CC BY
43
Protistology 9 (3/4), 117-126 (2015)
Protistology
Perspectives of microsporidia as human pathogens: clues from invertebrate research (minireview)1
Yuliya Y Sokolova
Institute ofCytology, Russian Academy of Sciences, St. Petersburg, Russia, and Louisiana State University, Baton Rouge LA, USA
Summary
Microsporidia (Phylum Microsporidia Balbiani 1882) are ubiquitous parasites within the Animal Kingdom. The phylum includes 1400 described species belonging to 200 genera. The host range, as well as molecular data, strongly suggest that microsporidia evolved as parasites of invertebrates and, to a lesser extent, fish. Only about 1% of microsporidia species have been found in endothermic vertebrates, birds and mammals. Microsporidiosis in humans has been observed worldwide mainly in patients with HIV infection and now increasingly in other groups such as children, immunosuppressed individuals (e.g. organ transplant recipients), contact lens wearers, travelers, and the elderly. Among AIDS patients, microsporidiosis is listed as the third important opportunistic infection causing gastrointestinal disorders, after Cytomegalovirus and Cryptosporidium. In fact, only four species belonging to two genera can be considered true mammalian parasites: Enterocytozoon bieneusi, Encephalitozoon cuniculi, E. intestinalis, and E. hellem. These represent a serious threat to human populations as zoonotic infections. Findings of other microsporidia (as a rule, parasites of arthropods or close relatives of those) in humans, are accidental. At the same time these records demonstrate the consecutive stages of microsporidia adaptation to parasitism in humans: from transient arthropod-related microsporidia known by sequences in stools of AIDS patients, through accidental surface infections in immunocompromised patients (Endoreticulatus-like Microsporidium sp., Tubulinosema) and development in immune privileged tissues of eyes (Vittaforma), skin, and muscles due to accidental exposure to spores of a "generalist" microsporidium (Trachipleistaphora, Anncaliia), to specialized infections of gut epithelium (Enterocytozoon), and systemic microsporidiosis disseminated by macrophages (Encephalitozoon). The review is addressing the following questions. What is special about the microsporidia that are able to infect warm-blooded animals? How high are the risks of acquisition of new microsporidia
1 This review is based on the materials presented at the invited talk at the satellite symposium sponsored by Organization for Economic Cooperation and Development - Cooperative Research Program (OECD-
CRP) as a part of the 48th Annual Meeting of the Society for Invertebrate Pathology, Vancouver, Canada, August 9-13, 2015 (http://www.sipweb.org/docs/Proceedings. pdf).
© 2015 The Author(s)
Protistology © 2015 Protozoological Society Affiliated with RAS
parasites by humans, given abundance of microsporidia in invertebrates, many of which may traverse food chains leading to humans and other mammals?
Key words: microsporidia, microsporidiosis, molecular phylogeny, opportunistic infections, parasitism, invertebrates
Introduction
Microsporidia (Phylum Microsporidia Balbiani 1882) are ubiquitous parasites within the Animal Kingdom. They have been recorded from nearly every major animal phylum. However, distribution of microsporidia among host taxa is far from even. Of about 1400 described species (200 genera), nearly 70% parasitize invertebrates, predominantly arthropods of the classes Insecta and Crustacea, 10% - fish. Only about 1% of known species have been found in endothermic ("warm-blooded") vertebrates - birds and mammals, including hominids (Becnel and Andreadis, 2014; Kent et al., 2014; Stentiford and Dunn, 2014; Vavra and Lukes, 2013). Microsporidiosis in humans has been observed worldwide mainly in patients with HIV infection and now increasingly in other groups such as children, immunosuppressed individuals (e.g. organ transplant recipients), contact lens wearers, travelers, and the elderly. Among AIDS patients, microsporidiosis is listed as the third important opportunistic infection causing gastrointestinal disorders, after Cytomegalovirus and Cryptosporidium (Sokolova et al., 2011).
The host range strongly suggests that the Phylum Microsporidia evolved as parasites of invertebrates and, to a lesser extent, fish. However, a few species managed to establish themselves in the cells of birds and mammals in spite of the temperature barrier and advanced immune defenses. In Homo sapiens, the best studied representative of warm-blooded animals, as many as 14 species belonging to 8 genera have been reported (Table 1)2. What is special about these species? How high are the risks of acquisition of new microsporidia parasites by humans, given abundance of microsporidia in invertebrates, many of which may traverse food chains leading to humans and other mammals?
2 Though the Table lists in fact 16 species, the records on Nosema ocularum and Microsporidium africanum do not contain molecular data or electron microscopy, and thus do not allow proper identification.
Origin of Microsporidia
The evolutionary origin of microsporidia has been significantly elucidated during the last 2-3 years. The consensus tree, based on phylogenies inferred from several genes with high statistical support places Microsporidia within the Aphelidea-Rozellamycota-Microsporidia (ARM) clade, a basal Fungi or sister-to-Fungi lineage (Karpov et al., 2013, Letcher et al., 2013). Aphelids are parasites of algae, and the Rozellamycota lineage comprises species parasitizing fresh-water chitrids and amoebas, as well as numerous "cryptic" species known only by their sequences. Within the ARM clade, microsporidia cluster with rozellids. Association with rozellids has been proved recently by genomic and proteomic studies on Paramicrosporidium spp. and Mitosporidium daphnia, the "missing links" between rozelids and microsporidia (Corsaro et al., 2014; Haag et al., 2014). The intranuclear parasites of free-living amoebae, Paramicrosporidium spp., bear striking morphological similarity with hyperparasitic metchnikovellids (subphylum Ru-dimicrosporidia), presumably a basal lineage of Microsporidia3 (Corsaro et al., 2014; Sokolova et al., 2013; Sokolova et al., 2014). Hence, the common ancestor of Paramicrosporidium and Microsporidia may have been an intranuclear parasite of a protist. This inference is supported by (i) obligatory intranuclear development in three genera (Nucleospora, Desmozoon (Paranucleospora) and Enterospora)4, (ii) occasional development of some species within host nucleoplasm (YS, unpublished observations), and (iii) existence of
3 So far no rDNA sequences for metchnikovellids are available through public database, making open three possible positions of Metchnikovellids: as a basal taxon of Microsporidia, as a close sister group to Microsporidia, and as a sister to Paramicrosporidium.
4 All intranuclear microsporidia are parasites of enterocytes, and ability to develop in the nucleus could be considered a rudimental trait, a "pre-adaptation" employed by nucleus-dwelling microsporidia to avoid degradation by the lysosome system of enterocytes.
Table 1. Microsporidia discovered in humans, their relatives and host groups1.
Genus and Species Tissue tropism Group of animal host Closest relatives and their host groups
Encephalitozoon Mockfordia xanthocaecilliae (79-81%)2, Insects, Psocoptera; E. romaleae (93-96%), Insects, Orthoptera; E. lacertae, E. pogonae (96-98%)3, Reptiles, Lacertilia
E. cuniculi Brain, IGT, disseminated. Mammals, birds, reptiles
E. hellem Disseminated
E. intestinalis IGT, gall-bladder, kidney, eye
Enterocytozoon Paranucleospora theridion (82%), Crustaceans, Copepoda, Fish; En. hepatopenaei (84%), Crustaceans, Decapoda; Nucleospora salmonis (80%), Fish
E. bieneusi IGT, gall-bladder, kidney, eye Mammals
Anncaliia Anncaliia spp. (97-99%), Insects, Diptera, Coleoptera; Crustaceans, Amphipoda4
A. vesicularum Surface, eye, skin, muscle, disseminated (A. connori) Insects (A.a); Primates (A.v., A.c.)
A. algerae
A. connori
Tubulinosema Tubulinosema spp (99%), Insects, Diptera, Lepidoptera, Coleoptera, Hymenoptera, Orthoptera
T. acridophagus Muscle, disseminated Insects, Primates
Trachipleistaphora T.extenrec (98%), Mammals, exp. infection in insects; Vavraia culicis (97%), V. oncoperae (96%), Insects, Diptera, Lepidoptera
T. hominis Eye, sinus, muscle Unknown; Exp. infection in insects
T. anthropopthera Eye, brain, disseminated
Vittaforma Endoreticulatus spp. (89%), Cystosporogenus sp.(98%), Insects, Lepidoptera, Coleoptera, Orthoptera
V. corneae Eye, bladder Unknown; Exp.infection in mammals
Endoreticulatus group Endoreticulatus spp. (83-91%), Insects, Lepidoptera
Microsporidium sp. Muscle Unknown
Pleistophora Pleistophora spp., Fish
P. ronneafiei* Muscle Unknown
Pleistophora sp *. Unknown
Nosema Identification under question No EM or molecular data available
N. ocularum* Eye
Microsporidium
M. africanus* Eye
M. ceilonenesis*
1 Sourses of information: Cali and Takvorian, 2003; Cali et al., 1998; Cali et al., 2005; Cheney et al., 2000; Docker et al., 1997; Franzen et al., 2006a; Franzen et al., 2006b; Koudela et al., 1998; Lange et al., 2009; Nylund et al., 2010; Pilarska et al., 2015; Plischuk et al., 2015; Richter et al., 2013; Sokolova et al., 2007, 2010; Sokolova et al., in press; Suankratay et al., 2012; Tourtip et al., 2009; Vavra et al., 2006; Vavra et al., 2011; Weiss, 2014.
2 Percent of identity ("relatedness") inferred from SSUrDNA-based pairwise distance analysis (in brackets).
3 YS, unpublished data.
4 Tokarev et al., unpublished data.
* No molecular data available.
unusual metabolic relationships of cytoplasmic microsporidia with the host nucleus, i.e. targeting microsporidian hexokinase to the host nucleus during intracellular development (Senderskiy et al., 2014). Paramirosporidium-like hyperparasites of Archigregarina5 infecting gut lumens of the common ancestor of annelids and arthropods, probably gave
rise to some lineages, like metchnikovellids and Chytridiopsis-like microsporidia.
5 Archigregarines that parasitize annelids and occasionally harbor metchnikovellids, is the earliest diverging lineages within Apicomplexa, a "polyphyletic stem" from which all other gregarines evolved (Leander, 2008).
Insects and annelids are the major host groups for both Gregarina (Perkins et al., 2000) and Microsporidia (Becnel and Andreadis, 2014), and a hypothesis that cannot be excluded posits that gregarines might have functioned as a "Trojan horse," enabling dispersal of microsporidia from marine and brackish water annelids to terrestrial arthropods and insects (Sokolova et al., 2013).
Distribution among invertebrates and fish
Estimates based on the analysis of distribution of microsporidia among hosts suggest that Microsporidia ancestors switched to parasitism in oligo-chaetes and polychaetes during their colonization of land, migrating from marine through brackish waters of river estuaries to fresh water basins during the Cambrian and Silurian. The radiation and flowering of Microsporidia likely took place during the Carboniferous and Triassic and was associated with diversification of Arthropods (Issi, 1986). Currently, 70% of microsporidia species parasitize aquatic hosts, mostly crustaceans and insects connected with aquatic habitats (Stentiford and Dunn, 2014). The distribution of microsporidia among groups of terrestrial and freshwater arthropods included numerous host switches via polyxenous life cycles, common parasites, and food chains. The result is the contemporary abundance of species, with evolutionary bonds that have been increasingly elucidated by SSUr-DNA-inferred phylogenies (Vossbrinck and Debrunner-Vossbrinck, 2005; Vossbrinck et al., 2014), though the whole puzzle is far from assembled.
Examples of adaptation of microsporidia to parasitism in invertebrates are numerous and exquisite, from bizarre ectospore appendages, multiple spore morphotypes, and polyxenous life cycles, to effects on host behavior, population dynamics and sex ratio (Becnel and Andreadis, 2014; Stentiford and Dunn, 2014; Vavra and Larsson, 2014). Microsporidia demonstrate an arsenal of adaptations to evade the innate immunity of invertebrate hosts including modification of the phenol-oxidase cascade, accumulation in specialized haemocytes and adipocytes, stimulating host cells to grow into gigantic cells with prolonged cell cycles (e.g., "cysts" in insects, and "xenomas" in fish), that form protected and nutrient-supplied niches for developing parasites. Biochemical and molecular studies have revealed that microsporidia are able to modulate host cell cycles by inhibition of
apoptosis, and influence host gene expression and metabolism by secreting diverse regulatory factors into the host cell (Senderskiy et al., 2014; Williams et al., 2014).
Tight ecological bonds within the aquatic habitats via numerous intersecting and overlapping food chains could have played a leading role in microsporidia host switches from invertebrates to fish (and in reciprocal host transfers), as suggested by phylogenetic analyses (Stentiford et al., 2013). Circumstantial evidence indicates that the most common parasite of the White Atlantic shrimp, Agmasoma penaei, cycles between shrimp and perci-form fish feeding on juvenile penaeids (Johnson, 1995; Overstreet, 1973; Pasharawipas and Flegel; 1994, Sokolova et al., 2015). The microsporidia parasites could have been spread among aquatic inhabitants also by parasites similar to sea fleas, Lepeophterius sp. (Copepoda), which are common fish ectoparasites related to free-living cyclopids. These crustaceans, like freshwater copepods, can be parasitized by several microsporidia species at least two of which display close evolutionary distances with a fish microsporidium, Nucleospora salmoni (Freeman and Sommerville, 2009; Jones et al., 2012). This suggests the presence ofpolyxenous fish-crustaceans life cycles now or in the past. Existence of such a cycle has been recently demonstrated for Desmozoon (Paranucleospora) theredion, a species that parasitizes simultaneously an Atlantic salmon and its copepod parasite (Nylund et al., 2010). The much broader distribution of microsporidia among fish versus birds and mammals can be explained by the fact that switching to parasitism in fish did not demand special adaptations to the elevated body temperatures, a major factor together with humoral immunity that has limited the spread of microsporidia among warm-blooded animals.
"Human microsporidia" and related species
The importance of the "clues from invertebrate research" for understanding the origin and accessing the threat of microsporidia to the human population are evident. Excluding two Pleistophora species, likely related to fish congeners, 12 species and 6 genera of microsporidia recorded as infectious to humans are either insect parasites themselves, like Tubulinosema acridophagus and Anncaliia algerae, or have close relatives among insect parasites (Table 1). The only exception, Enterocytozoon bieneusi, is likely derived from a microsporidium infecting
a marine arthropod given the broad distribution of enterocytozoonids among marine crustaceans (Stentiford et al., 2013).
Mammals are the very recent hosts for microsporidia parasites from the evolutionary perspective. Microsporidia were adapted to intracellular parasitism in invertebrates well before switching to mammals, and a few lineages were apparently more successful in expanding their host range to vertebrates than others. Hence, addressing phylogenetic bonds between the species infecting humans and those parasitizing invertebrates might help to answer questions posed in the first paragraph of this short review.
Cystosporogenes/ Endoreticulatus/ Vittaforma clade
Vittaforma corneae was once isolated from corneal stroma of immunocompetent HIV-negative patient, and was the first human microsporidium placed in culture. However, some authors maintain that this species cannot be considered a true human pathogen (Van Frankenhuyzen et al., 2004). It shares 98.2% identity of its rDNA sequence with the lepidopteran microsporidium Cystosporogenes legeri and most likely is an unknown isolate of a closely related Cystosporogenes species accidentally developing in the immune-privileged site. Infection with V. corneae in immunocompetent patients is associated with self-limited short-term conjunctivitis caused presumably by traumatic inoculation of environmental spores of the insect pathogen (Weiss, 2014). Recently another species clustering within the same Endoreticulatus-Cystosporogenes-Vittaforma clade was found to cause myositis in the immunocompetent patient (Suankratay et al., 2012). Comparatively low identity (83-91%) of this Microsporidium sp. precluded the authors from assigning it to Endoreticulatus. The Endoreticulatus-Cystosporogenes-Vittaforma clade is composed predominantly of parasites of Lepidoptera. Endoreticulatus spp. though also has been isolated from two other orders of insects, Coleoptera and Orthoptera (Pilarska et al., 2015). Cystosporogenus legeri, a common parasite of insect rearing facilities, infects as a many as 5 families of Lepidoptera demonstrating unusually broad host range (Van Frankenhuyzen et al., 2004). So, among representatives of this clade there are generalist parasites with broad host ranges among natural hosts. Another feature that might facilitate the transition to a new group of hosts is resistance of Cystosporogenes
spp. spores to high (up to 42o C) temperatures (Van Frankenhuyzen et al., 2004). Butterflies and moths, some of which are known as facultative blood and tear feeders (Plotkin and Goddard, 2013; Zaspel et al., 2014), could be vectors for transmission of microsporidia belonging to this clade.
Anncaliia/ Tubulinosema clade
Anncaliia/Tubulinosema clade is composed of two genera with extraordinarily broad host ranges for insect microsporidia. The host range of Anncaliia spp. includes representatives of at least two insect orders, Coleoptera, and Diptera (Franzen et al., 2006b), and also amphipod crustaceans (Tokarev et al., 2014). Tubulinasema spp. parasitize as many as 5 insect orders (Lepidoptera, Orthopteran, Coleoptera, Diptera and Hymenoptera) (Franzen et al., 2006a). Tubulinosema acridophagus from a grasshopper was found to cause myositis and disseminated infection in a patient with a bone marrow transplant (Weiss, 2014). This is a typical opportunistic infection, but as in the case involving the Endoreticulatus-related Microsporidium sp., it clearly demonstrates insignificance of temperature as a limiting factor for the parasite development. Anncaliia spp. probably diversified further as parasites of mammals. Anncaliia (Brachiola) algerae, a common mosquito parasite of several genera of mosquitoes (Andreadis, 2007), occasionally infects brain and eye tissues, and causes disseminated disease in immunocompromised individuals. It also may induce skin and muscle infections presumably transmitted by a mosquito vector in immunologically healthy humans (Weiss, 2014). Anncaliia algerae is known to develop infections in SCID mice (Koudela et al., 2001), and tolerate elevated temperatures; it can be cultivated in cell lines at >36o C (Trammer et al., 1999). Resistance of A. algerae to high temperatures could be an ecological adaptation, since this parasite in nature infects mosquito larvae inhabiting small pools heated during the summer period. The two other representatives of the genus, A. connori and A. vesicularum, have been recorded from humans with immunodeficiency, and their environmental source is unknown (Weiss, 2014).
Trachipleistophora/ Vavraia clade
Reperesentatives of this clade, Trachipleistaphora hominis and T. anthropophtera, the most widespread causative agents of human myositis due to micro-
sporidia, have been recorded from several immuno-defficient individuals (Suankratay et al., 2012). insect origin of these infections was suggested by successful experimental infection of insect larvae with human-isolated T. hominis (Weidner et al., 1999). One representative of this genus was described from a Madagascar insectivore, Hemicentatis semi-spinosus (Vavra et al., 2006). Interestingly, this mammal belongs to the peculiar family Tenrecidae (order Afrosoricida), which members are characterized by lower body temperatures. The spores isolated from the animal were also infectious to Spodoptora littoralis (Lepidoptera) larvae. It is unclear whether T. extenrec is a native parasite of tenrecs, or an insect pathogen. It may develop in both types of hosts, suggesting a potential transmission route from insects to insectivorous mammals. Trachipleistaphora hominis is a close relative of the mosquito microsporidium Vavraia culicis, sharing with the latter 98% of SSUrDNA sequence similarity. Vavraia culicis parasitizes mosquitoes belonging to 6 genera (Andreadis, 2007). Microsporidia from mosquitoes, as a rule, are species- or genus-specific "specialists." Among >30 mosquito-infecting microsporidia a similarly broad range of hosts is known only for the above mentioned human-pathogenic species, Anncaliia algerae (Andreadis, 2007). One more Vavraia (V oncoperae) was described from a lepidopteran host (Malone and Mclvor, 1995).
interestingly, all three lineages of insect micro-sporidia that contain forms known to infect humans, include taxa of generalist pathogens as well as species tolerating high temperatures. Such consistency may suggest specific biochemical pre-adaptations required for transmission to a foreign warm-blooded host for these groups. Genes and regulatory factors responsible for these adaptations are yet to be identified by genomic and proteomic analyses. Factors analogous to LRR proteins or products of the InterB multigene family (Williams et al., 2014) might potentially play a role in regulating limits of host specificity within certain lineages.
Enterocytozoon bieneusi and Encephalitozoon spp.
Enterocytozoon bieneusi, a specialized parasite of enterocytes, is the most common microsporidium known to cause diseases in humans, particularly in patients with AIDS. E. bieneusi is widely distributed among several orders of mammals, and also has been recorded in birds (Fayer and Santin-Duran, 2014).
The evolutionary history of E. bienusi parasitism in vertebrates is probably relatively short, since the taxon has not diversified into separate species, but is represented by numerous genotypes with different levels of host specificity (Fayer and Santin-Duran, 2014). The closest relatives of this microsporidium infect fish and crustaceans (Stentiford et al., 2013), so E.bieneusi ancestors probably transferred to parasitism in vertebrates from these hosts via food chains.
Five of six species of the genus Encephalitozoon — E.cuniculi, E. intestinalis, E. hellem, E. lacertae and E. pogonae, parasitize vertebrates — mammals, birds and reptiles, and one species, E.romaleae, has been found in an insect, the lubber grasshopper Romalea microptera (Lange et al., 2009). Encephalitozoon cuniculi is the most famous and ubiquitous microsporidium of mammals. It has diverged into at least three mammalian host-specific genotypes (Didier et al., 1995). In reptiles E.cuniculi or morphologically identical species causes multi-systemic granulomatous disease (Koudela et al., 1998; Richter et al., 2013). Infection discovered recently in a bearded dragon Pogona vitticeps (Richter et al., 2013), were caused by a yet unknown genotype that occurred to be a new species Encephalitozoon pogonae (Sokolova et al., in press). E. hellem is a natural pathogen of birds, and E. intestinalis is more restricted to humans (Snowden, 2014). Encephalitozoon spp, unlike E. bieneusi, are not confined to infecting gastrointestinal tracts, but often cause disseminated microsporidiosis (Weiss, 2014). In phylogenetic reconstructions the Encephalitozoon lineage clusters within the Clade 4 of "Terresporidia" (Vossbrinck and Debrunner-Vossbrinck, 2005) composed of predominantly insect microsporidia. The Encephalitozoon branch forms a dichotomy with Mockfordia xanthocaeciliae, a parasite of Xanthocaecilia sommermanae, order Psocoptera. Psocoptera is considered to be the most basal order of hemipteroids, originating during the Permian Period 295-248 million years ago. Psocoptera are closely related to Phthiraptera, sucking lice, which parasitize warm-blooded animals including humans. These two orders are placed in the infraorder Psocodea and share a common ancestor, based on robust morphological and molecular evidence (Johnson and Mockford, 2003; for other references, see Sokolova et al., 2010). Though the majority of barklice are free living species, various species of Psocoptera inhabit plumage of birds and the pelage of mammals, as well as their nests. This short-
term commensal-type relationship presumably gave rise to obligate parasitism characteristic to Phthiraptera (Johnson et al., 2004). Evidence of a close relationship of M. xanthocaecilliae to Ence-phalitozoon spp. (Sokolova et al., 2010), ubiquitous parasites of birds and mammals, supports the idea that the association of ancestral Psocodea with mammals and birds could be one of the avenues of transfer of Microsporidia from arthropods to warmblooded hosts. Within the Encephalitozoon clade the position of E. romaleae (Lange et al., 2009), which shares 96% of SSUrDNA similarity with E. hellem, certainly creates a problem. As an explanation of striking genetic relatedness of E.romaleae to E. hellem, perhaps this species evolved as a result of reciprocal transfer of the E. hellem-related bird-infecting microsporidium back to insects (Sokolova et al., 2010). The genomic survey revealed that the genomes of E. hellem and E. romaleae contained the gene for purine nucleotide phosphatase (PNP), a component of the purines salvage pathway, of insect origin (Pombert et al., 2012, Selman et al., 2011). This gene is absent in the genomes of E. intestinalis, E. cuniculi and other microsporidia with sequenced genomes, and likely was acquired from an insect host by a common ancestor of E. romaleae and E. hellem. The narrow distribution of this gene is most consistent with its recent gain (Selman et al., 2011) and conforms to the idea of reciprocal transfer that might have occurred relatively recently. Of note, the Encepahlitozoon spp.-derived PNP genes cluster with the orthologue from Pediculus humanus (Phthiraptera) (Fig. 1 in Selman et al., 2011). This suggests that a lice-related ectoparasite of birds harboring an ancestral encephalitozoonid could have been a source for the PNP gene transfer from insects to the E. hellem-E. romalea lineage. Further molecular studies based on broader sampling and robust analyses could test this hypothesis.
Concluding remarks and further directions
In fact, only four species belonging to two genera can be considered true mammalian parasites: Enterocytozoon bieneusi, Encephalitozoon cuniculi, E. intestinalis, and E. hellem. They have evolved as parasites of warm-blooded vertebrates and might represent a serious threat to human populations as zoonotic infections (Fayer and Santin-Duran, 2014). Records of other microsporidia in mammals, including humans, are more or less accidental.
However, it is hard to argue the opinion expressed by Irma V. Issi, that "now microsporidia represent a numerous and aggressive group of parasites expanding the range of their hosts" (Issi, 1986). Recorded cases of microsporidiosis demonstrate the consecutive stages of microsporidia transforming into parasites of humans: from transient arthropod-related microsporidia known by sequences in stools of AIDS patients (Genebank accessions CQ408913, CQ408914; Sokolova et al., 2011), through accidental surface infections in immunocompromised patients (Endoreticulatus-like Microsporidium sp., Tubulinosema) and development in immune privileged tissues of eyes (Vittaforma), skin, and muscles due to accidental exposure to spores of a "generalist" microsporidium (Trachipleistaphora, Anncaliia), to specialized infections of gut epithelium (Enterocytozoon), and systemic microsporidiosis disseminated by macrophages (Encephalitozoon). Potential sources of human infection with invertebrate microsporidia are likely associated with "generalist" parasites, and, particularly, with human (mammalian) hyperparasites. Further surveys of Microsporidia in Psocoptera, Phthiraptera and related orders, as well as in other ectoparasitic or bloodsucking insects (fleas, bed-bugs, dipterans and hematophagous lepidopterans) and acarines (ticks, mites and chiggers) will shed light on evolutionary routes of host transfers as well as on possible risks of infection.
Acknowledgements
I am thankful to Chris Carlton (Louisiana State University) for editing the manuscript and to Igor Sokolov for discussions and ideas.
References
Andreadis T.G. 2007. Microsporidian parasites of mosquitoes. J. Am. Mosquito Contr. Assoc. 23, 3-29.
Becnel J.J. and Andreadis T.G. 2014. Microsporidia in Insects. In: Microsporidia: pathogens of opportunity. 1st ed. John Wiley & Sons, Inc., pp. 521-570.
Cali A. and Takvorian P.M. 2003. Ultrastructure and development of Pleistophora ronneafiei n. sp., a microsporidium (Protista) in the skeletal muscle of an immune-compromised individual. J. Euk. Microbiol. 50, 77-85.
Cali A., Takvorian P.M., Lewin S., Rendel M., Sian C.S., Wittner M., Tanowitz H.B., Keohane E. and Weiss L.M. 1998. Brachiola vesicularum, n. g., n. sp., a new microsporidium associated with AIDS and myositis. J. Euk. Microbiol. 45, 240—251.
Cali A., Weiss L.M., and Takvorian P.M. 2005. A review of the development of two types of human skeletal muscle infections from microsporidia associated with pathology in invertebrates and coldblooded vertebrates. Folia Parasitol. 52, 51—61.
Cheney S.A., Lafranchi-Trist em N.J. and Canning E.U. 2000. Phylogenetic relationships of Pleistophora-Mke microsporidia based on small subunit ribosomal DNA sequences and implications for the source of Trachipleistophora hominis infections. J. Euk. Microbiol. 47, 280-287.
Corsaro D., Walochnik J., Venditti D., Steinmann J., Müller K.-D. and Michel R. 2014. Micro-sporidia-like parasites of amoebae belong to the early fungal lineage Rozellomycota. Parasitol. Res. 113, 1909-1918.
Didier E.S., Vossbrinck C.R., Baker M.D., Rogers L.B., Bertucci D.C., and Shadduck J.A. 1995. Identification and characterization of three Encephalitozoon cuniculi strains. Parasitology. 111, 411-421.
Docker M.F., Kent M.L., Hervio D.M.L. Khattra J.S., Weiss L.M., Call A., and Devlin R.H.
1997. Ribosomal DNA sequence of Nucleospora salmonis Hedrick, Groff and Baxa, 1991 (Micro-sporea: Enterocytozoonidae): Implications for phylogeny and nomenclature. J. Euk. Microbiol. 44, 55-60.
Fayer R. and Santin-Duran M. 2014. Epidemiology of microsporidia in human Infections. In: Microsporidia: pathogens of opportunity. First ed. (Eds: Weiss L.M. and Becnel J.J.). John Wiley & Sons, Inc., pp. 135-164.
Franzen C., Futerman P.H., Schroeder J., Salzberger B. and Kraaijeveld A.R. 2006a. An ultrastructural and molecular study of Tubulinosema kingi Kramer (Microsporidia: Tubulinosematidae) from Drosophila melanogaster (Diptera: Drosophili-dae) and its parasitoid Asobara tabida (Hymeno-ptera: Braconidae). J. Invertebr. Pathol. 91, 158-167.
Franzen C., Nassonova E.S., Schölmerich J. and Issi I.V. 2006b. Transfer of the members of the genus Brachiola (Microsporidia) to the genus Anncaliia based on ultrastructural and molecular data. J. Euk. Microbiol. 53, 26-35.
Freeman M.A. and Sommerville C. 2009. Desmo-zoon lepeophtherii n. gen., n. sp., (Microsporidia:
Enterocytozoonidae) infecting the salmon louse Lepeophtheirus salmonis (Copepoda: Caligidae). Parasites and Vectors. 2 (doi:10.1186/1756-3305-2-58).
Haag K.L., James T.Y., Pombert J.F., Larsson R., Schaer T.M.M. Refardt D. and Ebert D. 2014. Evolution of a morphological novelty occurred before genome compaction in a lineage of extreme parasites. Proc. Natl. Acad. Sci. USA. 111, 15480-15485.
Issi I.V. 1986. Microsporida as a type of parasitic Protozoa. In: Microsporidia. (Eds: Beyer T.V. and Issi I.V.). Nauka, Leningrad, pp. 6-136.
Johnson K.P. and Mockford E.L. 2003. Molecular systematics of Psocomorpha (Psocoptera). Syst. Entomol. 28, 409-416.
Johnson K.P., Yoshizawa K. and Smith V.S. 2004. Multiple origins of parasitism in lice. Proc. Royal Soc. B: Biological Sciences. 271, 1771-1776.
Johnson S.K. 1995. Handbook of shrimp diseases. College Station, Texas A&M University.
Jones S.R.M., Prosperi-Porta G. and Kim E. 2012. The diversity of microsporidia in parasitic copepods (Caligidae: Siphonostomatoida) in the northeast pacific ocean with description of Facilispora margolisi n. g., n. sp. and a new family Facilisporidae n. fam. J. Euk. Microbiol. 59, 206-217.
Karpov S.A., Mikhailov K.V., Mirzaeva G.S., Mirabdullaev I.M., Mamkaeva K.A., Titova N.N. and Aleoshin V.V. 2013. Obligately phagotrophic aphelids turned out to branch with the earliest-diverging fungi. Protist. 164, 195-205.
Kent M.L., Shaw R.W. and Sanders J.L. 2014. Microsporidia in fish. In: Microsporidia: pathogens of opportunity. 1st ed. John Wiley & Sons, Inc., pp. 493-520.
Koudela B., Didier E.S., Rogers L.B., Modry D. and Kucerovâ S. 1998. Intestinal microsporidiosis in African skink Mabuya perrotetii. Folia Parasitol. 45, 149-155.
Koudela B., Visvesvara G.S., Moura H. and Vâvra J. 2001. The human isolate of Brachiola algerae (Phylum Microspora): Development in SCID mice and description of its fine structure features. Parasitology. 123, 153-162.
Lange C.E., Johny S., Baker M.D., Whitman D.W. and Solter L.F. 2009. A new Encephalitozoon species (microsporidia) isolated from the lubber grasshopper, Romalea microptera (Beauvois) (Orthoptera: Romaleidae). J. Parasitol. 95, 976-986.
Larsson J.I.R. 2014. The primitive microsporidia. In: Microsporidia: pathogens of opportunity. First
ed. (Eds: Weiss L.M. and Becnel J.J.). John Wiley & Sons, Inc., pp. 605—635.
Leander B.S. 2008. Marine gregarines: evolutionary prelude to the apicomplexan radiation? Trends in Parasitol. 24, 60—67.
Letcher P.M., Lopez S., Schmieder R., Lee P.A., Behnke C., Powell M.J. and McBride R.C. 2013. Characterization of Amoeboaphelidium proto-coccarum, an algal parasite new to the Cryptomycota isolated from an outdoor algal pond used for the production of biofuel. PLoS ONE. 8 (doi:10.1371/ journal.pone.0056232).
Malone L.A. and Mclvor C.A. 1995. DNA probes for two microsporidia, Nosema bombycis and Nosema costelytrae. J. Invert. Pathol. 65, 269—273.
Nylund S., Nylund A., Watanabe K., Arnesen
C.E. and Karlsbakk E. 2010. Paranucleospora theridion n. gen., n. sp. (Microsporidia, Entero-cytozoonidae) with a life cycle in the salmon louse (Lepeophtheirus salmonis, Copepoda) and atlantic salmon (Salmo salar). J. Euk. Microbiol. 57, 95—114.
Overstreet R.M. 1973. Parasites of some penaeid shrimps with emphasis on reared hosts. Aquaculture. 1, 105-140.
Pasharawipas T. and Flegel T.W. 1994. A specific DNA probe to identify the intermediate host of a common microsporidian parasite of Penaeus merguensis and P. monodon. Asian Fish. Sci. 7, 157-167.
Perkins F.O., Barta J.R., Clopton R.E., Pierce M.A. and Upton S.J. 2000. Phyllum Apicomplexa. In: The Illustrated Guide to the Protozoa. 2nd ed. (Eds: Lee J.J. et al.). Society of Protozoologists, pp. 190-369.
Pilarska D.K., Radek R., Huang W.F., Takov
D.I., Linde A., and Solter L.F. 2015. Review of the genus Endoreticulatus (Microsporidia, Encephalitozoonidae) with description of a new species isolated from the grasshopper Poecilimon thoracicus (Orthoptera: Tettigoniidae) and transfer of Microsporidium itiiti Malone to the genus. J. Invert. Pathol. 124, 23-30.
Plischuk S., Sanscrainte N.D., Becnel J.J., Estep A.S. and Lange C.E. 2015. Tubulinosema pampeana sp. n. (Microsporidia, Tubulinosematidae), a pathogen of the South American bumble bee Bombus atratus. J. Invert. Pathol. 126, 31-42.
Plotkin D. and Goddard J. 2013. Blood, sweat, and tears: A review of the hematophagous, sudophagous, and lachryphagous Lepidoptera. J. Vector Ecol. 38, 289-294.
Pombert J.F., Selman M., Burki F., Bardell F.T., Farinelli L., Solter L.F., Whitman D.W., Weiss
L.M., Corradi N. and Keeling P.J. 2012. Gain and loss of multiple functionally related, horizontally transferred genes in the reduced genomes of two microsporidian parasites. Proc. Natl. Acad. Sci. USA. 109, 12638-12643.
Richter B., Csokai J., Graner I., Eisenberg T., Pantchev N., Eskens H.U. and Nedorost N. 2013. Encephalitozoonosis in two inland Bearded Dragons (Pogona vitticeps). J. Comp. Pathol. 148, 278-282.
Selman M., Pombert J.F., Solter L., Farinelli L., Weiss L.M., Keeling P. and Corradi N. 2011. Acquisition of an animal gene by microsporidian intracellular parasites. Curr. Biol. 21, R576-R577.
Senderskiy I.V., Timofeev S.A., Seliverstova E.V., Pavlova O.A. and Dolgikh, V.V. 2014. Secretion of Antonospora (Paranosema) locustae proteins into infected cells suggests an active role of microsporidia in the control of host programs and metabolic processes. PLoS ONE. 9 (doi:10.1371/ journal.pone.0093585).
Snowden K.F. 2014. Microsporidia in higher vertebrates. In: Microsporidia: pathogens of opportunity. 1st ed. John Wiley & Sons, Inc., pp. 469-491.
Sokolova Y.Y., Lange C.E. and Fuxa J.R. 2007. Establishment of Liebermannia dichroplusae n. comb. on the basis of molecular characterization of Perezia dichroplusae Lange, 1987 (Microsporidia). J. Euk. Microbiol. 54, 223-230.
Sokolova Y.Y., Sokolov I.M. and Carlton C.E. 2010. New microsporidia parasitizing bark lice (Insecta: Psocoptera). J. Invert. Pathol. 104, 186-194.
Sokolova O.I., Demyanov A.V., Bowers L.C., Didier E.S., Yakovlev A.V., Skarlato S.O. and Sokolova Y.Y. 2011. Emerging microsporidian infections in Russian HIV-infected patients. J. Clin. Microbiol. 49, 2102-2108.
Sokolova Y.Y., Paskerova G.G., Rotari Y.M., Nassonova E.S. and Smirnov A.V. 2013. Fine structure of Metchnikovella incurvata Caullery and Mesnil 1914 (microsporidia), a hyperparasite of gregarines Polyrhabdina sp. from the polychaete Pygospio elegans. Parasitology. 140, 855-867.
Sokolova Y.Y., Paskerova G.G., Rotari Y.M., Nassonova E.S. and Smirnov A.V. 2014. Description of Metchnikovella spiralis sp. n. (Microsporidia: Metchnikovellidae), with notes on the ultrastructure of metchnikovellids. Parasitology. 141, 1108-1122.
Sokolova Y., Pelin A., Hawke J. and Corradi N. 2015. Morphology and phylogeny of Agma-
soma penaei (Microsporidia) from the type host, Litopenaeus setiferus, and the type locality, Louisiana, USA. Int. J. Parasitol. 45, 1-16.
Sokolova Y.Y., Sakaguchi K., Paulsen D. B. Establishing a new species Encephalitozoon pogonae for the microsporidian parasite of inland bearded dragon Pogona vitticeps Ahl 1927 (Reptilia, Squama-ta, Agamidae). J. Euk. Microbiol. (in press).
Stentiford G.D. and Dunn A.M. 2014. Microsporidia in aquatic invertebrates. In: Microsporidia: pathogens of opportunity. 1st ed. John Wiley & Sons, Inc., pp. 579-604.
Stentiford G.D., Feist S.W., Stone D.M., Bateman K.S. and Dunn A.M. 2013. Microsporidia: Diverse, dynamic, and emergent pathogens in aquatic systems. Trends in Parasitol. 29, 567-578.
Suankratay C., Thiansukhon E., Nilaratanakul V., Putaporntip C., and Jongwutiwes S. 2012. Disseminated infection caused by novel species of Microsporidium, Thailand. Emerg. Infect. Dis. 18, 302-304.
Tokarev Y., Voronin V., Rusakovitch E. and Issi I. 2014. Detection of Microsporidia in Gammarids in the delta of the Kuban River (Azov Sea, Russia). Contributed paper, 47th Annual Meeting of the Society for Invertebrate Pathology, 3-7 August 2014, Mainz, Germany, Program and Abstracts, P. 86.
Tourtip S., Wongtripop S., Stentiford G.D., Bateman K.S., Sriurairatana S., Chavadej J. and Withyachumnarnkul B. 2009. Enterocytozoon hepatopenaei sp. nov. (Microsporida: Enterocyto-zoonidae), a parasite of the black tiger shrimp Penaeus monodon (Decapoda: Penaeidae): fine structure and phylogenetic relationships. J. Invert. Pathol. 102, 21-29.
Trammer T., Chioralia G., Maier W.A. and Seitz H.M. 1999. In vitro replication of Nosema algerae (Microsporidia), a parasite of anopheline mosquitoes, in human cells above 36° C. J. Euk. Microbiol. 46, 464-468.
Van Frankenhuyzen,K., Ebling P., McCron B., Ladd T., Gauthier D. and Vossbrinck C. 2004. Occurrence of Cystosporogenes sp. (Protozoa, Microsporidia) in a multi-species insect production facility and its elimination from a colony of the eastern spruce budworm, Choristoneura fumiferana
(Clem.) (Lepidoptera: Tortricidae). J. Invert. Pathol. 87, 16-28.
Vâvra, J. Horâk A., Modry D., Lukes J. and Koudela B. 2006. Trachipleistophora extenrec n. sp. a new microsporidian (Fungi: Microsporidia) infecting mammals. J. Euk. Microbiol. 53, 464-476.
Vâvra J., Kamler M., Modry D. and Koudela B. 2011. Opportunistic nature of the mammalian microsporidia: Experimental transmission of Trachipleistophora extenrec (Fungi: Microsporidia) between mammalian and insect hosts. Parasitology Research. 108, 1565-1573.
Vâvra J. and Larsson J.I.R. 2014. Structure of Microsporidia. In: Microsporidia: pathogens of opportunity. 1st ed. John Wiley & Sons, Inc. pp. 1-70.
Vâvra, J. and Lukes, J. (2013) Microsporidia and 'the art of living together'. Advances in Parasitology 82, 253-319.
Vossbrinck, C.R. and Debrunner-Vossbrinck, B.A. (2005) Molecular phylogeny of the Microsporidia: ecological, ultrastructural and taxonomic considerations. Folia Parasitologica 52, 131-142.
Vossbrinck C.R., Debrunner-Vossbrinck B.A. and Weiss L.M. 2014. Phylogeny of the Microsporidia. In: Microsporidia: pathogens of opportunity. 1st ed. John Wiley & Sons, Inc., pp. 203-220.
Weidner E., Canning E.U., Rutledge C.R. and Meek C.L. 1999. Mosquito (Diptera: Culicidae) host compatibility and vector competency for the human myositic parasite Trachipleistophora hominis (Phylum Microspora). J. Med. Entomol. 36, 522-525.
Weiss L.M. 2014. Clinical syndromes associated with microsporidiosis. In: Microsporidia: pathogens of opportunity. 1st ed. John Wiley & Sons, Inc., pp. 371-401.
Williams B.A.P., Dolgikh V.V. and Sokolova Y.Y. 2014. Microsporidian biochemistry and physiology. In: Microsporidia: pathogens of opportunity. 1st ed. John Wiley & Sons, Inc.,pp. 245-260.
Zaspel J.M., Scott C.H., Hill S.R., Ignell R., Kononenko V.S. and Weller S.J. 2014. Geographic distribution, phylogeny, and genetic diversity of the fruit-and blood-feeding moth Calyptra thalictri Borkhausen (Insecta: Lepidoptera: Erebidae). J. Parasitol. 100, 583-591.
Address for correspondence: Yuliya Sokolova. Microscopy Center, Department of Comparative Biomedical Sciences School of Veterinary Medicine, Louisiana State University, 1909 Skip Bertman Drive Baton Rouge LA 70803; e-mail: [email protected], [email protected]
