INTRODUCTION
The phylum Microsporidia Balbiani, 1882 is represented by at least 144 available genera. It is characterized by unicellular eukaryotic microorganisms living as obligate intracellular parasites, commonly infecting fishes, insects, crustaceans, and other invertebrate and vertebrate groups from different geographical areas (Lom and Dyková, Reference Lom, Dyková, Lom and Dyková1992; Sprague et al. Reference Sprague, Becnel and Hazard1992; Larsson, Reference Larsson1999; Lom, Reference Lom2002). In a recent paper, Lom and Nilsen (Reference Lom and Nilsen2003) described the following 15 microsporidian genera as infecting fish: Glugea Thélohan, 1891; Pleistophora Gurley, 1893; Ichthyosporidium Caullery and Mesnil, 1905; Heterosporis Schubert, 1969; Nosemoides Vinckier, 1975; Spraguea Weissenberg, 1976; Loma Morrison and Sprague, 1981; TetramicraMatthews and Matthews, Reference Matthews and Matthews1980; Microgemma Ralphs and Matthews, 1986; Microfilum Faye, Toguebaye and Bouix, Reference Faye, Toguebaye and Bouix1991; Nucleospora Hedrick, Graff and Baxa, 1991; Neonosemoides Faye, Toguebaye and Bouix, 1996; KabatanaLom, Dyková and Tonguthai, Reference Lom, Dyková and Tonguthai1999; Pseudoloma Matthews, Brown, Larison, Bishop-Stewart, Rogers and Kent, 2001; Ovipleistophora Pekkarinen, Lom and Nilsen, 2002. Recently 2 new genera were identified as infecting fish: Amazonspora in the gills of an Amazonian fish (Azevedo and Matos, Reference Azevedo and Matos2003) and Myosporidium in muscle of commercial hake (Merluccius sp.) from fisheries near Namibia (Baquero et al. Reference Baquero, Rubio, Moura, Pieniazek and Jordana2005).
There is very little knowledge about microsporidiosis in the ichthyological fauna of South America. Two Microsporidia were found in Amazonian fishes Loma myrophis (Azevedo and Matos, Reference Azevedo and Matos2002; Matos et al. Reference Matos, Corral and Azevedo2003) and Microsporidium brevirostris (Matos and Azevedo, Reference Matos and Azevedo2004) in Myrophis platyrhynchus and Brachyhypopomus brevirostris host species respectively.
In this paper, we described a new genus and new species of a microsporidian through morphological and ultrastructural observations, with special reference to the ultrastructural aspects of the xenoma wall and the spore differentiation. Phylogenetic relationships comparing the Potaspora morhaphis SSU rRNA gene with that of other fish infecting microsporidian species was also done. The morphological characteristics and taxonomic position are discussed.
MATERIALS AND METHODS
Fish, location of infection and prevalence
Thirty specimens of freshwater teleost fish Potamorhaphis guianensis Schomburgk, 1843 (Teleostei, Belonidae) (Brazilian common name ‘Peixe-Agulha’), were collected from the estuarine region of the Amazon River (01°11′ S/47°18′ W) near the city of Belém (Pará State), Brazil. The specimens were anaesthetized by MS 222 (Merck) and later measured (19–25 cm in length). Infection was determined by the presence of several xenomas located in the coelomatic cavity near the anal region. The prevalence of infection was 40% (12 fishes in 30 examined), in both sexes.
Light (LM) and transmission electron microscopy (TEM)
For LM smears of xenoma and free spores were observed directly without any fixation or stain by a light microscope equipped with Nomarski interference-contrast (DIC) optics.
For ultrastructural studies, the xenomas were excised and fixed in 3% glutaraldehyde in 0·2 m sodium cacodylate buffer (pH 7·2) at 4°C for 24 h. After washing overnight in the same buffer at 4°C and post-fixation in 2% osmium tetroxide in the same buffer and temperature for 3 h, the fragments were dehydrated through a graded ethanol ascending series, followed by propylene oxide (3 changes of 2 h each) and embedded in Epon (12 h in each change). Semi-thin sections were stained with methylene blue-Azur II and observed by DIC optics. Ultrathin sections were contrasted with aqueous uranyl acetate and lead citrate and observed with a JEOL 100CXII TEM, operated at 60 kV.
DNA isolation and PCR amplification
Several cysts were dissected from fishes, following homogenization to isolate the spores, and were then stored in 80% ethanol at 4°C. The genomic DNA of about 5×106 spores was extracted using a GenElute™ Mammalian Genomic DNA Miniprep Kit (Sigma) following the manufacturer's instructions for animal tissue, except for the incubation time. The DNA was stored in 50 μl of TE buffer at – 20°C until used. The DNA concentration was estimated with the Qubit™ Fluorometer (Invitrogen). The majority of the region coding for the small subunit (SSU) rRNA gene was amplified by PCR using the primers V1f (5′CACCAGGTTGATTCTGCC3′) and 1492r (5′GGTTACCTTGTTACGACTT3′) (Vossbrinck et al. Reference Vossbrinck, Baker, Didier, Debrunner-Vossbrinck and Shadduck1993; Nilsen, Reference Nilsen2000). To amplify the 3′-end of the SSU, internal transcribed spacer (ITS) and 5′-end of the large subunit (LSU) rRNA gene, HG4F (5′GCGGCTTAATTTGACTCAAC) and HG4R (5′TCTCCTTGGTCCGTGTTTCAA) primers were used (Gatehouse and Malone, Reference Gatehouse and Malone1998). To obtain the 5′-end of the SSU gene region a primer was designed (454r – 5′AATTAAGCCGCACACTCCAC). PCR was carried out in 50 μl reactions using 10 pmol of each primer, 10 nmol of each dNTP, 2 mm of MgCl2, 5 μl of 10X Taq polymerase buffer, 1·25 units Taq DNA polymerase (Invitrogen products), and 3 μl of the genomic DNA. The reactions were run on Hybaid PxE Thermocycler (Thermo Electron Corporation, Milford, MA). The amplification program consisted of 94°C denaturation for 5 min, followed by 35 cycles of 94°C for 1 min, 50°C for 1 min and 72°C for 2 min. A final elongation step was performed at 72°C for 10 min. Five μl aliquots of PCR products were visualized with ethidium bromide staining after running on a 1% agarose gel.
DNA sequencing
PCR products for the SSU gene and ITS region have approximate sizes of 1400 bp and 1100 bp respectively. They were cleaned using the MinElute PCR purification kit (QIAGEN) and then 3 purified PCR products were sequenced in both directions. Sequencing was done using BigDye Terminator v1.1 of Applied Biosytems Kit and the sequence reactions were run on an ABI3700 DNA analyser Perkin-Elmer, Applied Biosystems, Stabvida, Co., Oeiras, Portugal).
Distance and phylogenetic analysis
To evaluate the relationship of Potaspora morhaphis to other Microsporidia, we have used the 42 rDNA sequences, listed with their hosts in Table 1, obtained from GenBank data. The corresponding sequences and GenBank/NCBI Accession number of Endoreticulatus schubergi (L39109), Enterocytozoon bieneusi (L07123), Vairimorpha necatrix (Y00266) and Vittaforma corneae (L39112) were used as the outgroup. Sequences were aligned as described by Azevedo et al. (Reference Azevedo, Balseiro, Casal, Gestal, Aranguren, Stokes, Carnegie, Novoa, Burreson and Figueras2006). Alignment using Clustal W (Thompson et al. Reference Thompson, Higgins and Gilson1994), in MEGA 4 software (Tamura et al. Reference Tamura, Dudley, Nei and Kumar2007), with an opening gap penalty of 10 and a gap extension penalty of 4 was done for both pairwise and multiple alignments. Subsequent phylogenetic and molecular evolutionary analyses were conducted using MEGA 4, with the 42 rDNA sequences for microsporidian species and the outgroup species selected. Distance estimation was carried out using the Kimura-2 parameters model distance matrix for transitions and transversions. For the phylogentic tree reconstructions, maximum parsimony analysis was conducted using the close neighbour interchange (CNI) heuristic option with a search factor of 2 and random initial trees addition of 2000 replicates. Bootstrap values were calculated over 100 replicates.
Table 1. Hosts and GenBank Accession numbers for the SSU rRNA sequences of 42 microsporidian that parasite fishes species used in the phylogenetic analyses
RESULTS
Macroscopical and light microscopical observations
Some spherical to elipsoidal whitish cysts (xenomas) were macroscopically observed adherent to the internal wall of the coelomatic cavity of the teleost fish near the anal region. These xenomas with a variable number (up to 7) could reach dimensions of up to ~0·8 mm (Fig. 1A). In semi-thin section, the thick xenoma wall showed a lucent area surrounded by a layer of cells and inside was filled with numerous spores and other developmental stages (Fig. 1B). After rupture of the xenoma wall, the free spores were easily identified as belonging to the phylum Microsporidia (Fig. 4).
Fig. 1. (A–E) Light and transmission electron micrographs of the microsporidian Potaspora morhaphis n. gen., n. sp. (A) Some xenomas (arrowheads) on the abdominal cavity. (B) Semi-thin section of the xenoma periphery, showing the xenoma wall (Wa) and the matrix of the xenoma containing developmental stages including spores (*). The boxed area is enlarged in the figure C. (C) Ultrathin section of the xenoma wall (Wa) showing numerous filiform and anastomosed microvilli-like structures (Mv) projected toward the periphery and, externally, an erythrocyte nucleus (E) in contact with the wall. (D) Ultrathin section of the internal periphery of the xenoma, showing several host cell nuclei (Nu) and a dividing meront (Me), showing some nuclei (*), in direct contact with the host-cell cytoplasm. (E) Ultrathin section of a sporogonial plasmodium in division (Sr) showing the wall formation by a gradual deposition of the dense material on the membrane (arrows).
Ultrastructural observations
Xenoma
The xenoma wall was formed by numerous filiform and anastomosed microvilli-like structures, with a regular diameter, projected from the surface toward the periphery. The microvilli were intermingled by an amorphous, finely granular material. In favourable sections the microvilli were 4–5 μm long (Fig. 1C). Some zones towards the apical region of the microvilli were in contact with an external layer of erythrocytes (Fig. 1B, C). The asynchronous development was characterized by several merogonic and early sporogonic stages predominantly along the xenoma periphery, while immature and mature spores were more internally localized in the centre of the xenoma (Fig. 1B). Internally, the matrix of the xenoma possessed numerous host cells, their nuclei showing a prominent nucleolus, and great mass of peripheral heterochromatin (Figs 1D and 2D). Inside each host cell, the parasite was always in direct contact with the cytoplasm of host cells, without any surrounding membrane and frequently in the same stage of the developmental life cycle (Fig. 3B).
Description of the development stages
Meronts
These cells grow into multinucleate plasmodia. They appeared in ultrathin sections as round to elliptical uninucleated or binucleated cells always with the nuclei unpaired. In these cells, the chromatin was homogeneous in contrast to the nuclei of the host cells in which the chromatin was organized in dense masses. Their cytoplasm possessed numerous free ribosomes and was uniformly granular and poorly endowed with cytoplasmatic organelles (Fig. 1D). Meronts divided by binary fission and transformed into sporonts (Figs 1E and 2A).
Fig. 2. (A–D) Late sporogonic development of the microsporidian Potaspora morhaphis n. gen., n. sp. (A) Sporogonial plasmodium in division giving rise to 4 sporoblasts. (B) Some sporoblasts (Sb) in different developmental stages showing a dense globule (*) that gradually decreases in density and the polar filament in differentiation (arrowheads). (C) Detail of an immature spore showing the dense globule (*) strongly associated to the polar filament formation (PF). Nucleus (Nu). (D) Ultrastructure of a host cell showing the nucleus (Nu) and the nucleolus (*) with peripheral nucleolar heterochromatin (arrow) surrounding the nucleolus. The mature spores (S) are contained in the cytoplasm of the host cell.
Sporonts
The transition from merogony to sporogony is characterized by the acquisition of a thick and dense cell coat located on the outer surface of the plasmalemma (Figs 1E and 2A). Early on, the discontinuous coat of the sporogony stages appeared to be formed by isolated patches (Fig. 1E). They were rounded and uninucleated cells and in their cytoplasm several well-developed cisternae of rough endoplasmatic reticulum and small vesicles were observed. Before the sporont transformed in uninucleate sporoblasts they divided again by multiple fission giving rise to 4 sporoblasts (Figs 1E and 2A).
Sporoblasts
The sporoblasts do not have the capacity to divide further and gradually differentiate the organelles typical of the spores, composed of an anchorage disc, polaroplast, polar filament and posterior vacuole. In the sporoplasm a very electron-dense irregularly-shaped globule that persists until sporogenesis is concluded, was frequently observed (Fig. 2B). This structure is associated with the reticular body present in the sporoplasm during differentiation of the spores and later appears to be immersed into a posterior vacuole (Fig. 2C).
Systematic position
Phylum Microsporidia Balbiani, 1882; Class Haplophasea Sprague, Becnel and Hazard, 1992; Family Tetramicridae Matthews and Matthews, Reference Matthews and Matthews1980.
Description of the genus
Name: Potaspora n. gen.
Diagnosis: Xenoma formation has several nuclei and the plasmalemma differentiates numerous filiform and anastomosed microvilli-like structures projected externally. In all developmental stages the nuclei are monokaryotic and develop in direct contact with host cell cytoplasm. The merogony stages are binucleated and divide by binary fission. Each meront differentiates into a sporont by a gradual development of a thick electron-dense coat. The sporont divides by multiple fission into 4 sporoblasts. In this stage a very electron-dense irregular-shaped body differentiates. Monomorphic spores containing polaroplast with 2 distinct kinds of lamellae.
Description of the species
Name: Potaspora morhaphis n. gen., n. sp.
Type host: Potamorhaphis guianensis Schomburgk, 1843 (Teleostei, Belonidae).
Type Locality: Estuarine region of the Amazon river (01°11′S and 47°18′W) near the city of Belém (Pará State), Brazil.
Location in the host: Xenoma in the coelomatic cavity near the anal region.
Prevalence of infection: Twelve of 30 (40%).
Type specimens: One slide containing mature free spores and another with semi-thin sections of tissues containing spores and different developmental stages of hapantotype were deposited in the International Protozoan Type Slide Collection at Smithsonian Institution Washington, DC. 20560, USA with acquisition number (USNM 1113817). The histological semi-thin sections containing different developmental stages were deposited at the laboratory of the senior author.
Etymology: The genus name is the prefix from the name of the host genus and the specific name is derived from the suffix of the host genus name.
Description of the spores: Pyriform spores measuring 2·8±0·3×1·5±0·2 μm and containing all the typical characteristic structures of the Microsporidia (Figs 3A, C and 4). The spore wall was about 125 (114–131) nm thick (n=30), except for the anterior end where the anchoring disc contacted with the wall, which was ~50–70 nm thick (Fig. 3C, D). The spore wall consisted of an electron-lucent endospore and an electron-dense exospore each with just about the same thickness (Fig. 3C, D, E, F). The exospore was externally surrounded by a thin irregular layer of granular material (Fig. 3D).
Fig. 3. (A–F) Morphological and ultrastructural details of the microsporidian Potaspora morhaphis n. gen., n. sp. (A) Several isolated mature spores observed by DIC microscopy. (B) Some spores (S) in different stages of development in close contact with the cytoplasm of the host cell that shows the (Nu). (C) Ultrathin longitudinal and two transverse sections of a spore showing the typical microsporidian structures and organelles. Wa, wall; AD, anchoring disc; Pp, polaroplast; PF, polar filament; Va, vacuole. (D) Ultrastructural detail of the apical region of a spore showing anchoring disc (AD) in close contact with the wall (Wa) and the lamellar region of the polaroplast (Pp) containing dense material (arrowheads). (E) Ultrastructural detail of a transverse section of a spore showing the lamellar region of the polaroplast (Pp) containing dense material (arrowheads), the polar filament (PF) and the wall (Wa). (F) Ultrastructural detail of the wall (Wa), the polar filament coils (PF) showing the external membrane (arrowheads), as well as a central dense mass (arrows).
Fig. 4. Semi-schematic drawing of a spore of Potaspora morhaphis n. g., n. sp. showing specific characters, such as spore shape and dimensions, spore wall (Wa), polaroplast (Pp), anchoring disc (AD), polar filament (PF) coils (PF*), nucleus (Nu) and vacuole (Va).
The anchoring disc is located in the apical region of the spore in an eccentric position in relation to the spore axis, giving the spore bilateral asymmetry (Fig. 3C, D). The anterior part of the polar filament (FP) (manubrium) measured about 145 (140–149) nm (n=25) and the angle of tilt anterior PF to the spore axis was ~45° (Fig. 3C, D). The PF was isofilar arranged into 9–10 (rarely 11) coils in 2 layers and, when sectioned transversally, the PF exhibited concentric layers (Fig. 3F). The polaroplast (Pp) has 2 distinct lamellae folded around the PF. In the anterior zone the lamellae were without a lumen and were irregularly packed with a lucent space between them, while in the posterior lamellae the lumen was filled with electron-dense material approximately 35–40 nm thick (Fig. 3D, E). The nucleus, containing a moderately uniform nucleoplasm and surrounded by numerous ribosomes, was situated laterally between the polaroplast and the posterior vacuole. The posterior vacuole, situated at the basal part of the spore between the PF coils, was irregular and contained some masses of dense material (Fig. 3C).
Molecular analysis
Two bands of approximately 1·4 kb and 1·1 kb were obtained after amplification of the microsporidian genomic DNA. The primers used were V1f-1492r and HG4F-HG4R, respectively. The sequences were assembled and the resulting consensus DNA sequence of the complete SSU rRNA, ITS, and the 5'-end of the LSU rRNA gene was 1826 bp in length. This sequence with a GC content of 47% was deposited in GenBank (Accession number EU534408). In total, 42 SSU rDNA sequences, including those with the highest BLAST scores, were aligned with the Potaspora morhaphis SSU rDNA sequence. Only sequences belonging to species parasitizing fishes were included in the final analyses (Table 1). Trachipleistophora hominis found in muscle of humans, some Pleistophora spp. found in crustacean species and several Dictyocoela spp. parasitizing amphipods were excluded. The length of the aligned sequences used for phylogenetic analysis was 1527 bases after trimming the 3′end. Before phylogenetic analysis, only those sites which could be unambiguously aligned among all Microsporidia and outgroups were used, resulting in an alignment of 1321 bases long.
Based on pairwise comparisons among the SSU rDNA sequences, the maximal similarity was observed with Microgemma tincae, Microgemma caulleryi and Tetramicra brevifilum species, 87·3%, 87·2% and 87·2%, respectively (Table 2). Phylogenetic analyses using maximum parsimony placed Potaspora morhaphis clustered with the sequences of the Kabatana (AF356222, AJ295322, EF202572), Microgemma (AJ252952, AY651319, AY033054), Spraguea (AF104086, AF033197, AY465876, AF056014), Tetramicra (AF364303) genera and Microsporidium (AJ295323, AJ295324) collective group. This clade has 72% bootstrap support. Only Spraguea (68% bootstrap) clade suggested monophyly (Fig. 5). Neighbour-joining and maximum likehood analyses resulted in identical tree topology.
Fig. 5. Parsimony tree of SSU rDNA sequences to compare Potaspora morhaphis with selected sequences from other fish-infecting Microsporidia. The analysis was conducted using 1321 aligned nucleotide positions of the highest BLAST score microsporidian sequences and 4 more microsporidian sequences as outgroup. The bar indicates the equivalence between the distance and the number of changes. The numbers on the branches indicate bootstrap support from 100 replicates. Potaspora morhaphis is placed within group IV (Lom and Nilsen, Reference Lom and Nilsen2003) (highlighted box), which includes the sequences of the genera Kabatana, Microgemma, Spraguea, Tetramicra, and Microsporidium.
Table 2. Comparison of some SSU rDNA sequences: percentage of identity (top diagonal) and pairwise distance (bottom diagonal) obtained by Kimura-2 parameter analysis
DISCUSSION
Ultrastructural studies
The ultrastructural organization of the xenoma, as well as aspects of the developmental stages described in the present study, showed that all structures were typically from Phylum Microsporidia, Class Haplophasea and family Tetramicridae (Lom and Dyková, Reference Lom, Dyková, Lom and Dyková1992; Larsson, Reference Larsson1999; Lom and Nilsen, Reference Lom and Nilsen2003).
Of at least 156 fish microsporidian species distributed among 17 genera (Azevedo and Matos, Reference Azevedo and Matos2003; Lom and Nilsen, Reference Lom and Nilsen2003; Baquero et al. Reference Baquero, Rubio, Moura, Pieniazek and Jordana2005), only 12 develop xenoma. These formations are a characteristic consequence of the host cell defence to the parasite development having features specific to the genus and species (Lom and Nilsen, Reference Lom and Nilsen2003). Among these, the xenoma wall, of only 4 genera (Ichthyosporidium, Tetramicra, Microfilum and Amazonspora), possesses a structure characterized by numerous anastomosed microvilli-like structures, which could partially resemble the xenoma wall of the parasite reported by us. However, some ultrastructural aspects of the developmental stages of those genera are very distinct. In Ichthyosporidium, the xenoma wall presents microvilli-like ramified projections irregularly intermingled in the wall, but this parasite has the nuclei organized as a diplokaryon during all sporogonic stages and the polar filament (up to 46 coils) is the largest of the microsporidian group (Sprague and Vernick, Reference Sprague and Vernick1974; Casal and Azevedo, Reference Casal and Azevedo1995). In Microfilum, the xenoma wall was described as a very dense region covered by numerous apparently disorganized and ramified microvilli. This parasite gives rise to a spore characterized by a manubrium inserted on a laterally offset anchoring disc and extruding into a short non-coiled polar filament, which was very different from those of the present study (Faye et al. Reference Faye, Toguebaye and Bouix1991). A xenoma wall including a microvillous surface layer formed by anastomosed elongated cytoplasmic processes have been described in the genus Tetramicra (Matthews and Matthews, Reference Matthews and Matthews1980). Meronts located within a vacuole in the host cytoplasm and spores with conspicuous posterosomes surrounded by a membrane and located inside the posterior vacuole.
Recently in an Amazonian fish, a new genus and species (Amazonspora hassar) having a xenoma, strongly encapsulated, consisting of numerous anastomosed microvilli-like projections penetrating the 1–3 first layers of collagen fibres was described. Up to approximately 22 juxtaposed crossed layers of collagen fibres were observed (Azevedo and Matos, Reference Azevedo and Matos2003).
The presence of dense globules of unknown nature in the sporoplasm was also seen in other fish-infecting Microsporidia. Several electron-dense inclusion bodies, sometimes very large, measuring up to 1·38 μm in diameter, were described in sporoblasts and spores of Tetramicra brevifilum species (Matthews and Matthews, Reference Matthews and Matthews1980). In Kabatana arthuri (Lom et al. Reference Lom, Dyková and Tonguthai1999) and K. takedai (Lom et al. Reference Lom, Nilsen and Urawa2001) a very similar globule was reported, while in Loma acerinae (Lom and Pekkarinen, Reference Lom and Pekkarinen1999) 1–3 homogeneous dense globules occupying all the space of the posterior vacuole were observed. In our observations a large inclusion consisting of reticular material like that reported in Ichthyosporidium giganteum, was also found (Sprague and Vernick, Reference Sprague and Vernick1974; Casal and Azevedo, Reference Casal and Azevedo1995).
The polaroplast of Potaspora morhaphis has a bipartite structure comprising the anterior region having folds with a lamellar organization and the posterior region with larger lamellae (cisternae) with dense contents. A similar organization was reported by Lom et al. (Reference Lom, Dyková and Tonguthai1999) in the species Kabatana arthuri which infects the trunk muscles of fishes from the South-East Asia freshwater fish, Pangasius sutchi, as well as in Kabatana takedai (Lom et al. Reference Lom, Nilsen and Urawa2001). The polaroplast of the microsporidian, Spraguea americana, found in the nervous tissues of the Japanese anglerfish Lophius litulon (Freeman et al. Reference Freeman, Yokoyama and Ogawa2004) has a similar organization.
Phylogenetic relationships
The availability, in the public databases, of sequences from different species belonging to the phylum Microsporidia makes the SSU rRNA gene the most suitable not only for development of diagnostic tools and species identification, but also for molecular characterization of new parasites through phylogenetic studies (Weiss and Vossbrinck, Reference Weiss, Vossbrinck, Wittner and Weiss1999; Lom and Nilsen, Reference Lom and Nilsen2003).
In these studies we can see that there is 72% bootstrap support for a clade composed of microsporidian belonging to the Kabatana (Lom et al. Reference Lom, Dyková and Tonguthai1999, Reference Lom, Nilsen and Urawa2001; McGourty et al. Reference McGourty, Kinzger, Hendrickson, Goldsmith, Casal and Azevedo2007), Microgemma (Cheney et al. Reference Cheney, Lafranchi-Tristem and Canning2000; Leiro et al. Reference Leiro, Siso, Parama, Ubeira and Sanmartin2000; Mansour et al. Reference Mansour, Prensier, Jemaa, Hassine, Méténier, Vivarès and Cornillot2005), Spraguea (Freeman et al. Reference Freeman, Yokoyama and Ogawa2004), Tetramicra (Leiro et al. Reference Leiro, Siso, Parama, Ubeira and Sanmartin2000) genera, 2 unclassified species of Microsporidium group (Bell et al. Reference Bell, Aoki and Yokoyama2001) and the microsporidian reported in this study. This result is in concordance with cladograms previously obtained by Lom and Nilsen (Reference Lom and Nilsen2003) and designated as group IV. Our SSU rDNA sequence analysis also shows that Potaspora n. gen. does not have any sister taxa and the lineage is distantly related to the other species examined. Comparing SSU rRNA gene sequences between P. morhaphis with species Microgemma caulleryi and Tetramicra brevifilum (clade with bootstrap 81%) the genetic distances are 12·8% for both species. The smallest genetic distance was observed with the species Microgemma tincae (12·7%) but the SSU rRNA was not completely sequenced. The bootstrap support for this species and another Microgemma vivaresi is 81%. On the other hand, K. takedai and K. newberryi group in a clade with 83% bootstrap and all Spraguea species are clustered in the clade with 68% bootstrap.
Conclusion
When comparing the xenoma wall of the parasite described here with those fish Microsporidia which form xenoma some structural differences were found, such as the organization of the microvilli-like structures. In addition, the ultrastructural organization of the polaroplast and the presence of a dense globule were the most evident differences found compared with other mature spores of previously described species. Concerning this last aspect, the only exception is the spore of Kabatana genus which presents some similarities. However, they were found to parasitize only the muscle fishes and they do not develop inside of xenomas (Lom et al. Reference Lom, Dyková and Tonguthai1999, Reference Lom, Nilsen and Urawa2001; McGourty et al. Reference McGourty, Kinzger, Hendrickson, Goldsmith, Casal and Azevedo2007). As concerns molecular biology, the most parsimonious cladogram has shown that Potaspora morhaphis is placed in the same group as the Kabatana, Microgemma, Spraguea and Tetramicra genera, does not have any sister taxa and has the lowest percentage identity within the group.
So, our results suggest that this parasite does not fit into any of the known fish microsporidian genera, and for these reasons we propose a new genus Potaspora and a new species, Potaspora morhaphis.
This work was partially supported by the Eng°. A. Almeida Foundation (Porto, Portugal), Ph.D. grant from ‘CESPU’ (G. Casal), ‘CNPq’ and ‘CAPES’ – Brazil. We would like to thank the iconographic work of Mr João Carvalheiro. We would like to thank the anonymous reviewers for their helpful suggestions and comments.
