5 Microsporidia

Abstract

The phylum Microsporidia comprises an interesting group of intracellular fungal parasites that infect vertebrate and invertebrate hosts of commercial and medical significance. Microsporidia are unique for their mode of infection, whereby the spore contents are propelled through an inverting polar filament and injected into the host cell. Believed to branch among the earliest eukaryotes, microsporidia are now considered more highly evolved for exhibiting gene reduction and compaction, which also are signs of extreme parasite efficiency. This chapter highlights the unusual mechanism of infection and current classification of the microsporidia and describes the morphology, life cycle, and culture of microsporidia species of importance to arthropod, aquatic, and mammalian hosts.

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I. Introduction

The phylum Microsporidia Balbiani 1882 (Weiser 1977) is comprised of a diverse group of over 1,400 species (Didier and Weiss 2006). These organisms are obligate intracellular pathogenic protists uniquely characterized by a specialized invasion organelle, the polar tube, through which the cytoplasm and nucleus of these organisms pass during the infection of their host cells. Long considered early branching eukaryotes classified with the Archezoa, the microsporidia are now considered fungi based on accumulated data and more sophisticated analyses (Hibbett et al. 2007; James et al. 2006). Microsporidia infect commercially significant animals, such as honey bees, salmon, silkworms, farm animals, and companion pets, and are of medical importance because they cause emerging opportunistic infections in humans. A wide range of animals that are less commercially relevant can also be infected with microsporidia and thus pose a risk as environmental reservoirs of infection (Santín and Fayer 2009a). As a consequence of studies to better characterize the tree of life, the microsporidia have undergone a major transition from placement with the earliest diverging eukaryotes to now being classified with the deep-branching fungi, closely related to the zygomycetes (Hibbett et al. 2007; James et al. 2006; Keeling 2009; Lee et al. 2008, 2010b). Questions remain, however, about the true relationship between the microsporidia and fungi (Koestler and Ebersberger 2011). Particularly noteworthy is the evolution of gene compaction and reduction observed among the microsporidia (Corradi and Keeling 2009; Keeling 2009; Keeling et al. 2010; Lee et al. 2010a; Texier et al. 2010; Vossbrinck et al. 2004). In this regard, microsporidia are highly efficient parasites, to the point that at least one species, Enterocytozoon bieneusi, even lacks genes for core carbon metabolism and depends fully on host-cell ATP import (Keeling et al. 2010).

II. Occurrence and Distribution

Microsporidia infect hosts ranging from protists to invertebrates (mainly insects) and vertebrates (mainly fish and mammals, including humans). The only extracellular stage that survives is the spore, which is relatively resistant to environmental stress. Because of the wide host range and the environmentally resistant spore wall, it is not surprising that microsporidia exhibit a worldwide distribution and can be found in aqueous (fresh and salt water) and terrestrial environments.

A. Arthropod Hosts

An extensive knowledge base on microsporidia isolated from terrestrial arthropod hosts has been dominated by species from insects, but other host groups include, but are not restricted to, cestodes, trematodes, nematodes, oligochaetes, isopods, myriapods, arachnids, and anaplurans (Sprague 1977). One indicator of the importance of microsporidia isolated from this group is that nearly half of the approximately 186 described genera shown in what follows are from terrestrial arthropod hosts, with the majority of these from insects (Becnel and Andreadis 1999). Prior to classification as fungi, the genera of microsporidia were established by the International Code of Zoological Nomenclature, and new genera are listed below in boldface type (Sprague and Becnel 1999). A request to exclude microsporidia from the International Code of Botanical Nomenclature was submitted and accepted and is now part of the revised code (Redhead et al. 2009). As such, the microsporidia are considered fungi, but descriptions of species remain subject to the International Code of Zoological Nomenclature.

The most commonly encountered microsporidia from insects are from Lepidoptera (within the Nosema/Vairimorpha clade) and Diptera, including the well-known genera Amblyospora and Parathelohania (Becnel and Andreadis 1999). It is likely that the large number of genera and species described from Lepidoptera and Diptera is due to the extensive studies searching for biological control agents in these groups of pest insects. Microsporidia in insects and other terrestrial arthropods have been studied extensively where they are important as natural control factors, have potential as manipulated microbial control agents for pest species, and cause chronic infections in beneficial arthropods.

Microsporidia as important natural pest control agents have been most extensively studied in insect pests. Nosema pyrausta in the European corn borer, Ostrinia nubilalis, is one of the most important regulators of larval populations in the USA (Andreadis 1984; Siegel et al. 1986). Populations of an important forest pest, the spruce budworm, Choristoneura fumiferana, are suppressed by epizootics caused by Nosema fumiferanae (Wilson 1973, 1981). A more recent example of natural control is in the red imported fire ant, Solenopsis invicta, where Kneallhazia (syn. Thelohania) solenopsae has become widespread in US populations of this invasive species (Oi et al. 2004).

Perhaps the best example of a species of microsporidia used as a microbial pesticide was a program for the control of rangeland grasshoppers with Paranosema (syns. Nosema and Antonospora) locustae (Henry and Oma 1981). P. locustae infects over 90 species of grasshoppers in the family Acrididae (Brooks 1988) and was registered by the US Environmental Protection Agency in 1989 as a microbial insecticide. Whole-organism technology was used to produce large numbers of P. locustae spores in grasshoppers that were formulated into bait and applied by air (Henry and Oma 1981). P. locustae does not cause rapid mortality but has a debilitating effect on the host that can have long-term control implications when introduced.

1. Abelspora Azevedo 1987 2. Aedispora Kilochitskii 1997 3. Agglomerata Larsson & Yan 1988 4. Agmasoma Hazard & Oldacre 1975 5. Alfvenia Larsson 1983 6. Alloglugea Paperna & Lainson 1995 7. Amazonspora Azevedo & Matos 2003 8. Amblyospora Hazard & Oldacre 1975 9. Ameson Sprague 1977 10. Amphiacantha Caullery & Mesnil 1914 11. Amphiamblys Caullery & Mesnil 1914 12. Andreanna Simakova, Vossbrinck & Andreadis 2008 13. Anisofilariata Tokarev, Voronin, Seliverstova, Dolgikh, Pavlova, Ignatieva & Issi 2010 14. Anncaliia Issi, Krylova & Nicolaeva 1993 15. Antonospora Fries, Paxton, Tengo, Slemenda, da Silva & Pieniazek 1999 16. Auraspora Weiser & Purrini 1980 17. Bacillidium Janda 1928 18. Baculea Loubes & Akbarieh 1978 19. Becnelia Tonka & Weiser 2000 20. Berwaldia Larsson 1981 21. Binucleata Refardt, Decaestecker, Johnson & Vávra 2008 22. Binucleospora Bronnvall & Larsson 1995 23. Bohuslavia Larsson 1985 24. Brachiola Cali, Takvorian & Weiss 1998 25. Bryonosema Canning, Refardt, Vossbrinck & Curry 2002 26. Burenella Jouvenaz & Hazard 1978 27. Burkea Sprague 1977 28. Buxtehudea Larsson 1980 29. Campanulospora Issi, Radischcheva & Dolzhenko 1983 30. Canningia Weiser, Wegensteiner & Zizka 1995 31. Caudospora Weiser 1946 32. Caulleryetta Dogiel 1922 33. Chapmanium Hazard & Oldacre 1975 34. Chytridioides Trégouboff 1913 35. Chytridiopsis Schneider 1884 36. Ciliatosporidium Foissner & Foissner 1995 37. Coccospora Kudo 1925 38. Cougourdella Hesse 1935 39. Crepidulospora Simakova, Pankova & Issi 2004 40. Crispospora Tokarev, Voronin, Seliverstova, Pavlova & Issi 2010 41. Cristulospora Khodzhaeva & Issi 1989 42. Cryptosporina Hazard & Oldacre 1975 43. Cucumispora Ovcharenko, Bacela, Wilkinson, Ironside, Rigaud & Wattier 2009 44. Culicospora Weiser 1977 45. Culicosporella Weiser 1977 46. Cylindrospora Issi & Voronin 1986 47. Cystosporogenes Canning, Barker, Nicholas & Page 1985 48. Dasyatispora Diamant, Goren, Yokeş, Galil, Klopman, Huchon, Szitenberg & Karhan 2010 49. Desmozoon Freeman & Sommerville 2009 50. Desportesia Issi & Voronin 1986 51. Dimeispora Simakova, Pankova & Issi 2004 52. Duboscqia Perez 1908 53. Edhazardia Becnel, Sprague & Fukuda 1989 54. Encephalitozoon Levaditi, Nicolau & Schoen 1923 55. Endoreticulatus Brooks, Becnel & Kennedy 1988 56. Enterocytozoon Desportes, Le Charpentier, Galian, Bernard, Cochand-Priollet, Lavergne, Ravisse & Modigliani 1985 57. Enterospora Stentiford, Bateman, Longshaw & Feist 2007 58. Episeptum Larsson 1986 59. Euplotespora Fokin, Giuseppe, Erra & Dini 2008 60. Evlachovaia Voronin & Issi 1986 61. Fibrillanosema Galbreath, Smith, Terry, Becnel & Dunn 2004 62. Flabelliforma Canning, Killick-Kendrick & Killick-Kendrick 1991 63. Geusia Rühl & Korn 1979 64. Glugea Thélohan 1891 65. Glugoides Larsson, Ebert, Vávra & Voronin 1996 66. Golbergia Weiser 1977 67. Gurleya Doflein 1898 68. Gurleyides Voronin 1986 69. Hamiltosporidium Haag, Larsson, Refardt & Ebert 2010 70. Hazardia Weiser 1977 71. Helmichia Larsson 1982 72. Hepatospora Stentiford, Bateman, Dubuffet, Chambers & Stone 2011 73. Hessea Ormières & Sprague 1973 74. Heterosporis Schubert 1969 75. Heterovesicula Lange, Macvean, Henry & Streett 1995 76. Hirsutusporos Batson 1983 77. Holobispora Voronin 1986 78. Hrabyeia Lom & Dyková 1990 79. Hyalinocysta Hazard & Oldacre 1975 80. lchthyosporidium Caullery & Mesnil 1905 81. Inodosporus Overstreet & Weidner 1974 82. Intexta Larsson, Steiner & Bjørnson 1997 83. Intrapredatorus Chen, Kuo & Wu 1998 84. Issia Weiser 1977 85. Janacekia Larsson 1983 86. Jirovecia Weiser 1977 87. Jiroveciana Larsson 1980 88. Johenrea Lange, Becnel, Razafindratiana, Przybyszewski & Razafindrafara 1996 89. Kabatana Lom, Dyková & Tonguthai, 2000 90. Kinorhynchospora Adrianov & Rybakov 1991 91. Kneallhazia Sokolova & Fuxa 2008 92. Krishtalia Kilochitskii 1997 93. Lanatospora Voronin 1986 94. Larssonia Vidtman & Sokolova 1994 95. Larssoniella Weiser & David 1997 96. Liebermannia Sokolova, Lange & Fuxa 2006 97. Loma Morrison & Sprague 1981

98. Mariona Stempell 1909 99. Marssoniella Lemmermann 1900 100. Merocinta Pell & Canning 1993 101. Metchnikovella Caullery & Mesnil 1897 102. Microfilum Faye, Toguebaye & Bouix 1991 103. Microgemma Ralphs & Matthews 1986 104. Microsporidium Balbiani 1884 105. Microsporidyopsis Schereschewsky 1925 106. Mitoplistophora Codreanu 1966 107. Mockfordia Sokolova, Sokolov & Carlton 2010 108. Mrazekia Léger & Hesse 1916 109. Myospora Stentiford, Bateman, Small, Moss, Shields, Reece & Tuck 2010 110. Myosporidium Baquero, Rubio, Moura, Pieniazek & Jordana 2005 111. Myxocystis Mrazek 1897 112. Nadelspora Olson, Tiekotter & Reno 1994 113. Napamichum Larsson 1990 114. Nelliemelba Larsson 1983 115. Neoflabelliforma Morris & Freeman 2010 116. Neoperezia Issi & Voronin 1979 117. Neonosemoides Faye, Toguebaye & Bouix 1996 118. Nolleria Beard, Butler & Becnel 1990 119. Norlevinea Vávra 1984 120. Nosema Naegeli 1857 121. Nosemoides Vinckier 1975 122. Novothelohania Andreadis, Simakova, Vossbrinck, Shepard & Yurchenko 2011 123. Nucleospora Docker, Kent & Devlin 1996 124. Nudispora Larsson 1990 125. Octosporea Flu 1911 126. Octotetrasporea Issi, Kadyrova, Pushkar, Khodzhaeva & Krylova 1990 127. Oligosporidium Codreanu-Bălcescu, Codreanu & Traciuc 1981 128. Ordospora Larsson, Ebert & Vávra 1997 129. Ormieresia Vivares, Bouix & Manier 1977 130. Orthosomella Canning, Wigley & Barker 1991 131. Orthothelohania Codreanu & Bălcescu-Codreanu 1974 132. Ovavesicula Andreadis & Hanula 1987 133. Ovipleistophora Pekkarinen, Lom & Nilsen 2002 134. Pankovaia Simakova, Tokarev & Issi 2009 135. Paraepiseptum Hyliš, Oborník, Nebesářová & Vávra 2007 136. Paranosema Sokolova, Dolgikh, Morzhina, Nassonova, Issi, Terry, Ironside, Smith & Vossbrinck 2003 137. Paranucleospora Nylund, Nylund, Watanabe, Arnesen & Karlsbakk 2010 138. Parapleistophora Issi, Kadyrova, Pushkar, Khodzhaeva & Krylova 1990 139. Parastempellia Issi, Kadyrova, Pushkar, Khodzhaeva & Krylova 1990 140. Parathelohania Codreanu 1966 141. Paratuzetia Poddubnaya, Tokarev & Issi 2006 142. Pegmatheca Hazard & Oldacre 1975 143. Perezia Léger & Duboscq 1909 144. Pernicivesicula Bylén & Larsson 1994 145. Pilosporella Hazard & Oldacre 1975 146. Pleistophora Gurley 1893 147. Pleistophoridium Codreanu-Bălcescu & Codreanu 1982 148. Polydispyrenia Canning & Hazard 1982 149. Potaspora Casal, Matos, Teles-Grilo & Azevedo 2008 150. Pseudoloma Matthews, Brown, Larison, Bishop-Stewart, Rogers & Kent 2001 151. Pseudonosema Canning, Refardt, Vossbrinck, Okamura & Curry 2002 152. Pseudopleistophora Sprague 1977 153. Pulcispora Vedmed, Krylova & Issi 1991 154. Pyrotheca Hesse 1935 155. Rectispora Larsson 1990 156. Resiomeria Larsson 1986 157. Ringueletium Garcia 1990 158. Schroedera Morris & Adams 2002 159. Scipionospora Bylén & Larsson 1996 160. Semenovaia Voronin & Issi 1986 161. Senoma Simakova, Pankova, Tokarev & Issi 2005 162. Septata Cali, Kotler & Orenstein 1993 163. Simuliospora Khodzhaeva, Krylova & Issi 1990 164. Spherospora Garcia 1991 165. Spiroglugea Léger & Hesse 1924 166. Spraguea Weissenberg 1976 167. Steinhausia Sprague, Ormières & Manier 1972 168. Stempellia Léger & Hesse 1910 169. Striatospora Issi & Voronin 1986 170. Systenostrema Hazard & Oldacre 1975 171. Tardivesicula Larsson & Bylén 1992 172. Telomyxa Léger & Hesse 1910 173. Tetramicra Matthews & Mattews 1980 174. Thelohania Henneguy 1892 175. Toxoglugea Léger & Hesse 1924 176. Toxospora Voronin 1993 177. Trachipleistophora Hollister, Canning, Weidner, Field, Kench & Marriott 1996 178. Trichoctosporea Larsson 1994 179. Trichoduboscqia Léger 1926 180. Trichonosema Canning, Refardt, Vossbrinck, Okamura & Curry 2002 181. Trichotuzetia Vávra, Larsson & Baker 1997 182. Tricornia Pell & Canning 1992 183. Tubulinosema Franzen, Fischer, Schroeder, Schölmerich & Schneuwly 2005 184. Tuzetia Maurand, Fize, Fenwik & Michel 1971 185. Unikaryon Canning, Lai & Lie 1974 186. Vairimorpha Pilley 1976 187. Vavraia Weiser 1977 188. Vittaforma Silveira & Canning 1995 189. Weiseria Doby & Saguez 1964 190. Wittmannia Czaker 1997

Microsporidia are perhaps best known because several prominent species are the causative agents of chronic disease in beneficial insects such as silkworms ( Bombyx mori ) and honey bees ( Apis melifera ). The first named species of microsporidia was Nosema bombycis from B. mori and was the subject of landmark studies by Louis Pasteur, who established this pathogen as the etiological agent of “pébrine” or silkworm disease (Pasteur 1870). Pasteur proved that N. bombycis was transmitted from adult to progeny via the egg (transovarial transmission) and by the ingestion of spores, and he developed preventive methods that saved the silkworm industry worldwide. Adult honey bees worldwide are afflicted by nosemosis, which has been caused historically by Nosema apis, and more recently the Asian species, Nosema ceranae, has been implicated as playing a major role (Chen et al. 2008). Interactions of Nosema spp. with other bee pathogens have been implicated in contributing to colony collapse disorder and declines in honey bee colonies worldwide (Ratnieks and Carreck 2010). In addition, numerous microsporidia are implicated in reducing the effectiveness of commercially produced biological control agents. A few select examples are Nosema muscidifuracis, which reduced the fitness of the muscoid fly parasitoid Muscidifurax raptor (Geden et al. 1995), and Oligosporidium occidentalis from the predatory mite Metaseiulus occidentalis, which has a negative impact on the overall fitness of this predator (Becnel et al. 2002).

B. Aquatic Hosts (Marine and Freshwater)

Microsporidia infect a broad range of aquatic organisms, including crustaceans and amphipods, and freshwater, saltwater, and anadromous fish . The impacts of microsporidian parasites on fish in aquaculture , wild populations, and research have been documented on several occasions (Lom and Dyková 1992; Shaw and Kent 1999), and microsporidian species belonging to some 18 genera have been described in fishes (Lom 2002; Lom and Nilsen 2003). Most of these infections seem to be chronic, with minimal host mortality. Infections by some species, however, can have a profound economic impact on wild fish and aquaculture hosts in terms of mortality and commercial quality of fish. Several of these microsporidia have been shown to impact fish either by directly killing the host or indirectly by reducing fecundity (Ramsay et al. 2009; Wiklund et al. 1996) or decreasing the commercial quality of farmed fish. For example, Becko disease in yellowtail is caused by the formation of cysts in skeletal muscle by the microsporidium Kabatana seriolae Egusa 1982.

With the rapid growth of aquaculture, microsporidian pathogens in fish have increased in importance, and three species of microsporidia are the main causes of disease in seawater-reared salmon . Loma salmonae results in high mortality of salmonids reared in freshwater hatcheries and in seawater netpens in North America and Europe due to chronic gill infections (Kent and Poppe 1998). Nucleospora salmonis, also a microsporidian parasite of the Chinook salmon Oncorhynchus tshawytscha, is unique in that it infects the host cell nucleus and results in lymphoblastosis and a leukemia-like condition in fish (Chilmonczyk et al. 1991). Another intranuclear microsporidium, Paranucleospora theridion, infects the rainbow trout, Oncorhynchus mykiss, and salmonids (e.g., Atlantic salmon, Salmo salar), causing up to 80 % mortality in Atlantic salmon farms in Norway (Nylund et al. 2010). Additionally, P. theridion can infect the salmon louse L. salmonis, providing a potential reservoir for this parasite.

Microsporidia are also common pathogens of baitfish . The shiner, Notemigonus crysoleucas, and fathead minnow, Pimephales promelas, are frequently infected by Ovipleistophora ovariae, which generally does not result in acute mortalities but significantly affects the fecundity of spawning fish. Additionally, due to the increased use of fish in research, infections by microsporidia can have a confounding impact on experimental results using such fish (Kent et al. 2011). Zebrafish, Danio rerio, are also affected by microsporidia, with the first report describing infection of the spinal cord in fish purchased from a pet store for use in toxicological studies (de Kinkelin 1980). After further characterization, this microsporidian was assigned to a new genus and species, Pseudoloma neurophilia (Matthews et al. 2001). P. neurophilia infections are widespread in laboratory facilities (Kent et al. 2011) and are generally characterized as chronic and occasionally associated with spinal deformities and emaciation. Pleistophora hyphessobryconis, a muscle-infecting microsporidian, has also been identified in laboratory populations of zebrafish (Sanders et al. 2010). Commonly known as neon tetra disease for its type host, Paracheirodon innesi, this parasite is a frequent problem in the aquarium trade, often resulting in considerable mortality in a wide range of fishes. P. hyphessobryconis has a remarkably broad host range, infecting some 20 species of fishes in 4 orders (Lom and Dyková 1992; Schäperclaus 1991; Steffens 1962). As with P. neurophilia, P. hyphessobryconis can be harbored by otherwise healthy-appearing fish that may show clinical signs of the infection only after experiencing immunosuppressive events. Clinical presentation of the disease includes massive infections of myocytes resulting in liquefactive necrosis of the muscle tissue that almost invariably leads to the death of the fish. This example highlights the importance of obtaining fish used in research from reputable sources and the potential for introducing a microsporidian with a broad host range to new or accidental hosts.

In contrast to P. hyphessobryconis, many other microsporidia of fish are host-specific, at least at the family or genus level. One example, L. salmonae, infects all species of Pacific salmon, Onchorynchus spp., but does not infect the Atlantic salmon, S. salar, based on results from experimental exposure of fish to L. salmonae-infected gill tissue (Shaw et al. 2000). Using polymerase chain reaction to monitor infection of intestine, heart, spleen, and gill tissues, experimental exposure to L. salmonae showed an aberrant progression in Atlantic salmon, S. salar, compared to that seen in the rainbow trout, O. mykiss (Sanchez et al. 2001). Parasite DNA was detected in all tissues tested until week 3, at which point, rather than progressing to the gills to complete the life cycle by forming mature spores, the parasite was apparently cleared. This illustrates an abortive life cycle by L. salmonae infection in a nonpermissive host whereby the parasite was able to invade certain tissues and proliferate to some extent but was unable to progress to sporogony.

C. Mammalian and Avian Hosts

Members of the genus Encephalitozoon are the most common microsporidia infecting mammals and birds . The type species of this genus, Enc. cuniculi, was first identified in rabbits with motor paralysis in 1922 (Wright and Craighead 1922) and was also the first microsporidian genome to be sequenced (Katinka et al. 2001). There are now several sequenced microsporidian genomes, and the data can be found at http://microsporidiadb.org/micro/. Enc. cuniculi has an extraordinarily wide host range among mammals, such as rodents, lagomorphs, canines, equines, nonhuman primates, and humans. Enc. hellem Didier et al. 1991 and Enc. (syn. Septata) intestinalis Cali et al. 1993 were later isolated and identified from AIDS patients (Cali et al. 1993; Didier and Weiss 2006; Didier et al. 1991). Enc. intestinalis is still considered more common in humans, whereas Enc. hellem is more common in birds with humans believed to be zoonotic hosts. Since the Encephalitozoon species are indistinguishable by light microscopy, reports of Enc. cuniculi in birds prior to the AIDS pandemic may actually have been due to Enc. hellem (Didier et al. 1998; Snowden and Logan 1999; Snowden et al. 2000). Encephalitozoon species may infect enteric sites and contribute to diarrhea, but they more typically cause systemic infections to persist over the life of the host unless treated with effective drugs (e.g., albendazole). Disease occurs predominantly in immune-deficient hosts (e.g., AIDS patients, organ transplant recipients undergoing immunosuppressive therapy) and occurs sporadically in immune-competent hosts (Kotler and Orenstein 1998; Weber et al. 2000).

The most prevalent microsporidian in humans, Ent. bieneusi Desportes et al. 1985, was first identified in an AIDS patient in Haiti and is primarily associated with persistent and self-limiting diarrhea in immune-deficient and immune-competent humans, respectively (Desportes et al. 1985; Didier and Weiss 2006). The host range of Ent. bieneusi seems to be far wider than first believed and now includes wild, farm, and companion pet animals (Santín and Fayer 2009a, b). In addition, Ent. bieneusi has been increasingly identified in avian hosts such as chickens, pigeons, falcons, and exotic birds (Graczyk et al. 2008; Haro et al. 2005; Muller et al. 2008; Reetz et al. 2002). Currently the genus Enterocytozoon contains only a single species, Ent. bieneusi.

It is possible, however, that this organism is a species complex, and as additional information is obtained, it may be split into separate species, as was done with Cryptosporidium parvum. It should also be appreciated that the family Enterocytozoonidae contains the genus Nucleospora, which has several species, including N. salmonis, previously named Ent. salmonis.

The pathogenesis of Ent. bieneusi infections in immune-competent humans and nonhuman hosts has not been well characterized. For example, it is unknown whether Ent. bieneusi persists in otherwise healthy people and reactivates under conditions of immune deficiency. Additional species of microsporidia less frequently identified in mammals and birds include Vittaforma corneae, Trachipleistophora spp., Anncaliia algerae, Pleistophora ronneaifiei, Nosema ocularum, and Microsporidium spp. (Didier and Weiss 2006).

III. Morphology of the Microsporidian Spore

A. General Description and Common Features

Microsporidian spores are generally small and vary from 1 to 20 μm in length (Fig. 5.1). Spores of most species of microsporidia are oval in shape but may also exhibit pyriform, spherical, or rod shapes. The spore wall provides resistance to environmental influences and allows for the increase in hydrostatic pressure that causes spore discharge (see below; Frixione et al. 1997). The spore wall is surrounded by a glycoproteinaceous electron-dense exospore and electron-lucent endospore composed primarily of chitin (Vavrá and Larsson 1999). Ultrastructural studies of the genus Encephalitozoon using transmission electron microscopy, freeze fracture, and deep etching demonstrated that the exospore is very complex and consists of three layers: an outer spiny layer, an intermediate electron-lucent lamina endospore, and an inner fibrous layer (Bigliardi et al. 1996). The endospore is observed as a space crossed by bridges connecting the exospore to the plasma membrane. It has been suggested that chitin, a major component of the endospore, comprises the fibrils forming the bridges across the endospore and is part of the fibrillar system of the exospore (Bigliardi et al. 1996; Erickson and Blanquet 1969; Prigneau et al. 2000; Vavrá 1976). It is possible to distinguish subcompartments within the spore wall using polyclonal antisera against partially purified microsporidial proteins. A glycine- and serine-rich 51-kDa protein named SWP1 is localized to the exospore in Enc. cuniculi (Bohne et al. 2000) and Enc. intestinalis (Hayman et al. 2001). The corresponding gene, swp1, has been identified in Enc. cuniculi, Enc. hellem, and Enc. intestinalis (Bohne et al. 2000; Hayman et al. 2001). SWP1 is absent in meronts (proliferating stages) and first seen in early sporonts (stages that differentiate into spores) at a time when organisms translocate from the periphery to the center of the parasitophorous vacuole (PV) (Bohne et al. 2000). A 150-kDa glycoprotein in the spore wall named SWP2 was identified in Enc. intestinalis (Hayman et al. 2001). In addition, a putative glycophosphatidylinositol (GPI)-anchored chitin deacetylase has been localized to the plasmalemma endospore interface. Using proteomic techniques, a new spore wall protein, SWP3/EnP2, corresponding to ECU01_1270, was identified and localized to the endospore (Peuvel-Fanget et al. 2006; Xu et al. 2006). By immunoelectron microscopy this protein was found on the cell surface during sporogony and in the endospore in mature spores. SWP3 has several potential O-glycosylation sites and is likely a mannosylated protein like the major polar tube protein (PTP1). EnP1, corresponding to ECU01_0820, has also been localized to the endospore and demonstrated to be involved in adhesion of the spore to host cells (Peuvel-Fanget et al. 2006; Southern et al. 2007).

Fig. 5.1

Diagram of internal structure of a microsporidian spore (left) and a transmission electron micrograph of Vavraia culicis floridensis from Aedes albopictus (right). The spore coat has an outer electron-dense region called the exospore (Ex) and an inner thicker electron-lucent region known as the endospore (En). A unit membrane (P) separates the spore coat from the spore contents. The extrusion apparatus—anchoring disk (A), polar tubule (Pt), lamellar polaroplast (Lp), and tubular polaroplast (Tp)—dominates the spore contents and is diagnostic for microsporidian identification. The posterior vacuole (Pv) is a membrane-bound vesicle that sometimes contains a membrane whirl, a glomerular structure, flocculent material, or some combination of these structures. The spore cytoplasm is dense and contains ribosomes (R) in a tightly coiled helical array. The nucleation may consist of a single nucleus or pair of abutted nuclei, a diplokaryon (D). The size of the spore depends on the particular species and can vary from less than 1 μm to more than 10 μm. The number of polar tubule coils also varies from a few to 30 or more, again depending on the species observed. Reprinted with permission from Cali and Owen (1988)

Under light microscopy, viable spores are refractile, and after histochemical staining (e.g., chromotrope, Gram), a posterior vacuole may be observed. The unique structure that characterizes all microsporidia is the polar tube or filament that coils within the spore and is part of the germination and infection apparatus (see below). The arrangement and number of coils of the polar filament within the spore vary among the microsporidia species. Long considered to be amitochondriate, the microsporidia have been found to possess reduced mitochondria called mitosomes, as well as atypical Golgi that lack the classical stacked dictyosome structure but instead are comprised of vesicular tubules that connect with the endoplasmic reticulum, plasma membrane, and developing polar tube. An anchoring disk with a membranous lamellar polaroplast is located at the anterior end of the spore and functions to anchor and fuel the extruding polar filament during germination (Keeling 2009; Vavrá and Larsson 1999). Ribosomes found along the endoplasmic reticulum are smaller than those of most eukaryotes (70S rather than 80S), being more similar to those of bacteria, and the microsporidian nucleus may exist in a monokaryon or diplokaryon arrangement.

B. Species (Spores) Infecting Arthropod Hosts

Studies on microsporidia in insects have been instrumental in establishing many aspects of microsporidian biology, but perhaps none more important than basic information about spores and spore types. For more than 100 years following the first microsporidian species N. bombycis from silk worms was named, the one spore/one species concept was almost universally accepted. Over 100 years later, Hazard and Weiser (1968) discovered that some microsporidian species infecting mosquitoes were observed to exhibit spore dimorphism, where two spore types with distinctive morphology and function were formed over the course of the life cycle. They reported that a binucleate spore formed in the adult female was responsible for transmitting pathogens to progeny. These studies revealed that in infected male larval progeny, uninucleate spores (meiospores) were produced, while spore development was delayed until pupation and adult emergence in infected female progeny. Binucleate spores of the original type were produced in these infected females to repeat the cycle. This proved that the two morphologically distinctive spores found in larvae and adult hosts (formerly believed to belong to two genera) may represent a single species. The means by which these microsporidia were transmitted horizontally remained a mystery until the discovery that meiospores formed in larvae were infectious to a copepod intermediate host (Sweeney et al. 1985). When ingested by mosquito larvae, the spores from the copepod intermediate host initiate a sequence of development that ends with binucleate spores in the adult female mosquito. Multiple spore types within the same species have also been documented for K. solenopsae infecting the fire ant Solenopsis invicta where four different spore types have been reported (Sokolova and Fuxa 2008), suggesting that this trait may be common in many genera.

C. Species (Spores) Infecting Aquatic Hosts

Spores of microsporidia infecting fish are generally spherical or ovoid to pyriform in shape and contain a sporoplasm that is either monokaryotic or diplokaryotic. Most species form spores of relatively uniform size and shape, but highly variable spore sizes (macrospores and microspores) occur together in tissues of hosts infected with members of the genera Heterosporis , Pleistophora , and Ichthyosporidium . The genus Pleistophora comprises numerous species that generally infect skeletal muscle of fish. Several of these species have been found to produce two and sometimes three spore types of different sizes (Canning et al. 1986). The type species of this genus, Pleistophora typicalis, produces elongate macrospores averaging 7.5 × 3.0 μm and, more commonly, microspores that are ovoid and average 4.4 × 2.3 μm. Spores of Pleistophora spp. generally have a large posterior vacuole, occupying over half the total spore volume.

Other microsporidia of fish produce polymorphic spores that vary by host and even tissue within the same host in which they develop. For example, the intranuclear microsporidian P. theridion develops spherical monokaryotic spores of 2.2–2.5 μm in the salmon louse, Lepeophtheirus salmonis. In the fish host, P. theridion produces spherical, diplokaryotic spores of 0.9–1.2 μm in diameter in reticuloendothelial cells and ovoid spores of 2.4–2.7 × 2.0–2.1 μm in gill and skin epithelial cells (Nylund et al. 2010). Nucleospora salmonis, another intranuclear microsporidian, develops similarly small, ovoid spores (1 × 2 μm), but other spore forms have not been seen for this organism (Chilmonczyk et al. 1991). Spores of Glugea anomala are elongate and oval, and spore size varies minimally in the same host species. However, there are some variations in sizes of spores taken from different hosts such as Gasterosteus aculeatus (3–6 × 1.9–1.7 μm) or Pungitius pungitius (3.5–5.1 × 1.9–2.6 μm), and here also the posterior vacuole is relatively large, taking up approximately half of the spore volume.

D. Species (Spores) Infecting Mammalian and Avian Hosts

Encephalitozoon spores measure approximately 1–2 × 2–4 μm and exhibit a typical microsporidian spore configuration of a glycoproteinaceous electron-dense exospore, electron-lucent endospore composed of chitin, and a plasma membrane containing the cytoplasmic organelles. The polar filament typically coils five to seven times in single row arrangement, and the nucleus is monokaryotic. Mature spores usually contain a prominent posterior vacuole that often is visible by light microscopy of histochemically stained organisms. Additional organelles include the membranous anterior anchoring disk, lamellar polaroplast with Golgi-like vesicles, endoplasmic reticulum, and ribosomes.

Spores of Ent. bieneusi are among the smallest of the microsporidia measuring 1 × 1.5 μm and the chitinous endospore in Ent. bieneusi is somewhat thinner than found in Encephalitozoon spores. The polar filament coils five to seven times and commonly is observed to align in two rows. A prominent posterior vacuole may be observed, and the nucleus is monokaryotic.

IV. Microsporidian Invasion Apparatus

The invasion apparatus of the microsporidia consists of a polar tube , also referred to as the polar filament prior to discharge (Lom and Vávra 1963; Takvorian and Cali 1986; Weidner 1972, 1976, 1982), that consists of two domains: an anterior straight region surrounded by a lamellar polaroplast that is attached to the inside of the anterior end of the spore by an anchoring disk and a posterior coiled region that forms from 4 to approximately 30 coils around the sporoplasm in the spore, depending on the species (Wittner and Weiss 1999). In cross section, the polar filament inside the spore is composed of electron-dense and electron-lucent concentric layers that can range from as few as 3 to as many as 20 different layers (Cali et al. 2002; Chioralia et al. 1998; Lom 1972; Sinden and Canning 1974; Vavrá 1976; Weidner 1972, 1976). During germination the polar filament (tube) is discharged from the anterior of the spore and forms a hollow tube that remains attached to the spore and facilitates passage of its sporoplasm and nucleus (or diplokaryon) into its host cell (Frixione et al. 1992 ; Lom and Vávra 1963 ; Ohshima 1937 ; Walters 1958 ; Weidner 1972 ). Electron microscopy has demonstrated elongated sporoplasm in sections of extruded polar tube and the piercing of host cell membranes by the polar tube (Lom 1972; Weidner 1976). This process serves as a unique mechanism of infection, resulting in sporoplasm transfer directly into the host cell cytoplasm (Frixione et al. 1992; Lom and Vávra 1963; Ohshima 1937; Weidner 1972). In A. algerae, polar tube discharge is associated with the appearance of membrane infoldings surrounding the polar tube (Cali et al. 2002). These ultrastructural observations suggest that the polar tube is actually extracytoplasmic in the spore and explains how the sporoplasm can remain intact during the explosive germination reaction.

Polar tubes range from 50 to 100 μm in length and 0.1 to 0.15 μm in diameter (Frixione et al. 1992). A germinated spore is shown in Fig. 5.2. The polar tube discharges from the anterior pole of the spore in an explosive reaction occurring in less than 2 s (Frixione et al. 1992; Lom and Vávra 1963; Ohshima 1937; Weidner 1972). Spore discharge occurs through phases of (1) activation, (2) increase in intrasporal osmotic pressure, (3) eversion of the polar tube, and (4) passage of sporoplasm through the polar tube. The exact mechanism of this process is not well understood. Conditions that lead to spore germination vary widely among species, presumably reflecting the adaptation of each microsporidian to its host and external environment (Undeen and Epsky 1990 ; Wittner and Weiss 1999 ). Since microsporidia are found in a wide range of terrestrial and aquatic hosts, different species may require unique activation conditions for spore discharge. These specific conditions are also probably important to prevent accidental discharge in the environment (Undeen and Avery 1988; Undeen and Epsky 1990). It has been theorized that, regardless of the mode of activation, microsporidia exhibit the same response to stimuli by increasing the intrasporal osmotic pressure (Lom and Vávra 1963; Ohshima 1937; Undeen and Frixione 1990, 1991). The increase in osmotic pressure results in an influx of water into the spore accompanied by swelling of the polaroplast and posterior vacuole prior to spore discharge (Frixione et al. 1992; Lom and Vávra 1963; Weidner and Byrd 1982). This pressure forces the eversion of the polar tube and expulsion of sporoplasm (Undeen 1990). In hyperosmotic solutions, polar tube discharge is inhibited or slowed down, and sporoplasm passage does not occur, thus providing indirect support for the osmotic pressure theory (Frixione et al. 1992; Lom and Vávra 1963; Ohshima 1937; Weidner 1976; Undeen and Frixione 1990).

Fig. 5.2

Germinated spore of Edhazardia aedis from mosquito Aedes aegypti. The emptied spore (left) is shown attached to an extruded polar tube. The sporoplasm, or cytoplasmic contents of the spore, is propelled through the everting polar tube and is shown on the right side of the image. Under appropriate conditions, the sporoplasm is introduced into a new host cell to initiate infection. Note that the extruded polar filament is approximately 20 times the length of the spore

The polar tube has flexibility, varies in diameter from 0.1 to 0.25 μm during discharge, can increase to 0.4 μm in diameter during sporoplasm passage, and shortens in length by 5–10 % after sporoplasm passage (Frixione et al. 1992; Lom and Vávra 1963; Ohshima 1937; Weidner 1972). The hollow discharged tubes appear to be two to three times as long as the dense, coiled tube inside the spore, and it has been suggested that the internal contents of the tube are incorporated at its growing tip during discharge (Frixione et al. 1992; Weidner 1972, 1976, 1982). The evagination of the polar filament has been likened to reversing a finger of a glove (Lom and Vávra 1963; Ohshima 1937; Weidner 1972, 1982; Weidner and Byrd 1982; Weidner et al. 1995). The polar tube is essentially a delivery mechanism for transversing the intestinal lumen to deliver the spore contents into intimate association with the host cell. It is not clear whether the polar tube pierces the host cell or invagination and internalization are driven by an interaction of the sporoplasm at the tip of the polar tube with host cell membranes. Although it is accepted that the sporoplasm flows through the discharged polar tube and into the host cell, the mechanisms of activation and tube formation during discharge remain unclear.

Studies have demonstrated that the polar tube has unusual solubility properties and resists dissociation in 1–3 % sodium dodecyl sulfate (SDS), 1 % Triton X-100, 1–10 % H₂O₂, 5-8 N H₂SO₄, 1-2 N HCl, chloroform, 1 % guanidine HCl, 0.1 M proteinase K, 8–10 M urea, 50 mM NaCO₃, and 50 mM MgCl₂ (Weidner 1972, 1976; Weidner and Byrd 1982). The polar tube, however, dissociates in various concentrations of 2-mercaptoethanol (2-ME) and dithiothreitol (DTT) (Keohane et al. 1994; Weidner 1972, 1976). This has allowed proteomic investigations of the composition of the polar tube (Ghosh et al. 2011). A procedure was developed for the isolation and purification of the major polar tube protein (PTP1) from the spores of microsporidia (Keohane et al. 1994; Keohane and Weiss 1998). Soluble polar tube preparation of Glugea americanus, Enc. hellem, Enc. cuniculi, Enc. intestinalis, and A. algerae were prepared by sequentially extracting glass-bead-disrupted spores with 1 % SDS and 9 M urea, followed by solubilization of the residual polar tubes in 2 % DTT (Keohane et al. 1994, 1999b; Weiss 2001). PTP1 in the DTT-solubilized material was then purified to homogeneity using reverse phase high-performance liquid chromatography (HPLC) (Keohane and Weiss 1998; Keohane et al. 1996a). By SDS-PAGE and silver staining, this purified fraction migrated at 43 kDa for G. americanus, 45 kDa for Enc. cuniculi and Enc. intestinalis, and 55 kDa for Enc. hellem (Keohane et al. 1999a, b). Monoclonal or polyclonal antibodies raised against the purified PTP1 demonstrated reactivity with polar tubes by immunofluorescence (IF) and immunogold electron microscopy (EM) and demonstrated cross reactivity among the species by immunoblotting and immunogold EM (Keohane and Weiss 1998; Keohane et al. 1994, 1996a, b, c).

All of the major polar tube proteins (PTP1) purified to date demonstrate similarities in mass, hydrophobicity, high proline content, and immunologic epitopes. The major polar tube protein, PTP1, from both Enc. cuniculi and Enc. hellem was identified in 1998 (Delbac et al. 1998a; Keohane et al. 1998). It is somewhat surprising that the translated proteins have only limited identity in amino acid sequences (Weiss 2001). Further comparisons, however, strikingly reveal that these proteins are proline-rich and have a similar percentage of cysteine (Weiss 2001 ). PTP1 proteins have central amino acid repeat regions that are predominantly hydrophilic. However, the repeats differ in composition and number. It is possible that this region is not important for the assembly of the polar tube and may function as an immunologic mask. In the process of evolution a similar duplication of internal sequences has been noted in malaria and other protozoan genes, and this mechanism may be operative in the microsporidia PTP gene (Rich and Ayala 2000). Analysis of PTP1 from other isolates of Enc. hellem supports this view, as the number of repeats is variable (Weiss 2001). Post-translational o-linked mannosylation occurs on PTP1, and this modification is probably involved in the ability of PTP1 to interact with the surface of host cells (Xu et al. 2004).

While PTP1 is the major component of the polar tube, other polar-tube-associated proteins (PTPs) are clearly present in the DTT-solubilized polar tube fraction.

For example, several putative PTPs of 23, 27, and 34 kDa have been identified in G. americanus using monoclonal antibodies produced to the DTT-solubilized polar tube (Keohane et al. 1994). Using two-dimensional SDS-PAGE one can also demonstrate other PTPs in DTT-solubilized Enc. hellem polar tube (Weiss 2001). In addition, several polyclonal and monoclonal antibodies have localized to the polar tube by IFA and immunogold EM and recognized proteins of 34, 75, and 170 kDa in G. atherinae and 35, 52/55, and 150 kDa in Enc. cuniculi and 60 and 120 kDa in Enc. intestinalis (Beckers et al. 1996; Delbac et al. 1998b). This resulted in the identification of PTP2, a 35-kDa protein, in Enc. cuniculi (Delbac et al. 2001). Enc. cuniculi PTP2 exists as a single copy per haploid genome and is located on the same chromosome as the Ecptp1 gene, i.e., chromosome VI (Delbac et al. 2001), and has been found as a PTP1–PTP2 gene cluster in several other microsporidia (Delbac et al. 2001). By immunoscreening of a cDNA library of Enc. cuniculi, another polar tube protein, PTP3, was found (Peuvel et al. 2002). This protein, predicted to be synthesized as a 1,256-amino-acid precursor (136 kDa) with a cleavable signal peptide, is encoded by a single transcription unit (3,990 bp) located on chromosome XI of Enc. cuniculi (Peuvel et al. 2002). PTP3 is solubilized in the presence of SDS alone (Peuvel et al. 2002). Considering that PTP3 is extractable from Enc. cuniculi spores in the absence of thiol-reducing agent, lacks cysteine, but is rich in charged residues, it has been suggested that PTP3 interacts with PTP1 or PTP2 via ionic bonds and may play a role in the control of the conformational state of PTP1–PTP2 polymers (Peuvel et al. 2002). For example, when the polar tube exists as a coiled structure inside a spore, interactions with PTP3 may permit the maintenance of PTP1–PTP2 polymers in a condensed form (Peuvel et al. 2002).

It was found that DSP, a chemical cross linker that creates disulfide linkages between proteins, could mediate the purification of a large multimolecular complex from polar tubes that contained PTP1, PTP2, and PTP3 (Peuvel et al. 2002). Studies using yeast two hybrid vectors have confirmed the interaction of PTP1, PTP2, and PTP3 and determined that both the N-terminal and C-terminal regions of PTP1 are involved in these interactions, but that the central repeat region of PTP1 is not involved in these protein–protein interactions (Bouzahzah et al. 2010). It is likely that the regular multilayered organization of the microsporidian polar tube is dependent on specific interactions between its protein components.

V. Life Cycle

Microsporidia generally undergo three phases of development (Cali and Takvorian 1999). The infective phase occurs following the release of spores into the environment or tissues where, under suitable conditions, spores germinate and inject their spore contents through the everting polar filament to infect the host cell. Organisms then continue through a proliferative phase within the host cells (often referred to as merogony ), which is followed by sporogony, during which organisms commit to maturation and spore formation. The modes of transmission, intracellular sites of development, number of proliferation cycles, and maturation vary widely among the species of microsporidia. Selected examples of the more common species infecting arthropod, aquatic, mammalian, and avian hosts are described in what follows.

A. Species Infecting Arthropod Hosts

The complete life cycles of many microsporidia in insects are well documented and have shown great diversity from the very simple to the complex, with some involving an intermediate host (Becnel et al. 2005 ). Microsporidia with simple life cycles are generally characterized as having a single sporulation sequence (sometimes with a second sporulation sequence) that occurs in a single host or host group. Vavraia culicis in mosquitoes is an example of a species that has only uninucleate stages throughout the life cycle and produces only one spore type (Vavrá and Becnel 2007). Nosema apis in honey bees has only binucleate (diplokaryotic ) stages throughout the life cycle but is a bit more complex with the production of a primary (early) binucleate spore in the midgut epithelium that serves to spread the infection (autoinfection) to other midgut cells (de Graaf et al. 1994). These infections lead to the production of a second thick-walled (environmental) spore that can be released into the environment to infect a new host (Fries 1993).

Some species are characterized by complex life cycles involving multiple spore types responsible for horizontal and vertical transmission. They often affect two generations of the definitive host and some involve an obligate intermediate host. These microsporidia (often termed polymorphic or heterosporous) are generally very host-specific with complex developmental sequences that can be characterized by specialized stages and high levels of tissue specificity, as well. Amblyospora californica (Kellen and Lipa 1960) parasitizes the mosquito Culex tarsalis and is representative of a species with a complex life cycle that involves an intermediate host (Becnel 1992). Binucleate spores are formed in oenocytes of adult female C. tarsalis following a blood meal. These oenocytes invade the ovaries, where germinations occur, to infect the developing eggs. Developmental sequences in progeny are sex-dependent where females carry benign infection throughout larval development, which leads to the formation of binucleate spores in adults capable of initiating another round of transovarial transmission. Male progeny from infected adults undergo a distinctively different development where the pathogen invades fat bodies with rapid vegetative reproduction that terminates with meiosis and the production of meiospores. These male larvae die, releasing massive numbers of spores into the larval habitat. Meiospores are not infectious to mosquito larvae, but they are horizontally transmitted when ingested by females of the copepod intermediate host Macrocyclops albidus. Uninucleate stages replicate in ovaries of the female copepods, which terminates with the production of uninucleate spores and the death of the host. These spores are infectious when ingested by C. tarsalis mosquito larvae where the uninucleate stages invade larval oenocytes and remain dormant until pupation and adult emergence. In female adult mosquitoes, binucleate spores that are responsible for transovarial transmission are produced to complete the life cycle. To date, it has been determined that the involvement of an intermediate host in the life cycles of microsporidia in insects is restricted to mosquitoes and copepod intermediate hosts and has been documented in a number of species and genera such as Amblyospora dyxenoides (Sweeney et al. 1985), Amblyospora connecticus (Andreadis 1988), Parathelohania anophelis (Avery and Undeen 1990), Culicospora magna (Becnel et al. 1987), Hyalinocysta chapmani (Andreadis 2002), and Edhazardia aedis (Becnel et al. 1989).

B. Species Infecting Aquatic Hosts

Several hundred described species of microsporidia infect fish , but the life cycles of only a few have been described. Many of the fish microsporidia that have been investigated to date can be transmitted directly by ingestion of free spores or spores from infected tissues (Baxa-Antonio et al. 1992; Kent and Bishop-Stewart 2003; Kent and Speare 2005; McVicar 1975; Sanders et al. 2010; Weissenberg 1968). Autoinfection has been suggested or demonstrated for some Loma species, in which spores within tissues establish new infections (Matos et al. 2003; Rodriguez-Tovar et al. 2003; Shaw et al. 1998). The potential for maternal transmission, either transovum or transovarial, has been reported for L. salmonae (Sanchez et al. 2001) and P. neurophilia (Kent and Bishop-Stewart 2003). Phelps and Goodwin (2008) provided the most conclusive evidence for vertical transmission, showing the presence of the Ovipleistophora ovariae DNA by polymerase chain reaction from within the eggs of infected golden shiners, and similar results were obtained with P. neurophilia (Sanders and Kent 2011).

The sequential development from early infection to the site of sporulation is poorly understood for almost all microsporidia of fish. The development of L. salmonae has been elucidated and serves as an example for the development of microsporidia in fish (Kent and Speare 2005). The initial site of infection by L. salmonae is the mucosal epithelium of the stomach and intestine. Within 4 h of exposure, spores are seen in close association with the stomach epithelium and parasite DNA is present in the cytoplasm of epithelial cells and lamina propria of the small intestine by 12 h post exposure (Sprague and Hussey 1980). Two days post exposure, infected cells with dividing stages of L. salmonae can be visualized in the endocardium of the heart by in situ hybridization. After 2–3 weeks, uninucleate or binucleate merogonial stages can be seen developing within the endothelial cells or pillar cells of the blood vessels in the gills, the primary site of sporulation. Meronts are located at the periphery of the host cytoplasm, and the parasite (meront) cell membrane is in close proximity with the surrounding host cell membrane.

The sporogonic stages of L. salmonae occur in hypertrophic host cells of gills to generate a xenoparasitic complex or xenoma . Xenomas are host cells with a radically altered structure in which the microsporidia have integrated into the host cell cytoplasm to undergo massive proliferation while isolated from the body’s defense mechanisms (Lom and Dyková 2005 ). Other xenoma-forming species, such as members of the genus Glugea, can produce very large xenomas (up to 3 mm) in many organs, especially in the subepithelium of the intestine, resulting in grossly visible tumorlike structures that are derived from a single hypertrophic host cell. Another type of xenoma develops with infection by Ichthyosporidium giganteum, which forms a large syncytium from the coalescence of several host cells, resulting in a large, lobular cyst (Rodriguez-Tovar et al. 2003). In contrast to xenoma-forming species, skeletal-muscle-infecting microsporidia of the genus Pleistophora develop within host cells, replacing the sarcoplasm and destroying infected cells without inducing the hypertrophy characteristic of xenomas.

Early during the first 2–3 weeks of xenoma formation, meronts of L. salmonae occupy the periphery of the host cell, and by 5 weeks this area begins to be occupied by mature spores (Lom and Dyková 2005). Eventually, spores can be seen throughout the xenoma that can reach a diameter of up to 0.4 mm. With other genera, sporogony occurs asynchronously throughout the xenoma (Morrison and Sprague 1983). A parasite-derived sporophorous vesicle forms prior to sporogonial division, and sporogony proceeds by binary fission, resulting in two uninucleate sporoblasts per vacuole (Chilmonczyk et al. 1991). The formation of this vacuole is absent in the genus Spraguea, which instead develops in direct contact with the host cytoplasm. The intranuclear microsporidia N. salmonis (Rodriguez-Tovar et al. 2003) and P. theridion (Nylund et al. 2010) develop in direct contact with the host cell nucleoplasm. Eventually, sporoblasts develop into mature spores. The intact xenoma is surrounded by numerous inflammatory cells but seems to elicit little, if any, response by those cells (Rodriguez-Tovar et al. 2003 ). Eventual rupture of the xenoma results in the release of mature spores, elicitation of a severe proliferative inflammatory reaction, and uptake of spores by infiltrating phagocytes.

C. Species Infecting Mammalian and Avian Hosts

The most common modes of transmission of Encephalitozoon species in mammals, and presumably birds, are by ingestion and inhalation of spores shed from urine, feces, or other fluids. Transmission may occur through direct contact (e.g., trauma), and vector-borne, sexual, and horizontal routes have also been reported to occur in mammals. The life cycle of Encephalitozoon is relatively simple in comparison to that of other microsporidia. After germination and introduction of the spore cytoplasmic contents into the host cell, Encephalitozoon species undergo multiple cycles of binary division within a PV, the membrane of which seems to be host-cell-derived (Rönnebäumer et al. 2008). The proliferative stages or meronts tend to be larger than the mature spore and appear to adhere to the inner PV membrane. In some cases, karyokinesis occurs slightly faster than cytokinesis such that ribbons of dividing multinucleated meronts can be observed. Sporogony or spore maturation occurs as the parasite plasma membrane thickens and differentiates to form the exospore and endospore layers. These stages separate from the PV membrane and may continue to undergo a limited number of cell divisions. During this phase, the polar filament develops and the organisms become smaller and more electron-dense. Eventually, the PV becomes full of organisms leading to host cell and PV rupture and release of organisms. Among the sites of infection in mammals and birds are kidney, small intestine, and liver, so spores are commonly shed with urine and feces.

Ent. bieneusi infections typically occur in cells lining the small intestine in which organisms replicate in direct contact with the host cell cytoplasm (Cali and Owen 1990). Merogony is characterized by nuclear division without cytokinesis to generate a multinucleated plasmodium. During sporogony, electron-dense disks are observed to stack and eventually fuse to form the polar filaments in association with each nucleus. The individual nuclei become more defined, and the plasmalemma of the plasmodium begins to thicken and invaginate to surround the individual nucleus and polar filament units. Maturation continues with the thickening and differentiation of the spore wall, release of spores into the intestinal lumen, and shedding with feces.

VI. Systematics and Evolution

Microsporidia possess prokaryote-sized 70S ribosomes and lack typical mitochondria and Golgi. Early molecular biology studies also demonstrated fusion of the 5.8S and large subunit rRNAs similar to that in prokaryotes. These observations and the initial phylogenetic analyses of microsporidian small subunit rRNA genes supported divergence of the microsporidia prior to the symbiotic origin of mitochondria and placed microsporidia at the earliest and deepest branch of the eukaryotic tree (Vossbrinck et al. 1987). Evidence began to mount, however, that microsporidia are more highly evolved. Nuclear-encoded genes that target mitochondrial proteins (e.g., mHSP70, alpha and beta subunits of pyruvate dehyrdrogenase E1) were discovered and antibodies to mHSP70 identified membrane-bound organelles called mitosomes that function as mitochondrial remnants for iron–sulfur cluster assembly (Fast and Keeling 2001; Germot et al. 1997; Hirt et al. 1997; Williams and Keeling 2005; Williams et al. 2002). Genome-sequence studies of several microsporidia species (e.g., Enc. cuniculi, Enc. intestinalis, Ent. bieneusi, P. locustae, A. algerae) and improved phylogenetic analyses on additional genes have shed further light in demonstrating a close relationship between the microsporidia and the fungi (Akiyoshi et al. 2009; Burri et al. 2006; Cornman et al. 2009; Corradi and Slamovits 2011; Corradi et al. 2007; Katinka et al. 2001; Keeling et al. 2010; Williams et al. 2008). Microsporidia are highly efficient parasites and have undergone significant gene reduction and compaction (Keeling 2009; Keeling et al. 2010). In addition, microsporidia seem to have evolved relatively quickly and exhibit a high degree of gene sequence divergence, so phylogenetic analyses to address their evolution are problematic. However, a strong conservation of gene order, or synteny , among several gene clusters, including the sex locus, was reported among distantly related microsporidia and the zygomycetes (zygomycete mating type, MAT) (Dyer 2008; Lee et al. 2008, 2010b; Corradi and Keeling 2009; Corradi and Slamovits 2011). These findings were used to support a deep-branching fungal origin of the microsporidia from a zygomycete ancestor and suggest that microsporidia may have a genetically controlled sexual cycle. Concerns have been raised, however, about whether this shared syntenic relationship truly supports a microsporidial–fungal relationship because the gene cluster of the microsporidia that resemble the zygomycete sex-related loci traces back to an ancient gene cluster in the common ancestor of plants, animals, and fungi (Koestler and Ebersberger 2011).

VII. Classification

Newer molecular biology analyses and approaches being applied to better understanding evolution of the microsporidia also have impacted their taxonomy and classification. The microsporidia are now fairly well accepted for classification with Kingdom Fungi (Corradi and Keeling 2009; Hibbett et al. 2007; James et al. 2006), but some analyses question this association (Koestler and Ebersberger 2011), suggesting that further studies are required. Classification of fungi is based on the International Code of Botanical Nomenclature, but microsporidia had been described using the International Code of Zoological Nomenclature. To avoid nullification, a formal request to accept the current nomenclature of the microsporidia was presented at the last taxonomy meeting at the International Botanical Congress and was approved (Redhead et al. 2009). As a result, microsporidia are considered fungi but remain subject to the rules of the International Code for Zoological Nomenclature. A broad-based consensus classification (Hibbett et al. 2007) did not subdivide the microsporidia within the fungi due to a lack of well-sampled multilocus analyses at that time. More recently, gene order (i.e., synteny) between several unrelated microsporidia and the zygomycetes was highly conserved (Corradi and Keeling 2009; Dyer 2008; Lee et al. 2008), but again, others suggest that synteny was not more similar between microsporidia and the zygomycetes than with any other fungal taxon (Koestler and Ebersberger 2011). The phylum name Zygomycota is considered invalid because the interrelationships among the major clades are still unresolved, and it was named without a Latin description, so further classification of the basal fungi to relate microsporidia to the zygomycetes or another fungal group is still in progress (Hibbett et al. 2007). Based on the complexity of microsporidian evolution, it is also possible that the microsporidia might represent a sister group to the fungi. As newer analytical tools incorporate additional genomic and proteomic information, a better picture will emerge regarding the classification of the microsporidia.

Primary classification of organisms into the phylum Microsporidia was based on the presence of the polar tube. Further classification was based on morphological and ultrastructural features, as well as host and habitat (Larsson 1986 ; Sprague et al. 1992 ). More specific characteristics used to classify the microsporidia include host cell, spore size, nucleus configuration (i.e., monokaryon, diplokaryon), number and configuration of the polar filament coils, type of nuclear and cellular division (e.g., binary division, plasmotomy), interface with the host cell (e.g., replication within a PV, direct contact with host cell cytoplasm), and whether a sporophorous vesicle is formed. Microsporidia initially fell into two groups based on the presence or absence of a sporoblast vesicle (Pansporoblastina and Apansporoblastina, respectively) and then were divided into groups based on nuclear configuration as single (Haplophasea) or double (Dihaplophasea) nuclear arrangement, the latter being grouped on the basis of diplokaryon formation through meiosis or nuclear dissociation.

With the advancement of molecular biology technology, classifications within the phylum incorporated phylogenetic analyses (Vossbrinck and Debrunner-Vossbrinck 2005), and genera accepted to date are found in Sect. II.A. A comparative molecular phylogenetic analysis using ssrDNA sequences of 125 species in relation to host and habitat led to a proposal for grouping microsporidia into five clades among three new classes: the Aquasporidia (clades I, II, and V), the Marinosporidia (clade III), and the Terresporidia (clade IV). This new classification, however, is considered to be under development due to the relatively small representation for analyzing only 125 of over 1,200 species of microsporidia, the as yet undescribed microsporidia that are likely to be found, and a need to account for features related to morphology, life cycle, and host–parasite relationship (Larsson 2005; Vossbrinck and Debrunner-Vossbrinck 2005).

VIII. Maintenance and Culture

A. Species Infecting Arthropod Hosts

Brooks (1988) presents an excellent review of spore storage and maintenance of microsporidia infecting arthropods, but the optimal storage conditions must be determined experimentally for each isolate. There are no standard guidelines on the best practices to preserve spore viability. There is general agreement, however, that many microsporidian spores from terrestrial hosts will tolerate freezing or desiccation, whereas spores from aquatic hosts do not but in some cases can be maintained long term under other conditions. Spores to be stored are most commonly handled as intact infected cadavers or as purified suspensions.

Many species of microsporidia can be maintained for extended periods (months to years) as highly purified spores held in deionized water at 5 °C (±3). Antibiotics and fungicides are routinely added to the suspensions to retard microbial growth, which can reduce spore viability. Highly purified spores of A. algerae cannot survive freezing but have maintained viability after being held at 5 °C (±3) for more than 10 years. Many terrestrial species of microsporidia can be frozen (–30 to –20 °C) as cadavers, or purified spores can be placed into liquid nitrogen for long-term storage. The addition of 50 % glycerol to the pure spore suspensions as a cryoprotectant is often beneficial. Some spores can also be stored in the dried host cadaver for extended periods, such as Nosema whitei, a pathogen of flour beetles. Spores that can be dried can often be successfully lyophilized. If information on storage parameters for a species is not available, it is suggested that highly purified spores be held in deionized water at 5 °C (±3).

In vitro culture of microsporidia in insects has a long history and began with the successful infection of a B. mori cell line with N. bombycis Trager 1937, but few additional species have been established in cell culture. Until the mid to late 1980s, only about eight species of microsporidia from insects had been cultured in insect cell systems that included A. algerae, N. apis, N. bombycis, N. disstriae, N. heliothidis, N. mesnili, Vairimorpha necatrix, and Vavraia culicis (Brooks 1988; Jaronski 1984). More recently, a few additional species have been cultured, including Cystosporogenes operophterae, N. furnacalis, and a Vairimorpha sp. (Becnel and Andreadis 1999). The species with the broadest host range and ability to grow in both invertebrate and vertebrate cell lines is A. algerae. It has been grown in many insect cell lines (Brooks 1988) and in pig kidney cells (Undeen 1975), rabbit kidney cells (Lowman et al. 2000), several warm-water fish cell lines (Monaghan et al. 2011), and human muscle fibroblasts (Trammer et al. 1999). A. algerae has also been grown at elevated temperatures (37 °C), which is unique for insect microsporidia (Lowman et al. 2000).

B. Species Infecting Aquatic Hosts

Few microsporidia that infect aquatic organisms have been successfully propagated in long-term cell culture, and these generally depend upon maintaining groups of infected hosts or obtaining infected hosts from the wild. The presence of microsporidia in nonmammalian model organisms, such as the zebrafish and the nematode Caenorhabditis elegans , provides researchers the opportunity to study these parasites in well-described systems with numerous genetic tools (Troemel 2011 ) to elucidate host responses to microsporidia infections. The in vitro propagation of microsporidia infecting fish has proven difficult, and the use of fish cell cultures in the long-term maintenance of fish microsporidia was recently reviewed (Monaghan et al. 2009).

For example, the intranuclear microsporidium N. salmonis has been successfully maintained in a long-term primary culture of salmonid mononuclear leukocytes grown in supplemented Iscove’s modified Dulbecco’s medium by adding small numbers of infected leukocytes to uninfected leukocytes (Wongtavatchai et al. 1994). Infected cultures can be preserved long term by freezing in liquid nitrogen with cryoprotectant.

A continuous cell line, EP-1, derived from the Japanese eel, Anguilla japonica, is persistently infected with Heterosporis anguillarum. Whereas this cell line was passaged over 223 times in vitro for maintaining intracellular merogonic stages of the parasite, no spore stages were observed to develop, yet eels inoculated with cells from this culture system became infected and exhibited intramuscular cysts consistent with H. anguillarum infection (Kou et al. 1995). To date, this remains the only cell line developed to be persistently infected with a microsporidian parasite of fish.

Four fish cell lines—channel catfish ovary, zebrafish caudal fin fibroblast, carp epithelioma, and fathead minnow—have been shown to support limited growth of the microsporidian parasite of zebrafish, P. neurophilia. Whereas sporogony occurs in all cell lines, development to the spore stages is limited, and the parasites could not be passaged into new cultures (Watral et al. 2006). Similarly, spores of Glugea spp. were internalized by Chinook salmon embryo cells and, while meronts were detected, development ceased by 48 h and no sporogony was observed (Lores et al. 2003). The same parasite did develop in a mosquito cell line (ECACC 90100401), producing spores within 72 h post inoculation, illustrating the potential for insect cell lines in the propagation of fish microsporidia in vitro.

C. Species Infecting Mammalian and Avian Hosts

Enc. cuniculi was the first mammalian microsporidian to be isolated from a rabbit and grown in long-term tissue culture (Shadduck 1969). Since then, Enc. hellem, Enc. intestinalis, A. algerae, V. corneae, and Trachipleistophora hominis isolates from humans have been grown in culture, but unfortunately long-term culture of Ent. bieneusi still has not been accomplished (Braunfuchsová et al. 1999; Didier et al. 1991, 1996; Juarez et al. 2005; Lafranchi-Tristem et al. 2001; Monaghan et al. 2009; Trammer et al. 1999; Visvesvara 2002).

Cultures are typically initiated via coculture of source specimen (tissue biopsy or fluids such as urine, feces, or sputum) and host cells such as Vero, RK-13, MDCK, and other epithelial cells. Examples of tissue culture media that support the growth of the host cells and facilitate propagation of the microsporidia include RPMI 1604 or D-MEM supplemented with 2 mM l-glutamine, 5–10 % fetal bovine serum, and antibiotics (e.g., penicillin, streptomycin, and amphotericin B). The medium is typically changed twice a week and the supernatants can be collected in sterile bottles for short-term storage at 4 °C.

Encephalitozoon-infected cells appear to contain vacuoles filled with organisms. V. corneae replicates in the cytoplasm of the host cell, and infected cells may appear larger and multinucleated when filled with organisms. Individual microsporidia suspended in the supernatants after release from ruptured host cells can be observed approximately 2–4 weeks after initiation of coculture, but sometimes longer periods of time are required if the initial inoculum dose of organisms is low. In the case of V. corneae, large aggregates of parasite-laden host cells are often also observed in the culture supernatants and can be separated by vortexing or washing the collected culture supernatants. Host cells tend to replicate and replace the ruptured infected cells, but if overgrowth of microsporidia occurs, fresh host cells can be added to the culture flasks.

To enrich microsporidia from host cell debris, the sedimented culture supernatants can be washed sequentially in distilled water, tris-buffered saline (TBS)\ containing 0.3 % Tween 20 (TBS-TW), and TBS (400× g for 15 min). The pellets can be further enriched by centrifugation through 50 % Percoll (i.e., mixing equal volumes of spores in TBS and 100 % Percoll) at 400 × g for 30–45 min. Extraneous host cell debris remains in the top layers, and the spores centrifuge to the pellet (Didier et al. 1996). Microsporidia to be used for extracting DNA or RNA require additional washing with mild ionic detergent (e.g., 0.5–1 % sodium dodecyl sulfate) to remove host cell DNA that can adhere to the spore surface (Corradi et al. 2010).

Cryopreservation of mammalian microsporidia can be accomplished most efficiently by “scraping” or trypsinizing infected host cells, centrifuging at 400 × g (15 min at 4 °C), and resuspending in fetal bovine serum (FBS) containing 10 % dimethyl sulfoxide (DMSO). The vials are then frozen slowly (1 °C per minute) using commercially available cryopreservation containers, followed by final storage of vials in liquid nitrogen. To reestablish culture from cryopreserved spores, flasks of host cells at approximately 50 % confluence should be prepared. The frozen vial of microsporidia should be thawed quickly and added directly to the host cells; a few hours later, after the microsporidia have had an opportunity to infect the host cells, the culture medium should be changed to remove the DMSO. Alternatively, the inoculum of cryopreserved spores can be washed (i.e., centrifuged) and the pellet resuspended in a small volume of medium to remove the DMSO prior to inoculation of the culture flasks. If host cells other than those used to generate the cyropreserved spores are used, the spore inoculum will need to be washed with mild detergent (e.g., 0.5 % SDS) to prevent growth of cryopreserved host cells in the new culture.

IX. Conclusions

The microsporidia comprise a fascinating group of organisms that infect their hosts through an unusual spore germination process of polar filament extrusion and direct inoculation of the sporoplasm into the cell. They are an extremely successful group of organisms that are widespread in both vertebrate and invertebrate hosts and are highly efficient parasites, as noted by their gene compaction and reduction. Over the past 10 years, molecular studies have reshaped our understanding of phylogeny and led to the classification of microsporidia as fungi, although knowledge of the exact relationship is still in flux (Hibbett et al. 2007; Koestler and Ebersberger 2011). Whereas previously the microsporidia had been recognized pathogens of agriculturally and commercially relevant insects, fish, companion pets, domestic animals, and food-producing animals, it was only recently, during the AIDS pandemic, that these organisms came to be seen as common causes of opportunistic and emerging infections in humans. The tremendous increase in the recognition of new species of microsporidia in such a wide host range and the application of newer molecular tools will now need to be applied to improving diagnostics, developing intervention and chemotherapeutic strategies, and learning more about the basic biology and phylogeny of the microsporidia.