Skip to main content
Download PDF

Clinical signs, morphological and phylogenetic characterization of Myxozoan spp. infecting Nile tilapia, Oreochromis niloticus and African catfish, Clarias gariepinus in Qalyubia Governorate, Egypt

BMC Veterinary Research volume 20, Article number: 530 (2024) Cite this article

Abstract

Context

Myxosporean endoparasites (phylum cnidarian) are critical pathogens that affect both wild and cultured freshwater and marine water fishes globally causing huge economic losses and high mortalities.

Study objective

The present study investigated myxosporean infections in Nile tilapia and African catfish collected from the natural resources.

Methods

A total of four hundred Nile tilapia with an average weight (60 ± 5 g) and two hundred African catfish with an average weight (185 ± 30 g) were collected seasonally from Qalyubia Governorate, Egypt for parasitological and molecular diagnosis of isolated myxozoan species.

Results

Microscopic examination revealed Myxobolus heterosporous, Myxobolus brachysporus, Myxobolus tilapiae, and Myxobolus amieti in Nile tilapia and Henneguya suprabranchiae, and Myxobolus brachysporus in African catfish. Sequencing of 18S rDNA gene for isolated Myxozoan spp. from Nile tilapia revealed Myxobolus tilapiae deposited in GenBank under accession numbers (OR766325 and OR766326). In African catfish, the isolated Myxobolus brachysporus sequence was deposited under accession numbers (OR766327 and OR766328). Henneguya suprabranchiae was also identified in African catfish (accession. No. OR763724 and OR763433).

Conclusion

Overall, these results indicate a high prevalence of myxozoan infection in naturally inhabiting Nile tilapia and African catfish. Curiously, Henneguya suprabranchiae was detected in the digestive tract and kidneys of African catfish, which is considered a rare form.

Implication

This study highlighted the importance of parasitic surveys in natural resources that impact fish production.

Peer Review reports

Introduction

Myxozoans are the most significant microscopic obligate cnidarian parasites that cause severe diseases in wild and cultured fishes all over the world [1, 2]. The complex myxozoans life cycle includes annelids and bryozoans as final invertebrate hosts and fishes as intermediate vertebrate hosts [3, 4]. Transmission is accomplished by two definite types of waterborne spores: actinospores in invertebrate hosts and myxospores in vertebrate hosts [5]. Myxozoans are a highly diverse group of parasites consisting of 2554 species [6] and their identification is mainly based on spore morphology [7, 8]. In addition, molecular characterization has been developed as a crucial tool to diagnose a wide range of new myxozoan species in combination with spore morphology [9, 10] and to ascertain its phylogenetic position within metazoans [11].

In Egypt, different myxozoan species have been recorded from different organs of wild and cultured Nile tilapia, such as Myxospora sp. [12], M. agolus, M. heterosporus type 2, M. clarri, M. heterosporous (type 3) [13, 14], Myxobolus dermatobius [15], Myxobolus cerebralis [16], Myxobolus brachysporus [17, 18], and Myxobolus tilapiae [19, 20].

Henneguya Thèlohan, 1892 is one of the most important genera of the subphylum Myxozoa and contains more than 200 described species [21]. This genus is distinguished from other members of myxobolidae based on the morphology of its symmetrical spores along with the presence of paired polar capsules and two caudal projections [22]. Although the presence of two caudal projections is an important characteristic of this genus, the previous molecular studies indicate that Henneguya is polyphyletic and that the character has independently arisen several times in Myxobolidae [23, 24]. Henneguya suprabranchiae, has been found to cause economic losses in catfish farms [25,26,27,28]. In addition, H. exilis causes serious mortality in mature catfish [29] and Henneguya ictaluri causes a proliferative gill disease [30]. Microscopic examination of Henneguya infection is the most common step in diagnosis as its spores appear elongated with a two-rounded polar capsule and long tail (spermatozoa like) [31].

Nile tilapia is the main fish species inhabiting the Nile River and one of the least expensive and most readily available fish for people [32]. It is also the most cultured freshwater species due to its many benefits including the fact that it survives in low environmental conditions, disease resistance, fast growth, and high meat quality [33]. African catfish, Clarias gariepinus, are also a popular species in the Nile River and its tributaries and have been cultured with other fish species to enhance productivity [31, 33]. In Egypt, the capture fish production from the Nile River, lakes, and sea (Red Sea and Mediterranean Sea) accounts for 21.27% of total Egyptian production in 2021. The largest percentage of caught fish arose from the northern lakes [33] which represents 12.77% of the total catch [34]. Meanwhile, production in the Nile River decreased to 3.72% compared to 3.96% in 2020 [34]. Decreases in fish production from natural resources may be due to several factors including environmental pollution and global warming which affect fish immunity and increase their susceptibility to infectious diseases. In addition, rising water temperatures and high carbon dioxide represent a critical threat to ecosystems because they accelerate the parasite life cycle and are consequently able to multiply rapidly in aquatic environments [35]. Disease outbreaks in fish are the most devastating challenge for fish production. Many freshwater fish species are seriously afflicted with various parasites, resulting in high fish mortality, reduced productivity, and negative influence on the economy [2, 36]. Nearly 80% of disease outbreaks in Egypt are due to parasitic infections [37]. Therefore, the current study was designed to investigate the prevalence of Myxozoan species infecting Nile tilapia and African catfish naturally from the El-Riah El-Tawfiki canal in Toukh, Qalyubia Governorate, lower Egypt.

Materials and methods

Samples collection

A total of four hundred Nile tilapia weighing an average of (55–65 g) and two hundred African catfish weighing an average of (150–210 g) were examined between December 2020 and September 2021. Fish were purchased from fishermen who caught them naturally from the El-Riah El-Tawfiki streams, in Toukh, Qalyubia Governorate, Egypt (Fig. 1). All fish samples were immediately transported to the Diagnostic Laboratory of Aquatic Animal Medicine Department, Faculty of Veterinary Medicine, Benha University, Egypt for clinical and parasitological examinations.

Fig. 1
figure 1

A map showing the location of studying area in Toukh, Qalyubia Governorate, Egypt

Full size image

Clinical and parasitological examination

Fish were subjected to external examination of the body surface, euthanized by an overdose of MS-222 (Sigma) at a dose of 150 mg/L, and sacrificed for internal examination according to Austin et al. [38]. Wet mount slides were prepared from naturally infected fishes showing white nodules or cysts externally or internally. These slides were prepared by gently opening the nodules using a fine needle on a clean glass slide, and the milky fluid was mixed with a drop of distilled water for microscopic examination. The positive slides were air-dried, fixed with absolute methyl alcohol (Sigma-Aldrich) and stained with Giemsa according to the method described by Abdel Ghaffar et al. [17]. The photographed slides were imported to Image J software for morphometric dimensions of spores and polar capsules of each observed spore according to Lom and Arthur [39] criteria.

Molecular identification and phylogenetic analysis

The Genomic DNA from the recovered Myxozoan spp. was extracted following manufacturer’s instructions for GF-1® Tissue DNA extraction Kit (Vivantis, Malaysia). A Nano Drop™ ND-1000 Spectrophotometer (Thermo Scientific, Germany) was used to measure the quality and concentration of the extracted DNA. The extracted DNA was stored at -20 ◦C for further molecular analysis.

A fragment of 18S ribosomal DNA (18S rDNA) genes from the retrieved Myxozoan spp. was amplified using the universal primer pair 18e and 18 g and universal primers, MX5 and MX3 for Henneguya spp. as described by Andree et al. [40]. The amplification assay was performed as described by Eszterbauer [41] and Abdel-Gaber et al. [42]. The second nested PCR amplification was performed using the primer pair MX5 and MX3 [40] for Myxozoan spp. and the myxozoan-specific primers; MC5 and MC3 [43] for Henneguya spp. (Table 1).

Table 1 List of PCR primers used in the present study with amplicons size and references
Full size table

Polymerase Chain Reaction (PCR) was conducted in a final volume of 25 µl reaction mixture comprising 12.5 µl of 2x MyTaq™ Red Mix (Cat. BIO-25043, Meridian Life Science Inc., USA), 0.5 µl of each primer (10 Mmol), and (2 µl) of aim DNA. The PCR conditions were as follows: initial denaturation for 4 min at 94 ◦C, followed by 40 cycles of 94 ◦C for 30s, 56 ◦C for 60s, and 72 ◦C for 90 s, with a final extension at 72 ◦C for 7 min. PCR products were subjected to electrophoresis in 1.5% agarose gel in a TAE buffer, stained with ethidium bromide stain (Merck, Germany), and then analyzed with a gel documentation system. For sizing and approximate quantification of double-stranded DNA, a 1 kb plus DNA Ladder (Vivantis, Malaysia) was used. The Gene JET Gel Extraction Kit (K0691, Thermo Fisher, USA) was used to clean two of the positive PCR products targeting 18S rDNA genes from Myxozoan spp. in Nile tilapia and African catfish. The sequences were then run by Macrogen Company (Korea). Two-way sequencing using the specific primers used in PCR confirmed the accuracy of the data. The programs Bioedit 7.0.4.1 and MUSCLE were used to examine the nucleotide sequences acquired in this work. Using a neighbor-joining technique for the aligned sequences implemented in the application CLC 6, the obtained sequences were aligned with reference sequences genes of Myxozoan species (Table 2).

Table 2 18S rDNA gene sequences of Myxozoan spp. from GenBank used for phylogenetic tree construction
Full size table

Statistical analysis

The seasonal and total prevalence of myxozoan infection among the examined fishes were assessed based on the obtained data using the following formula [44]; prevalence (%) = (∑ infected fish/ ∑ fish examined × 100).

Results

Clinical signs of naturally infected fishes

Nile tilapia infected with Myxozoan spp. showed slight abdominal distension, uni- and bilateral exophthalmia and white nodules in the eye (inner wall of cornea) (Fig. 2A), posterior part of the kidneys (Fig. 2B), and gills (Fig. 2C). In African catfish, yellowish-white nodules were observed in the dendritic organs, intestine, kidneys, and liver (Fig. 3A-D).

Fig. 2
figure 2

Nile tilapia infected with Myxozoan spp. showing multi white nodules in the eyes (inner wall of cornea) (arrow) (A), in the kidneys (B) and gill filaments (C)

Full size image
Fig. 3
figure 3

African catfish infected with Myxozoan spp. showed yellowish white nodules, in the dendritic organs (arrow) (A), intestine (arrows) (B), kidneys (arrow) (C). dendritic organ ( arrow) and in liver (arrow) (D)

Full size image

Microscopical observation

Myxosporean spores isolated from different organs of Nile tilapia are presented in Fig. 4. Myxobolus heterosporous spores from the kidneys were ovoid, measuring (14.84–14.05 μm) × (9.65–10.11 μm). The two polar capsules were ovoid unequal in size occupying one-third of the spore and measuring (3.71–4.68 μm) × (3.04–4.12 μm) (Fig. 4A). Myxobolus amieti spores recorded from the kidneys were pyriform with pointed anterior end and rounded posterior end, ranging (10.19–12.64 μm) in length and (7.48–7.87 μm) in width. Their two polar capsules were elongated, and equal in size occupying more than half the length of spores measuring (6.83 ± 0.3 μm) in length and (2.92 ± 0.4 μm) in width (Fig. 4B).

Fig. 4
figure 4

(A) Wet-mount preparation from nodules found in the kidneys of Nile tilapia showing M. heterosporous spores (x40); (B) Giemsa-stained smear from nodules recovered in the kidneys of Nile tilapia showing M. amieti spore (x40); (C) Giemsa-stained smear from nodules recovered in the eyes showing M. brachysporus spore (x40); (D) Giemsa-stained smear from nodules recorded in the kidneys showing M. tilapiae spore (x40)

Full size image

Myxobolus brachysporus spores isolated from the eye and gills of Nile tilapia and kidneys of African catfish were ellipsoidal and well known for their width (42. 0 μm) exceeded than length (28.42 μm) with ovoid polar capsules measuring (9.84 ± 1.06 × 7.48 ± 0.35) µm (Fig. 4C). Myxobolus tilapiae spore recorded from the kidneys was relatively large ovoid (21.34 × 16.11 μm), with rounded anterior end and nearly wide posterior end Its polar capsules were ovoid, equal in size (6.33 ± 0.24 × 5.29 ± 0.36) µm, and sporoplasm occupying almost the spore size (Fig. 4D).

In African catfish, the examined nodules from dendritic organs, kidneys, and intestines showed mature spores of Henneguya spp. which resemble spermatozoon. The spores were elongated anteriorly, had rounded anterior ends, were fusiform and had clear vacuoles (Fig. 5A.B). Moreover, M. brachysporus was also isolated from the kidneys of African catfish, similar to that recorded in Nile tilapia.

Fig. 5
figure 5

A: Wet-mount preparation from nodules recorded in the dendritic organ of African catfish showing Henneguya spp. spores resembles spermatozoon (X40), B: Giemsa-stained smear from nodules in the intestine of African catfish showing Henneguya spp. spores (X40)

Full size image

Total and seasonal prevalence rate

The total and seasonal prevalence of Myxozoan infections among the examined Nile tilapia are presented in Table 3. Myxozoan infection in Nile tilapia revealed total prevalence 12% (48/400), representing 8.75% infection with M. brachysporus 2.75% infection with M. amieti, 0.25% infection with M. heterosporous, and 0.25% infection with M. tilapiae. The highest seasonal prevalence was 21% in spring, including (15% M. brachysporus, 5% M. amieti, and 1% M. tilapiae). In winter, the infection rate reaches 13%, including infection with M. brachysporus and M. amieti at the rate of 9% and 4%, respectively. Meanwhile, the seasonal prevalence in autumn was 9%, involving (7% M. brachysporus, 1% M. amieti, and 1% M. heterosporous). The lowest prevalence was 5% in summer, including infection with M. brachysporus at a rate of 4% and M. amieti at prevalence rate of 1% (Table 3. The total tissue susceptibility to Myxozoan spp. infection in Nile tilapia was as follows: from the eyes (M. brachysporus and, M. amieti) at a rate of 11%, from the kidneys (M. amieti,M. heterosporous and M. tilapiae) at a rate of 0.75%, and from gill filaments (M. brachysporus) at a rate of 0.25%.

Table 3 Total and seasonal prevalence of myxozoan infection in Nile tilapia
Full size table

In African catfish, total prevalence of Henneguya spp. infection was 24%, with seasonal prevalences 32%, 22%, 24 and 18% in winter, spring, summer, and autumn, respectively. M. brachysporus was recorded at a total prevalence of 2.5% and the seasonal infection rate was high in winter and autumn (4%), followed by spring (2%), with no record in winter (0%) (Table 4).

Table 4 Total and seasonal prevalence of myxozoan infection in African catfish
Full size table

Molecular and phylogenetic tree building

PCR amplification of the 18S rDNA for Myxozoan spp. isolated from Nile tilapia revealed 1300 bp and that isolated from African catfish showed 1000 bp. Phylogenetic analysis recorded M. tilapiae firmly embedded within the family Myxobolidae and showed 99% similarity to that of M. tilapiae (KX632950) and 89% similarity to that of M. tilapiae (MZ090095). Also, revealed 72% similarity with M. brachysporus (KX632949) and 68% similarity with that of Triangula egyptica (KX632951) (Fig. 6.M. tilapiae identified from Nile tilapia was deposited in GenBank under accession numbers (OR766325, OR766326). M. brachysporus sequences isolated from African catfish were deposited in the GenBank under accession numbers (OR766327, OR766328). It showed 100% similarity to that of M. brachysporus (KX632949),64% similarity to that of M. agolus (KX632949), and 78% with M. cerebralis (AY479924) (Fig. 7). The assembled sequence of 18S rDNA of Myxozoan spp. isolated from African catfish showed H. suprabranchiae that deposited in the GenBank under the accession number (OR763724, OR763433). H. suprabranchiae of this study showed 100% similarity to that of H. suprabranchiae described from Clarias gariepinus (JN201199) (Fig. 8).

Fig. 6
figure 6

Phylogenetic tree was re-constructed based on the 18S rDNA sequences of M. tilapiae (OR766325 and OR766326) and its closest Myxobolus species. and Henneguya spp. using the neighbor-joining method

Full size image
Fig. 7
figure 7

Phylogenetic tree was re-constructed based on the 18S rDNA sequences of M. brachypterous (OR766327 - OR766328) and its closest Myxobolus spp. and Henneguya species using the neighbor-joining method

Full size image
Fig. 8
figure 8

Phylogenetic tree was re-constructed based on the 18S rDNA sequences of Henneguya suprabranchiae (OR763724 and OR763433) and its closest Henneguya species using the neighbor-joining method

Full size image

Discussion

Freshwater fishes are susceptible to parasitic diseases, including Myxosporean infection. In the present study, Nile tilapia infected with Myxozoan spp. showed slight abdominal distension, uni- or bilateral exophthalmia, and creamy white nodules of variable size around the iris, liver, gills, and kidneys. Similarly, macroscopic creamy whitish nodules in the eyes and gills of wild Nile tilapia infected with myxobolus spp. were recorded [12, 14, 45]. In addition, numerous white cysts of Myxobolus dermatobius have been recorded in the eyes of cultured Nile tilapia in Sharkia Governorate, Egypt [15]. In the present study, the clinical and postmortem lesions of henneguyosis, appeared as oval to round yellowish to white nodules in the dendritic organs, intestine, and kidneys of African catfish. Similar signs and lesions have been previously recorded [12, 17, 46,47,48].

Microscopic examination in this study revealed different Myxozoan spp. in different tissues. M. brachysporus spores isolated from the eyes and gills were ellipsoidal (42. 0 μm width, 28.42 μm in length), with ovoid polar capsules (9.84 ± 1.06 × 7.48 ± 0.35 μm). This species has been recorded in Nile tilapia in the kidneys, liver, and spleen [14, 18, 42, 49, 50]. M. tilapiae spores were also recorded in our study from kidneys which appear large ovoid with rounded anterior end and nearly wide posterior end measuring (21.34 × 16.11 μm). Mohammed et al. [13] recorded M. tilapiae from the gills of Tilapia zilli collected from the Nile River in Qena Governorate, Egypt. Abdel-Baki et al. [19] and Abdel- Gaber et al. [42] reported M. tilapiae from the kidneys measuring (15.3 ± 0.2 μm and 15.0 ± 0.5 μm) in length and (10.3 ± 0.1 μm and 11 ± 0.3 μm) in width, respectively. Moreover, M. tilapiae spores of an average (12.5 ± 0.67 μm in length × (5.7-± 1.2 μm in width) were observed in the gills of cultured Nile tilapia [20].

Moreover, in our study, M. heterosporous spores from the kidneys measuring (14.84–14.05) × (9.65–10.11) µm were recorded. Baker [51] recorded M. heterosporous spores (13 − 16 × 7–9.2 μm) from the gills of Nile tilapia, East Africa. Soror et al. [14] revealed M. heterosporous spores (18.52 μm length×11.54 μm width) from kidneys of Nile tilapia collected from El-Riah El-Tawfiki, Qaliobia Governorate, Egypt, and Eissa et al. [16] isolated the spore from the cornea of Nile tilapia at Abbasa Fish Farm, Sharkiya Governorate, Egypt. M. heterosporous spores (9.7 μm in length× 7 μm in width) were also recorded in the liver and intestines of naturally collected Nile tilapia from the Nile River [50]. In the current study the recorded spores of M. amieti from the kidneys appeared pyriform ranging from (10.19–12.64) µm in length and (7.48–7.87) µm in width with two equal elongated polar capsules (6.83 ± 0.3 × 2.92 ± 0.4 μm) occupying more than half the length of spore. M. amieti spores were previously recorded in the kidneys of Nile tilapia measuring (16.09 × 9.42 μm) with elongated polar capsule (9.91 × 3.09 μm) [14]. Our results were similar to the morphological description of mature spores of Henneguya spp. recorded in the organs of African catfish [12, 17, 31, 48, 52].

The overall prevalence of Myxozoan infections among the examined Nile tilapia in the current study was 12%, with the highest infection rate of 12% in spring and the lowest rate of 5% in summer. Shehab El-Din [45] recorded the total myxozoan infection of 10.6% in wild Nile tilapia with a seasonal prevalence of 3.03%, 2.08%, 0%, and 5% in winter, spring, summer, and autumn, respectively. Our results have been found to be lower than the data published by Soror et al. [14] who recorded 83.46% prevalence of Myxosporean infections in Nile tilapia, with the highest infection rate in autumn (95.08%) and the lowest rate was in summer (76%). In addition, Matter et al. [12] recorded that total prevalence of myxosporidiosis in Nile tilapia was 24.0%, and the highest seasonal prevalence was observed in winter (43.4%), whereas the lowest rate was recorded in summer (8%). Abdel-Baki et al. [19] and El-Asely et al. [37] respectively also reported higher infection rates with 61% and 100%. This difference could be attributed to differences in localities and water sources. Ali [53] reported that the highest prevalence of myxosporidiosis was in spring, explaining that parasites begin to form nodules in winter, the maximum number reaches in spring, and then starts to decrease after rupture of cysts to release spores in the environment.

In the present study, the infection with M. brachysporus reached 8.75% in Nile tilapia, and 2.5% in African catfish. On the other hand, Abdel-Baki et al. [18] recorded total infection rate of M. brachysporus in Nile tilapia collected from the Nile River was 51.9% and Georges et al. [49] observed 12.29% infection rate in Nile tilapia from Adamawa-Cameroon. M. brachysporus at prevalence of (12%) with the highest incidence rate of 14% in winter and 10% in summer was recorded by Abdel-Gaber et al. [42]. The examined Nile tilapia in the present study revealed total prevalence 0.25% for M. heterosporous. This result is lower than that reported by Georges et al. [49].

For M. tilapiae recorded in the present study, the total infection rates reached 0.25%, which only isolated in spring season. In contrast, Abdel-Gaber et al. [42] recorded that M. tilapiae among the examined Nile tilapia exhibited overall prevalence of 6%, representing 8% incidence in winter and 4% occurrence in summer. In addition, Georges et al. [49] observed a 15.14% infection rate with M. tilapiae. These results could be attributed to the availability of intermediate hosts and the increase in fish feeding activity at warm temperatures [54, 55]. Moran et al. [56] attributed the variation between the seasonal prevalence of myxosporean infection, which may be due to variation in the environmental conditions, and the time of exposure which may extended to 5–6 month.

The total prevalence of henneguyasis was 24%, with seasonal prevalence of 32%, 22%, 24%, and 20% in winter, spring, summer, and autumn, respectively. The overall prevalence of infection with Henneguya spp. which have been previously reported was 18% [57], 40% [58], and 43.65% [12]. Furthermore, Henneguya spp. in Nile tilapia was recorded by Ramadan et al. [59] at a rate of 8.4% [12]. The difference in infection rates could be attributed to fish feeding behavior as a carnivorous species that assists in the transmission of more enteric parasites by feeding on aquatic animals that harbor the infective stage of these parasites or even fed on small infected fish [23].

The identification of diverse species of myxosporidian spores on a morphological and morphometric basis, is more difficult due to their abundance. Therefore, molecular assays based on small subunit ribosomal DNA gene sequences are the most sensitive method for accurate identification [60]. Molecular identification using 18s rDNA gene sequences of Myxozoan spp. in the present study, recorded M. tilapiae from the kidneys of Nile tilapia which deposited in the GenBank (OR766325, OR766326) and showed 89 − 99% similarity to that of M. tilapiae,72% similarity with M. brachysporus, and 68% similarity to that of Triangula egyptica. Eissa et al. [20] recorded that M. tilapiae (MZ090095) showed 99.46% identity with M. tilapiae (KX632950), 98.85% similarity with M. brachysporus, 98.77% similarity with Triangula egyptica, and 97.45% similarity with M. cerebralis. Abdel-Gaber et al. [42] reported that the amplified and sequenced SSU rDNA gene regions for the recovered Myxospora sp. from Nile tilapia had 95% identity with all Myxobolus species available in GenBank. In this study, M. brachysporus that isolated from the kidneys of African catfish with accession numbers (OR766327, OR766328) showed 100% similarity to that of M. brachysporus (KX632949),64% similarity to that of M. agolus (KX632949), and 78% identity with M. cerebralis (AY479924) in Nile tilapia in Egypt. In the current study, molecular assays of Henneguya spp. recovered from the dendritic organs and kidneys of African catfish recorded H. suprabranchiae sequences deposited in GenBank (OR763724, OR763433), which exhibited 100% similarity to that of H. suprabranchiae (JN201199) reported by Morsy et al. [31].

Conclusion

The results of the present study revealed that Nile tilapia and African catfish collected from natural water resources were infected with myxozoan spp. The prevalence was higher in African catfish than that in Nile tilapia. Molecular and phylogenetic tree building as an important diagnostic tool, confirmed M. tilapiae in Nile tilapia, and M. brachysporus and H. suprabranchiae in African catfish. Further investigations are needed to estimate the economic impacts of myxozoan infection on natural captures and total productivity.

Data availability

https://www.ncbi.nlm.nih.gov/nuccore/OR766325 https://www.ncbi.nlm.nih.gov/nuccore/OR766326 https://www.ncbi.nlm.nih.gov/nuccore/OR766327 https://www.ncbi.nlm.nih.gov/nuccore/OR766328 https://www.ncbi.nlm.nih.gov/nuccore/OR763724 https://www.ncbi.nlm.nih.gov/nuccore/OR763433.

References

  1. Shafiq A, Abbas F, Hafeez-ur-Rehman M, Khan BN, Aihetasham A, Amin I, Hmidullah, Mothana RA, Alharbi MS, Khan I, Khalil AA. Parasite diversity in a freshwater ecosystem. Microorganisms. 2023;11(8):1940.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  2. Maor-Landaw K, Smirnov M, Brekhman V, Ofek-Lalzar M, Yahav T, Lotan T. Infection by the Parasite Myxobolus bejeranoi (Cnidaria: Myxozoa) suppresses the immune system of hybrid tilapia. Microorganisms. 2022;10(10):1893.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  3. Holzer AS, Bartošová-Sojková P, Born‐Torrijos A, Lövy A, Hartigan A, Fiala I. The joint evolution of the Myxozoa and their alternate hosts: a cnidarian recipe for success and vast biodiversity. Mol Ecol. 2018;27(7):1651–66.

    Article  PubMed  Google Scholar 

  4. Holzer AS, Piazzon MC, Barrett D, Bartholomew JL, Sitjà-Bobadilla A. To react or not to react: the dilemma of fish immune systems facing myxozoan infections. Front Immunol. 2021;12:734238.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  5. Kent ML, Margolis L, Whitaker DJ, Hoskins GE, McDonald TE. Review of Myxosporea of importance in salmonid fisheries and aquaculture in British Columbia. Folia Parasitol. 1994;41(1):27–38.

    CAS  Google Scholar 

  6. Chinh NN, Ha NT, Doanh PN, Eiras JC, Whipps CM, Shirakashi S. Synopsis of myxosporean species (Cnidaria: Myxozoa) parasitizing fishes from Vietnam. Syst Parasitol. 2023;100(4):325–44.

    Article  PubMed  Google Scholar 

  7. Abdel-Ghaffar F, Abdel-Gaber R, Maher S, El Deeb N, Kamel R, Al Quraishy S, Mehlhorn H. Morphological and ultrastructural characteristics of Myxobolus Ridibundae n. sp. (Myxosporea: Bivalvulida) infecting the testicular tissue of the marsh frog Rana ridibunda (Amphibia: Ranidae) in Egypt. Parasitol Res. 2017;116:133–41.

    Article  PubMed  Google Scholar 

  8. Mathews PD, Naldoni J, Adriano EA. Morphology and small subunit rDNA-based phylogeny of a new Henneguya species, infecting the ornamental fish Corydoras leucomelas from the Peruvian Amazon. Acta Trop. 2017;176:51–7.

    Article  PubMed  Google Scholar 

  9. Iwanowicz DD, Iwanowicz LR, Howerth EW, Schill WB, Blazer VS, Johnson RL. Characterization of a new myxozoan species (Myxozoa: Myxobolidae: Myxosporea) in largescale stonerollers (Campostoma oligolepis) from the Mobile River Basin (Alabama). J Parasitol. 2013;99(1):102–11.

    Article  CAS  PubMed  Google Scholar 

  10. Alama-Bermejo G, Hernández-Orts JS, García-Varela M, Oceguera-Figueroa A, Pecková H, Fiala I. Diversity of myxozoans (Cnidaria) infecting neotropical fishes in southern Mexico. Sci Rep. 2023;13(1):12106.

    Article  PubMed  PubMed Central  Google Scholar 

  11. Mallatt J, Craig CW, Yoder MJ. Nearly complete rRNA genes from 371 Animalia: updated structure-based alignment and detailed phylogenetic analysis. Mol Phylogenet Evol. 2012;64(3):603–17.

    Article  CAS  PubMed  Google Scholar 

  12. Matter AF, Abbass AA, Abd El-Gawad EA, El-Asely AM, Shaheen AA. Studies on myxosporidiosis in some freshwater fishes. Benha Veterinary Med J. 2013;25(2):316–25.

    Google Scholar 

  13. Mohammed NI, Rabie SA, Hussein AN, Hussein NM. Infestation of Oreochromis niloticus and Tilapia zilli fresh-water fishes with myxosporean parasites, Qena Province, Egypt. Egypt Acad J Biol Sci B Zool. 2012;4(1):235–46.

    Google Scholar 

  14. Soror EI, Mahrous KF, Ismail AM, Abbass AA. Epizotiological studies on proliferative kidney disease in tilapia (Oreochromis niloticus) and African catfish (Clarias gariepinus). Benha Vet Med J. 2012;23(1):150–8.

    Google Scholar 

  15. ElSayed NM. Ultrastructural morphology of the Myxobolus Dermatobius Ishii 1915 (Mixosporea: Myxobolideae) microspores infecting eyes of Nile tilapia (Oreochromis niloticus) in Egypt. Vet Ital. 2020;56(4):251–5.

    PubMed  Google Scholar 

  16. Eissa AE, Mourad A, Borhan T. A contribution on Myxosoma infection in cultured Oreochromis niloticus in Lower Egypt. Nat Sci. 2006;4:40–6.

    Google Scholar 

  17. Abdel-Ghaffar F, Morsy K, El-Ganainy S, Ahmed M, Gamal S, Bashtar AR, Al Quraishy S, Mehlhorn H. Twelve myxosporean species of the family Myxobolidae infecting freshwater fishes of the River Nile, Egypt, with the description of four novel species. Parasitol Res. 2015;114:2985–98.

    Article  PubMed  Google Scholar 

  18. Abdel-Baki AA, Zayed E, Sakran T, Al-Quraishy S. A new record of Myxobolus Brachysporus and M. Israelensis in the tilapia (Oreochromis niloticus) collected from the Nile River, Egypt. Saudi journal of biological sciences. 2015;22(5):539–42.

  19. Abdel-Baki AA, Abdel-Haleem HM, Sakran T, Zayed E, Ibrahim KE, Al-Quraishy S. Two Myxobolus spp. infecting the kidney of Nile tilapia (Oreochromis niloticus) in the River Nile at Beni-Suef governorate, Egypt, and the associated renal changes. Parasitol Res. 2015b;114:1107–12.

    Article  PubMed  Google Scholar 

  20. Eissa AE, Attia MM, Elgendy MY, Ismail GA, Sabry NM, Prince A, Mahmoud MA, El-Demerdash GO, Abdelsalam M, Derwa HI, Streptococcus. Centrocestus formosanus and Myxobolus tilapiae concurrent infections in farmed Nile tilapia (Oreochromis niloticus). Microbial Pathogenesis. 2021; 158:105084.

  21. Fiala I, Bartoˇsov´a-Sojkov´a P, Whipps CM. Classification and phylogenetics of Myxozoa. In: Okamura B, Gruhl A, Bartholomew JL, editors. Myxozoan Evolution, Ecology and Development. Springer Cham; 2015. pp. 85–110.

  22. Lom J, Dykova I. Protozoan parasites of fishes. Developments in aquaculture and fisheries Science. Amsterdam): Elsevier; 1992. p. 315.

    Google Scholar 

  23. Kent ML, Andree KB, Bartholomew JL, EL-Matbouli MA, Desser SS, Devlin RH, Feist SW, Hedrick RP, Hoffmann RW, Khattra J, Hallett SL. Recent advances in our knowledge of the Myxozoa. J Eukaryot Microbiol. 2001;48(4):395–413.

  24. Liu XH, Yuan S, Zhao YL, Fang P, Chen H, Zhang JY. Morphological and molecular characterization of Myxobolus sheyangensis n. sp. s(Myxosporea: Myxobolidae) with intralamellar sporulation in allogynogenetic gibel carp, Carassius auratus Gibelio (Bloch) in China. Parasitol Res. 2016;115:3567–74.

    Article  CAS  PubMed  Google Scholar 

  25. McCRAREN JP, Landolt ML, Hoffman GL, Meyer FP. Variation in response of channel catfish to Henneguya sp. infections (Protozoa: Myxosporidea). J Wildl Dis. 1975;11(1):2–7.

    Article  Google Scholar 

  26. Landsberg JH. Myxosporean parasites of the catfish, Clarias lazera (Valenciennes). Syst Parasitol. 1987;9(1):73–81.

    Article  Google Scholar 

  27. Ashmawy KI, SA AE, Imam EA, El-Otifi YZ. Description of newly recorded myxosporidian protozoa of freshwater fishes in Behera province, Egypt. J Egypt Vet Med Ass. 1989;49(1–2):43–53.

    Google Scholar 

  28. Abdel Ghaffar F, El-Shahawi G, Naas S. Myxosporidia infecting some Nile fishes in Egypt. Parasitol Res. 1995;81:163–6.

    Article  CAS  PubMed  Google Scholar 

  29. CURRENT WL, JANOVY JO JR. Comparative study of ultrastructure of interlamellar and intralamellar types of Henneguya exilis Kudo from channel catfish. J Protozoology. 1978;25(1):56–65.

  30. Pote LM, Hanson LA, Shivaji R. Small subunit ribosomal RNA sequences link the cause of proliferative gill disease in channel catfish to Henneguya n. sp. (Myxozoa: Myxosporea). J Aquat Anim Health. 2000;12(3):230–40.

    Article  Google Scholar 

  31. Morsy K, Abdel-Ghaffar F, Bashtar AR, Mehlhorn H, Al Quraishy S, Abdel-Gaber R. Morphology and small subunit ribosomal DNA sequence of Henneguya suprabranchiae (Myxozoa), a parasite of the catfish Clarias gariepinus (Clariidae) from the River Nile, Egypt. Parasitol Res. 2012;111:1423–35.

    Article  PubMed  Google Scholar 

  32. FAO. Food and Agriculture Organization of the United Nations Fisheries and Aquaculture Division. Rome. (2022).

  33. Shaalan M, El-Mahdy M, Saleh M, El-Matbouli M. Aquaculture in Egypt: insights on the current trends and future perspectives for sustainable development. Reviews Fisheries Sci Aquaculture. 2018;26(1):99–110.

    Article  Google Scholar 

  34. LFRPDA. Lakes and Fish Resources Protection and Development Agency. Statistical report on Fish Production. Egypt: Cairo; 2021.

    Google Scholar 

  35. Brander K, Cochrane K, Barange M, Soto D. Climate change implications for fisheries and aquaculture. In: Phillips BF, Pérez-Ramírez M, editors. Climate Change impacts on fisheries and Aquaculture. Wiley; 2017. pp. 45–62.

  36. Shinn AP, Dong HT, Vinh NT, Wongwaradechkul R, Lio-Po GD. Infectious diseases of warmwater fish in fresh water. Climate Change on diseases and disorders of Finfish in Cage Culture. GB: CABI; 2023. pp. 202–77.

    Chapter  Google Scholar 

  37. El-Asely AM, Abd El-Gawad EA, Soror EI, Amin AA, Shaheen AA. Studies on some parasitic diseases in Oreochromis niloticus fish hatchery with emphasis to life stages. J Adv Veterinary Res. 2015;5(3):99–108.

    Google Scholar 

  38. Austin B, Austin DA, Munn CB. Bacterial fish pathogens: disease of farmed and wild fish. Dordrecht, The Netherlands: Springer; 2007.

    Google Scholar 

  39. Lom J, Arthur JR. A guideline for the preparation of species descriptions in Myxosporea. J Fish Dis. 1989;12(2):151–6.

    Article  Google Scholar 

  40. Andree KB, Székely C, Molnár K, Gresoviac SJ, Hedrick RP. Relationships among members of the Genus Myxobolus (Myxozoa: Bilvalvidae) based on small subunit ribosomal DNA sequences. J Parasitol. 1999: 68–74.

  41. Eszterbauer E. Genetic relationship among gill-infecting Myxobolus species (Myxosporea) of cyprinids: molecular evidence of importance of tissue-specificity. Dis Aquat Organ. 2004;58(1):35–40.

    Article  PubMed  Google Scholar 

  42. Abdel-Gaber R, Abdel-Ghaffar F, Maher S, El-Mallah AM, Al Quraishy S, Mehlhorn H. Morphological re-description and phylogenetic relationship of five myxosporean species of the family Myxobolidae infecting Nile tilapia. Dis Aquat Organ. 2017;124(3):201–14.

    Article  CAS  PubMed  Google Scholar 

  43. Molnár K, Eszterbauer E, Székely C, Dán Á, Harrach B. Morphological and molecular biological studies on intramuscular Myxobolus spp. of cyprinid fish. J Fish Dis. 2002;25(11):643–52.

    Article  Google Scholar 

  44. Margolis L, Esch GW, Holmes JC, Kuris AM, Schad GA. The use of ecological terms in parasitology (report of an adhoc committee of the American society of parasitologists). J Parasitol. 1982;68(1):131–3.

    Article  Google Scholar 

  45. Shehab El-Din MTE. Studies on some nodular parasitic diseases of fish. Ph.D thesis (Fish Diseases and Management), Faculty of Veterinary Medicine, Benha University, Egypt. 2008.

  46. Sabri DM, Eissa IA, Danasoury MA, Khouraiba HM. Prevalence of Henneguya branchialis in catfish (Clarias gariepinus) in Ismailia, Egypt. Int J Agric Biology. 2010;12:897–900.

    Google Scholar 

  47. Woodyard ET, Rosser TG, Stilwell JM, Camus AC, Khoo LH, Waldbieser G, Lorenz WW, Griffin MJ. New data on Henneguya Postexilis Minchew, 1977, a parasite of channel catfish Ictalurus punctatus, with notes on resolution of molecular markers for myxozoan phylogeny. Syst Parasitol. 2022:1–22.

  48. Emeish WF, Fawaz MM, Hussein NM, Al-Amgad Z, Abd-ElHafeez HH, Rutland CS, Bakry KA. Characterizing two New Henneguya Species in the respiratory organs of African Sharptooth Catfish. Microsc Microanal. 2023;29(5):1774–90.

    Article  PubMed  Google Scholar 

  49. Georges FO, Marc KK, Timoléon TC, Minette TE, Joseph TC. Myxobolus (Myxosporea: Myxobolidae) polyinfection patterns in Oreochromis niloticus in Adamawa-Cameroon. Int J Fish Aquat Stud. 2021;9:123–30.

    Article  Google Scholar 

  50. El-Khayat HM, Sayed SS, Mohammed WA, Sadek AS. Protozoan and helminths infestation of Nile tilapia Oreochromis Niloticus and its correlation with certain water quality variables along river Nile in the area of Greater Cairo. Environ Pollut. 2024;345:123459.

    Article  CAS  PubMed  Google Scholar 

  51. Baker JR. Three new species of Myxosoma (Protozoa: Myxosporidia) from east African freshwater fish. Parasitology. 1963;53(1–2):285–92.

    Article  Google Scholar 

  52. Figueredo RT, Müller MI, Arana S, Long PF, Adriano EA. Phylogenetic and host-parasite relationship analyses of Henneguya caquetaia sp. nov (Myxosporea: Myxobolidae) infecting an amazonian cichlid fish. Microb Pathog. 2023;179:106116.

    Article  PubMed  Google Scholar 

  53. Ali MA. Henneguya Ghaffari sp. n. (Myxozoa: Myxosporea), infecting the Nile perch lates niloticus (Teleostei: Centropomidae). Dis Aquat Organ. 1999;38(3):225–30.

  54. Golomazou E, Athanassopoulou F, Karagouni E, Kokkokiris L. The effect of seasonality on the health and growth of a newly recorded Myxobolus species infecting cultured sharp snout seabream (Diplodus puntazzo C). Turkish J Veterinary Anim Sci. 2009;33(1):1–5.

    Google Scholar 

  55. Ousman A, Félix BB, Thomas N, Abraham F. Structure and dynamics of myxosporean parasites component communities in two freshwater cichlids in the Chari River (Republic of Chad). Pakistan J Biol Sciences: PJBS. 2007;10(5):692–700.

    Article  Google Scholar 

  56. Moran JD, Whitaker DJ, Kent ML. A review of the myxosporean genus Kudoa Meglitsch, 1947, and its impact on the international aquaculture industry and commercial fisheries. Aquaculture. 1999;172(1–2):163–96.

    Article  Google Scholar 

  57. Abd rabo ER, Ezz AE, Abbass AE, Abdel-Gawad AA, El-Asely EA, El-Abd AM, Shaheen H. Isolation of some parasitic diseases from African catfish (Clarias gariepinus) in downstream El-Rahawy drain. Benha Veterinary Med J. 2017;33:233–43.

    Google Scholar 

  58. ALY ME, Dyab AK, Arafa MI, Darwish M, Gareh A. Prevalence and morphological characterization of parasitic infection in freshwater fish in Sohag Province, Egypt. Assiut Veterinary Med J. 2024;70(182):340–50.

    Article  Google Scholar 

  59. Ramadan RM, Mahdy OA, El-Saied MA, Mohammed FF, Salem MA. Novel insights into immune stress markers associated with myxosporeans gill infection in Nile tilapia (molecular and immunohistochemical studies). PLoS ONE. 2024;19(6):e0303702.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  60. Iwanowicz LR, Iwanowicz DD, Pote LM, Blazer VS, Schill WB. Morphology and 18s rDNA of Henneguya Gurlei (myxosporea) from Ameiurus nebulosus (siluriformes) in North Carolina. J Parasitol. 2008;94(1):46–57.

    Article  CAS  PubMed  Google Scholar 

Download references

Acknowledgements

The authors would like to thank their respective universities and institutes for their support and assistance.

Funding

Open access funding provided by The Science, Technology & Innovation Funding Authority (STDF) in cooperation with The Egyptian Knowledge Bank (EKB).

Author information

Authors and Affiliations

  1. Aquatic Animal Medicine Department, Faculty of Veterinary Medicine, Benha University, Moshtohor, Toukh, Qalyubia, 13736, Egypt

    Doaa A. Yassen, Eman A. Abd El-Gawad & Amany A. Abbass

  2. Animal Reproduction Department, Veterinary Research Institute, National Research Centre (NRC), Dokki, Cairo, 12622, Egypt

    Khaled A. Abd El-Razik

  3. Cell Biology Department, Biotechnology Research Institute, National Research Centre (NRC), Dokki, Cairo, 12622, Egypt

    Karima F. Mahrous

Authors
  1. Doaa A. Yassen
    View author publications

    You can also search for this author inPubMed Google Scholar

  2. Eman A. Abd El-Gawad
    View author publications

    You can also search for this author inPubMed Google Scholar

  3. Khaled A. Abd El-Razik
    View author publications

    You can also search for this author inPubMed Google Scholar

  4. Karima F. Mahrous
    View author publications

    You can also search for this author inPubMed Google Scholar

  5. Amany A. Abbass
    View author publications

    You can also search for this author inPubMed Google Scholar

Contributions

D.A.Y. Methodology and investigation, writing the original draft. E.A.A and A.A.A conceptualization, methodology, review, editing, supervision, and correspondence. K.A.A and K.F.M. methodology, investigation, review, and editing. All the authors have read and approved the final manuscript.

Corresponding authors

Correspondence to Eman A. Abd El-Gawad or Amany A. Abbass.

Ethics declarations

Ethical approval

The study was conducted following the protocol involving the use of animals that was approved by the Benha University, Faculty of Veterinary Medicine Animal Care and Use Committee (BUFVTM 13-10-22). All fish-handling procedures and regulations followed the ARRIVE guidelines for Animal Care and Use. All applicable institutional and governmental rules relating to the ethical use of animals were followed. No written consent was obtained from the fishermen for used fish in the study.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.

Additional information

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Electronic supplementary material

Below is the link to the electronic supplementary material.

Supplementary Material 1

Supplementary Material 2

Supplementary Material 3

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Yassen, D.A., Abd El-Gawad, E.A., Abd El-Razik, K.A. et al. Clinical signs, morphological and phylogenetic characterization of Myxozoan spp. infecting Nile tilapia, Oreochromis niloticus and African catfish, Clarias gariepinus in Qalyubia Governorate, Egypt. BMC Vet Res 20, 530 (2024). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s12917-024-04378-0

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s12917-024-04378-0

Keywords

BMC Veterinary Research

ISSN: 1746-6148

Contact us