Skip Navigation

ICES Journal of Marine Science: Journal du Conseil 2006 63(1):169-175; doi:10.1016/j.icesjms.2005.08.010
This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrowRequest Permissions
Google Scholar
Right arrow Articles by Pascual, S.
Right arrow Articles by Azevedo, C.
Right arrow Search for Related Content
PubMed
Right arrow Articles by Pascual, S.
Right arrow Articles by Azevedo, C.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?

© 2005 International Council for the Exploration of the Sea

Host–parasite interaction of a muscle-infecting didymozoid in the Atlantic mackerel Scomber scombrus L.

S. Pascuala,*, E. Abollob and C. Azevedoc

a Instituto de Investigaciones Marinas (CSIC), ECOBIOMAR Eduardo Cabello 6, 36208, Vigo, Spain
b Centro de Investigacións Mariñas, CIMA Pedras do Corón s/n., 36620 Vilanova de Arousa, Pontevedra, Spain
c Department of Cell Biology, Institute of Biomedical Sciences and CIIMAR, University of Oporto Largo Prof. Abel Salazar, 4099-003 Porto, Portugal

*Correspondence to S. Pascual: tel: +34 986 23 1930, ext 183; fax: +34 986 29 2762. e-mail: spascual{at}iim.csic.es.

Numerous ribbon-like aggregations of eggs of a trematode didymozoid were found embedded in the connective tissue, mostly between the epoxial skeletal muscle fibres, of mature mackerel Scomber scombrus collected from post-recruits of the eastern stock. Egg masses contained numerous developmental stages, including pre-vitellogenous oocytes to fully embryonated eggs. Genomic DNA isolation, PCR amplification, cloning, and sequencing strongly supported the view that the eggs belong to a yet unidentified didymozoid. There was minor muscular damage to the host, namely focal distortion of host fibre architecture to accommodate the parasite, and the mild host response consisted of different stages of encapsulation. Concurrent heavy infection makes the mackerel flesh unappealing to eat, although the quality of the fish is not significantly affected and there is no public health danger.

Keywords: didymozoid, infected muscle, mackerel, trematode

Received 17 June 2005; accepted 28 August 2005.


   Introduction
Top
 Introduction
Material and methods
 Results
 Discussion
 References
 
Flesh quality is a concern to any industry that sells a fresh meat product. Poor muscle quality of fish has been attributed to many factors, including processing problems, high fat feeds, stress, and quality of the fish at harvest (Talbot, 1994). Parasites have also been considered a factor critical in influencing conventional seafood quality and price (Malouf, 1986; Alvarez-Pellitero and Sitja-Bobadilla, 1993). Infections by some microsporidians, especially multivalvulid myxosporidians, have been associated with post-harvest autolysis in several marine fish species (Lom and Dyková, 1992). Among the largest, most numerous, and most diversified assembly of muscle-invading marine parasites are the free or encysted stages of didymozoid digenean trematodes and anisakid nematodes. Both have a long history of infection of large areas of muscle in a variety of fish species worldwide (Lester, 1980; Koie and Lester, 1985; Korotaeva, 1985; Abollo et al., 2001; Karlsbakk, 2001). Deterioration of fish condition and loss of marketability have been related to extensive fish muscle degeneration, parasite invasion, and chronic inflammatory reactions. Further, Thompson (2000) recorded emerging zoonoses and allergies associated with the presence of these helminth parasites in fish muscle, underlining the economic and public health importance of instituting control measures for edible seafood during inspections.

The Atlantic mackerel Scomber scombrus is an epipelagic and mesodemersal species that is most abundant in cold and temperate shelf sea areas. Its total catch reported to the FAO for 1995 was 610 947 t (www.fao.org), most of which was traded fresh, frozen, smoked, or canned for human consumption. Despite its importance mostly to local economies, little is known about agent–host interactions involving it. This study provides data on the host–parasite interactions of a muscle-infecting didymozoid in the eastern stock of Atlantic mackerel (Gibson et al., 1981), addressing the molecular characterization of the parasite, analysing prevalence data, and investigating the ultrastructure of host–parasite interactions.


   Material and methods
Top
 Introduction
 Material and methods
Results
 Discussion
 References
 
Sampling
During 2002, 200 post-recruiting fish of the eastern stock of Atlantic mackerel were collected by fishers operating on a groundfish spawning area near Vigo on the North Atlantic coast of Spain (ICES Division IXa North). In all, 50 were collected each season, winter (January–March), spring (April–June), summer (July–September), and autumn (October–December). The study was carried out over a full year, the duration of a fish cohort, to explore potential seasonal differences in infection. The entire musculature of each market-size fish was carefully examined ashore for dot-, nodule-, or cyst-like parasites. Each fish was longitudinally filleted into muscle sections of about 4 mm, and each fillet was examined over a candling table with a dissecting microscope.

Genomic DNA isolation, PCR amplification, cloning, and sequencing
The eggs, preserved in 70% ethanol, were lysed in 500 µl Tris–HCl 20 mM pH 8.0, EDTA 5 mM, NaCl 400 mM, and SDS 1% with proteinase K (20 mg ml–1) overnight at 37°C. Subsequently, the DNA was purified with one phenol:chloroform:isoamyl alcohol extraction, followed by one chloroform:isoamyl alcohol extraction. DNA was precipitated with ethanol and sodium acetate 3 M pH 5.2 overnight at –20°C. The precipitated pellet was dried and resuspended in 50 µl of Tris–EDTA (TE) buffer (Sambrook and Russell, 2001).

Two conserved primers, U1 forward (5' AAC CTG GTT GAT CCT GCC AGT 3') and U2 reverse (5' TGA TCC TTC TGC AGG TTC ACC TAC 3'), derived from known conserved regions of the 18S rRNA gene, were used in PCR to amplify the small subunit ribosomal DNA gene (Medlin et al., 1988; Figueras et al., 2000). PCR reactions were performed in a total volume of 25 µl containing PCR buffer at 1x concentration, MgCl2 at 1.25 mM, nucleotides at 0.4 mM, primers at 0.1 µM, Taq DNA polymerase at 0.025 units µl–1, and 10–100 ng of DNA. The cycling protocol was 94°C for 3 min, 35 cycles of 94°C for 1 min, 55°C for 1 min, and 72°C for 2 min, then 72°C for 10 min. PCR products were separated on a 1% agarose (in 1x Tris-acetic EDTA buffer) gel stained with ethidium bromide and analysed using a UV image analysis system.

Fresh PCR products were ligated into cloning vector pCR2.1 (Invitrogen) at 14°C overnight, then transformed into E. coli One Shot Top 10F' Chemically Competent (Invitrogen). Transformed cells were screened by PCR using the vector's M13 forward (5' GTA AAA CGA CGG CCA G 3') and reverse (5' CAG GAA ACA GCT ATG AC 3') primers. The positive clones with a fragment length of about 2000 bp were directly sequenced. DNA sequencing was performed by the dideoxy-chain termination method using a BigDye terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems) and an automated DNA sequencer ABI PRISMTM 3100 (Applied Biosystems). Each product was sequenced employing the M13F and M13R primers.

All sequences generated were searched for similarity using the basic local alignment search tool (BLAST) at Web servers of the National Centre of Biotechnology Information.

Ultrastructure
Fresh squash preparations of small, infected pieces of muscle were examined under Nomarski differential interference contrast (DIC) optics, and the parasites were photographed. Histological preparations were made from infected muscle fixed in 10% formaldehyde for 24 h, then embedded in paraffin, sectioned at 4 µm, and stained with haematoxylin and eosin, following standard methods (Culling et al., 1985). Semithin sections stained with toluidine blue were also examined by light microscopy. For scanning electron microscopy (SEM), an egg suspension was fixed for 4 h in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.3 at 4°C), then washed for 30 min in the same buffer. The sample was finally dehydrated in an ethanol series, critical-point-dried in CO2 using a Polaron E3000, and sputter-coated in a Polaron SC500, using 60% gold–palladium. The parasites were examined with a Philips XC30 SEM operating at 10–20 kV. For transmission electron microscopy (TEM), small fragments of the infected muscle were fixed in 3.0% glutaraldehyde in 0.2 M sodium cacodylate buffer, pH 7.2, for 12 h at 4°C, washed in the same buffer for 4 h at 4°C, then post-fixed in buffered 2.0% osmium tetroxide for 4 h at the same temperature. After dehydration in a graded ethanol series, the fragments of infected muscle were embedded in Epon, sectioned by diamond knife, double-stained with uranyl acetate and lead citrate, and observed in a JEOL 100CXII TEM operated at 60 kV.

Prevalence and mean intensity of infection were determined according to Bush et al. (1997). A non-parametric test was used to generate statistical confidence in analysing seasonal and host–parasite relationships.


   Results
Top
 Introduction
 Material and methods
 Results
Discussion
 References
 
Parasite
During necropsy, examination by eye of some filleted muscle sections revealed a total of four to eight parallel yellowish threads about 8 mm long by 200–290 µm wide. They conformed to dorsoventrally flattened filamentous worm fragments plus numerous ribbon-like egg masses of an unidentified trematode coiled inside the fish muscle in a zigzag fashion. Loop masses contained large numbers of eggs, with developmental stages ranging from the untanned smaller pre-vitellogenous oocytes in the proximal region to fully embryonated eggs in the distal part of the uterus (Figures 1–5). These larger reniform eggs characteristically showed internal masses of undifferentiated cellular material or germ balls of the miracidium (Figure 5). A large operculum is located 1.8–4 mm from the top of the generally bean-shaped eggs. The eggs measured 25–30 µm long and had a smooth, thick (0.25 µm), scleratin-like shell (Figures 4–6).


Figure 1
View larger version (77K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Figures 1–3 Aggregations of eggs. (1) Pre-vitellogenous oocytes (oc) and embryonated eggs. Note that the residual egg mass retains the shape of the residual adult worm (DIC x310). (2) Isolated egg stages, including stages between pre-vitellogenous oocytes (arrows) and fully embryonated eggs (DIC x625). (3) A miracidium (m) in the process of being released from the egg (e) (DIC x625).

 


Figure 4
View larger version (53K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Figures 4–5 Ultrastructure of eggs. (4) SEM observation of a bean-shaped embryonated egg. Note the distal operculum (arrow) (x5960). (5) Ultrathin section of a developing embryo, showing the thick-sclerotin egg wall (arrow) and internal germ balls (g) of embryo (x8000).

 


Figure 6
View larger version (70K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Figures 6–8 Host–parasite interaction. (6–7) Ultrathin sections showing early stages of encapsulation within the musculature (m), with pockets of cell debris and amorphous material in the capsule wall (asterisks). The capsule is closely applied to the integumentary surface of the didymozoid, which is wrinkled and cobblestone-like with spines (arrow) (Figure 6, x8000; Figure 7, x10 600). (8) Ultrathin section showing an intermediate stage of encapsulation, with numerous flocculent electron-dense material (arrow). A leukocyte (Le) with phagocytic vacuoles is also visible (x16 000).

 
The universal U1 and U2 primers successfully amplified over 1900 bp from every sample examined. The DNA sequence of PCR was registered in GenBank, with accession number DQ087251 [GenBank] . BLAST searches using the obtained sequence confirmed that it belonged to 18S rDNA and bore close similarity to the family Didymozoidae. The highest nucleotide identity values were obtained with Didymozoon scombri (AJ287500 [GenBank] ) and Didymozoid sp. (AY222102 [GenBank] , AY222103 [GenBank] , AY222104 [GenBank] ), at 95% sequence identity.

Host–parasite interaction
Histological and semithin sections revealed egg masses both superficially and in deep connective tissue layers between muscular fibres, through the epoxial and hypaxial musculature in the cranio-dorsal (50%), medio-lateral (25%), and caudal (25%) regions of mackerel 244–335 mm long and 115–394 g in weight. Single egg masses were common in the fish muscle, but multiple infections (up to eight residual aggregations of eggs in a 335-mm, 303-g mackerel) were found in 14.3% of the infected fish. Overall prevalence of infection reached 18.5% (11.6–25.4 CI). No statistically significant seasonal changes in infection values were evident in a year-long fish cohort (U = 362; p = 0.5696), but prevalence was greater in late spring (20%) and summer (24%) than in autumn (14%) or winter (16%). No significant relationships were found between prevalence of infection and fish body length (U = 435; p = 0.7265) or weight (U = 362.5; p = 0.0625).

The parasites always followed the orientation of the muscle fibres and did not traverse the myosepta. Focal damage to the constricted muscle fibres surrounding the parasite could be seen in some histological and semithin sections by increasing the mechanical pressure, which locally distorted the fish muscular architecture. Nevertheless, myofibres remote from the parasite always retained their structural integrity.

Ultrastructure examination of host–parasite interaction revealed early and intermediate stages of parasite encapsulation. An early host reaction consisted of a 1.13–1.32-µm-wide capsule with pockets of amorphous material and cell debris, which in intermediate stages of encapsulation resulted in an inner, closely applied capsule of infiltrated leukocytes surrounding the egg masses (Figures 6 and 7). Numerous membranes of variable size were seen in the integument of the adult didymozoid, in direct contact with the inner capsule. The parasite integument was intact, and the sub-integumental cells were normal. Accumulation of flocculent electron-dense material not normally present on the integument and its surface was observed near the basement membrane and the surface of the integument (Figure 8). Late stages of encapsulation were associated with residual aggregations of embryonated eggs, consisting of numerous fibroblasts and large amounts of collagen (Figures 9 and 10). In this case, the residual integument covering the egg masses was degraded and not easily identifiable.


Figure 9
View larger version (68K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Figures 9–10 Histopathology. (9) Longitudinal histological section showing a focal parting of skeletal muscular fibres to accommodate a residual aggregation of eggs. Note that the parasite follows the orientation of the muscle fibres (x100). (10) Higher magnification than in Figure 9, showing marked fibrosis (arrow) (x1000).

 

   Discussion
Top
 Introduction
 Material and methods
 Results
 Discussion
References
 
The molecular analyses presented here support the view that the eggs found in the musculature of Atlantic mackerel belong to a yet unidentified didymozoid. According to Gibson et al. (1981), live adult didymozoids of Halvorsenius exilis commonly found in eastern Atlantic mackerel occur only in young fish, whereas egg aggregations (which are retained in utero until the death of the worm) are found in older fish (3-group) from the western English Channel. Eggs seem to remain in the host tissue and are released into the water only when the fish is eaten and digested, or dies and decomposes. Death of the fish is therefore apparently necessary to make larvae available to the first intermediate host. Our finding is in full agreement with this thesis, because we only found worm fragments and residual aggregations of eggs in the musculature of post-recruit market-size mackerel. Based on cohort infection analysis, Gibson et al. (1981) suggested that most infections in 2-group and older mackerel could be identified by the occurrence of eggs rather than live worms. We also agree with this statement, because in our material, 80% of the residual egg-containing mackerel were ≥30 cm long (i.e. age ≥2-group; FAO-FIGIS –www.fao.org). Moreover, as the parasite recruits to the reproductive stock all year round at about 20% prevalence, and no significant relationship was found between parasite prevalence and fish length or weight, it is likely that residual egg aggregations really mark the earlier position of dead worms. If so, then it is also likely that metacercariae and adult stages would be common in fish from the pre-recruiting young stock, whereas residual egg aggregates would be predominant in mature fish from the target reproductive stock. Overall, unfortunately, no sequences of PCR products of H. exilis are registered in GenBank.

Although trematodes are the most prevalent and conspicuous helminth parasites that infect mackerel (Arandas-Rego and Santos, 1983; Arandas-Rego et al., 1985), neither histopathological nor host tissue reactions have been studied. Tissue infected by didymozoids usually reveals little or even no tissue reaction (Lester, 1980; Karlsbakk, 2001). Gibson et al. (1981) documented no sign of host reaction associated with live H. exilis in very close contact with fish tissues. Although our findings were sometimes similar, in most cases we did see a host cellular encapsulation response. It is likely that primarily non-specific mackerel responses react against invading metacercariae but that these responses do not reach the adults. Our ultrastructure data have proved that exposed integumentary surfaces that are common in early encapsulated adult worms were always undamaged. It is likely that an evasion mechanism of host defence may be associated with the release of lytic or toxic substances to host cells enclosed in the numerous electron-dense structures secreted by the parasite. This ultrastructural observation corresponds with early and intermediate phases of encapsulation. Once the adult parasite leaves the fish muscle tissue or dies, late stages of encapsulation are common, in association with worm fragments and residual eggs. Such a strategy developed to avoid destruction by the host response has been suggested for other helminths (Ubelaker et al., 1970; Bogitsh and Wikel, 1974), and for trematode infections of long duration in fish hosts (Mathews and Mathews, 1993; Buchmann et al., 2001). Clearly, more research on cytochemical identification and biochemical characterization of such toxic secretions from our material is required.

Numerous egg masses of a muscle-infecting didymozoid are certainly consumed by man, but this does not seem to pose a public health problem. However, cases of concurrent heavy infection in the edible part can reduce the commercial value of mackerel, mostly because of the unappetizing "yellowish presence" of numerous residual ribbon-like eggs along with quinone-staining shell components that are visible to the naked eye.


   Acknowledgements
 
This work was supported by Marie Curie Fellowship contract No. QLK5-CT2000-52101 under the Fifth Framework Programme of the European Community. S. D'Amelio and an anonymous reviewer are thanked for their constructive comments on the draft submission.


   References
Top
 Introduction
 Material and methods
 Results
 Discussion
 References
 

    Abollo E., Gestal C., Pascual S. (2001) Anisakis infestation in marine fish and cephalopods from Galician waters: an updated perspective. Parasitology Research 87:492–499.[CrossRef][Web of Science][Medline]

    Alvarez-Pellitero P. and Sitja-Bobadilla A. (1993) Pathology of Myxosporea in marine fish culture. Diseases of Aquatic Organisms 17:229–238.[Web of Science]

    Arandas-Rego A., Carvalho-Varela M., Mendonça M.M., Afonso-Roque M.M. (1985) Helmintofauna da Sarda (Scomber scombrus L.), peixe da costa continental Portuguesa. Memorias do Instituto Oswaldo Cruz 80:97–100.

    Arandas-Rego A. and Santos C.P. (1983) Helmintofauna de Cavalas, Scomber japonicus Houtt, do Rio de Janeiro. Memorias do Instituto Oswaldo Cruz 78:443–448.

    Bogitsh B.J. and Wikel S.K. (1974) Schistosoma mansoni: ultrastructural observations on the small intestine of the marine host. Experimental Parasitology 35:68–79.[CrossRef][Web of Science][Medline]

    Buchmann K., Lindenstrøm T., Bresciani J. (2001) Defence mechanisms against parasites in fish and the prospect for vaccines. Acta Parasitologica 46:271–81.[Web of Science]

    Bush A.O., Lafferty K.D., Lotz J.M., Shostak A.W. (1997) Parasitology meets ecology on its own terms: Margolis et al, revisited. Journal of Parasitology 83:575–583.[CrossRef][Medline]

    Culling C.F.A., Allison R.T., Barr W.T. (1985) Cellular Pathology Techniques(Butterworth & Co., London).

    Figueras A., Lorenzo G., Ordás M.C., Gouy M., Novoa B. (2000) Sequence of the small ribosomal RNA gene of Perkinsus atlanticus-like isolated from carpet shell clam in Galicia, Spain. Marine Biotechnology 2:419–428.[Medline]

    Gibson D.I., MacKenzie K., Cottle J. (1981) Halvorsenius exilis gen. et. sp. nov, a new didymozoid trematode from the mackerel Scomber scombrus L. Journal of Natural History 15:917–929.[CrossRef][Web of Science]

    Karlsbakk E. (2001) Didymozoid larvae (Trematoda) in the body musculature of cultured Atlantic herring larvae (Clupea harengus). Acta Parasitologica 46:3164–170.[Web of Science]

    Koie M. and Lester R.J.G. (1985) Larval didymozoids (Trematoda) in fishes from Moreton Bay, Australia. Proceedings of the Helminthological Society of Washington 52:196–203.[Web of Science]

    Korotaeva V.D. (1985) Trematodes of commercial fish of the Pacific of practical importance. NOAA Technical Report, NMFS 25:63–64.

    Lester R.J.G. (1980) Host–parasite relationships in some didymozoid trematodes. Journal of Parasitology 66:527–531.[CrossRef]

    Lom J. and Dyková I. (1992) Protozoan Parasites of Fish(Elsevier, Amsterdam) 315 pp.

    Malouf A. H. (1986) Report of the Royal Commission on seals and sealing in Canada, Ottawa vol. 3: 679 pp.

    Mathews R.A. and Mathews B.F. (1993) Cryptocotyle lingua in mullet, Chelon labrosus; significance of metacercarial excretory proteins in the stimulation of the immune response. Journal of Helminthology 67:1–9.[Web of Science][Medline]

    Medlin L., Elwood H.L., Stickel S., Sogin M.L. (1988) The characterization of enzymatically amplified eukaryotic 16S-like rRNA-coding regions. Gene 71:491–499.[CrossRef][Web of Science][Medline]

    Sambrook J. and Russell D.W. (2001) Molecular Cloning. A Laboratory Manual(Cold Spring Harbor Laboratory Press, New York).

    Talbot C. (1994) Soft flesh. Food for Thought 2:316.

    Thompson R.C.A. (2000) Emerging parasite zoonoses. International Journal for Parasitology 30:12–13.

    Ubelaker J.E., Cooper N.B., Allison V.F. (1970) Possible defensive mechanisms of Hymenolepis diminuta cysticercoids to hemocytes of the beetle, Tribolium confusum. Journal of Invertebrate Pathology 16:310–312.[CrossRef][Web of Science][Medline]


Add to CiteULike CiteULike   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us    What's this?



This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrowRequest Permissions
Google Scholar
Right arrow Articles by Pascual, S.
Right arrow Articles by Azevedo, C.
Right arrow Search for Related Content
PubMed
Right arrow Articles by Pascual, S.
Right arrow Articles by Azevedo, C.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?