Open Access
Issue
Parasite
Volume 31, 2024
Article Number 22
Number of page(s) 14
DOI https://doi.org/10.1051/parasite/2024021
Published online 10 April 2024

© M.B. Ebert et al., published by EDP Sciences, 2024

Licence Creative CommonsThis is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Introduction

Cichlids belonging to genera Oreochromis Günther, 1889 and Coptodon Gervais, 1853, commonly known as tilapias, are subtropical to tropical freshwater fishes that are native to Africa and the southwestern Middle East [11]. Over the past nine decades, tilapias have been intentionally dispersed worldwide to be used as biological control of aquatic weeds and insects, as baitfish in fisheries, as aquarium species, and as a source of food protein [11].

Tilapias have been translocated outside their native range and currently they are farmed in 79 countries; tilapia production in countries such as China, Indonesia and Bangladesh accounts for 66.76% of all tilapia grown [68]. In Brazil, two tilapia species are commonly found primarily on fish farms, although they have been fortuitously released to wild environments. The red-breasted tilapia, Coptodon rendalli (Boulenger, 1897) (= Chromis rendalli Boulenger, 1897; Tilapia rendalli (Boulenger, 1897)) was the first to be introduced to the country in the 1950s with the purpose of populating dams of electric energy companies in São Paulo state [13]; and a while later, the Nile tilapia Oreochromis niloticus (Linnaeus, 1758) (= Perca nilotica Linnaeus, 1758) was introduced as part of a federal government program against hunger [13]. Since then, the two species have been intensely farmed on fish farms and “fish and pay” establishments around the country, to promote developing economies and recreation [6, 7]. Even though Brazil is one of the world’s richest in the diversity of native ichthyofauna [64], tilapias are presently the main commercially cultivated species in national aquaculture, and the most consumed fishes in the country [13, 55, 74].

Since their introduction for aquacultural purposes, C. rendalli and O. niloticus have been intentionally and accidentally spread into several lakes, dams, and reservoirs [2, 54], and also into the basins of several rivers throughout the country, namely: Tietê [54], Paraná [9], Doce [72], Paranapanema [12, 44], Grande [79], Itajaí [23, 29], Uruguay [46], and even the Amazon [5, 55, 73]. The presence of non-native species such as C. rendalli and O. niloticus is a major threat to local biodiversity [70] since they can affect native biota not only through direct interactions, e.g., competition [15] but also through indirect interactions such as parasites which are usually co-introduced with their hosts [71]. A serious problem of co-introducing parasites is that they might switch hosts and successfully infect native species [24], increasing the risk of decline and/or extinction of native fauna [24, 71].

African cichlid fishes are known to harbour species contained in six genera of monogenean parasites; two of them are mesoparasites (i.e., Enterogyrus Paperna, 1963 and Urogyrus Bilong Bilong, Birgi & Euzet, 1994) which live in a host’s body cavity, and four are ectoparasites found on the gills or the body surface [21, 56] feeding on mucus, skin and possibly the blood of their hosts [25]. The monogenean gill parasites include three genera belonging to the Dactylogyridae Bychowsky, 1933, i.e., Cichlidogyrus Paperna, 1960, Onchobdella Paperna, 1968, and Scutogyrus Pariselle & Euzet, 1995, and one genus belonging to the Gyrodactylidae Cobbold, 1864, i.e., Gyrodactylus von Nordmann, 1832 [21, 56]. Cichlidogyrus is one of the most diverse genera of monogeneans parasitizing cichlid fishes and is found in more than 40 fish species across 11 different genera [60]. To date, 130 species of Cichlidogyrus are considered valid [76]. In contrast, Scutogyrus is presently represented by only seven species and known to be restricted to cichlids of the genera Sarotherodon Rüppell, 1852 and Oreochromis [43, 69], although S. longicornis has been reported parasitizing C. rendalli in Brazil [24, 77].

Among other monogenean parasites (see [18, 55, 68, 72]), species of Cichlidogyrus and Scutogyrus have been commonly reported parasitizing tilapias in Brazil: O. niloticus was found infested with Cichlidogyrus tilapiae Paperna, 1960 [4, 8, 19, 41, 49], Cichlidogyrus halli (Price & Kirk, 1967) (= Cleidodiscus halli Price & Kirk, 1967) [8, 18, 29, 34, 79], Cichlidogyrus thurstonae Ergens, 1981 [8, 18, 23, 29, 34, 46, 79] Cichlidogyrus sclerosus [3, 8, 18, 23, 26, 29, 34, 44, 45, 69], Cichlidogyrus rognoni Pariselle, Bilong Bilong & Euzet, 2003 [8, 18], and Scutogyrus longicornis (Paperna & Thurston, 1969) (= Cichlidogyrus longicornis Paperna & Thurston, 1969) [8, 18, 29, 34, 45, 79]. As for C. rendalli, only C. tilapiae, Cichlidogyrus sp. and S. longicornis were reported to parasitize this species in Brazil [26, 73]. These monogeneans are native to the African continent [56] and the presence of these species in wild populations in the freshwaters of Brazil is strong evidence of their co-introduction into the country along with their fish hosts.

This study aimed to report the monogenean parasites found on the gills of the introduced cichlid fishes C. rendalli and O. niloticus from the Pardo River, Paranapanema River basin, São Paulo state, Brazil. The morphology and phylogenetic relationships using the LSU rDNA gene of four species of Cichlidogyrus and one species of Scutogyrus found on the gills of the fishes were investigated. The ecological consequences of the presence of exotic species such as C. rendalli and O. niloticus in a river spring are also discussed.

Material and methods

Host sampling, parasitological procedures and ethics

During an inventory of fishes and parasites in the Pardo River, Paranapanema River basin, São Paulo state, Brazil, 20 specimens of red-breasted tilapia C. rendalli and 10 specimens of Nile tilapia O. niloticus were collected near the river spring (23°0′19.00″ S; 48°22′34.25″ W), municipality of Botucatu, in June 2021. Tilapias were introduced into the region by farm owners who, without proper technical knowledge, attempted to produce fishes for commercial purposes or other reasons.

The fishes were collected using casting nets and euthanized with sodium thiopental (Thiopentax®). The specimens were then individually stored in plastic bags and frozen to later conduct necropsy at the laboratory, while others were examined in situ to collect fresh monogeneans, which were placed into 96% molecular-grade ethanol for molecular analyses. At the laboratory, gills were removed, placed in Petri dishes with tap water, and analyzed for monogeneans under a stereomicroscope. Monogeneans were counted, separated from the gills, and mounted on slides with Hoyer and Gray and Wess’ medium [41].

The morphology of the sclerotized structures of the haptor (bar, anchors, and hooks) and copulatory complex of the monogeneans was analyzed using a V3 Leica Application Suite (LAS) computerized system for image analysis adapted to a microscope with differential interference contrast. Measurements of the specimens (total body length, haptoral characters (bars, anchors, and hooks), heel, penis, accessory piece of the male copulatory organ, and vagina) were made following Euzet & Prost [16] and Jorissen et al. [32]. Ecological data such as the prevalence, abundance, and mean intensity of infestation, were calculated following Bush et al. [10].

Representative specimens of the monogeneans were deposited in the Helminthological Collection of the Institute of Biosciences (CHIBB), Botucatu, São Paulo state, Brazil, under numbers: S. longicornis 740L; C. mbirizei 741L -744L; C. thurstonae 745L-749L; C. tilapiae 750L-754L; C. papernastrema 755L-759L. Representative fish hosts were deposited in the Fish Collection of the Laboratory of Fish Biology and Genetics (LBP) at the São Paulo State University (UNESP), Botucatu, Brazil, under numbers 33636–33638. Fishes were collected under the authorization of the Instituto Chico Mendes de Conservação da Biodiversidade (SISBIO #60640-1). All procedures followed the recommendations and approval of the Ethics Committee for Animal Experimentation of the São Paulo State University (UNESP), Institute of Biosciences, Botucatu, Brazil (CEUA 9415260520). According to Brazilian laws, species registration for scientific research purposes was carried out at SisGen (AD05367).

Molecular and phylogenetic analyses

Genomic DNA of the monogeneans was extracted using a DNeasy Blood & Tissue Kit (QIAGEN, Valencia, CA, USA), following the manufacturer’s protocol. Fragments of the LSU rDNA gene were amplified using the primers 382F (5′-AGCTGGTGGAGTCAAGCTTC-3′) and 1289R (5′-TGCTCACGTTTGACGATCGA-3′) [17], using the following cycling conditions: initial denaturation of 5 min at 95 °C followed by 40 cycles of 95 °C for 30 s, 56 °C for 30 s, 72 °C for 2 min, and a final extension of 10 min at 72 °C [17]. Conventional polymerase chain reaction (PCR) amplifications were performed on a final volume of 25 μL containing 12.5 μL of 2× MyFiTM Mix (Bioline, Taunton, MA, USA), 3.0 μL of extracted DNA, 7.5 μL of pure water and 1.0 μL of each PCR primer. PCR products (2.0 μL) were run on an agarose gel (1%) using GelRedTM fluorescent nucleic acid dye and loading buffer to confirm amplicon size and yield. PCR amplicons were purified using the QIAquick PCR Purification Kit (QIAGEN), following the manufacturer’s instructions. Automated sequencing was performed directly on purified PCR products using a BigDye v.3.1 Terminator Cycle Sequencing Ready Reaction kit on an ABI 3500 DNA genetic sequencer (Applied Biosystems, Waltham, MA, USA). Forward and reverse sequences were assembled and edited using Sequencher v. 5.2.4 (Gene Codes, Ann Arbor, MI, USA).

To investigate the phylogenetic position of our newly generated partial LSU rDNA sequences, a dataset was created and complemented with published sequences of Cichlidogyrus spp. and Scutogyrus spp. retrieved from GenBank (Table 1). Cichlidogyrus pouyaudi Pariselle & Euzet, 1994, Cichlidogyrus berrebi Pariselle & Euzet, 1994 and Cichlidogyrus kothiasi Pariselle & Euzet, 1994 were used as outgroups because of their basal position assumed in the phylogenetic tree delivered by Mendlová et al. [49], Cruz-Laufer et al. [14]. Alignment of the dataset was performed using the MUSCLE algorithm implemented on Geneious 7.1.3 [36] with default settings. The best-fitting model of nucleotide substitution for the aligned dataset was selected in the JModelTest software [58] using the Akaike information criterion, as GTR+I+G.

Table 1

List of monogeneans included in the phylogenetic analyses, with details of the host, locality, GenBank accession numbers of sequences from the LSU rDNA gene, and their references. New sequences obtained for the present study are in bold.

Phylogenetic trees were obtained using Bayesian Inference (BI) and Maximum Likelihood (ML). The BI analysis was performed using MrBayes 3.2 [66] on the online platform CIPRES [52]. The Markov chain Monte Carlo (MCMC) was run with 106 generations saving one tree every 100 generations, with a burn-in set to the first 25% of the trees. Only nodes with posterior probabilities (pp) greater than 0.90 were considered well-supported. The ML analysis was run in RAxML [27] at the online platform CIPRES [52] with 1000 bootstrap replicates. Only nodes with bootstrap values greater than 70 were considered well-supported. The BI and ML trees were visualized in FigTree v. 1.3.1 software [62] and edited in CorelDRAW X6. Pairwise genetic distances among and between sequences were calculated using the Kimura-2-parameter (K2P) model and a bootstrap procedure with 1000 replicates in the MEGA11 program [37].

Results

Morphological analyses

The 10 analyzed O. niloticus (mean length: 7.26 [7.4–19.7] cm; mean weight: 10.8 [6.97–164.28] g) were infested with three species of Cichlidogyrus, i.e., Cichlidogyrus mbirizei Muterezi Bukinga, Vanhove, Van Steenberge & Pariselle, 2012 (Fig. 1), C. thurstonae (Fig. 2), and C. tilapiae (Fig. 3), and one species of Scutogyrus, i.e., S. longicornis (Fig. 4). As for the 20 analyzed C. rendalli (mean length: 7.18 [3.2–13.0] cm; mean weight: 13.54 [4.5–23.0] g), we only found Cichlidogyrus papernastrema Price, Peebles & Bamford, 1969 (Fig. 5). All monogeneans were identified based on the morphology of the reproductive organs and the sclerotized parts of the haptor, which is characterized by baring two pairs of anchors, a V-shaped ventral transversal bar, and a dorsal transversal bar with two auricles and seven pairs of marginal hooks. Measurements of the collected specimens are provided in Table 2.

thumbnail Figure 1

Cichlidogyrus mbirizei Muterezi Bukinga, Vanhove, Van Steenberge & Pariselle, 2012 from the gills of Oreochromis niloticus (Linnaeus, 1758) from Pardo River, Paranapanema River basin, São Paulo state, Brazil. A. Haptor; B. Male copulatory organ; C. Vagina. Hoyer’s mounting medium. Legend: ap, accessory piece; da, dorsal anchor, db, dorsal bar; I–VII, hook pairs; pe, penis; va, ventral anchor; vb, ventral bar; and vg, vagina.

thumbnail Figure 2

Cichlidogyrus thurstonae Ergens, 1981 from the gills of Oreochromis niloticus (Linnaeus, 1758) from Pardo River, Paranapanema River basin, São Paulo state, Brazil. A. Haptor; B. Male copulatory complex. Hoyer’s mounting medium. Legend: ap, accessory piece; da, dorsal anchor; db, dorsal bar; I–VII, hook pairs; pe, penis; va, ventral anchor; vb, ventral bar; and vg, vagina.

thumbnail Figure 3

Cichlidogyrus tilapiae Paperna, 1960 from the gills of Oreochromis niloticus (Linnaeus, 1758) from Pardo River, Paranapanema River basin, São Paulo state, Brazil. A. Haptor; B. Male copulatory complex. Hoyer’s mounting medium. Legend: ap, accessory piece; da, dorsal anchor; db, dorsal bar; I–VII, hook pairs; pe, penis; va, ventral anchor; and vb, ventral bar.

thumbnail Figure 4

Scutogyrus longicornis (Paperna & Thurston, 1969) from the gills of Oreochromis niloticus (Linnaeus, 1758) from Pardo River, Paranapanema River basin, São Paulo state, Brazil. A. Haptor; B. Male copulatory complex; C. Vagina. Hoyer’s mounting medium. Legend: ap, accessory piece; da, dorsal anchor; db, dorsal bar; I–VII, hook pairs; pe, penis; va, ventral anchor; vb, ventral bar; and vg, vagina.

thumbnail Figure 5

Cichlidogyrus papernastrema Price, Peebles & Bamford, 1969 from the gills of Coptodon rendalli (Boulenger, 1897) from Pardo River, Paranapanema River basin, São Paulo state, Brazil. A. Haptor; B. Male copulatory complex. Hoyer’s mounting medium. Legend: ap, accessory piece; da, dorsal anchor; db, dorsal bar; I–VII, hook pairs; pe, penis; va, ventral anchor; and vb, ventral bar.

Table 2

Measurements of Cichlidogyrus spp. Paperna, 1960 and Scutogyrus longicornis (Paperna & Thurston, 1969) collected from Oreochromis niloticus (Linnaeus, 1758) and Coptodon rendalli (Boulenger, 1897) (Cichlidae) in a river spring in Brazil. Measurements are represented in μm as the average, the range in parentheses, and the count in brackets.

Cichlidogyrus papernastrema reached the highest prevalence (P = 60.0%), although it was found only in C. rendalli, followed by C. thurstonae (P = 40.0%) found in O. niloticus. However, greater abundance (A = 2.0) and intensity of infestation (MII = 10.0) were observed for C. tilapiae found in O. niloticus. The ecological parameters of infection of the monogeneans found on the gills of C. rendalli and O. niloticus are presented in Table 3.

Table 3

Ecological data of monogeneans found on the gills of Coptodon rendalli (Boulenger, 1897) and Oreochromis niloticus (Linnaeus, 1758) in ponds from a river spring in Brazil. P%, prevalence as percentage; A, mean abundance; MII, mean intensity of infestation.

Molecular analyses

We successfully obtained 15 partial sequences of the LSU rDNA gene: nine from specimens of C. papernastrema (GenBank accession numbers PP477261PP477269), four from specimens of C. tilapiae (GenBank accession numbers PP477257PP477260), one from a specimen of C. thurstonae (GenBank accession number PP477271), and one from a specimen of S. longicornis (GenBank accession number PP477270). Unfortunately, we were not able to obtain good quality sequences from C. mbirizei.

The final partial LSU rDNA alignment comprised a total of 74 sequences of Cichlidogyrus spp. and Scutogyrus spp. and, after trimming to the shortest sequence, it was 405 bp long. The BI and ML analyses of the partial LSU rDNA alignment produced phylograms with similar topologies in which the differences were on the level of poorly supported clades, especially on basal branches, and/or unresolved interspecific relationships. The BI topology is shown (Fig. 6), with the BI posterior probabilities (pp) followed by ML bootstrap support values.

thumbnail Figure 6

Bayesian topology based on partial LSU rDNA sequences of Cichlidogyrus Paperna, 1960 and Scutogyrus Pariselle & Euzet, 1995 species. GenBank accession numbers are after species names. Newly sequenced species are in bold. The support values are included above the nodes as follows: posterior probabilities for BI analysis, followed by bootstraps for the ML analysis. Only nodes with posterior probabilities >0.90 and bootstrap scores >70 are considered well-supported. Dashes before nodes represent clades that were not recovered by both analyses. Branch length scale bar indicates the number of substitutions per site.

Both BI and ML analyses recovered Cichlidogyrus spp. as a non-monophyletic group as it included the sequences of Scutogyrus spp., which were recovered as monophyletic within Cichlidogyrus (pp = 0.95; bootstrap = 76). The newly generated sequences of C. tilapiae (pp = 1; bootstrap = 100), C. thurstonae (pp = 1; bootstrap = 89), and S. longicornis (pp = 0.22; bootstrap = 95) were each grouped together with their conspecific sequences available in the database. Our newly generated sequences of C. papernastrema also grouped in a monophyletic clade together with the sequences of C. papernastrema (MW580347) collected from C. rendalli and C. papernastrema (OM720076) collected from Tilapia sparrmanii Smith, 1840 (pp = 1; bootstrap = 93) (referred to here as C. papernastrema “sensu stricto” clade); however, the sequence of C. papernastrema (OM720075) collected from T. sparrmanii was placed as the sister taxon to the clade formed by C. papernastrema “sensu stricto” + Cichlidogyrus zambezensis Douëllou, 1993 (MW580361) + Cichlidogyrus philander Douëllou, 1993 (MG279691) (pp = 1; bootstrap = 99).

Among the sequences of C. papernastrema “sensu stricto”, the maximum intraspecific LSU rDNA genetic divergence was 1.4%. The minimum LSU rDNA genetic divergence between the sequence C. papernastrema OM720075 and C. papernastrema “sensu stricto” was 2.4% (C. papernastrema OM720075 × C. papernastrema OM720076), while the maximum LSU rDNA genetic divergence was 3.4% (C. papernastrema OM720075 × C. papernastrema PP477268).

The interspecific LSU rDNA genetic divergences found between the sequence of C. papernastrema OM720075 and C. zambezensis was 2.4%, whereas the divergence with C. philander was 2.0%. In contrast, the interspecific LSU rDNA genetic divergence between C. papernastrema “sensu stricto”, and C. zambezensis ranged from 3.6% to 4.3%, and concerning C. philander, it ranged from 4.0% to 4.7%. The interspecific LSU rDNA genetic divergence between C. zambezensis and C. philander was 2.1%.

The LSU rDNA genetic divergence within the sequences of C. papernastrema “sensu stricto” and the sequences of C. tilapiae was 0.3%. No intraspecific genetic variation for the LSU rDNA region was found within the sequences of C. thurstonae and S. longicornis.

Discussion

The individuals of O. niloticus analyzed in this study were infested with four monogenean species; three of them, i.e., C. tilapiae, C. thurstonae, and S. longicornis, are commonly reported parasitizing the Nile tilapia in Brazil [3, 4, 23, 26, 29, 34, 44, 45, 55, 69, 79]. In contrast, C. mbirizei is reported for the first time in the country. Cichlidogyrus mbirizei was first described by Muterezi Bukinga et al. [53] infesting the gills of O. tanganicae (Günther, 1894) in the Democratic Republic of the Congo. Later, the species was found in the Nile tilapia and its hybrids O. niloticus × mossambicus in Thailand [42], in O. niloticus and Oreochromis sp. in Malaysia [1, 43] and Oreochromis mweruensis Trewavas, 1983 in Africa [35, 75]. In C. rendalli, the only species of Cichlidogyrus found was C. papernastrema, which is also reported for the first time in Brazil. Cichlidogyrus papernastrema was first described from Tilapia sparrmanii in South Africa [60]. After that, a few recent studies have registered C. papernastrema on the gills of C. rendalli [14, 31, 33], T. sparrmanii [31], and O. mweruensis [31, 33], all in the Democratic Republic of Congo. The fact that C. mbirizei and C. papernastrema have mostly been registered on the African continent reinforces the hypothesis of their co-introduction into Brazil along with their fish hosts brought from Africa, despite some C. mbirizei also having been registered in Thailand and Malaysia. In light of that, here we extend the geographical range of two monogenean species, i.e., C. mbirizei and C. papernastrema and confirm the assessment by Shinn et al. [68] regarding the translocation of parasites along with tilapias from their native range in Africa.

Our phylogenetic results agree with previously published analyses on Cichlidogyrus spp. and Scutogyrus spp., in which Cichlidogyrus is non-monophyletic, with Scutogyrus forming a monophyletic group nested within it [14, 33, 49, 50, 59, 78]. The representative sequences of Cichlidogyrus and Scutogyrus used in our analyses formed several supported/unsupported monophyletic groups, showing that the relationships between congeners are still unresolved and require further analyses. Cruz-Laufer et al. [14] provided a phylogeny based on a four-locus multiple alignment (partial ITS1, SSU rDNA, LSU rDNA, and COI mtDNA sequence data) along with different comparative methods and parameters (see Cruz-Laufer et al. [14] for specific methodology), which recovered similar topology compared to ours albeit with moderate to well-supported clades within Cichlidogyrus. This difference was probably generated by the increased set of molecular markers and methods used in those authors’ analyses. It is advisable that future research encompasses datasets with an increased number of DNA sequences from a more complete taxon coverage and molecular markers to provide a detailed picture of the evolution of the genus. Despite that, our phylogenetic results were useful for the correct identification of our Cichlidogyrus and Scutogyrus specimens, in combination with a detailed morphological analysis. The newly generated sequences are the first from the American continent and the sequences of C. tilapiae, C. thurstonae, and S. longicornis formed reciprocally monophyletic clades with their correspondent conspecific sequences available in the GenBank database from other localities. Interestingly, our molecular results recovered sequences of C. papernastrema as a non-monophyletic assemblage, casting doubts on the correct identification of some of the sequences deposited in GenBank; this requires further verification (see “Results” section; Fig. 6).

The maximum intraspecific LSU rDNA genetic divergence found among our sequences of C. papernastrema “sensu stricto” was 1.4%, which is in agreement with the maximum intraspecific LSU rDNA distance values found among specimens of Cichlidogyrus spp. (1.4%) shown by Jorissen et al. [33]. Our results also demonstrated that the minimum interspecific genetic distance between the sequences of C. papernastrema “sensu stricto” and C. papernastrema OM720075 (2.4%) is larger than the genetic distance found between C. zambezensis and C. philander (2.1%). Therefore, it is possible that the specimens from which the sequences of C. papernastrema “sensu stricto” were generated and the specimen from which the sequence C. papernastrema OM720075 was obtained might represent separate species. This hypothesis, previously suggested by Jorissen et al. [33], is confirmed here considering both our phylogenetic trees and genetic distances.

The morphological measurements from our C. papernastrema specimens are congruent with the data presented by Price [60] for the holotype of C. papernastrema collected in T. sparrmanii from South Africa, and by Jorissen et al. [32] for C. papernastrema collected in T. sparrmanni and C. rendalli from the Democratic Republic of Congo (Supplementary Table 1). Therefore, we propose that all these specimens are conspecific. Nevertheless, Jorissen et al. [32] noted that the C. papernastrema specimens collected in O. mweruensis from the Democratic Republic of Congo presented some morphological differences mostly related to the size of anchors and bars and the total length of the body, which the authors considered as intraspecific variation possibly related to host specificity. Future studies should include sequences of C. papernastrema collected from different host localities to test this hypothesis. Moreover, Jorissen et al. [31, 32] noted a large variation in the thickness of the copulatory tube in specimens of C. papernastrema, suggesting that such variation should be further investigated as a possible diagnostic character to delineate species along with genetic analyses [33].

Furthermore, several studies addressing monogenean parasites of C. rendalli and O. niloticus in Brazil have reported unidentified Cichlidogyrus sp. [3, 23, 26, 44, 45, 63, 73, 79] which could not be identified based on the morphology. Difficulties related to feasible and/or correct identification of monogeneans, such as the minute size of the parasites and/or inefficient collection and preservation methods, could be compromising the specific identification of those specimens since these monogeneans are predominantly diagnosed based on the morphology of their sclerotized parts from the reproductive organs and the attachment organ (haptor). Therefore, genetic information is very useful to confirm their taxonomic status. We propose that many Cichlidogyrus sp. found in previous studies could now be correctly identified at the species level, especially with the aid of molecular approaches. This is the case of C. papernastrema and C. mbirizei, which are added here to the bulk list of Cichlidogyrus species found in Brazil. Also, the fact that only one monogenean species, i.e., C. papernastrema, was found infesting C. rendalli, which is in contrast with the scarce literature on parasites of this cichlid species in Brazil [26, 73], highlights that possible misidentifications on both hosts and parasites might still represent a challenge yet to be overcome regarding parasitological studies in Brazil. The addition of new molecular data is still highly appreciated and contributes substantially to the phylogenetic interrelationships of the group.

The tilapias C. rendalli and O. niloticus have been translocated around the world, including to Brazil, for different purposes [11, 14, 68]. Despite the benefits to human society, tilapia aquaculture and open-water introductions can be very prejudicial to the native environment [11], since these fishes are highly invasive species and exist under feral conditions. The tilapias C. rendalli and O. niloticus may disrupt ecological native fish communities by competing and feeding on their resources, maintaining a strong territorial behaviour with multiple and expanded spawning, and presenting strong parental care [2, 5, 11, 57]. Moreover, as fishes with great food plasticity and omnivorous feeding habits, C. rendalli and O. niloticus can feed on zooplankton, phytoplankton, detritus, sediments, insect larvae, eggs, and even young fish of native species [2]. This can be extremely problematic when it comes to an area near a river spring, as is the case of our study. River springs are known as important breeding sites and refuges for several species of native biota (fishes, insects, snails, etc.), especially in the early stages of their development. The presence of C. rendalli and O. niloticus in an area of a river spring can adversely impact ecosystem services and cause a drastic decline in native biodiversity. In fact, many studies have already demonstrated the negative impact of the introduction of tilapias on the native fauna [2, 5, 11, 61]. For example, the invasion of Oreochromis sp. in lakes of Nicaragua reduced greatly the native cichlid populations due to environmental competition [47]. In Panama, the introduction of O. niloticus led to the extinction of two endemic species of cichlids, due to several factors including diseases and loss of habitat [65]. Lastly, in Brazilian public reservoirs, the dominance of the Nile tilapia O. niloticus has drastically altered the composition of the native fish populations, jeopardizing local artisanal fishing [2, 54].

Another serious risk to native freshwater fishes is the introduction of exotic fishes such as C. rendalli and O. niloticus which may also introduce alien parasites associated with their hosts [63, 68]. The problem with alien parasites is that they might switch hosts and successfully infect native species [24], increasing the risk of extinction of native fauna. Tilapia parasite host-switching events to native hosts (spillover effect) have already been reported for cichlid, poeciliid, and goodeid fishes in Mexico (Cichlidogyrus, Gyrodactylus and Enterogyrus species) [19, 20, 30, 67, 71], for cichlid fishes from America (C. sclerosus, C. tilapiae, S. longicornis, and Enterogyrus malmbergi Bilong Bilong, 1988) [30, 67] and aplocheilid fishes from Madagascar (C. tilapiae) [71]. There is also evidence of native parasite fauna infecting introduced tilapias Oreochromis mossambicus (Peters, 1852) in Colombia [40]. Moreover, there are documented host-switching events of dactylogyrids from the Nile tilapia being transferred towards native tilapias on their native continent [22]. Although the transmission of monogenean parasites to new hosts is poorly reported, it is evident that dactylogyrids that manage to establish outside their native range can exploit a phylogenetically broad host range [68].

The scenario presented here regarding the introduction of tilapias into wild environments and the potential spillover of their parasites may cause further damage to native fish species and biodiversity in the studied area, unless the growth of tilapia populations is controlled. We strongly recommend additional ecological and parasitological studies to better understand the role of tilapia and their parasites. There is also an imminent need for the implementation of a management plan to control this alien species, to prevent the extinction of native species in an area of a river spring due to horizontal transmission events among fishes.

Acknowledgments

This study was supported by the Pro-Rectory of Research (PROPG/PROPe – UNESP, 04/2022 granted to M.B.E); FAPESP – São Paulo Research Foundation (M.B.E.: 2021/12779-9; R.B.N.: 2019/26831-2; D.H.M.D.V.: 2019/19060-0; R.J.S.: 2020/05412-9); CNPq – National Council for Scientific and Technological Development (M.M.P.O.: 161838/2021-9; M.J.: 161839/2021-5; R.J.S.: 311635-2021/0). GPPL was supported by the Consejo Nacional de Ciencia y Tecnología (CONAHCYT A1-S-21694) and the Programa de Apoyo a Proyectos de Investigación e Innovación Tecnológica (PAPIIT-UNAM IN212621). RJS and GPPL were also supported by CAPES/PRINT (#88887.839573/2023-00 and #88887.839159/2023-00, respectively). Thanks to UNESP Pro-Rectory of Research for the financial support for the manuscript publication fees.

Conflict of interests

The authors declare that they have no conflict of interest.

Ethics approval

All procedures followed the recommendations and approval of the Ethics Committee for Animal Experimentation of the São Paulo State University (UNESP), Institute of Biosciences, Botucatu, Brazil (CEUA 9415260520).

Supplementary material

Supplementary Table 1: Measurements of Cichlidogyrus papernastrema Price, Peebles & Bamford, 1969. Measurements are represented in μm as the average, the range in parentheses, and the count in brackets († = doubled measurements of both structures of the same individual). * = Holotype. Access here

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Cite this article as: Ebert MB, Narciso RB, Dias DHMV, Osaki-Pereira MM, Jorge M, Pérez-Ponce de León G & Silva RJ. 2024. Parasites (Monogenea) of tilapias Oreochromis niloticus and Coptodon rendalli (Cichlidae) in a river spring in Brazil. Parasite 31, 22.

All Tables

Table 1

List of monogeneans included in the phylogenetic analyses, with details of the host, locality, GenBank accession numbers of sequences from the LSU rDNA gene, and their references. New sequences obtained for the present study are in bold.

Table 2

Measurements of Cichlidogyrus spp. Paperna, 1960 and Scutogyrus longicornis (Paperna & Thurston, 1969) collected from Oreochromis niloticus (Linnaeus, 1758) and Coptodon rendalli (Boulenger, 1897) (Cichlidae) in a river spring in Brazil. Measurements are represented in μm as the average, the range in parentheses, and the count in brackets.

Table 3

Ecological data of monogeneans found on the gills of Coptodon rendalli (Boulenger, 1897) and Oreochromis niloticus (Linnaeus, 1758) in ponds from a river spring in Brazil. P%, prevalence as percentage; A, mean abundance; MII, mean intensity of infestation.

All Figures

thumbnail Figure 1

Cichlidogyrus mbirizei Muterezi Bukinga, Vanhove, Van Steenberge & Pariselle, 2012 from the gills of Oreochromis niloticus (Linnaeus, 1758) from Pardo River, Paranapanema River basin, São Paulo state, Brazil. A. Haptor; B. Male copulatory organ; C. Vagina. Hoyer’s mounting medium. Legend: ap, accessory piece; da, dorsal anchor, db, dorsal bar; I–VII, hook pairs; pe, penis; va, ventral anchor; vb, ventral bar; and vg, vagina.

In the text
thumbnail Figure 2

Cichlidogyrus thurstonae Ergens, 1981 from the gills of Oreochromis niloticus (Linnaeus, 1758) from Pardo River, Paranapanema River basin, São Paulo state, Brazil. A. Haptor; B. Male copulatory complex. Hoyer’s mounting medium. Legend: ap, accessory piece; da, dorsal anchor; db, dorsal bar; I–VII, hook pairs; pe, penis; va, ventral anchor; vb, ventral bar; and vg, vagina.

In the text
thumbnail Figure 3

Cichlidogyrus tilapiae Paperna, 1960 from the gills of Oreochromis niloticus (Linnaeus, 1758) from Pardo River, Paranapanema River basin, São Paulo state, Brazil. A. Haptor; B. Male copulatory complex. Hoyer’s mounting medium. Legend: ap, accessory piece; da, dorsal anchor; db, dorsal bar; I–VII, hook pairs; pe, penis; va, ventral anchor; and vb, ventral bar.

In the text
thumbnail Figure 4

Scutogyrus longicornis (Paperna & Thurston, 1969) from the gills of Oreochromis niloticus (Linnaeus, 1758) from Pardo River, Paranapanema River basin, São Paulo state, Brazil. A. Haptor; B. Male copulatory complex; C. Vagina. Hoyer’s mounting medium. Legend: ap, accessory piece; da, dorsal anchor; db, dorsal bar; I–VII, hook pairs; pe, penis; va, ventral anchor; vb, ventral bar; and vg, vagina.

In the text
thumbnail Figure 5

Cichlidogyrus papernastrema Price, Peebles & Bamford, 1969 from the gills of Coptodon rendalli (Boulenger, 1897) from Pardo River, Paranapanema River basin, São Paulo state, Brazil. A. Haptor; B. Male copulatory complex. Hoyer’s mounting medium. Legend: ap, accessory piece; da, dorsal anchor; db, dorsal bar; I–VII, hook pairs; pe, penis; va, ventral anchor; and vb, ventral bar.

In the text
thumbnail Figure 6

Bayesian topology based on partial LSU rDNA sequences of Cichlidogyrus Paperna, 1960 and Scutogyrus Pariselle & Euzet, 1995 species. GenBank accession numbers are after species names. Newly sequenced species are in bold. The support values are included above the nodes as follows: posterior probabilities for BI analysis, followed by bootstraps for the ML analysis. Only nodes with posterior probabilities >0.90 and bootstrap scores >70 are considered well-supported. Dashes before nodes represent clades that were not recovered by both analyses. Branch length scale bar indicates the number of substitutions per site.

In the text

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