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Mammalian Microsporidiosis

Office of Laboratory Animal Resources, University of Illinois, Urbana, IL

	Abstract

Top Abstract Mammalian Microspora: Life... Diagnosis of Mammalian... Natural and Experimental... Microsporidiosis as an Emerging... Animal Models of Human... Comments References

 
The phylum Microspora contains a diverse group of single-celled, obligate intracellular protozoa sharing a unique organelle, the polar filament, and parasitizing a wide variety of invertebrate and vertebrate animals, including insects, fish, birds, and mammals. Encephalitozoon cuniculi is the classic microsporidial parasite of mammals, and encephalitozoonosis in rabbits and rodents has been and continues to be recognized as a confounding variable in animal-based biomedical research. Although contemporary research colonies are screened for infection with this parasite, E. cuniculi remains a cause of morbidity and mortality in pet and conventionally raised rabbits. In addition, E. cuniculi is a potential pathogen of immature domestic dogs and farm-raised foxes. The recent discovery and identification of Encephalitozoon intestinalis, Encephalitozoon hellem, and Enterocytozoon bieneusi, in addition to E. cuniculi, as opportunistic pathogens of humans have renewed interest in the Microspora. Veterinary pathologists, trained in the comparative anatomy of multiple animal species and infectious disease processes, are in a unique position to contribute to the diagnosis and knowledge of the pathogenesis of these parasitic diseases. This review article covers the life cycle, ultrastructure, and biology of mammalian microsporaidia and the clinical disease and lesions seen in laboratory and domestic animals, particularly as they relate to Encephalitozoon species. Human microsporidial disease and animal models of human infection are also addressed. Often thought of as rabbit pathogens of historical importance, E. cuniculi and the related mammalian microsporidia are emerging as significant opportunistic pathogens of immunocompromised individuals.

Key words: Animal model; Encephalitozoon cuniculi; Encephalitozoon hellem; Encephalitozoon intestinalis; Enterocytozoon bieneusi; HIV; Microspora; opportunistic pathogens; review.

Encephalitozoon cuniculi has had a colorful and confounding history in animal-based biomedical research, even after its initial identification as the etiologic agent of "infectious motor paralysis" in rabbits by Wright and Craighead in 1922.¹⁵² The use of rodents or rabbits endemically infected with E. cuniculi with for xenodiagnoses and experimental models of human disease resulted in the clinical manifestations and histologic lesions of encephalitozoonosis being mistaken for poliomyelitis and human syphilis.¹⁷ Endemic encephalitozoonosis in research animals has also been mistaken for animal models of herpes simplex and "human epidemic encephalitis," B-virus infection (cercopithecine herpesvirus 1), toxoplasmosis, "human viral hepatitis," and multiple sclerosis.^{52,59,64,65,92,113,140} Although less well documented, E. cuniculi has also been mistaken for or implicated as the agent of rabies, scrub typhus, psittacosis, murine leukemia, experimental allergic encephalomyelitis, and chemical carcinogenesis.¹²⁴ In addition, E. cuniculi has been transmitted through rodent tumor lines, altering the biologic behavior of certain experimental murine tumors.^5,104

E. cuniculi is a member of the phylum Microspora, which includes single-celled, spore-forming, obligate intracellular parasites with a wide host distribution.²¹ Microsporidia are traditionally considered protozoal organisms, although there is some genetic and molecular evidence to suggest a closer phylogenetic relationship to fungi, including the possession of a mitochondrial heat shock protein more closely related to that of fungi; - and ß-tubulins with compositions more closely resembling those of fungal tubulins; and the presence of chitin and trehalose, which are also components of fungi.⁶⁹ E. cuniculi is the most extensively studied mammalian microsporidium, and spontaneous infections have been documented in rabbits, mice, rats, muskrats, guinea pigs, hamsters, ground shrews, goats, sheep, pigs, horses, domestic dogs, wild and captive foxes, domestic cats, a variety of exotic carnivores, and nonhuman primates.¹²⁴

Interest in mammalian microsporidiosis was renewed with the identification of Enterocytozoon bieneusi as the cause of chronic diarrhea and wasting in an immunocompromised person that was positive for the human immunodeficiency virus (HIV).³⁵ Identification and molecular differentiation of Encephalitozoon intestinalis (originally named Septata intestinalis), Encephalitozoon hellem, and E. cuniculi in this immunocompromised population followed.^18,38,51Vittaforma corneum, Nosema spp., Pleistophora spp., and Trachipleistophora spp. have also been diagnosed in humans, although less frequently.^{24,61,128,138} While Enterocytozoon bieneusi remains the most frequently diagnosed microsporidial pathogen of humans, it has yet to be cultivated in vitro and, unlike the Encephalitozoon spp., lacks convenient rodent models of infection.^97,134 In addition, increasing reports of encephalitozoonosis in pet animals suggests that Encephalitozoon spp. may be zoonotic.^33,43,58,85 For these reasons, encephalitozoonosis will continue to be of interest to research-oriented veterinary pathologists and anatomic veterinary pathologists. This review article covers the life cycle of mammalian microsporidia and the basic ultrastructural and biological features shared by all members of the Microspora, and will discuss the clinical disease and lesions seen in spontaneously and experimentally infected laboratory and domestic animals, particularly as they relate to Encephalitozoon spp. Lastly, human microsporidial disease and animal models of human infection will be addressed.

	Mammalian Microspora: Life Cycle, Ultrastructure, and Cellular Biology

Top Abstract Mammalian Microspora: Life... Diagnosis of Mammalian... Natural and Experimental... Microsporidiosis as an Emerging... Animal Models of Human... Comments References

 
All microsporidia have an obligate intracellular life cycle, existing as environmentally resistant spores outside the host.²¹ Their life cycle in mammals is postulated to be simple and direct, and most infections are acquired through ingestion or inhalation, transplacentally, or rarely, through trauma to epithelium (Fig. 1). ⁴¹ The defining criterion for inclusion in the phylum Microspora is possession of a unique organelle known as the polar filament (also referred to as a polar tube) found coiled within the spore.²¹ Other ultrastructural features common to microsporidia include a corrugated, proteinaceous exospore (spore wall), a chitinous, radiolucent endospore, an anchoring disc at the anterior pole, and an electron-lucent posterior vacuole (Fig. 2). The spore nucleus can be difficult to discern from the remainder of the electron-dense sporoplasm. Stimulation by an uncharacterized environmental signal causes extrusion of the polar filament from the spore and "injection" of the infectious sporoplasm into the host cell.¹⁴⁸ Although polar tube extrusion is postulated to be the primary mechanism by which all microsporidia establish intracellular infection, in vitro cultivation of Encephalitozoon spp. suggests that phagocytosis of spores by host cells may also establish patent intracellular infection.^21,100 Once inside the host cell, the deposited sporoplasm divides to yield proliferative forms (also known as meronts) with simple plasma membranes, small prokaryotic-like ribosomes, and sparse organelles. As the proliferative forms differentiate into sporoblasts and then sporonts, a dense spore wall is laid down over the plasma membrane, the plasma membrane uniformly thickens, spore organelles (posterior vacuole, coiled polar tube, anchoring disk) are formed, and the cytoplasm becomes increasingly electron dense. Depending on the genera of mammalian microsporidia, the developing spores either become packaged within a parasitophorous vacuole (i.e., Encephalitozoon spp.) or remain in direct contact with the cytoplasm (i.e., Enterocytozoon bieneusi, Nosema spp.). There is a distinct lack of mitochondria, peroxisomes, and classical stacked Golgi membranes in all stages of parasite development. A progression of developing parasite forms is typically found in each infected cell (Fig. 3), and the appearance of an electron-lucent endospore, a clear posterior vacuole, and cross-sections of the coiled polar filament characterize mature spores. Enlargement of the parasitophorous vacuoles and/or host cell with parasites results in host cell rupture and release of spores into extracellular spaces. Infection is disseminated by direct extension of spores into surrounding cells, by introduction into the vascular system, or by both methods.⁴¹ Parasites are difficult to identify in tissue sections by light microscopy with hematoxylin and eosin stains but may appear as clusters of refractile intracytoplasmic structures. Parasites are gram positive and rod shaped and may appear to have a clear vacuole at one pole. Although not appreciable at the light microscopic level, the presence or absence of the parasitophorous vacuole and the polar filament cross-sectional number and arrangement, in addition to spore size, aid in differentiating parasite genera at the electron microscopic level (Table 1).¹⁸

View larger version (29K): [in this window] [in a new window]   Fig. 1. Ultrastructural features common to most mammalian microsporidia, and simplified life cycle of Encephalitozoon spp.

View larger version (137K): [in this window] [in a new window]   Fig. 2. Fetal intestine; rabbit. Tissue was implanted into athymic nude mouse and experimentally inoculated with E. hellem.143 Ultrastructural features of the spore include electron-dense exospore (EX), electron-lucent endospore (EN), a faint spore nucleus (N), and four cross-sections of the polar tubule (arrows) typical of E. hellem. Bar = 0.1 µm. Fig. 3. Fetal intestine; rabbit. Tissue was implanted into athymic nude mouse and experimentally inoculated with E. hellem.143 Asynchronously developing parasite forms are present within a parasitophorous vacuole: proliferative forms (P), sporont (ST), sporoblast (SB), mature spore (S). Note host cell mitochondrion (m). Bar = 0.6 µm.

Molecular analysis of microsporidia has also revealed several interesting characteristics. The presence of genes in E. cuniculi coding for a mitochondrially expressed heat shock protein suggests that microsporidia may have evolved from a mitochondria-containing ancestor that subsequently lost this organelle through evolutionary or selective pressure.¹⁰⁵ The significance and function of this heat shock protein is not known. The nuclear genome of E. cuniculi is quite small, 2.9 x 10⁶ nucleotide base pairs (as compared with 4.7 x 10⁶ base pairs in Escherchia coli and 6.0 x 10⁹ base pairs in human somatic cells).¹¹ In addition, intervening sequences (or introns) in microsporidial DNA are relatively rare.¹⁴⁹ Thus, microsporidia appear to encode information for infection, replication, and dissemination in an extremely efficient manner, suggesting that these parasites are highly adapted to survival in their hosts.

	Diagnosis of Mammalian Microsporidiosis

Top Abstract Mammalian Microspora: Life... Diagnosis of Mammalian... Natural and Experimental... Microsporidiosis as an Emerging... Animal Models of Human... Comments References

 
The approach for diagnosing microsporidiosis in mammals will depend on the origin, use, and immune status of the animals in question. Laboratory colonies of immunocompetent rabbits and rodents are routinely screened for infection with E. cuniculi by serology; histopathology is used with selected animals to confirm positive enzyme-linked immunosorbent assays or indirect fluorescent antibody tests. Dogs suspected of being infected with E. cuniculi may also be tested initially by serology.¹²³ Immunodeficient animals do not mount reliable antibody responses, and microsporidial infections in these animals must be diagnosed by other methods. For animals displaying clinical microsporidial disease (e.g., seizures), examination of urine, feces, or cerebrospinal fluid may yield an antemortem diagnosis of encephalitozoonosis. These clinical samples are often screened with a calcofluor white stain for chitin, and Encephalitozoon infection is confirmed with modified trichrome stain, immunofluorescent antibodies, polymerase chain reaction (PCR), transmission electron microscopy, or some combination of methods.^{23,39,81,93,145} Light microscopic examination of tissue samples and the use of special stains will aid the pathologist in detecting infection and differentiating microsporidial infection from other common protozoal infections of animals (Table 2). However, even with the use of appropriate stains light microscopy is not sufficient for species-specific identification of microsporidia, particularly in humans, who may be infected with a number of different microsporidia. PCR tests have been developed and validated for Encephalitozoon spp. and Enterocytozoon bieneusi infections in humans and animals, and although these tests are not yet routinely available to clinical veterinary diagnostic laboratories, they are specific and sensitive enough for diagnosis and identification of microsporidial infections.^14,68,83

	Natural and Experimental Microsporidiosis in Laboratory and Domestic Animals

Top Abstract Mammalian Microspora: Life... Diagnosis of Mammalian... Natural and Experimental... Microsporidiosis as an Emerging... Animal Models of Human... Comments References

E. cuniculi was first identified in the brains and kidneys of rabbits exhibiting lethargy, tremors, and local or generalized paresis.¹⁵² The organisms were shed in urine, and the authors speculated on their transmission and protozoal nature of these organisms. It has been suggested that E. cuniculi may spread transplacentally in rabbits.²⁶ Study of the pathogenesis of encephalitozoonosis in rabbits was initially hampered by the lack of non-subclinically infected rabbits and an in vitro assay to cultivate E. cuniculi.¹²¹ These limitations have been overcome, and it has been shown that experimental infection in rabbits can be established by a variety of routes (intravenous, oral, intratracheal, rectal); serum antibodies develop by day 21 postinoculation; spores are excreted in the urine for approximately 63 days; and early infection (day 30 postinoculation) is largely confined to the lung, kidney, and liver, while later infection (day 98 postinoculation) is most commonly localized to heart, brain, and kidney.^27,53,150 The presence or absence of encephalitozoon spores in rabbit fluids or tissues may yield useful epidemiologic clues for the clinician investigating a disease outbreak.

In naturally infected rabbits, E. cuniculi is primarily acquired through ingestion of infected urine. Occasionally, an infected rabbit will manifest neurologic signs of ataxia, opisthotonos, torticollis, hyperaesthesia, or paralysis; other less specific signs may include weight loss and failure to thrive.^77,99,111 Gross lesions are usually absent in infected rabbits, although chronic infection may manifest as focal, irregular, depressed areas in the renal cortex.⁵⁰ Histologically, lesions are most frequently identified in the brain and kidney, consisting of focal nonsuppurative meningoencephalitis with astrogliosis and perivascular lymphocytic infiltration and focal to segmental lymphocytic-plasmacytic interstitial nephritis with variable amounts of fibrosis; parasites may not be evident in areas of inflammation.²⁵ Other sections of the same tissue may contain intracellular clusters of parasites with no associated inflammatory response (Figs. 4, 5). Rarely, parasites are identified within the lens, lung, liver, and cardiac tissues of infected rabbits.^27,131 Parasites are best demonstrated with Gram's stain as gram-positive rod-shaped organisms free within a focus of inflammation or contained within intracellular parasitophorous vacuoles of endothelial cells, epithelial cells, or macrophages. Differential diagnoses for encephalitozoonosis in rabbits include central nervous system infection with Pasteurella multocida, Listeria monocytogenes, and Toxoplasma gondii.^{52,77,103,144}

View larger version (152K): [in this window] [in a new window]   Fig. 4. Cerebellum; domestic rabbit spontaneously infected with E. cuniculi. A focus of mononuclear cell infiltration and gliosis is present within the Purkinje cell layer. A small focus of intracellular parasites is present (arrow). Gram's stain. Bar = 50 µm. Fig. 5. Cerebrum and meninges; domestic rabbit spontaneously infected with E. cuniculi. Parasites are present within an intracellular parasitophorous vacuole, with no associated inflammation (arrow); meninges are thickened. Gram's stain. Bar = 50 µm.

Infection in mice

E. cuniculi infection of immunocompetent mice, as in rabbits, is usually subclinical.^55,102 Histologic lesions of mononuclear inflammatory foci in the liver, lungs, and brain have been attributed to subclinical E. cuniculi infection.⁶³ Experimental infection in athymic nude or severe combined immunodeficient (SCID) mice however, is typically widespread, with parasites disseminating by direct extension or hematogenously, depending on the route of inoculation. Clinical signs include lethargy, wasting, and death. Gross necropsy lesions include hepatosplenomegaly and multifocal miliary white spots throughout the liver, spleen, lung, and cardiac tissue.^42,60,76 Infection of the liver, spleen, pancreas, lungs, heart, kidneys, brain, peritoneum, and pleura is characterized by random, multifocal miliary granulomas with various amounts of cell debris and suppurative necrosis.^42,76 The gram-positive parasites may be found free within foci of inflammation or intracellularly in macrophages, epithelium, or endothelium. Mice inoculated orally may have focal ulcerative intestinal mucosal lesions that often extend to the submucosa and tunica muscularis.⁷⁶ Differential diagnoses for E. cuniculi infection in mice include infection with Clostridium piliforme (Tyzzer's disease), Corynebacterium kutscheri (pseudotuberculosis), Pseudomonas aeruginosa, salmonellosis, mouse hepatitis virus, ectromelia virus (mousepox), and hematopoietic/lymphoreticular neoplasms.^{3,36,46,54,57,62,127}

In addition to E. cuniculi infection, experimental E. intestinalis and E. hellem, but not Enterocytozoon bieneusi, infections have been documented in immunocompromised mice. Intestinal and systemic E. intestinalis infections have been documented in several strains of immunosuppressed mice.^1,49,143 E. hellem administered intraperitoneally to athymic nude mice results in similar gross and histologic lesions as seen in E. cuniculi-infected immunoincompetent mice.⁴² Spontaneous infections with E. intestinalis, E. hellem, or Enterocytozoon bieneusi have not been described in mice.

Infection in guinea pigs

As with immunocompetent mice, E. cuniculi infection in guinea pigs is subclinical. Vertical transmission of the organism is speculated to occur; E. cuniculi infection was detected serologically in hysterectomy-derived gnotobiotic guinea pigs.¹³ A recent, small serologic survey of inbred and outbred guinea pigs from a variety of sources revealed a seroprevalence of almost 50% to E. cuniculi. Most of these animals lacked gross and histologic evidence of infection.¹⁴² Multifocal necrosis of the cerebrum with granulomatous encephalitis and interstitial nephritis with tubular necrosis, regeneration, and fibrosis have been described in infected guinea pigs.¹⁴² Spontaneous and experimental infections with other Encephalitozoon species or other microsporidia have not been reported in guinea pigs.

Infection in carnivores

Natural infection of carnivores with E. cuniculi has been most thoroughly described in farm-raised foxes and domestic dogs. High mortality among blue fox kits, resulting in economic losses for the fur industry in Scandinavian countries, occurred as a result of endemic encephalitozoonosis.⁹⁵ E. cuniculi infection in domestic dogs has been reported from South Africa and the United States.^8,106,123 Puppies in these outbreaks had histologic and serologic evidence of encephalitozoonosis, and organisms were successfully reisolated in vitro from infected tissues. Serologic results in the parents were suggestive of recent E. cuniculi exposure.¹²³

Unlike the subclinical asymptomatic infection typically seen in rabbits and immunocompetent rodents, encephalitozoonosis is a sporadic, fulminating disease in neonatal carnivores.¹²³ In utero infection is suspected as the mode of transmission in fox kits.⁸⁹ Clinical signs in pups and kits include reduced appetite and stunted growth, ataxia, tremors, posterior weakness, and blindness with progression to circling, aggressive behavior, and convulsions.^87,132 Gross lesions include pale radial streaks extending from the cortex to the renal pelvis on cross sections of kidney and edematous, distended meninges. Thickened, necrotic, and tortuous medium-size to small arteries (consistent with polyarteritis nodosa) are often observed at necropsy in cardiac, intestinal, and central nervous system tissues in blue foxes.⁹⁵ Histologic lesions in dogs and foxes include multifocal microgranulomatous hepatitis, interstitial lymphocytic-plasmacytic nephritis, interstitial pneumonia, and lymphocytic-plasmacytic meningoencephalitis with perivascular cuffing and occasional intracellular parasitic cysts containing gram-positive spores.¹³² Surviving animals shed parasites in the urine.⁸⁷ Morbidity and mortality vary within a litter of pups or kits, and the parents typically remain unaffected. Differential diagnoses for neonatal canine encephalitozoonosis include canine herpesvirus and canine distemper virus infections, Neospora caninum, gram-negative septicemia, and hydrocephalus or other intracranial abnormalities.^{4,6,94,112,126,133} Infection of carnivores with other species of Encephalitozoon or other microsporidia has not been reported.

Infection in nonhuman primates

Reports of spontaneous infection with E. cuniculi in nonhuman primates have been best documented in squirrel monkeys (Saimiri sciureus).^122,154 In these studies, affected monkeys were classified as either neonates (<4 weeks old) or infants (<9 months old) suspected of acquiring the infection in utero. Antemortem signs were usually absent. There is a single case report of spontaneous intestinal microsporidiosis, presumed due to E. cuniculi, in an adult, wild-caught dusky titi monkey (Callicebus moloch) that was likely immunocompromised.¹²⁰

E. cuniculi infection in squirrel monkeys is typically subclinical, and gross lesions are not apparent.⁷ Neonatal, and presumably immunosuppressed, squirrel monkeys most frequently have microscopic lesions in the central nervous system. These lesions include multifocal granulomas within the white and gray matter, nonsuppurative meningitis, and vasculitis.^7,154 Parasites are gram-positive, rod-shaped organisms free within foci of inflammation or contained within parasitophorous vacuoles. Nonsuppurative interstitial nephritis and pneumonia with intralesional parasites also occur.^7,154 Placentitis, characterized by a mononuclear inflammatory infiltrate and organizing granulomas, has been described.⁷ The primary differential diagnosis for encephalitozoonosis in squirrel monkeys is infection with Toxoplasma gondii.⁷ Subclinical encephalitozoonosis has also been demonstrated histologically in vervet monkeys (Cercopithecus pygerythrus) experimentally infected with E. cuniculi isolated from domestic dogs.¹³⁷

There is a single report of experimental infection of immunocompetent and immunosuppressed rhesus macaques (Macaca mulatta) with E. cuniculi and E. hellem.⁴² Immunosuppressed macaques that received E. hellem intraveneously demonstrated spores in suppurative lesions of the liver and kidneys, whereas macaques inoculated orally with E. hellem or E. cuniculi shed spores periodically in the urine and feces but lacked gross and histologic evidence of parasite infection. Experimental and spontaneous Enterocytozoon bieneusi infections have recently been reported in several macaques (Macaca mulatta, M. nemestrina, and M. cyclopsis) infected with simian immunodeficiency virus (SIV).^22,83,134 Infection was associated with chronic hepatobiliary disease in these macaques and was characterized histologically by nonsuppurative and proliferative cholecystitis.⁸³ Parasites were also identified in the upper gastrointestinal tract mucosa.⁸³ E. bieneusi has also been diagnosed in SIV-negative, clinically healthy captive rhesus macaques, suggesting that this parasite may be a common or transient inhabitant of the digestive system of otherwise healthy macaques.⁸² Spontaneous infection with Encephalitozoon intestinalis or E. hellem infections have not been reported in macaques, squirrel monkeys, or other Old or New World primates.

Infection in other animals

There are relatively few reports in the literature describing natural or experimental infection with E. cuniculi in rats; most of these reports are limited to historical surveys of rodent populations.^70,102,104 Infections with other microsporidia have not been reported in rats. However, there is an increasing number of reports of spontaneous microsporidial infection in psittacines, including lovebirds (Agapornis spp.), budgerigars (Melopsittacus undulatus), and eclectus parrots (Eclectus roratus).^12,71,96,108 In many of the early reports, the etiologic agent was identified as E. cuniculi, but molecular analysis has revealed that the currently reported psittacine infections are caused by E. hellem.¹² Infected birds are young and frequently are coinfected with other pathogens. Clinical signs include anorexia, lethargy, and failure to thrive.¹² Lesions include focal hepatic necrosis with intralesional parasites. Parasites can also be identified in intestinal mucosa, crypts, and lamina propria and in renal tubular and bile duct epithelial cells, without an associated inflammatory response.¹² There are no published reports of experimental infections of psittacines with E. cuniculi or E. hellem nor are there reports of spontaneous or experimental infections with E. intestinalis or Enterocytozoon bieneusi. Although less widely reported, spontaneous infection of pigs with E. bieneusi and Encephalitozoon intestinalis and infection of a donkey, dog, cow, and goat with E. intestinalis have been described.^14,34 Infection in these animals was determined by PCR identification of spores from fecal samples. Additional diagnostic and epidemiologic work is needed to determine whether infection in these mammals results in clinical disease and whether they serve as potential zoonotic reservoirs of Microspora.

	Microsporidiosis as an Emerging Human Disease

Top Abstract Mammalian Microspora: Life... Diagnosis of Mammalian... Natural and Experimental... Microsporidiosis as an Emerging... Animal Models of Human... Comments References

 
Until the mid-1980s, human encephalitozoonosis was infrequently reported and was usually associated with immunoincompetent children manifesting neurologic signs.^10,84,86 The etiologic agent in these cases was presumed to be E. cuniculi. With the advent of HIV, acquired immunodeficiency disease (AIDS), and the development of more sophisticated diagnostic techniques, the spectrum of microsporidia capable of infecting humans has broadened and reports of human infections have increased. The majority of these infections result in disorders of the digestive tract (Table 3); Enterocytozoon bieneusi is most frequently identified.⁹⁷ E. bieneusi was first described in 1985 as the cause of chronic diarrhea and wasting in an AIDS patient.³⁵ It is primarily a pathogen of the small intestine and has been implicated as the etiologic agent responsible for 30–70% of the cases of AIDS-related diarrhea and wasting.¹⁵ More recent epidemiologic surveys of AIDS patients with chronic diarrhea suggest that the prevalence of microsporidiosis in this patient population is closer to 15%.¹⁶ This organism is also recognized as a cause of self-limiting "traveler's diarrhea" in immunocompetent individuals.² E. bieneusi infects mucosal enterocytes, resulting in duodenitis, but has also been implicated as the cause of sclerosing cholangitis and cholecystitis.^97,107 There is also some suggestion that E. bieneusi can infect the upper respiratory tract through aspiration of intestinal contents, resulting in sinusitis.³⁰ Clinical signs of diarrhea and wasting associated with E. bieneusi are reported to improve with albendazole, thalidomide, fumagillin, or furazolidone therapy, but elimination of infection has not been confirmed with these compounds.^45,47,90,125 In a more recent study, clindamycin was suggested to be efficacious against E. bieneusi infection.⁷²

E. hellem was isolated in 1990 and was first described as the cause of keratoconjunctivitis in HIV-positive patients.^38,153 Although morphologically and ultrastructurally identical to E. cuniculi, E. hellem was identified by immunologic and molecular analysis as a separate species.^38,44,141 E. hellem has since been shown to infect the respiratory and urogenital tracts of immunocompromised individuals, causing lesions of necrotizing tracheobronchitis, necrosuppurative prostatitis, lymphocytic-plasmacytic interstitial nephritis, and ulcerative cystitis.^114,119,147 Ocular infections with E. hellem have been eradicated with topical fumagillin or parental itraconazole, and disseminated infections have responded to treatment with albendazole.^40,153

It was not until 1995 that E. cuniculi infection, differentiated from E. hellem with monoclonal antibodies, was identified in an HIV-positive patient with clinical signs of chronic sinusitis, rhinitis, and keratoconjunctivitis.⁵¹ E. cuniculi has since been shown to have a tissue distribution in immunocompromised humans similar to that in rabbits, infecting renal endothelial and epithelial cells, microglia, and myocytes.^88,146 Electron microscopy remains the gold standard for diagnosing and differentiating all human microsporidial infections, although PCR primers have been developed and hold promise of a faster, less invasive means of detection that can be adapted to large scale epidemiologic studies.¹⁴⁹ Reports of human microsporidiosis will continue to increase as recognition of these parasites and the spectrum of diseases they cause increase.

	Animal Models of Human Microsporidiosis

Top Abstract Mammalian Microspora: Life... Diagnosis of Mammalian... Natural and Experimental... Microsporidiosis as an Emerging... Animal Models of Human... Comments References

 
The ease with which the Encephalitozoon spp. can be cultivated in vitro has facilitated the development of several experimental animal models of infection for E. cuniculi, E. intestinalis, and E. hellem (Table 4). Mice are most frequently used because of their small size, ease of handling and infecting, and simple housing requirements, and because they are available in a variety of genetically defined immunocompromised strains. None of these murine models reproduces the clinical signs of diarrhea commonly observed in humans infected with E. intestinalis nor the respiratory signs most frequently associated with human E. hellem infection (see Table 2). Tissue distribution of E. cuniculi in the brain and kidneys in infected mice (and rabbits) appears similar to that seen in infected humans.^88,146 Systemic disease can be reproduced in a number of immunosuppressed mouse strains by a variety of inoculation routes with E. cuniculi, E. hellem, and E. intestinalis, and these models will aid in examining the mechanisms of microsporidial dissemination.^42,76,143 Mouse models have also been valuable in elucidating the immunologic response to infection with Encephalitozoon spp.⁷³ Foreshadowing the diagnosis of microsporidial infection in T-cell-deficient AIDS patients, early work with E. cuniculi demonstrated that antibody production and T lymphocytes contribute to the control of encephalitozoonosis in mice and that lymphokine-activated macrophages were responsible for killing the parasite, as occurs with other intracellular parasites.^55,115,116 The course of E. cuniculi infection in immunocompetent mice also represents a model that will aid in answering questions regarding the host-parasite relationship.⁴² This information may be important as reports of transient microsporidial infection in healthy humans increase.¹⁰⁹ Despite the number of case reports documenting treatment of human microsporidial infection with various therapeutic agents, there are relatively few reports of drug efficacy against this disease in animals.^28,131 Future work with the mouse models should address host stimuli that initiate polar tube extrusion, identification of potential host cell receptors for parasite invasion, mechanisms of intracellular survival and immune system evasion, and drug screening and efficacy testing. Mouse models will continue to be valuable for filling the gaps in our knowledge of human and mammalian encephalitozoonosis.

	Comments

Top Abstract Mammalian Microspora: Life... Diagnosis of Mammalian... Natural and Experimental... Microsporidiosis as an Emerging... Animal Models of Human... Comments References

 
Recognized as a mammalian protozoal parasite for nearly 80 years, Encephalitozoon cuniculi and several additional microsporidia have been identified as opportunistic pathogens in AIDS patients only in the last 15 years. Very little is understood regarding the cellular and molecular biology, pathogenicity, and epidemiology of these organisms, particularly in comparison with other human parasites such as Plasmodia, Trypanosoma, Leishmania, and Toxoplasma gondii. Although not causing the same global morbidity and mortality in the human population as some other protozoal pathogens, microsporidia are recognized as significant secondary invaders in immunosuppressed people. Additional information is needed regarding animal and environmental reservoirs of the microsporidia and the zoonotic potential of microsporidiosis for humans. Methods of preventing infection and identifying effective therapeutic agents against Encephalitozoon and other microsporidia will also be important for immunocompromised humans and for affected companion and production animals. A variety of animal models for human E. cuniculi, E. hellem, and E. intestinalis infection exist, but there are still gaps in our knowledge regarding parasite transmission and life cycle, the host–parasite relationship, modes of cellular infection and dissemination, and the molecular signals that govern these events. Although nonhuman primates are capable of sustaining infection with Enterocytozoon bieneusi, development of a reproducible rodent model of infection would facilitate study of this microsporidium. Future work should be directed at refining current models and developing new animal models of human encephalitozoonosis and other microsporidial infections. In addition, the differences in pathogenicity and disease mechanisms among Encephalitozoon cuniculi, E. hellem, E. intestinalis, and Enterocytozoon bieneusi have yet to be determined. Veterinary pathologists are uniquely situated to assist in answering these questions because of their exposure to a diverse number of animal species and their knowledge of infectious disease processes. Considered perhaps a disease of historical importance in the field of laboratory animal medicine, encephalitozoonosis has reemerged as a parasitic infection to be added to the expanding list of human microsporidial diseases and other HIV-associated opportunistic infections. Microsporidiosis of humans and animals will continue to be of scientific interest well into the next century.

	Acknowledgments

 
The editorial comments of Dr. Howard B. Gelberg are gratefully acknowledged.

	Footnotes

 
¹ Present address: Department of Pathology, Microbiology and Immunology, School of Veterinary Medicine, University of California, Davis, CA.

	References

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1. Achbarou A, Ombrouck C, Gneragbe T, Charlotte F, Renia L, Desportes-Livage I, Mazier D: Experimental model for human intestinal microsporidiosis in interferon gamma receptor knockout mice infected by Encephalitozoon intestinalis. Parasit Immunol 18:387-392, 1996[CrossRef][Medline]
2. Albrecht H, Sobottka I: Enterocytozoon bieneusi infection in patients who are not infected with human immunodeficiency virus. Clin Infect Dis 25:344, 1997[Medline]
3. Amao H, Komukai Y, Sugiyama M, Takahashi KW, Sawada T, Saito M: Natural habitats of Corynebacterium kutscheri in subclinically infected ICGN and DBA/2 strains of mice. Lab Anim Sci 45:6-10, 1995[Medline]
4. Anvik JO: Clinical considerations of canine herpesvirus infection. Vet Med 86:394-403, 1991
5. Arison RN, Cassaro JA, Pruss MP: Studies on a murine ascites-producing agent and its effect on tumor development. Cancer 26:1915-1920, 1966
6. Bagley RS: Intracranial disease. Vet Clin North Am Small Anim Pract 26:667-980, 1996[Medline]
7. Baskin G: Encephalitozoon cuniculi infection, squirrel monkey. In: Nonhuman Primates II, ed. Jones TC, Mohr U, and Hunt RD, pp 193-196, Springer-Verlag, Berlin, Germany 1993
8. Basson PA, McCully RM, Warnes WEJ: Nosematosis: report of a canine case in the Republic of South Africa. J S Afr Vet Med Assoc 37:3-9, 1966
9. Behm CA: The role of trehalose in the physiology of nematodes. Int J Parasitol 27:215-229, 1997[CrossRef][Medline]
10. Bergquist NR, Stintzing G, Smedman L, Waller T, Andersson T: Diagnosis of encephalitozoonosis in man by serological tests. Br Med J 288:902, 1984
11. Biderre C, Pages M, Metenier G, Canning EU, Vivares CP: Evidence for the smallest nuclear genome (2.9 Mb) in the microsporidium Encephalitozoon cuniculi. Mol Biochem Parasitol 74:229-231, 1995[CrossRef][Medline]
12. Black SS, Steinohrt LA, Bertucci DC, Rogers LB, Didier ES: Encephalitozoon hellem in budgerigars (Melopsittacus undulatus). Vet Pathol 34:189-198, 1997[Abstract]
13. Boot R, van Knapen F, Kruijt BC, Walvoort HC: Serological evidence for Encephalitozoon cuniculi infection (nosemiasis) in gnotobiotic guinea pigs. Lab Anim 22:337-342, 1988[Abstract/Free Full Text]
14. Bornay-Llinares F, da Silva A, Moura H, Schwartz DA, Visvesvara GS, Pieniazek NJ, Cruz-Lopez A, Hernandez-Jauregui P, Guerrero J, Enriquez FJ: Immunologic, microscopic and molecular evidence of Encephalitozoon intestinalis (Septata intestinalis) infection in mammals other than humans. J Infect Dis 178:820-826, 1998[Medline]
15. Bryan RT: Microsporidiosis as an AIDS-related opportunistic infection. Clin Infect Dis 21:S62-S65, 1995
16. Bryan RT, Schwartz DA: Epidemiology of microsporidiosis. In: The Microsporidia and Microsporidiosis, ed. Wittner M, pp 502-516, ASM Press, Herdon, VA 1999
17. Bull CG: The pathologic effects of streptococci from cases of poliomyelitis and other sources. J Exp Med 25:557-580, 1917[Abstract]
18. Cali A, Kolter DP, Orenstein JM: Septata intestinalis n. g., n. sp., an intestinal microsporidian associated with chronic diarrhea and dissemination in AIDS patients. J Eukaryot Microbiol 40:101-112, 1993[Medline]
19. Cali A, Owen RL: Intracellular development of Enterocytozoon, a unique microsporidian found in the intestine of AIDS patients. J Protozool 37:145-155, 1990[Medline]
20. Candy DJ, Becker A, Wegener G: Coordination and integration of metabolism in insect flight. Comp Biochem Physiol 117:497-512, 1997[CrossRef]
21. Canning EU, Lom J: The Microsporidia of Vertebrates. Harcourt Brace Jovanovich, New York, NY 1986
22. Chalifoux LV, MacKey J, Carville A, Shvetz D, Lin KC, Lackner A, Mansfield KG: Ultrastructural morphology of Enterocytozoon bieneusi in biliary epithelium of rhesus macaques (Macaca mulatta). Vet Pathol 35:292-296, 1998[Abstract]
23. Chioralia G, Trammer T, Kampen H, Seitz HM: Relevant criteria for detecting microsporidia in stool specimens. J Clin Microbiol 36:2279-2283, 1998[Abstract/Free Full Text]
24. Chupp GL, Alroy J, Adelman LS, Breen JC, Skolnik PR: Myositis due to Pleistophora (Microsporidia) in a patient with AIDS. Clin Infect Dis 16:15-21, 1993[Medline]
25. Cox JC, Gallichio HA: Serological and histological studies on adult rabbits with recent, naturally acquired encephalitozoonosis. Res Vet Sci 24:260-261, 1978[Medline]
26. Cox JC, Gallichio HA, Pye D, Walden NB: Application of immunofluorescence to the establishment of an Encephalitozoon cuniculi-free rabbit colony. Lab Anim Sci 27:204-209, 1977[Medline]
27. Cox JC, Hamilton RC, Attwood HD: An investigation of the route and progression of Encephalitozoon cuniculi infection in adult rabbits. J Protozool 26:260-265, 1979[Medline]
28. Coyle C, Kent M, Tanowitz HB, Wittner M, Weiss LM: TNP-470 is an effective antimicrosporidial agent. J Infect Dis 177:515-518, 1998[Medline]
29. De Groote MA, Visvesvara GS, Wilson ML, Pieniazek NJ, Slemenda SB, da Silva AJ, Leitch GJ, Bryan RT, Reves R: Polymerase chain reaction and culture confirmation of disseminated Encephalitozoon cuniculi in a patient with AIDS: successful therapy with albendazole. J Infect Dis 171:1375-1378, 1995[Medline]
30. del Aguila C, Lopez-Velez R, Fenoy S, Turrientes C, Cobo J, Navajas R, Visvesvara GS, Croppo GP, da Silva AJ, Pieniazek NJ: Identification of Enterocytozoon bieneusi spores in respiratory samples from an AIDS patient with a 2-year history of intestinal microsporidiosis. J Clin Microbiol 35:1862-1866, 1997[Abstract]
31. Delbac F, Duffieux F, David D, Metenier G, Vivares C: Immunocytochemical identification of spore proteins in two microsporidia, with emphasis on extrusion apparatus. J Eukaryot Microbiol 45:224-231, 1998[Medline]
32. Delbac F, Peyret P, Metenier G, David D, Danchin A, Vivares CP: On proteins of the microsporidian invasive apparatus: complete sequence of a polar tube protein of Encephalitozoon cuniculi. Mol Microbiol 29:825-834, 1998[CrossRef][Medline]
33. Deplazes P, Mathis A, Baumgartner R, Tanner I, Weber R: Immunologic and molecular characteristics of Encephalitozoon-like microsporidia isolated from humans and rabbits indicate that Encephalitozoon cuniculi is a zoonotic parasite. Clin Infect Dis 22:557-559, 1996[Medline]
34. Deplazes P, Mathis A, Muller C, Weber R: Molecular epidemiology of Encephalitozoon cuniculi and first detection of Enterocytozoon bieneusi in faecal samples of pigs. J Eukaryot Microbiol 43:93S, 1996[Medline]
35. Desportes I, Le Charpentier Y, Galian A, Bernard F, Cochand-Proillet B, Lavergne A, Ravisse P, Modigliani R: Occurrence of a new microsporidian: Enterocytozoon bieneusi n. g., n. sp., in the enterocytes of a human patient with AIDS. J Protozool 32:250-254, 1985[Medline]
36. Dick EJ, Kittell CL, Meyer H, Farrar PL, Ropp SL, Esposito JJ, Buller RML, Neubauer H, Kang YH, McKee AE: Mousepox outbreak in a laboratory mouse colony. Lab Anim Sci 46:602-611, 1996[Medline]
37. Didier ES: Effects of albendazole, fumagillin, and TNP-470 on microsporidial replication in vitro. Antimicrob Agents Chemother 47:1541-1546, 1997
38. Didier ES, Didier PJ, Friedberg DN, Stenson SM, Orenstein JM, Yee RW, Tio FO, Davis RM, Vossbrinck C, Millichamp N, Shadduck JA: Isolation and characterization of a new human microsporidian, Encephalitozoon hellem (n. sp.), from three AIDS patients with keratoconjunctivitis. J Infect Dis 163:617-621, 1991[Medline]
39. Didier ES, Orenstein JM, Aldras A, Bertucci D, Rogers LB, Janney FA: Comparison of three staining methods for detecting microsporidia in fluids. J Clin Microbiol 33:3138-3145, 1995[Abstract]
40. Didier ES, Rogers LB, Brush AD, Wong S, Traina-Dorge V, Bertucci D: Diagnosis of disseminated microsporidian Encephalitozoon hellem infection by PCR–Southern analysis and successful treatment with albendazole and fumagillin. J Clin Microbiol 34:947-952, 1996[Abstract]
41. Didier ES, Snowden K, Shadduck JA: Biology of microsporidian species infecting mammals. In: Advances in Parasitology, ed. Baker JR, Muller R, and Rollinson D, pp 283-320, Academic Press, San Diego, CA 1998
42. Didier ES, Varner PW, Didier PJ, Aldras A, Millichamp NJ, Murphey-Corb M, Bohm R, Shadduck JA: Experimental microsporidiosis in immunocompetent and immunodeficient mice and monkeys. Folia Parasitol 41:1-11, 1994
43. Didier ES, Visvesvara GS, Baker MD, Rogers LB, Bertucci D, De Groote MA, Vossbrinck CR: A microsporidian isolated from an AIDS patient corresponds to Encephalitozoon cuniculi III, originally isolated from domestic dogs. J Clin Microbiol 34:2835-2837, 1996[Abstract]
44. Didier PJ, Didier ES, Orenstein JM, Shadduck JA: Fine structure of a new human microsporidian, Encephalitozoon hellem, in culture. J Protozool 38:502-507, 1991[Medline]
45. Dieterich DT, Lew EA, Kolter DP, Poles MA, Orenstein JM: Treatment with albendazole for intestinal disease due to Enterocytozoon bieneusi in patients with AIDS. J Infect Dis 169:178-183, 1994[Medline]
46. DiGiacomo RF, Garlinghouse LE, Giddens WE, Dennis MB: Outbreaks of salmonellosis in mice. Lab Anim Sci 33:496, 1983 [abstract]
47. Dionisio D, Manneschi LI, Di Lollo S, Orsi A, Sterrantino G, Meli M, Gabbrielle M, Tani A, Papucci A, Leoncini F: Enterocytozoon bieneusi in AIDS: symptomatic relief and parasite changes after furazolidone. J Clin Pathol 50:472-476, 1997[Abstract/Free Full Text]
48. Dore GJ, Marriott DJ, Hing MC, Harkness JL, Field AS: Disseminated microsporidiosis due to Septata intestinalis in nine patients infected with the human immunodeficiency virus: response to therapy with albendazole. Clin Infect Dis 21:70-76, 1995[Medline]
49. Fakhry YE, Achbarou A, Desportes-Livage I, Mazier D: Encephalitozoon intestinalis: humoral responses in interferon- receptor knockout mice infected with a microsporidium pathogenic in AIDS patients. Exp Parasitol 89:113-121, 1998[CrossRef][Medline]
50. Flatt RE, Jackson SJ: Renal nosematosis in young rabbits. Vet Pathol 7:492-497, 1970
51. Franzen C, Schwartz DA, Visvesvara GS, Muller A, Schwenk A, Salzberger B, Fatkenheuer G, Hartmann P, Mahrle G, Diehl V, Schrappe M: Immunologically confirmed disseminated, asymptomatic Encephalitozoon cuniculi infection of the gastrointestinal tract in a patient with AIDS. Clin Infect Dis 21:1480-1484, 1995[Medline]
52. Frenkel JK: Pathogenesis of toxoplasmosis and of infections with organisms resembling Toxoplasma. Ann NY Acad Sci 64:215-231, 1956[CrossRef]
53. Fuentealba IC, Mahoney NT, Shadduck JA, Harvill J, Wicher V, Wicher K: Hepatic lesions in rabbits infected with Encephalitozoon cuniculi administered per rectum. Vet Pathol 29:536-540, 1992[Abstract]
54. Ganaway JR, Allen AM, Moore TD: Tyzzer's disease. Am J Pathol 64:717-732, 1971[Medline]
55. Gannon J: The course of infection of Encephalitozoon cuniculi in immunodeficient and immunocompetent mice. Lab Anim 14:189-192, 1980[Abstract/Free Full Text]
56. Gardiner CH, Fayer R, Dubey JP: An Atlas of Protozoan Parasites in Animal Tissues, 2nd ed. American Registry of Pathology, Washington, DC 1998
57. Gardner MB, Henderson BE, Rongey RW, Estes JD, Huebner RJ: Spontaneous tumors of aging wild house mice: incidence, pathology, and C-type virus expression. J Natl Cancer Inst 50:719-734, 1973
58. Glaser CA, Angulo FJ, Rooney JA: Animal-associated opportunistic infections among persons infected with the human immunodeficiency virus. Clin Infect Dis 18:14-24, 1994[Medline]
59. Goodpasture EW: Spontaneous encephalitis in rabbits. J Infect Dis 34:428-432, 1924
60. Hermanek J, Koudela B, Kucerova Z, Ditrich O, Travnicek J: Prophylactic and therapeutic immune reconstitution of SCID mice infected with Encephalitozoon cuniculi. Folia Parasitol 40:287-291, 1993
61. Hollister WS, Canning EU, Weidner E, Field AS, Kench J, Marriott DJ: Development and ultrastructure of Trachipleistophora hominis n.g., n.sp. after in vitro isolation from an AIDS patient and inoculation into athymic mice. Parasitology 112:143-154, 1996
62. Homberger FR, Zhang-LiNong, Barthold SW: Prevalence of enterotropic and polytropic mouse hepatitis virus in enzootically infected mouse colonies. Lab Anim Sci 48:50-54, 1998[Medline]
63. Innes JRM, Zeman W, Frenkel JK: Occult endemic encephalitozoonosis of the central nervous system of mice. J Neuropathol Exp Neurol 21:519-533, 1962[Medline]
64. Jordan J, Mirick GS: An infectious hepatitis of undetermined origin in mice: I. Description of the disease. J Exp Med 102:601-616, 1965
65. Jordan J, Mirick GS: An infectious hepatitis of undetermined origin in mice: II. Characteristics of the infective agent. J Exp Med 102:617-630, 1965
66. Jorge JA, De-Lourdes M, Polizeli TM, Thevelein JM, Terenzi HF: Trehalases and trehalose hydrolysis in fungi. FEMS Microbiol Lett 154:165-171, 1997[CrossRef][Medline]
67. Joste NE, Rich JD, Busam KJ, Schwartz DA: Autopsy verification of Encephalitozoon intestinalis (microsporidiosis) eradication following albendazole therapy. Arch Pathol Lab Med 120:199-203, 1996[Medline]
68. Katzwinkel-Wladarsch S, Lieb M, Heise W, Loscher T, Rinder H: Direct amplification and species determination of microsporidian DNA from stool specimens. Trop Med Int Health 1:373-378, 1996[CrossRef][Medline]
69. Keeling PJ, McFadden GI: Origins of microsporidia. Trends Microbiol 6:19-23, 1998[CrossRef][Medline]
70. Kellett BS, Bywater JEC: A modified india-ink immunoreaction for the detection of encephalitozoonosis. Lab Anim 12:59-60, 1978[Abstract/Free Full Text]
71. Kemp RL, Kluge JP: Encephalitozoon sp. in the blue-masked lovebird, Agapornis personata (Reichenow): first confirmed report of microsporidian infection in birds. J Protozool 22:489-491, 1975[Medline]
72. Kester KE, Turiansky GW, McEvoy PL: Nodular cutaneous microsporidiosis in a patient with AIDS and successful treatment with long-term oral clindamycin therapy. Ann Intern Med 128:911-914, 1998[Abstract/Free Full Text]
73. Khan IA, Moretto M: Role of gamma interferon in cellular immune response against murine Encephalitozoon cuniculi infection. Infect Immun 67:1887-1893, 1999[Abstract/Free Full Text]
74. Kolter D, Orenstein JM: Clinical syndromes associated with microsporidiosis. In: Advances in Parasitology, ed. Baker JR, Muller R, and Rollinson D, pp 321-349, Academic Press, San Diego, CA 1998
75. Kondova I, Mansfield K, Buckholt MA, Stein B, Widmer G, Carville A, Lackner A, Tzipori S: Transmission and serial propagation of Enterocytozoon bieneusi from humans and rhesus macaques in gnotobiotic piglets. Infect Immun 66:5515-5519, 1998[Abstract/Free Full Text]
76. Koudela B, Vitovec J, Kucerova Z, Ditrich O, Travnicek J: The severe combined immunodeficient mouse as a model for Encephalitozoon cuniculi microsporidiosis. Folia Parasitol 40:279-285, 1993
77. Kunstyr I, Naumann S: Head tilt in rabbits caused by pasteurellosis and encephalitozoonosis. Lab Anim 19:208-213, 1985[Medline]
78. Leitch GJ, He Q, Wallace S, Visvesvara GS: Inhibition of the spore polar filament extrusion of the microsporidium Encephalitozoon hellem, isolated from an AIDS patient. J Eukaryot Microbiol 40:711-717, 1993[Medline]
79. Leitch GJ, Scanlon M, Visvesvara GS, Wallace S: Calcium and hydrogen ion concentrations in the parasitophorous vacuoles of epithelial cells infected with the microsporidian Encephalitozoon hellem. J Eukaryot Microbiol 42:445-451, 1995[Medline]
80. Lowder CY, McMahon JT, Meisler DM, Dodds EM, Calabrese LH, Didier ES, Cali A: Microsporidial keratoconjunctivitis caused by Septata intestinalis in a patient with acquired immunodeficiency syndrome. Am J Ophthalmol 121:715-717, 1996[Medline]
81. Luna V, Stewart BK, Bergeron DL, Clausen CR, Plorde JJ, Fritsche TR: Use of the fluorochrome calcofluor white in the screening of stool specimens for spores of microsporidia. Am J Clin Pathol 103:656-659, 1995[Medline]
82. Mansfield KG, Carville A, Hebert D, Chalifoux L, Shvetz D, Lin KC, Tzipori S, Lackner AA: Localization of persistent Enterocytozoon bieneusi infection in normal rhesus macaques (Macaca mulatta) to the hepatobiliary tree. J Clin Microbiol 36:2336-2338, 1998[Abstract/Free Full Text]
83. Mansfield KG, Carville A, Shvetz D, MacKey J, Tzipori S, Lackner AA: Identification of an Enterocytozoon bieneusi-like microsporidian parasite in simian-immunodeficiency-virus-inoculated macaques with hepatobiliary disease. Am J Pathol 150:1395-1405, 1997[Abstract]
84. Margileth AM, Strano AJ, Chandra R, Neafie R, Blum M, McCully RM: Disseminated nosematosis in an immunologically compromised infant. Arch Pathol 95:145-150, 1973[Medline]
85. Mathis A, Michel M, Kuster H, Muller C, Weber R, Deplazes P: Two Encephalitozoon cuniculi strains of human origin are infectious to rabbits. Parasitol 114:29-36, 1997
86. Matsubyashi M, Kioke T, Mikata I, Takei H, Hagiwara S: A case of encephalitozoon-like body infection in man. Arch Pathol 67:73-79, 1959
87. McInnes EF, Stewart CG: The pathology of subclinical infection of Encephalitozoon cuniculi in canine dams producing pups with over encephalitozoonosis. J S Afr Vet Med Assoc 62:51-54, 1991
88. Mertens RB, Didier ES, Fishbein MC, Bertucci D, Rogers LB, Orenstein JM: Encephalitozoon cuniculi microsporidiosis: infection of the brain, heart, kidneys, trachea, adrenal glands, and urinary bladder in a patient with AIDS. Mod Pathol 10:68-77, 1997[Medline]
89. Mohn SF, Nordstoga K, Krogsrud J, Helgebostad A: Transplacental transmission of Nosema cuniculi in the blue fox (Alopex lagopus). Acta Pathol Microbiol Scand 82:299-300, 1974
90. Molina J, Goguel J, Sarfati C, Chastang C, Desportes-Livage I, Michiels J, Maslo C, Katlama C, Cotte L, Leport C, Raffi F, Derouin F, Modai J: Potential efficacy of fumagillin in intestinal microsporidiosis due to Enterocytozoon bieneusi in patients with HIV infection: results of a drug screening study. AIDS 11:1603-1610, 1997[CrossRef][Medline]
91. Molina J, Oksenhendler E, Beauvais B, Sarfati C, Jaccard A, Derouin F, Modai J: Disseminated microsporidiosis due to Septata intestinalis in patients with AIDS: clinical features and response to albendazole therapy. J Infect Dis 171:245-249, 1995[Medline]
92. Morris JA, McCown JM, Blount RE: Ascites and hepatosplenomegaly in mice associated with protozoan-like cytoplasmic structures. J Infect Dis 98:306-311, 1956[Medline]
93. Moura H, Sodre FC, Bornay-Llinares F, Leitch GJ, Navin T, Wahlquist SP, Bryan RT, Meseguer I, Visvesvara GS: Detection by an immunofluorescence of Encephalitozoon intestinalis spores in routinely formalin-fixed stool samples stored at room temperature. J Clin Microbiol 37:2317-2322, 1999[Abstract/Free Full Text]
94. Muller C, Fatzer RS, Beck K, Vandevelde M, Zurbriggen A: Studies on canine distemper virus persistence in the central nervous system. Acta Neuropathol 89:438-445, 1995[Medline]
95. Nordstoga K, Westbye K: Polyarteritis nodosa associated with nosematosis in blue foxes. Acta Pathol Microbiol Scand 84:291-296, 1976
96. Norton JH, Prior HC: Microsporidiosis in a peach-faced lovebird (Agapornis roseicollis). Aust Vet J 71:23-24, 1994[Medline]
97. Orenstein JM: Microsporidiosis. In:Pathology of Infectious Diseases, ed. Connor DH, Chandler FW, Schwartz DA, Manz HJ, and Lack EE, pp 1223-1239, Appleton & Lange, Stamford, CT 1997
98. Orenstein JM, Tenner M, Cali A, Kolter DP: A microsporidian previously undescribed in humans, infecting enterocytes and macrophages, and associated with diarrhea in an acquired immunodeficiency syndrome patient. Hum Pathol 23:722-728, 1992[CrossRef][Medline]
99. Pakes SP, Gerrity LW: Protozoal diseases. In: The Biology of the Laboratory Rabbit, ed. Manning PJ, Ringler DH, and Newcomer CE, 2nd ed., pp 205-229, Academic Press, San Diego, CA 1994
100. Pakes SP, Shadduck JA, Cali A: Fine structure of Encephalitozoon cuniculi from rabbits, mice and hamsters. J Protozool 22:481-488, 1975[Medline]
101. Percy DH, Barthold SW: Pathology of Laboratory Rodents and Rabbits. Iowa State University Press, Ames, IA 1993
102. Perrin TL: Spontaneous and experimental encephalitozoon infection in laboratory animals. Arch Pathol 36:559-567, 1943
103. Perrin TL: Toxoplas