Hostname: page-component-89b8bd64d-rbxfs Total loading time: 0 Render date: 2026-05-07T11:11:40.658Z Has data issue: false hasContentIssue false
  • English
  • Français

First report of a dactylogyrid, Cichlidogyrus sclerosus Paperna & Thurston, 1969 (Monogenoidea) infecting Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) (Cichliformes: Cichlidae) in the United States, with a review of host and locality records in its invasive range and a phylogenetic analysis

Published online by Cambridge University Press:  18 February 2025

J.H. Brule Open the ORCID record for J.H. Brule [Opens in a new window] ,
M.B. Warren  and
S.A. Bullard
J.H. Brule*
Affiliation:
Southeastern Cooperative Fish Parasite and Disease Laboratory, School of Fisheries, Aquaculture, and Aquatic Sciences, College of Agriculture, Auburn University, Auburn, Alabama, 36849, USA
M.B. Warren
Affiliation:
Southeastern Cooperative Fish Parasite and Disease Laboratory, School of Fisheries, Aquaculture, and Aquatic Sciences, College of Agriculture, Auburn University, Auburn, Alabama, 36849, USA
S.A. Bullard
Affiliation:
Southeastern Cooperative Fish Parasite and Disease Laboratory, School of Fisheries, Aquaculture, and Aquatic Sciences, College of Agriculture, Auburn University, Auburn, Alabama, 36849, USA Department of Zoology, School for Environmental Sciences and Development, North-West University, Private Bag X6001, Potchefstroom, 2520, South Africa
*
Corresponding author: J.H. Brule; Email: jhb0088@auburn.edu
Rights & Permissions [Opens in a new window]

Abstract

The parasites of Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) are poorly documented in the United States despite the economic importance and global introduction of this African fish. Only one metazoan parasite (Gyrodactylus cichlidarum Paperna, 1968; Gyrodactylidae) reportedly infects Nile tilapia in the United States. Examining Nile tilapia from a flow-through aquaculture system hydrologically linked to Sougahatchee Creek (Tallapoosa River, Auburn, Alabama), we observed a gill infection by Cichlidogyrus sclerosus Paperna & Thurston, 1969 (Dactylogyridae). This monogenoid was originally described from the gill of Mozambique tilapia, Oreochromis mossambicus (Peters, 1852) from Lake Victoria, Uganda. Specimens of C. sclerosus were studied for morphology and phylogenetic analyses using the 28S and ITS1. We identified our specimens as C. sclerosus because they had the following combination of morphological features: marginal hooks shorter than dorsal anchor length; anchor roots reduced; dorsal anchor point bent; dorsal bar pyriform projections approximately half as long as dorsal bar width; penis short (<100 μm), not coiled, tubular, lacking swelling, having irregularly surfaced heel; and accessory piece straight and bifid. Our 28S and ITS1 phylogenies recovered our C. sclerosus sequences in a clade with conspecific sequences and showed no obvious biogeographic pattern. Cichlidogyrus sclerosus reportedly infects 21 fishes of 11 genera and 3 families from 36 countries in Africa, Asia, North America, South America, and Europe. The study of Nile tilapia parasites, especially those exhibiting direct life cycles and low host specificity, is important because they comprise potential invasive species.

Keywords

Exoticmorphologyparasitesystematicstaxonomy

Information

Type
Research Paper
Information
Journal of Helminthology , Volume 99 , 2025 , e17
Creative Commons
Creative Common License - CCCreative Common License - BY
This is an Open Access article, distributed under the terms of the Creative Commons Attribution licence (http://creativecommons.org/licenses/by/4.0), which permits unrestricted re-use, distribution and reproduction, provided the original article is properly cited.
Copyright
© The Author(s), 2025. Published by Cambridge University Press

Species of Cichlidogyrus Paperna, 1960 (Monogenoidea: Polyonchoinea: Dactylogyridae Bychowsky, 1933) primarily infect African cichlids, have evidently been introduced beyond the natural geographic distribution of their tilapia hosts, have been distributed globally due to production aquaculture (FAO 2022) and the aquarium trade, and have been infrequently reported to spill over into native fish communities (Table 1) (Jiménez-García et al. Reference Jiménez-García, Vidal-Martínez and López-Jiménez2001; Zhang et al. Reference Zhang, Zhi, Xu, Zheng, Bilong Bilong, Pariselle and Yang2019). The introduction of non-native Cichlidogyrus spp. should cause alarm among natural resource managers and aquaculturists because they have direct life cycles, can infect numerous fish hosts, and can cause disease (Kabata Reference Kabata1985; Khalil Reference Khalil1971; Jiménez-García et al. Reference Jiménez-García, Vidal-Martínez and López-Jiménez2001; Shinn et al. Reference Shinn, Avenant‐Oldewage, Bondad‐Reantaso, Cruz‐Laufer, García‐Vásquez, Hernández‐Orts, Kuchta, Longshaw, Metselaar, Pariselle and Pérez‐Ponce de León2023). Little is known about the parasites of Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) (Cichliformes: Cichlidae) in the United States despite it being an invasive species and aquaculture product. Brule et al. (Reference Brule, Warren, Dutton, Truong, Ksepka, Curran, Shurba, Lawson and Bullard2024) reported the African gyrodactylid, Gyrodactylus cichlidarum Paperna, 1968 (Monogenoidea: Polyonchoinea: Gyrodactylidae Cobbold, 1864) from the skin and gill of Nile tilapia from the same locality we report herein, Sougahatchee Creek (Tallapoosa River, Mobile-Tensaw Basin, Auburn, Alabama). We herein report the second exotic monogenoid species (Cichlidogyrus sclerosus Paperna & Thurston, Reference Paperna and Thurston1969) from Nile tilapia in the United States. This also comprises the first record of a dactylogyrid infecting Nile tilapia in the United States.

Table 1.

Host and locality records for infection by Cichlidogyrys sclerosus

Materials and methods

During November 2021, hatchery-reared Nile tilapia fingerlings were stocked (200 fish/raceway) into raceways (0.71 m × 0.61 m × 3.00 m; ~ 1 m3 volume) supplied with flow-through water (single pass) at the E.W. Shell Fisheries Center (EWSFC), Auburn, Alabama. This facility comprises a series of earthen aquaculture ponds comprising that segment of Sougahatchee Creek (Tallapoosa River) as well as an indoor flow-through aquaculture facility fed by surface water and having an effluent connected to the creek. Ten (10) Nile tilapia were removed from the raceways one day after stocking, transported to the laboratory alive in buckets containing raceway water, euthanized using tricaine methanesulphonate (MS-222), and necropsied for parasitological examination.

The gill from each Nile tilapia was removed intact, and each gill arch was separated and examined in a petri dish with raceway water with the aid of a stereo-dissecting microscope (Meiji RZ 3288) with fiber optic light sources as well as light and dark field sub-stage illumination. Live monogenoids intended for morphology were removed from the gill filaments with minutien pins affixed to wooden dowel rods and pipetted onto a glass slide, cover-slipped (no pressure), flame-killed on the slide using a hand lighter, fixed for several days in 10% neutral buffered formalin (n.b.f.), washed in deionized water to remove n.b.f., stained overnight in Van Cleave’s hematoxylin, dehydrated using a graded EtOH series, made basic in 70% EtOH with lithium carbonate and n-butylamine, dehydrated in 100% EtOH, cleared in clove oil, and permanently whole-mounted on glass slides using Canada balsam. Whole mounted specimens were drawn (Figures 1−4) with the aid of an Olympus BX51 compound microscope (Olympus, Tokyo, Japan) equipped with DIC optical components and a drawing tube. Measurements were obtained by using a Jenoptik Gryphax camera (Jenoptik AG, Jena, Germany). Measurements are reported as a range in micrometers (μm) followed by the mean, standard deviation, and sample size. Voucher specimens were deposited in the National Museum of Natural History’s Invertebrate Zoology Collection (Smithsonian Institution, USNM Collection Nos. 1743701–1743707). Hook numbering follows Llewellyn (Reference Llewellyn1963). Sclerite measurements follow Douëllou (Reference Douëllou1993). Scientific names for fishes, including taxonomic authorities and dates, follow Eschmeyer et al. (Reference Eschmeyer, Fricke and Van der Laan2016; online version updated 2023). Common names for fishes follow Froese and Pauly (Reference Froese and Pauly2023). The hosts were identified as Nile tilapia by having 23 gill rakers and 18 dorsal fin spines as per Boschung and Mayden (Reference Boschung and Mayden2004).

Figure 1.

Cichlidogyrus sclerosus Paperna & Thurston, Reference Paperna and Thurston1969 (Monogenoidea: Polyonchoinea: Dactylogyridae) (USNM No. 1743705) infecting the gill filaments of Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) (Cichliformes: Cichlidae) from the E.W. Shell Fisheries Center, Auburn, Alabama (hydrologically linked to Tallapoosa River, Mobile-Tensaw Basin). Scale value aside bar. Abbreviations: accessory piece (ap), anchor (a), distal vagina (dv), eyespot (e), haptor (h), haptoral gland (hg), head organ (ho), intestine (i), marginal hook (mh), mouth (m), ovary (o), peduncle muscle (pm), penis (p), pharynx (ph), proximal vagina (pv), seminal recepticle (sr), seminal vesicle (sv), uterus (u), vitellarium (v).

Two EtOH-preserved specimens intended for DNA extraction were wet-mounted and cover-slipped on a glass slide to confirm the presence of morphological features diagnostic for the species before being digested to extract complete genomic DNA using the Qiagen DNeasy Blood and Tissue Kit (Qiagen, Hilden, Germany) following the manufacturer’s protocol. Primer choice and PCR thermocycling follows Bullard et al. (Reference Bullard, Warren and Dutton2021). All PCR reactions were performed with a ProFlex PCR System (Applied Biosystems, Waltham, Massachusetts). Amplification of PCR product was checked in a 5% agarose gel stained with 1% ethidium bromide using 8μl of PCR product. PCR product was purified using the QIAquick PCR Product Purification Kit (Qiagen). Sequencing was performed by Genewiz, Incorporated (South Plainfield, New Jersey). Chromatograms were assembled based on sequence overlap, were proofread by eye, and had low-quality read ends trimmed in Geneious version 2023.2.1 (http://www.geneious.com), resulting in two 28S rDNA segments of 770 base pairs (bp) and ITS1 segments of 541 bp and 542 bp. All sequences were deposited in GenBank under accession numbers PQ728084–7.

Our phylogenetic analysis consisted of 28S and ITS1 phylogenies. Our sequences were aligned with conspecific sequences and appropriate, respective sequences of Cruz-Laufer et al. (Reference Cruz‐Laufer, Pariselle, Jorissen, Muterezi Bukinga, Al Assadi, Van Steenberge, Koblmüller, Sturmbauer, Smeets, Huyse and Artois2022) with the multiple alignment using fast Fourier transform (MAFFT) tool (Katoh and Standley Reference Katoh and Standley2013). Each alignment was trimmed to the length of our included sequence (770 bp, GenBank No. PQ728086 [28S]; 542 bp, GenBank No. PQ728084 [ITS1]). Aligned sequences were exported as a .phy file to run maximum likelihood (ML) tree inference. The ML trees were inferred with IQTREE v.1.16.12 (Nguyen et al. Reference Nguyen, Schmidt, von Haeseler and Minh2015). Substitution model testing was done with ModelFinder (Kalyaanamoorthy et al. Reference Kalyaanamoorthy, Minh, Wong, von Haeseler and Jermiin2017) as implemented in IQTREE. After model testing, tree inference was done using best-fitting substitution models (Chernomor et al. Reference Chernomor, von Haeseler and Minh2016). Default tree search parameters were used, except perturbation strength was set to 0.2, and 500 iterations had to be unsuccessful to stop the tree search. Tree inference was preformed 20 times with only the tree with the best log-likelihood score reported. Support for relationships was measured with 1,000 ultrafast bootstrap replicates (UFBoot) (Hoang et al. Reference Hoang, Chernomor, von Haeseler, Minh and Vinh2018). The recovered phylogenies were visualized using FigTree v1.4.4 (Rambaut et al. Reference Rambaut, Suchard, Xie and Drummond2014) and further edited for visualization purposes with Adobe Illustrator (Adobe Systems) (Figures 5 and 6).

Results

Cichlidogyrus sclerosus Paperna & Thurston, Reference Paperna and Thurston1969

Supplemental description (Figures 1–4)

Based on 10 stained, whole-mounted adult specimens: Body (including haptor) elongate, 444–708 (604 ± 110; 6) long, 123–172 (149 ± 19; 6) wide at maximum width at level immediately posterior to body midpoint, having 4 eyespots; eyespots dorsal; posterior pair of eyespots dense; anterior pair of eyespots diffuse (Figure 1). Mouth, subterminal, at level of anterior eyespot pair (Figure 1). Head organs in 3 pairs, containing string-like substance; gland ducts extending anteriad 78–126 (101 ± 16; 6) before connecting to terminal pores; gland ducts branching immediately anterior to level of mouth (Figure 1). Pharynx circular in outline, 36–43 (39 ± 3; 6) long, 33–41 (37 ± 3; 6) wide, extending dorsally and posteriad before joining with intestine (Figure 1). Intestine bifurcating at level of base of head organs, with cyclocoel, extending posteriad approximately in parallel with respective lateral body margin, looping in hind body (Figure 1).

Haptor irregular in outline, its connection to body indistinct, receiving glandular material from peduncle, associated with 2 muscle cords; cords extending anteriad into peduncle, approximately 44–93 (64 ± 10; 6) long, 74–93 (84 ± 7; 6) wide (Figure 1). Haptoral sclerites comprising 1 pair of ventral anchors, 1 pair of dorsal anchors, a ventral and a dorsal transverse bar, and 7 pairs of marginal hooks (Figures 1 and 2); ventral and dorsal anchors each comprising deep root, superficial root, base, shaft, and point (Figures 2a, b); ventral anchors 24–30 (28 ± 2; 8) long (Figures 1 and 2a); ventral anchor deep root 3–4 (4 ± 1; 8) long (Figure 2a); ventral anchor superficial root 7–9 (8 ± 1; 8) long (Figure 2a); ventral anchor point 11–15 (13 ± 1; 8) long (Figure 2a); ventral anchor length from peak of shaft to notch 23–31 (28 ± 3; 8) long (Figure 2a); dorsal anchors 22–27 (26 ± 2; 4) long (Figures 1 and 2b); deep root of dorsal anchor 3–4 (4 ± 1; 4) long (Figure 2b); superficial root of dorsal anchor 6–7 (6 ± 1; 4) long (Figure 2b); point of dorsal anchor bent, thin, 8–12 (9 ± 2; 4) long (Figure 2b); dorsal anchor length from peak of shaft to notch 23–28 (26 ± 2; 4) long (Figure 2b); ventral transverse bar spanning breath between bases of ventral anchors, lacking associated plate, comprising two projections forming the outline of a V, having a median groove extending along anterior margin, 3–7 (6 ± 2; 6) long at midpoint, 34–36 (35 ± 1; 3) wide at maximum width, 17–21 (20 ± 2; 6) tall (Figure 2c); ventral bar projections 21–27 (25 ± 2; 6) long (Figure 2c); dorsal transverse bar 6–8 (7 ± 1; 2) long at midpoint, 27–37 (32 ± 7; 2) wide, 23–26 (25 ± 2; 2) tall, spanning breath between bases of dorsal anchors, H-shaped in outline, having two pyriform projections ‘auricles’ sensu Pariselle and Euzet (Reference Pariselle and Euzet2009) and two peg-like projections (Figure 2d); dorsal bar pyriform projections 10–12 (11 ± 1; 3) long; distance between dorsal bar pyriform projections at base 10–11 (10 ± 1; 3) (Figure 2d); marginal hook pairs: pair I 8–12 (11 ± 2; 5) long; pair II 4–6 (5 ± 1; 3) long; pair III 11–16 (15 ± 2; 6) long; pair IV 11–17 (14 ± 3; 4) long; pair V 13–15 (14 ± 1; 5) long; pair VI 12–16 (13 ± 2; 3) long; pair VII 10–15 (13 ± 2; 4) long (Figures 2e–k).

Figure 2.

Haptoral sclerites of Cichlidogyrus sclerosus Paperna & Thurston, Reference Paperna and Thurston1969 (Monogenoidea: Polyonchoinea: Dactylogyridae) infecting the gill filaments of Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) (Cichliformes: Cichlidae) from the E.W. Shell Fisheries Center, Auburn, Alabama (hydrologically linked to Tallapoosa River, Mobile-Tensaw Basin). Scale values aside bars. (a) Ventral anchor of voucher (USNM No. 1743702), (b) Dorsal anchor of voucher (USNM No. 1743702), (c) ventral transverse bar of voucher (USNM No. 1743702), (d) dorsal transverse bar of voucher (USNM No. 1743702), (e) marginal hook I of voucher (USNM No. 1743704), (f) marginal hook II of voucher (USNM No. 1743703), (g) marginal hook III of voucher (USNM No. 1743704), (h) marginal hook IV of voucher (USNM No. 1743706), (i) marginal hook V of voucher (USNM No. 1743704), (j) marginal hook VI of voucher (USNM No. 1743707), (k) marginal hook VII of voucher (USNM No. 1743706).

Testis dorsal to ovary, margin indistinct, approximately 32 long, approximately 11 wide, entirely intercaecal (Figure 3). Vas deferens emerging from anterior margin of testis and extending anteriad (Figure 3). Seminal vesicle comprised of proximal portion and distal tube (Figure 3); proximal portion of seminal vesicle 28–37 (33 ± 4; 4) long, 14–21 (19 ± 3; 4) wide, containing sperm (Figure 3); distal tube of seminal vesicle extending mediad to male copulatory organ (MCO), connecting to MCO dorsally (Figures 1, 3, and 4). MCO medial, comprising penis and accessory piece, 88–136 (119 ± 18; 6) from anterior body end (Figures 1, 3, and 4). Penis ‘copulatory tube’ sensu Douëllou (Reference Douëllou1993) tubular, tapering distally, lacking swelling, recurved, not coiled, 59–68 (63 ± 4; 4) long, containing irregularly surfaced heel ‘serrated plate’ sensu Douëllou (Reference Douëllou1993) with elevated rim; heel 8–11 (10 ± 1; 5) long, 12–14 (14 ± 1; 5) wide (Figures 1, 3, and 4). Accessory piece bifid, straight, 50–61 (50 ± 5; 7) long or 74–83% of penis length (Figures 1, 3, and 4).

Figure 3.

Male and female reproductive genitalia of Cichlidogyrus sclerosus Paperna & Thurston, Reference Paperna and Thurston1969 (Monogenoidea: Polyonchoinea: Dactylogyridae) (USNM No. 1743701) infecting the gill filaments of Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) (Cichliformes: Cichlidae) from the E.W. Shell Fisheries Center, Auburn, Alabama (hydrologically linked to Tallapoosa River, Mobile-Tensaw Basin). Scale value aside bar. Abbreviations: accessory piece (ap), distal tube (dt), distal vagina (dv), heel (he), ovary (o), penis (p), proximal portion (pp), proximal vagina (pv), seminal receptacle (sr), seminal vesicle (sv), testis (t), transverse vitelline duct (tv), uterus (u), vaginal pore (vp), vas deferens (vd).

Figure 4.

Male copulatory organ of Cichlidogyrus sclerosus Paperna & Thurston, Reference Paperna and Thurston1969 (Monogenoidea: Polyonchoinea: Dactylogyridae) (USNM No. 1743702) infecting the gill filaments of Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) (Cichliformes: Cichlidae) from the E.W. Shell Fisheries Center, Auburn, Alabama (hydrologically linked to Tallapoosa River, Mobile-Tensaw Basin). Scale value aside bar. Abbreviations: accessory piece (ap), penis (p), heel (he).

Ovary irregular in shape, thin-walled, containing many fully developed ova, intercaecal, 47–64 (52 ± 7; 5) long, 24–43 (36 ± 8; 5) wide, 217–271 (251 ± 24; 4) or 37–43% of total body length (including haptor) from anterior body end, 280–368 (358 ± 38; 4) or 48–55% of total body length from posterior body end (including haptor) (Figures 1 and 3). Oviduct extending anteriad, dorsal to seminal receptacle (Figure 4). Transverse vitelline duct intersecting oviduct between seminal vesicle and seminal receptacle (Figure 4). Oötype not observed. Vitellarium extending posteriad from pharynx and slightly surpassing cyclocoel, dorsal and ventral to intestine, spanning the maximum breadth of body (Figure 1). Vagina funnel-shaped, originating ventral to seminal receptacle, comprised of proximal and distal portions (Figures 1 and 4); proximal vagina thin-walled, extending anterodextrad from seminal receptacle, 12–17 (14 ± 3; 4) in maximum width (Figures 1 and 3); distal vagina sclerotized, looped, surrounded by muscle distally, 33–37 (35 ± 2; 4) long (Figure 3). Vaginal pore slightly dextral, 57–59 (58 ± 1; 3) or 34–45% of body width from dextral body margin, 79–104 (92 ± 13; 3) or 56–79% of body width from sinistral body margin (Figure 1). Uterus thick-walled, dorsal to seminal vesicle, curving ventrally around dorsal portion of seminal vesicle, having crenulated luminal surface, tapering distally, extending sinistrad towards MCO, 34–64 (54 ± 17; 3) long, 2–3 (3 ± 1; 3) thick (Figure 3).

Taxonomic summary

Host: Oreochromis niloticus (Linnaeus, 1758) (Cichliformes: Cichlidae), Nile tilapia.

Locality: E. W. Shell Fisheries Center, Auburn, Alabama, USA (Sougahatchee Creek, Tallapoosa River, Mobile-Tensaw Basin).

Specimens and sequences deposited: Voucher specimens (USNM Nos. 1743701–7);

28S rDNA (GenBank Nos. PQ728086–7); ITS1 (GenBank Nos. PQ728084–5)

Site on host: Gill filaments.

Taxonomic remarks

Cichlidogyrus sclerosus is most easily differentiated from its congeners by having the combination of marginal hooks that are shorter than the dorsal anchor length, reduced anchor roots, a bent dorsal anchor point, dorsal bar pyriform projections that are approximately half as long as the dorsal bar width, a short penis (<100 μm) that is tubular (not coiled, lacking swelling) and having an irregularly surfaced heel, and an accessory piece that is straight (not curved) and bifid. Our specimens matched the original (Paperna and Thurston Reference Paperna and Thurston1969) and supplemental descriptions (Douëllou Reference Douëllou1993; Vidal-Martínez et al. Reference Vidal-Martínez, Aguirre-Macedo, Scholz, González-Solís and Mendoza-Franco2001) of C. sclerosus by having that combination of features. This monogenoid was originally described from the gill of Mozambique tilapia, Oreochromis mossambicus (Peters, 1852) (as Tilapia mossambica [Peters, 1852], Cichlidae, type host) from Lake Victoria, Kajjansi, Uganda (type locality); Nile tilapia (as Tilapia nilotica [Linnaeus, 1758]) and Haplochromis sp. (Cichlidae) from Lake George, Uganda; blue spotted tilapia, Oreochromis leucostictus (Trewavas, 1933) (as Tilapia leucosticta Trewavas, 1933) from Lake Victoria, Jinja, Uganda; and redbelly tilapia, Coptodon zillii (Gervais, 1848) (as Tilapia zillii [Gervais, 1848]; Cichlidae; experimental infection, locality unspecified) (Table 1).

Herein, we collected specimens of C. sclerosus and a congener. These congeneric specimens generally resemble C. thurstonae (type-host: Nile tilapia; type-locality: Nile River, Cairo, Egypt) by having marginal hook pairs III–VII that are approximately twice as long as the marginal hook pair I; an s-shaped accessory piece with a proximal finger-like process; and a straight vaginal sclerite having a jagged margin (Ergens Reference Ergens1981; Pariselle et al. Reference Pariselle, Bilong Bilong and Euzet2003). However, these specimens were poorly fixed, and we cannot formally detail them further herein.

Phylogenetic results

The two 28S sequences of C. sclerosus (GenBank Nos. PQ728086–7) produced herein each comprised 770 bp and were identical. Our included 28S sequence was recovered in a clade of other sequences of C. sclerosus. Our sequences differ from the other 12 extant 28S sequences ascribed to C. sclerosus (GenBank Nos. DQ157660, MH767401, MK524728, MK524729 MN078060, MT994744, MW580351, MW580352, OL415084, OR338287, OR528026, PP191180) by 0 (0%) to 8 bp (1%) (Figure 5). The 28S sequences of C. sclerosus used herein were recovered sister to that of Cichlidogyrus amphoratus Pariselle & Euzet, 1996 (GenBank No. HE792772) infecting Guinean tilapia, Coptodon guineensis (Gunther, 1862) from Senegal (Mendlová et al. Reference Mendlová, Desdevises, Civáňová, Pariselle and Šimková2012), and our sequence differed from it by 37 bp (4.5%) (Figure 5).

Figure 5.

Phylogenetic relationships of species within Cichlidogyrys Paperna, 1960 reconstructed using maximum likelihood tree inference using the large ribosomal subunit (28S) gene. Numbers aside nodes indicate bootstrap values. New sequence of Cichlidogyrus sclerosus Paperna & Thurston, Reference Paperna and Thurston1969 is shown in bold. GenBank numbers are in parentheses following each taxon.

Figure 6.

Phylogenetic relationships of species within Cichlidogyrys Paperna, 1960 reconstructed using maximum likelihood tree inference using the internal transcribed spacer 1 (ITS1). Numbers aside nodes indicate bootstrap values. New sequence of Cichlidogyrus sclerosus Paperna & Thurston, Reference Paperna and Thurston1969 is shown in bold. GenBank numbers are in parentheses following each taxon.

The two ITS1 sequences of C. sclerosus (GenBank Nos. PQ728084–5) produced herein comprised 541 bp and 542 bp and were identical. Our included ITS1 sequence was recovered in a clade of other sequences of C. sclerosus (Figure 5). Our sequences differed from the other 9 extant ITS1 sequences ascribed to C. sclerosus (GenBank Nos. DQ537359, KX869722, MH767390, MN905938, ON819193, ON819218, ON819279, ON819301, OR335571) by 0 (0%), 1 (0.3%), or 3 bp (0.6%) except for GenBank No. ON819218, which differed by 29 bp (4.7%). The ITS1 sequences of C. sclerosus used herein were recovered sister to C. amphoratus (GenBank No. HE792782) infecting Guinean tilapia from Senegal (Figure 5) (Mendlová et al. Reference Mendlová, Desdevises, Civáňová, Pariselle and Šimková2012), and our sequence differed from it by 48 bp (10.4%).

The ITS1 and 28S phylogenetic analyses failed to recover clades of C. sclerosus from the same geographic locality (i.e., sequences of C. sclerosus from the same continent are not monophyletic).

Discussion

Cichlidogyrus sclerosus exhibits characteristics of a potential invasive pathogen, especially to endemic cichlids. It has a direct life cycle, a geographically broad (worldwide) introduced/invasive range, and a relatively low degree of host specificity, infecting cichlids of various genera as well as a carp (Cyprinidae) and a splitfin (Goodeidae) (Table 1). It has spread globally probably because of African cichlid aquaculture (FAO 2022; Table 1). Since its original description from cichlids in Uganda, C. sclerosus has been reported from 35 other countries in Africa, Asia, North America, South America, and Europe. Our phylogenetic analysis showed that isolates of C. sclerosus do not clade by geographic locality, which could indicate that this parasite has been introduced and reintroduced repeatedly. Cichlidogyrus sclerosus has been reported from 21 host species belonging to 11 genera and 3 families (Cichlidae, Cyprinidae, Goodeidae), including non-African hosts (Table 1). Many of these records need confirmation and are not accompanied by a voucher specimen or published morphological diagnosis. Cichlidogyrus sclerosus has been identified as a primary pathogen in fish culture facilities by Kabata (1986) and Paperna (Reference Paperna1996). A congener, Cichlidogyrus philander Douëllou, Reference Douëllou1993, causes gill hyperplasia, lamellar fusion, ablation of epithelium, and filament malformation in high intensity infections among wild southern mouthbrooders, Pseudocrenilabrus philander (Weber, 1897) (Cichlidae) from a manmade South African reservoir (Igeh and Avenant-Oldewage Reference Igeh and Avenant‐Oldewage2020). More experimental work that documents intensity-dependent pathological change and studies that demonstrate Koch’s postulates for C. sclerosus would help determine its threat level as an invading, primary pathogen. Rather alarming from the perspective of fisheries and conservation management, infections of C. sclerosus have been reported from local, non-African, non-cichlid fishes in Mexico (blackfin goodea, Goodea atripinnis Jordan, 1880) and Iraq (common carp, Cyprinus carpio Linnaeus, 1758) (Table 1). Considering all host records and phylogenetic host specificity (Table 1), it seems relatively unlikely that non-cichlid endemic fishes would be at high risk for infection and disease caused by Cichlidogyrus spp. However, the fact that non-cichlids can be infected (Abd Al-Khenifsawy and Al-Mayali Reference Abd Al-Khenifsawy and Al-Mayali2022; Salgado-Maldonado and Rubio-Godoy Reference Salgado-Maldonado, Rubio-Godoy, Alfaro and Osorio2014) is cause for concern. We also acknowledge that probably few workers are examining the gill of native stream fishes for infections by these minute ectoparasites, and perhaps infections among non-cichlid fishes are underreported.

Acknowledgements

We thank Anna Phillips, Chad Walter, Kathryn Ahlfeld, Amanda Robinson, and William Moser (all Department of Invertebrate Zoology, National Museum of Natural History, Smithsonian Institution, Washington, DC) for curating our museum specimens.

Financial support

This study was supported by the Southeastern Cooperative Fish Parasite and Disease Project (Auburn University), U.S. Fish and Wildlife Service (Department of Interior), National Sea Grant (National Oceanic and Atmospheric Administration), United States Department of Agriculture (National Institute of Food and Agriculture), Federal Aid in Sport Fish Restoration (Alabama Department of Conservation and Natural Resources, Inland and Marine Resources Divisions), and the Alabama Agricultural Experiment Station (Auburn University, College of Agriculture).

Competing interest

The authors declare that they have no conflict of interest.

Ethical standard

All applicable institutional, national, and international guidelines for the care and use of animals were followed.

References

Abd Al-Khenifsawy, RH and Al-Mayali, HMH (2022) New hosts records for some fish parasites from Al-Dalmaj Marsh/Iraq. International Journal ofHhealth Sciences 6(S8), 5,180–185,191.Google Scholar
Abdulmalik, IA, Shariman, YZ and Hazrin, HZ (2023) Freshwater fish parasites in Temengor, Chenderoh and Bersia Reservoirs, Perak, Malaysia. Biological and Environmental Sciences Journal for the Tropics 20(3), 17–26.Google Scholar
Aguirre-Fey, D, Benítez-Villa, GE, de León, GPP and Rubio-Godoy, M (2015) Population dynamics of Cichlidogyrus spp. and Scutogyrus sp. (Monogenea) infecting farmed tilapia in Veracruz, México. Aquaculture 443, 11–15.CrossRefGoogle Scholar
Akoll, P, Fioravanti, ML, Konecny, R and Schiemer, F (2012) Infection dynamics of Cichlidogyrus tilapiae and C. sclerosus (Monogenea, Ancyrocephalinae) in Nile tilapia (Oreochromis niloticus L.) from Uganda. Journal of Helminthology 86(3), 302–310.CrossRefGoogle Scholar
Anshary, H, Azra, K and Sukarni, N (2022) Occurrence of ectoparasites on Nile Tilapia (Oreochromis niloticus) from South Sulawesi Lakes, and Aquaculture facility. IOP Conference Series: Earth and Environmental Science 1119(1), 012013.Google Scholar
Anshary, H, Sriwulan, S and Amriana, A (2023) High prevalence and mean intensity of trichodinids and monogeneans on Nile tilapia (Oreochromis niloticus) in Indonesian hatcheries. Veterinary Parasitology: Regional Studies and Reports 43, 100898.Google ScholarPubMed
Boschung, HT and Mayden, RL (2004) Fishes of Alabama. Washington, DC: Smithsonian Books.Google Scholar
Britto, YCT and Silva-Souza, ÂT (2017) Temporal variation of monogenoideans component community in the gills of Oreochromis niloticus (Cichlidae) in fish farming in northern Parana state, Brazil. Pan-American Journal of Aquatic Sciences 12(4), 333–342.Google Scholar
Brule, JH, Warren, MB, Dutton, HR, Truong, TN, Ksepka, SP, Curran, SS, Shurba, JA, Lawson, LL and Bullard, SA (2024) First taxonomic description of a gyrodactylid, Gyrodactylus cichlidarum Paperna, 1968 (Monogenoidea) infecting Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) (Cichlidae) in the United States. BioInvasions Record 13(1), 281–291.CrossRefGoogle Scholar
Bullard, SA, Warren, MB, and Dutton, HR (2021) Redescription of Cathariotrema selachii (MacCallum, 1916) Johnston and Tiegs, 1922 (Monogenoidea: Monocotylidae), emendation of monotypic Cathariotrema Johnston and Tiegs, 1922, and proposal of Cathariotrematinae n. subfam. based on morphological and nucleotide evidence. The Journal of Parasitology 107(3), 481–513.CrossRefGoogle ScholarPubMed
Cabrera, M (2019) Identificación y caracterización ultraestructural de monogeneos presentes en las branquias de Tilapia del Nilo Oreochromis niloticus asociados a las lesiones histológicas procedentes de cultivos de la provincia de San Martín. Universidad Peruana Cayetano Heredia. 101 p.Google Scholar
Cáceres-Farías, L, Cruz-Quintana, Y, Muñoz-Chumo, LG and Santana-Piñeros, AM (2022) First evidence of Cichlidogyrus Paperna, 1960 (Monogenea: Ancyrocephalidae) in cultures of Oreochromis spp. from Ecuador. Journal MVZ Córdoba 27(2), e2402.Google Scholar
Cavalcanti, LD, Gouveia, EJ, Leal, FC, Figueiró, CSM, Rojas, SS and Russo, MR (2020) Responses of monogenean species to variations in abiotic parameters in tilapiculture. Journal of Helminthology 94, e186.CrossRefGoogle ScholarPubMed
Chernomor, O, von Haeseler, A and Minh, BQ (2016) Terrace aware data structures for phylogenomic inference from supermatrices. Systematic Biology 65, 997–1008.CrossRefGoogle ScholarPubMed
Christison, KW (2002) Branchial monogenean parasites (Monogenea: Dactylogyridae) of fishes from the Okavango River and Delta, Botswana. University of the Free State. 172 p.Google Scholar
Cruz‐Laufer, AJ, Pariselle, A, Jorissen, MW, Muterezi Bukinga, F, Al Assadi, A, Van Steenberge, M, Koblmüller, S, Sturmbauer, C, Smeets, K, Huyse, T and Artois, T (2022) Somewhere I belong: Phylogeny and morphological evolution in a species‐rich lineage of ectoparasitic flatworms infecting cichlid fishes. Cladistics 38(4), 465–512.CrossRefGoogle Scholar
Dayoub, AI and Salman, HM (2015) Study of using Monogenea parasites on free – living fishes in the lake of 16 Tishreen Dam as bio indicators of environment pollution. International Journal of Biomedical Engineering and Clinical Science 1(1), 15–22.Google Scholar
de Azevedo, TMP, Martins, ML, Bozzo, FR and Moraes, FRD (2006) Haematological and gill responses in parasitized tilapia from Valley of Tijucas River, SC, Brazil. Scientia Agricola 63, 115–120.CrossRefGoogle Scholar
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 25(3), 159–186.CrossRefGoogle Scholar
Duncan, BL (1973) Cichlidogyrus sclerosus Paperna and Thurston from cultured Tilapia mossambica. Kalikasan, the Philippine Journal of Biology 2, 154–158.Google Scholar
El-Naggar, MM and Khidr, AA (1986) Population dynamics of Cichlidogyrid monogeneans from the gills of three Tilapia spp. from Damietta branch of the River Nile in Egypt. Proceedings of the Zoological Society A.R. Egypt 12, 275–286.Google Scholar
Ergens, R (1981) Nine species of the genus Cichlidogyrus Paperna, 1960 (Monogenea: Ancyrocephalinae) from Egyptian fishes. Folia Parasitologica 28, 205–214.Google Scholar
Eschmeyer, WN, Fricke, R and Van der Laan, R (2016) Catalog of Fishes: Genera, Species, References. Available at: http://researcharchive.calacademy.org/research/ichthyology/fishcatmain.asp. Accessed 22 January 2024.Google Scholar
FAO (2022) The State of World Fisheries and Aquaculture 2022. Towards Blue Transformation. Rome: FAO.Google Scholar
Ferdousi, UK and Chandra, KJ (2002) New monogenean gill parasites of Oreochromis niloticus (Linnaeus) and Oreochromis mossambicus (Peters) (Osteichthyes, Cichlidae) from Mymensingh, Bangladesh. Rivista di parassitologia 19(63), 49–60.Google Scholar
Firmat, C, Alibert, P, Mutin, G, Losseau, M, Pariselle, A and Sasal, P (2016) A case of complete loss of gill parasites in the invasive cichlid Oreochromis mossambicus. Parasitology Research 115, 3657–3661.CrossRefGoogle ScholarPubMed
Flores-Crespo, J, Flores-Crespo, R, Ibarra-Velarde, F, Vera-Montenegro, Y and Vasquez-Pelaez, C (1995) Evaluación de Quimioterapéuticos contra la Ciclidogiriasis de la Tilapia (Oreochromis hornorum) en México. Revista Latinoamericana de Microbiologia (Mexico) 37, 179–187.Google Scholar
Froese, R and Pauly, D (eds.). (2023) FishBase. Available at www.fishbase.org (version 10/2023) (accessed January 22, 2024).Google Scholar
Garcia, DAZ, Orsi, ML and Silva-Souza, ÂT (2019) From Africa to Brazil: Detection of African Oreochromis niloticus parasites in Brazilian fish farms. Acta Limnologica Brasiliensia 31, e202.CrossRefGoogle Scholar
Geraerts, M, Huyse, T, Barson, M, Bassirou, H, Bilong Bilong, CF, Nyom, ARB, Manda, AC, Cruz-Laufer, AJ, Kabalika, CK, Kasembele, GK, Bukinga, FM, Njom, S, Steenberge, MV, Artois, T and Vanhove, MPM (2023) Sharing is caring? Barcoding suggests co-introduction of dactylogyrid monogeneans with Nile tilapia and transfer towards native tilapias in sub-Saharan Africa. International Journal for Parasitology 53(13), 711–730.CrossRefGoogle ScholarPubMed
Ghiraldelli, L, Martins, ML, Jerônimo, GT, Yamashita, MM and Adamante, WDB (2006) Ectoparasites communities from Oreochromis niloticus cultivated in the State of Santa Catarina, Brazil. Journal of Fisheries and Aquatic Science 1(2), 181–190.CrossRefGoogle Scholar
Harrison, FNM (1983) Monogenetic trematodes (Dactylogyridae: Ancyrocephalinae) on the gills of tilapia (a warm-water cultured fish) with special reference to Cichlidogyrus sclerosus Paperna and Thurston 1969. Institute of Aquaculture, University of Stirling, Scotland.Google Scholar
Hernández-Ocampo, D, Pienda-López, RF, Ponce-Palafox, JT and Arredono-Figuero, JL (2012) Parasitic helminth infection in tropical freshwater fishes of commercial fish farms, in Morelos State, Mexico. International Journal of Animal and Veterinary Advances 4(5), 338–343.Google Scholar
Hind, H and Suhad, J (2023) Review of fish endoparasites in Tigris River, Iraq. GSC Biological and Pharmaceutical Sciences 25(3), 196–200.CrossRefGoogle Scholar
Hoang, DT, Chernomor, O, von Haeseler, A, Minh, BQ and Vinh, LS (2018) UFBoot2: Improving the ultrafast bootstrap approximation. Molecular Biology and Evolution 35, 518–522.CrossRefGoogle ScholarPubMed
Igeh, PC and Avenant‐Oldewage, A (2020) Pathological effects of Cichlidogyrus philander Douëllou, 1993 (Monogenea, Ancyrocephalidae) on the gills of Pseudocrenilabrus philander (Weber, 1897) (Cichlidae). Journal of Fish Diseases 43(2), 177–184.CrossRefGoogle ScholarPubMed
Jerônimo, GT, Speck, GM, Cechinel, MM, Gonçalves, ELT and Martins, ML (2011) Seasonal variation on the ectoparasitic communities of Nile tilapia cultured in three regions in southern Brazil. Brazilian Journal of Biology 71, 365–373.CrossRefGoogle ScholarPubMed
Jiménez-García, MI, Vidal-Martínez, VM and López-Jiménez, S (2001) Monogeneans in introduced and native cichlids in Mexico: evidence for transfer. Journal of Parasitology 87(4), 907–909.Google ScholarPubMed
Jorissen, MWP, Huyse, T, Pariselle, A, Lunkayilakio, SW, Bukinga, FM, Manda, AC, Kasembele, GK, Vreven, EJ, Snoeks, J, Decru, E, Artois, T and Vanhove, MPM (2020) Historical museum collections help detect parasite species jumps after tilapia introductions in the Congo Basin. Biological Invasions 22, 2825–2844.CrossRefGoogle Scholar
Jorissen, MWP, Pariselle, A, Huyse, T, Vreven, EJ, Snoeks, J, Volckaert, FAM, Manda, AC, Kasembele, GK, Artois, T and Vanhove, MPM (2018) Diversity and host specificity of monogenean gill parasites (Platyhelminthes) of cichlid fishes in the Bangweulu-Mweru ecoregion. Journal of Helminthology 92(4), 417–437.CrossRefGoogle ScholarPubMed
Jorissen, M, Vanhove, MP, Pariselle, A, Snoeks, J, Vreven, E, Šimková, A, Lunkayilakio, SW, Manda, AC, Kasembele, GK, Bukinga, FM and Artois, T (2022) Molecular footprint of parasite co-introduction with Nile tilapia in the Congo Basin. Organisms Diversity & Evolution 22(4), 1003–1019.CrossRefGoogle Scholar
Kabata, Z (1985) Parasites and Diseases of Fish Cultured in the Tropics. London: Taylor & Francis.Google Scholar
Kasembele, GK, Manda, AC, Abwe, E, Pariselle, A, Bukinga, FM, Huyse, T, Jorissen, MWP, Vreven, EJ, Luus-Powell, WJ, Smit, WJ and Sara, JR (2023) First record of monogenean fish parasites in the Upper Lufira River Basin (Democratic Republic of Congo): Dactylogyrids and gyrodactylids infesting Oreochromis mweruensis, Coptodon rendalli and Serranochromis macrocephalus (Teleostei: Cichlidae). Parasites & Vectors 16(1), 48.CrossRefGoogle ScholarPubMed
Kalyaanamoorthy, S, Minh, BQ, Wong, TKF, von Haeseler, A and Jermiin, LS (2017) ModelFinder: Fast model selection for accurate phylogenetic estimates. Nature Methods 14, 587–589.CrossRefGoogle ScholarPubMed
Katoh, K and Standley, DM (2013) MAFFT Multiple sequence alignment software version 7: Improvements in performance and usability. Molecular Biology and Evolution 30, 772–780.CrossRefGoogle ScholarPubMed
Khalil, LF (1971) Checklist of the helminth parasites of African freshwater fishes. In Technical Communication of the Commonwealth Institute of Helminthology, vol. 42. St. Albans: Commonwealth Agricultural Bureaux.Google Scholar
Kritsky, DC and Thatcher, VE (1974) Monogenetic trematodes (Monopisthocotylea: Dactylogyridae) from freshwater fishes of Colombia, South America. Journal of Helminthology 48(1), 59–66.CrossRefGoogle ScholarPubMed
Kritsky, DC, Vidal-Martínez, VM and Rodríguez-Canul, R (1994) Neotropical Monogenoidea. 19. Dactylogyridae of Cichlids (Perciformes) from the Yucatán Peninsula, with descriptions of three new species of Sciadicleithrum Kritsky, Thatcher, and Boeger, 1989. Journal of the Helminthological Society of Washington 61(1), 26–33.Google Scholar
Lehmann, NB, Owatari, MS, Furtado, WE, Cardoso, L, Tancredo, KR, Jesus, GFA, Lopes, GR and Martins, ML (2020) Parasitological and histopathological diagnosis of a non-native fish (Oreochromis sp.) with a noticeable presence in a natural Brazilian river environment. Journal of Parasitic Diseases 44, 201–212.CrossRefGoogle Scholar
Lerssutthichawal, T (2008) Diversity and distribution of external parasites from potentially cultured freshwater fishes in Nakhonsithammarat, Southern Thailand. In Bondad-Reantaso, MG, Mohan, CV, Crumlish, M and Subasinghe, RP (eds), Diseases in Asian Aquaculture VI. Manila, Philippines: Asian Fisheries Society, 235–244.Google Scholar
Lerssutthichawal, T, Maneepitaksanti, W and Purivirojkul, W (2016) Gill monogeneans of potentially cultured tilapias and first record of Cichlidogyrus mbirizei Bukinga et al., 2012, in Thailand. Walailak Journal of Science and Technology 13(7), 543–553.Google Scholar
Lerssutthichawal, T and Supamattaya, K (2005) Diversity and distribution of parasites from potentially cultured freshwater fish in Nakhon Si Thammarat. Songklanakarin Journal of Science and Technology 27(Suppl. 1), 333–334.Google Scholar
Lizama, MAP, Takemoto, RM, Ranzani-Paiva, MJT, Ayroza, LMA and Pavanelli, GC (2007) Relação parasito-hospedeiro em peixes de pisciculturas da região de Assis, Estado de São Paulo, Brasil. 1. Oreochromis niloticus (Linnaeus, 1757). Acta Scientiarum Biological Sciences 29(2), 223–231.CrossRefGoogle Scholar
Llewellyn, J (1963) Larvae and larval development of monogeneans. Advances in Parasitology 1, 287–326.CrossRefGoogle ScholarPubMed
Lopez, NC (1991) Metazoan ectoparasites of some cultured fishes from Laguna Lake and vicinities. Transactions of the National Academy of Science and Technology (Philippines) 13, 499–516.Google Scholar
Madanire-Moyo, GN, Matla, MM, Olivier, PAS and Luus-Powell, WJ (2011) Population dynamics and spatial distribution of monogeneans on the gills of Oreochromis mossambicus (Peters, 1852) from two lakes of the Limpopo River System, South Africa. Journal of Helminthology, 85(2), 146–152.CrossRefGoogle ScholarPubMed
Maneepitaksanti, W and Nagasawa, K (2012) Monogeneans of Cichlidogyrus Paperna, 1960 (Dactylogyridae), gill parasites of tilapias, from Okinawa Prefecture, Japan. Biogeography 14(2), 111–119.Google Scholar
Maneepitaksanti, W, Worananthakij, W, Sriwilai, P and Laoprasert, T (2014) Identification and distribution of gill monogeneans from Nile Tilapia and red tilapia in Thailand. Veterinary Integrative Sciences 12(1), 57–68.Google Scholar
Martins, ML, de Azevedo, TM, Ghiraldelli, L and Bernardi, N (2010) Can the parasitic fauna on Nile tilapias be affected by different production systems? Anais da Academia Brasileira de Ciências 82, 493–500.CrossRefGoogle ScholarPubMed
Martins, ML, de Sá, ARS, Jerônimo, GT, Tancredo, KR, Gonçales, ELT, Bampi, D, Speck, GM and Sandin, AM (2014) Microhabitat preference and seasonality of gill monogeneans in Nile Tilapia reared in Southern Brazil. Neotropical Helminthology 8(1), 47–58.Google Scholar
Matla, MM (2012) Helminth ichthyo-parasitic fauna of a South African sub-tropical lake. University of Limpopo (Turfloop Campus).Google Scholar
Méndez, O, Salgado-Maldonado, G, Caspeta-Mandujano, JM and Cabanas-Carranza, G (2010) Helminth parasites of some freshwater fishes from Baja California Sur, Mexico. Zootaxa 2327, 44–50.CrossRefGoogle Scholar
Mendlová, M, Desdevises, Y, Civáňová, K, Pariselle, A and Šimková, A (2012) Monogeneans of West African cichlid fish: Evolution and cophylogenetic interactions. PLoS One 7(5), e37268.CrossRefGoogle ScholarPubMed
Mendoza-Franco, EF, Caspeta-Mandujano, JM and Osorio, MT (2018) Ecto-and endo-parasitic monogeneans (Platyhelminthes) on cultured freshwater exotic fish species in the state of Morelos, South-Central Mexico. ZooKeys 776, 1–12.CrossRefGoogle Scholar
Mendoza-Franco, EF, Vidal-Martínez, VM, Cruz-Quintana, Y and León, FP (2006) Monogeneans on native and introduced freshwater fishes from Cuba with the description of a new species of Salsuginus Beverley-Burton, 1984 from Limia vittata (Poeciliidae). Systematic Parasitology 64(3), 181–190.CrossRefGoogle ScholarPubMed
Modi, AK, Thummala, C and Vankara, AP (2021) Water quality assessment based on gill monogenean parasites of Oreochromis niloticus (Linnaeus, 1758) and Labeo calbasu (Hamilton, 1822) fishes of River Penna (India). Applied Biological Research 23(3), 235–244.CrossRefGoogle Scholar
Morales-Serna, FN, Medina-Guerrero, RM, Pimentel-Acosta, C, Ramírez-Tirado, JH and Fajer-Ávila, EJ (2018) Parasite infections in farmed Nile tilapia Oreochromis niloticus in Sinaloa, Mexico. Comparative Parasitology 85(2), 212–216.CrossRefGoogle Scholar
Mhaisen, FT (2021) Checklist of parasites of the redbelly tilapia, Coptodon zillii (Gervais, 1848) (Cichliformes: Cichlidae) in Iraq. Aalborg Academy Journal of Pure Sciences 2(1), 1–17.Google Scholar
Natividad, JM, Bondad-Reantaso, MG and Arthur, JR (1986) Parasites of Nile tilapia (Oreochromis niloticus) in the Philippines. In Maclean, JL, Dizon, LB and Hosillos, LV (eds), The First Asian Fisheries Forum. Manila, Philippines: Asian Fisheries Society, 255–259.Google Scholar
Nguyen, LT, Schmidt, HA, von Haeseler, A and Minh, BQ (2015) IQ-TREE: A fast and effective stochastic algorithm for estimating maximum-likelihood phylogenies. Molecular Biology and Evolution 32, 268–274.CrossRefGoogle ScholarPubMed
Olivier, PAS, Luus-Powell, WJ and Saayman, JE (2009) Report on some monogenean and clinostomid infestations of freshwater fish and water bird hosts in Middle Letaba Dam, Limpopo Province, South Africa. Onderstepoort Journal of Veterinary Research 76, 187–199.CrossRefGoogle Scholar
Outa, JO, Dos Santos, QM, Avenant-Oldewage, A and Jirsa, F (2021) Parasite diversity of introduced fish Lates niloticus, Oreochromis niloticus and endemic Haplochromis spp. of Lake Victoria, Kenya. Parasitology Research 120(5), 1583–1592.CrossRefGoogle Scholar
Paredes-Trujillo, A, Mendoza-Carranza, M, del Río-Rodriguez, RE and Cerqueda-García, D (2022) Comparative assessment of metazoans infestation of Nile tilapia (Oreochromis niloticus) (L.) (Perciformes: Cichlidae) in floating cages and ponds from Chiapas, Mexico. Veterinary Parasitology: Regional Studies and Reports 34, 100757.Google ScholarPubMed
Paperna, I (1996) Parasites, Infections and Diseases of Fishes in Africa – An Update. CIFA Technical Paper No. 31. FAO: Rome.Google Scholar
Paperna, I and Lahav, M (1971) New records and further data on fish parasites in Israel. Bamidgeh 23, 43–52.Google Scholar
Paperna, I and Thurston, JP (1969) Monogenetic trematodes collected from cichlid fish in Uganda; including the descriptions of five new species of Cichlidogyrus. Revue de zoologie et de botanique africanines 79, 15–33.Google Scholar
Pariselle, A, Bilong Bilong, CF and Euzet, L (2003) Four new species of Cichlidogyrus Paperna, 1960 (Monogenea, Ancyrocephalidae), all gill parasites from African mouthbreeder tilapias of the genera Sarotherodon and Oreochromis (Pisces, Cichlidae), with a redescription of C. thurstonae Ergens, 1981. Systematic Parasitology 56, 201–210.CrossRefGoogle ScholarPubMed
Pariselle, A and Euzet, L (2009) Systematic revision of dactylogyridean parasites (Monogenea) from cichlid fishes in Africa, the Levant and Madagascar. Zoosystema 31(4), 849–898.CrossRefGoogle Scholar
Pinacho-Pinacho, CD, Sánchez-Angeles, J, Martínez-Ramírez, E, Calixto-Rojas, M and Rubio-Godoy, M (2023) Feral parasites: African monogeneans recorded in ferine Nile tilapia in Oaxaca, Mexico. Aquaculture 577, e739974.CrossRefGoogle Scholar
Prieto, A and Fajer, E (1987) Género Cichlidogyrus (Monogenea: Ancyrocephalinae) en tilapias en cultivo intensivo: características de la invasión. Revista de Salud Animal 9(4), 280–287.Google Scholar
Prieto, A, Fajer, E and Vinjoy, M (1985) Cichlidogyrus sclerosus (Monogenea: Ancyrocephalinidae) en Tilapia hornorum × Tilapia mossambica (perca dorada) en cultivo intensivo. Revista de Salud Animal 7, 291–295.Google Scholar
Rambaut, A, Suchard, MA, Xie, D and Drummond, AJ (2014) FigTree v1.4.3. Available at http://tree.bio.ed.ac.uk/software/figtree (accessed October 4, 2023).Google Scholar
Rindoria, NM, Mungai, LK, Yasindi, AW and Otachi, EO (2016) Gill monogeneans of Oreochromis niloticus (Linnaeus, 1758) and Oreochromis leucostictus (Trewavas, 1933) in Lake Naivasha, Kenya. Parasitology Research 115, 1501–1508.CrossRefGoogle ScholarPubMed
Roberts, RJ and Sommerville, C (1982) Diseases of tilapias, In Pullin, RSV and Lowe-McConnell, RH (eds), The Biology and Culture of Tilapias. ICLARM Conference Proceedings. Manila, Philippines: International Center for Living Aquatic Resources Management, 247–263.Google Scholar
Salgado-Maldonado, G and Rubio-Godoy, M (2014) Helmintos parásitos de peces de agua dulce introducidos. In Alfaro, REM and Osorio, PK (eds), Especies acuáticas invasoras en México, 269–285 Comisión Nacional para el Conocimiento y Uso de la Biodiversidad, México.Google Scholar
Salgado-Maldonado, G, Mercado-Silva, N, Cabañas-Carranza, G, Caspeta-Mandujano, JM, Aguilar-Aguilar, R and Iñiguez-Dávalos, LI (2004) Helminth parasites of freshwater fishes of the Ayuquila River, Sierra de Manantlan Biosphere Reserve, west central Mexico. Comparative Parasitology 71(1), 67–72.CrossRefGoogle Scholar
Sanchez-Ramirez, C, Vidal-Martinez, VM, Aguirre-Macedo, ML, Rodriguez-Canul, RP, Gold-Bouchot, G and Sures, B (2007) Cichlidogyrus sclerosus (Monogenea: Ancyrocephalinae) and its host, the Nile tilapia (Oreochromis niloticus), as bioindicators of chemical pollution. Journal of Parasitology 93(5), 1097–1106.CrossRefGoogle ScholarPubMed
Santacruz, A, Barluenga, M and Pérez-Ponce de León, G (2022) The macroparasite fauna of cichlid fish from Nicaraguan lakes, a model system for understanding host–parasite diversification and speciation. Scientific Reports 12(1), 3944.CrossRefGoogle Scholar
Shinn, AP, Avenant‐Oldewage, A, Bondad‐Reantaso, MG, Cruz‐Laufer, AJ, García‐Vásquez, A, Hernández‐Orts, JS, Kuchta, R, Longshaw, M, Metselaar, M, Pariselle, A, Pérez‐Ponce de León, G (2023) A global review of problematic and pathogenic parasites of farmed tilapia. Reviews in Aquaculture 15, 92–153.CrossRefGoogle Scholar
Sriwongpuk, S (2018) A study of ectoparasites of Red Tilapia (Oreochromis niloticus × O. mossambicus) in cage culture from Klong 13, Amphoe Nong-Sua, Pathum Thani Province. Journal of Agricultural Research and Academic Promotion 35(2) (special issue 2), 1044–1050.Google Scholar
Thongbumrung, W and Lerssutthichawal, T (2014) Monogeneans in cage-cultured red Tilapia (Oreochromis niloticus × O. mossambicus) in Tapi River, Nakhonsithammarat. Ubon Ratchathani University Journal of Science and Technology 16, 32–40.Google Scholar
Thongdon-A, R, Limsuwan, C and Chuchird, N (2012) Seasonality of gill monogeneans in red hybrid tilapia (Oreochromis sp.) cage culture systems in central Thailand. Journal of Fisheries and Environment 36(3), 43–53.Google Scholar
Thurston, JP (1970) The incidence of Monogenea and parasitic Crustacea on the gills of fish in Uganda. Revue de zoologie et de botanique africanines 82, 111–130.Google Scholar
Vidal-Martínez, VM, Aguirre-Macedo, ML, Scholz, T, González-Solís, D and Mendoza-Franco, EF (2001) Atlas of the helminth parasites of cichlid fish of Mexico. Academia (Academy of Sciences of the Czech Republic), Prague.Google Scholar
Wu, XY, Zhu, XQ, Xie, MQ and Li, AX (2007) The evaluation for generic-level monophyly of Ancyrocephalinae (Monogenea, Dactylogyridae) using ribosomal DNA sequence data. Molecular Phylogenetics and Evolution 44(2), 530–544.CrossRefGoogle ScholarPubMed
Wunderlich, A, Simioni, W, Zica, É and Siqueira, T (2022) Experimental evidence that host choice by parasites is age-dependent in a fish-monogenean system. Parasitology Research 121(1), 115–126.CrossRefGoogle Scholar
Zhang, S, Zhi, T, Xu, X, Zheng, Y, Bilong Bilong, CF, Pariselle, A and Yang, T (2019) Monogenean fauna of alien tilapias (Cichlidae) in south China. Parasite 26(4) https://www.parasite-journal.org/articles/parasite/full_html/2019/01/parasite180107/parasite180107.html.CrossRefGoogle ScholarPubMed
Zhi, T, Xu, X, Chen, J, Zheng, Y, Zhang, S, Peng, J, Brown, CL and Yang, T (2018) Expression of immune-related genes of Nile tilapia Oreochromis niloticus after Gyrodactylus cichlidarum and Cichlidogyrus sclerosus infections demonstrating immunosupression in coinfection. Fish & Shellfish Immunology 80, 397–404.CrossRefGoogle ScholarPubMed
Figure 0

Table 1. Host and locality records for infection by Cichlidogyrys sclerosus

Figure 1

Figure 1. Cichlidogyrus sclerosus Paperna & Thurston, 1969 (Monogenoidea: Polyonchoinea: Dactylogyridae) (USNM No. 1743705) infecting the gill filaments of Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) (Cichliformes: Cichlidae) from the E.W. Shell Fisheries Center, Auburn, Alabama (hydrologically linked to Tallapoosa River, Mobile-Tensaw Basin). Scale value aside bar. Abbreviations: accessory piece (ap), anchor (a), distal vagina (dv), eyespot (e), haptor (h), haptoral gland (hg), head organ (ho), intestine (i), marginal hook (mh), mouth (m), ovary (o), peduncle muscle (pm), penis (p), pharynx (ph), proximal vagina (pv), seminal recepticle (sr), seminal vesicle (sv), uterus (u), vitellarium (v).

Figure 2

Figure 2. Haptoral sclerites of Cichlidogyrus sclerosus Paperna & Thurston, 1969 (Monogenoidea: Polyonchoinea: Dactylogyridae) infecting the gill filaments of Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) (Cichliformes: Cichlidae) from the E.W. Shell Fisheries Center, Auburn, Alabama (hydrologically linked to Tallapoosa River, Mobile-Tensaw Basin). Scale values aside bars. (a) Ventral anchor of voucher (USNM No. 1743702), (b) Dorsal anchor of voucher (USNM No. 1743702), (c) ventral transverse bar of voucher (USNM No. 1743702), (d) dorsal transverse bar of voucher (USNM No. 1743702), (e) marginal hook I of voucher (USNM No. 1743704), (f) marginal hook II of voucher (USNM No. 1743703), (g) marginal hook III of voucher (USNM No. 1743704), (h) marginal hook IV of voucher (USNM No. 1743706), (i) marginal hook V of voucher (USNM No. 1743704), (j) marginal hook VI of voucher (USNM No. 1743707), (k) marginal hook VII of voucher (USNM No. 1743706).

Figure 3

Figure 3. Male and female reproductive genitalia of Cichlidogyrus sclerosus Paperna & Thurston, 1969 (Monogenoidea: Polyonchoinea: Dactylogyridae) (USNM No. 1743701) infecting the gill filaments of Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) (Cichliformes: Cichlidae) from the E.W. Shell Fisheries Center, Auburn, Alabama (hydrologically linked to Tallapoosa River, Mobile-Tensaw Basin). Scale value aside bar. Abbreviations: accessory piece (ap), distal tube (dt), distal vagina (dv), heel (he), ovary (o), penis (p), proximal portion (pp), proximal vagina (pv), seminal receptacle (sr), seminal vesicle (sv), testis (t), transverse vitelline duct (tv), uterus (u), vaginal pore (vp), vas deferens (vd).

Figure 4

Figure 4. Male copulatory organ of Cichlidogyrus sclerosus Paperna & Thurston, 1969 (Monogenoidea: Polyonchoinea: Dactylogyridae) (USNM No. 1743702) infecting the gill filaments of Nile tilapia, Oreochromis niloticus (Linnaeus, 1758) (Cichliformes: Cichlidae) from the E.W. Shell Fisheries Center, Auburn, Alabama (hydrologically linked to Tallapoosa River, Mobile-Tensaw Basin). Scale value aside bar. Abbreviations: accessory piece (ap), penis (p), heel (he).

Figure 5

Figure 5. Phylogenetic relationships of species within Cichlidogyrys Paperna, 1960 reconstructed using maximum likelihood tree inference using the large ribosomal subunit (28S) gene. Numbers aside nodes indicate bootstrap values. New sequence of Cichlidogyrus sclerosus Paperna & Thurston, 1969 is shown in bold. GenBank numbers are in parentheses following each taxon.

Figure 6

Figure 6. Phylogenetic relationships of species within Cichlidogyrys Paperna, 1960 reconstructed using maximum likelihood tree inference using the internal transcribed spacer 1 (ITS1). Numbers aside nodes indicate bootstrap values. New sequence of Cichlidogyrus sclerosus Paperna & Thurston, 1969 is shown in bold. GenBank numbers are in parentheses following each taxon.