1X PCR Buffer

1.5 mM MgCl₂

0.2 mM dNTP mix

0.4 µM F-primer

0.4 µM R-primer

15 µl ddH₂O

1.2 µl Template

3 Units/reaction Taq

S1 – IR8 (18S + ITS-1)

94 °C: 2 min

94 °C: 60 s

50 °C: 60 s 39 x

72 °C: 90 s

72 °C: 10 min

1X PCR Buffer

1.5 mM MgCl₂

0.2 mM dNTP mix

0.4 µM F-primer

0.6 µM R-primer

14.8 µl ddH₂O

1.2 µl Template

3 Units/reaction Taq

C1 – D2 (28S)

94 °C: 2 min

94 °C: 20 s

50 °C: 30 s 39 x

72 °C: 90 s

72 °C: 10 min

1X PCR Buffer

1.5 mM MgCl₂

0.2 mM dNTP mix

0.4 µM F-primer

0.4 µM R-primer

15 µl ddH₂O

1.2 µl Template

3 Units/reaction Taq

Table 4. Reference sequences of Cichlidogyrus species used in the phylogenetic analyses.

Parasite	Host	Genbank Accession 18S+ITS-1	Genbank Accession 28S	Locality	Reference
Cichlidogyrus acerbus	Sarotherodon galilaeus	HE792780.2	HQ010036.2	Senegal	Mendlová et al. 2012, 2010
Cichlidogyrus aegypticus	Coptodon guineensis	HE792781.1	HQ010021.1	Senegal	Mendlová et al. 2012, 2010
Cichlidogyrus agnesi	Coptodon guineensis	AJ920286.1		Ebrie Lagoon, Ivory Coast	Pouyaud et al. 2006
Cichlidogyrus amphoratus	Coptodon guineensis	HE792782.1	HE792772.1	Senegal	Mendlová et al. 2012
Cichlidogyrus bilongi	Coptodon guineensis	AJ920287.1		Ebrie Lagoon, Ivory Coast	Pouyaud et al. 2006
Cichlidogyrus arthracantus	Coptodon guineensis	HE792783.1	HQ010022.1	Senegal	Mendlová et al. 2012, 2010
Cichlidogyrus casuarinus	Bathybates fasciatus	KX007788.1	KX007814.1	Uvira, Lake Tanganyika, DRC	Kmentová et al. 2016a
Cichlidogyrus cirratus	Oreochromis niloticus	HE792784.1	HE792773.1	Senegal	Mendlová et al. 2012
Cichlidogyrus cubitus	Coptodon guineensis	HE792785.1	HQ010037.1	Senegal	Mendlová et al. 2012, 2010
Cichlidogyrus digitatus	Coptodon guineensis	HE792786.1	HQ010023.1	Senegal	Mendlová et al. 2012, 2010
Cichlidogyrus douellouae	Sarotherodon galilaeus	HE792787.1	HE792774.1	Senegal	Mendlová et al. 2012
Cichlidogyrus ergensi	Coptodon guineensis	HE792788.1	HQ010038.1	Senegal	Mendlová et al. 2012, 2010
Cichlidogyrus falcifer	Hemichromis fasciatus	HE792789.1	HQ010024.1	Senegal	Mendlová et al. 2012, 2010
Cichlidogyrus flexicolpos	Coptodon guineensis	AJ920283.1		Ebrie Lagoon, Ivory Coast	Pouyaud et al. 2006
Cichlidogyrus gallus	Coptodon guineensis	AJ920285.1		Ebrie Lagoon, Ivory Coast	Pouyaud et al. 2006
Cichlidogyrus halli 1	Oreochromis niloticus x Oreochromis mweruensis	MG973075.1	MG973075.1	Mweru-Luapula	Vanhove et al. 2018
Cichlidogyrus halli 2	Sarotherodon galilaeus	HE792790.1	HQ010025.1	Senegal	Mendlová et al. 2012, 2010
Cichlidogyrus halli 3	Oreochromis niloticus	MH767392	MH767403.1	Madagascar	Šimkov et al. 2019
Cichlidogyrus irenae	Gnathochromis pfefferi	KT692939.1	MH708145.1	Mukuruka, Lake Tanganyika, Burundi	Kmentová et al. 2016b
Cichlidogyrus longicirrus	Hemichromis fasciatus	HE792791.1	HQ010026.1	Senegal	Mendlová et al. 2012, 2010
Cichlidogyrus njinei	Sarotherodon galilaeus	HE792792.1	HE792775.1	Senegal	Mendlová et al. 2012
Cichlidogyrus pouyaudi	Tylochromis intermedius	HE792793.1	HQ010039.1	Senegal	Mendlová et al. 2012, 2010
Cichlidogyrus sclerosus	Oreochromis niloticus	MH767390.1	MH767401.1	National Park Ankafarantsika Madagascar	Šimkov et al. 2019
Cichlidogyrus thurstonae	Oreochromis niloticus	MH767394.1	MH767406.1	Anjingo River, Antsohihy, Madagascar	Šimkov et al. 2019
Cichlidogyrus tiberianus	Coptodon guineensis	HE792796.1	HE792776.1	Senegal	Mendlová et al. 2012
Cichlidogyrus tilapiae	Hemichromis fasciatus	HE792797.1	HQ010029.1	Senegal	Mendlová et al. 2012, 2010
Cichlidogyrus yanni	Coptodon guineensis	HE792798.1	HE792777.1	Senegal	Mendlová et al. 2012
Scutogyrus bailloni	Sarotherodon galilaeus	HE792799.1	HE792778.1	Senegal	Mendlová et al. 2012
Scutogyrus longicornis	Oreochromis niloticus	HE792800.1	HQ010035.1	Senegal	Mendlová et al. 2012, 2010
Scutogyrus minus	Sarotherodon melanotheron	HE792801.1	HE792779.1	Senegal	Mendlová et al. 2012

Download PDF

Version 1

posted

You are reading this latest preprint version

Molecular Footprint of Parasite Co-introduction with Nile Tilapia in the Congo Basin

Status:

Version 1

Abstract

Figures

Introduction

Material & Methods

Results

Co-introduced parasites of Nile tilapia

Phylogenetic relationships among native and introduced tilapia parasites and evaluation of species status

Discussion

Co-introduced parasites of Nile tilapia

Marker performance and barcoding gap

Phylogenetic relationships among native and introduced tilapia parasites and evaluation of species status

Conclusion

Declarations

References

Tables

Supplementary Files

Status:

Version 1