49 Microsporidiosis

The class or order Microsporidia was elevated in to the phylum Microspora by Sprague and Vávra (1997), and Sprague and Becnel (1998) subsequently suggested that the term Microsporidia instead be used for the phylum name. Microsporidia, i.e. Nosema bombycis, were first described about 150 years ago as the cause of the disease pebrine in silkworms. In 1922, there were descriptions of gram-positive spores consistent with microsporidiosis in the brain of rabbits that were being used for investigations on poliomyelitis (Wright and Craighead 1922). From 1923 to 1926, Levaditi and colleagues studied the organisms seen by Wright and Craighead, which they named Encephalitozoon cuniculi, recognizing them as Microsporidia and demonstrating their lack of host specificity by transmitting infections from rabbits to mice, rats and dogs (Levaditi et al. 1923). Microsporidia were clearly confirmed of being a cause of human disease in 1959 (Matsubayashi et al. 1959), when they were isolated from the cerebrospinal fluid of a 9 year old boy with encephalitis with seizures, coma, and fever lasting about 25 days. Bergquist et al. (1984) reported a 2 year old child with encephalitis and seizures who had Encephalitozoon spores in urine and Margileth et al. (1973) isolated the microsporidium Anncaliia (Nosema) connori from a 4 month old athymic male infant who died with severe diarrhoea and malabsorption. Microsporidia can produce a wide range of clinical diseases. A diarrhoeal syndrome associated with microsporidiosis and HIV infection was reported by Desportes et al. (1985) and the number of articles describing human disease increased rapidly after 1990. In addition to gastrointestinal tract involvement, it has been recognized that Microsporidia can infect virtually any organ system; and patients with encephalitis, ocular infection, sinusitis, myositis, and disseminated infection are well described in the literature.

The Microsporidia are obligate intracellular unicelluar eukaryotes containing a nucleus with a nuclear envelope, an intracytoplasmic membrane system, chromosome separation on mitotic spindles, and vesicular Golgi (Desportes-Livage 2000). While originally believed to be lacking mitochondria, it is now appreciated that they have a mitochondrial ‘remnant’ organelle named the mitosome (Williams et al. 2002). The nuclei are typical of eukaryotes, but electron-dense centriolar plaques in nuclear pores are present at the poles of division spindles in place of centrioles. In several genera, including Vittaforma and Nosema, which have been isolated from humans, the nuclei are unusual in being paired as diplokarya which are closely appressed and divide synchonously. In some genera this diplokaryotic condition is maintained throughout the life cycle but, in others, there is an alternation between unpaired and paired nuclei. In a few genera alternation of hosts is an obligate part of the life cycle but, as far as is known, these complexities do not occur in the genera infecting humans.

The Microsporidia are defined by their characteristic unicellular spores (Fig. 49.1) which are resistant to the environment (Wittner and Weiss 1999). The spore size and shape vary depending on the species. The spore coat consists of an electron-dense, proteinaceous exospore, an electron-lucent endospore composed of chitin and protein, and an inner membrane or plasmalemma (Vavra 1976). Spore coat proteins have adhesion domains that may facilitate the binding of spores to host cells or gastrointestinal track mucous (Southern et al. 2007). A defining characteristic of all microsporidia is an extrusion apparatus that consists of a polar filament (tube) attached to the inside of the anterior end of the spore by an anchoring disk which coils around the sporoplasm in the spore. Proteomic and genetic studies have defined some of the proteins of the polar tube and spore wall (Xu et al. 2006) as well as the presence of O-mannosylatation on these proteins (Xu et al. 2004). When the spore is exposed to appropriate environmental conditions it germinates and the polar filament rapidly everts, forming the hollow polar tube which forms a bridge delivering the sporoplasm into intimate contact with the host cell (Fig. 49.2) (Lom 1972; Weidner 1972). The polar tube has been described as a hypdermic needle, but the mechanism by which the polar tube interacts with the host cell membrane is not known (Foucault and Drancourt 2000). Conditions that promote germination vary widely among species, presumably reflecting the organisms’ adaptation to their host and external environment (reviewed by Keohane and Weiss 1999). Interestingly, if a spore is phagocytosed by a host cell, germination occurs; and the polar tube can pierce the phagocytic vacuole, delivering the sporoplasm into the host cell cytoplasm (Franzen 2005).

The general features of microsporidian life cycles are as follows:

After subsequent division the sporonts give rise to sporoblasts, which go on to form mature spores without additional multiplication. Once a host cell becomes distended with mature spores, the cell ruptures, releasing mature spores into the environment, thereby completing the life cycle. The combination of multiplication during merogony and sporogony results in a large number of spores being produced from a single infection and illustrates the enormous reproductive potential of these organisms.

Microsporidia have prokaryotic-size ribosomes (Curgy et al. 1980) that do not have a 5.8S ribosome subunit but do have sequences homologous to the 5.8S region in the 23S subunit (Vossbrinck and Woese 1986). The small-subunit rRNA of many Microsporidia have been sequenced and found to be significantly shorter than both eukaryotic and prokaryotic small-subunit rRNA (Vossbrinck et al. 1987; Weiss and Vossbrinck 1998). These rRNA genes are in a subtelometeric location on each chromosome of E. cuniculi (Brugere et al. 2000a; Vivares and Metenier 2000) and lack the paromomycin binding site seen in protozoa and animals (Katiyar et al. 1995). The karyotype of several members of the phylum Microspora has been determined by pulsed-field electrophoresis. The genome size of the microsporidia varies from 2.3 to 19.5 Mb (Weiss and Vossbrinck 1999) with that of the Encephalitizoonidae being less than 3.0 Mb, making them among the smallest eukaryotic nuclear genomes so far identified (Vivares and Metenier 2000; Katinka et al. 2001). In the compact genomes of the Encephalitozoonidae, there are almost no introns, the gene density is high and proteins are shorter than the corresponding genes in Saccharomyces cervisiae. There appears to be a high degree of gene conservation among the Microsporidia (Corradi et al. 2007). Genome data on the Microsporidia is available at EuPathdB (http://microsporidiadb.org/micro/). Chromosomal analysis of E. cuniculi suggests that it is diploid (Brugere et al. 2000b).

Analysis of the rRNA genes of a variety of Microsporidia highlights the polyphyletic nature of the Microsporidia and brings into doubt the use of any single character for developing higher taxonomic groupings. For example, E. hellem and E. cuniculi are indistinguishable at the ultrastructural level, and E. intestinalis has a distinct extracellular matrix surrounding the sporoblasts and spores; based on rDNA analysis, E. intestinalis and E. cuniculi are more similar to each other than to E. hellem (Baker et al. 1995).

Microsporidia are classified by their ultrastructural features, including the size and morphology of the spores, number of coils of the polar tube, developmental life cycle, and host-parasite relationship. Overviews of the the microsporidian taxa have been reported a number of times (reviewed by Sprague et al. 1992). Molecular phylogenetic data indicates that the Microsporidia are related to fungi and are not ‘primitive eukaryotes’ (Keeling and McFadden 1998; Hirt et al. 1999; Weiss et al. 1999). Based on tubulin sequence analysis, it has been suggested that the Microsporidia are related to the fungi (Keeling and Doolittle 1996). Analysis of hsp70 for various microsporidia provided confirmatory evidence of this relationship (Germot et al. 1997; Hirt et al. 1997). Keeling (2003), in an analysis of β-tubulin data that included additional species of Microsporidia and more fungal phyla, suggested that the Microsporidia were a sister group to the Zygomycota. Additional evidence for the relationship of Microsporidia to the fungi includes the following:

Although initially regarded as rare, Microsporidia appear to be common enteric pathogens causing self-limited infections in immune competent hosts (Weber and Bryan 1994; Deplazes et al. 2000). Serosurveys in humans have demonstrated a high prevalence of antibodies to E. cuniculi and E. hellem, suggesting that asymptomatic infection may be common (Bergquist et al. 1984a,b; van Gool et al. 1995). E. intestinalis antibodies were found in 5% of pregnant French women and 8% of Dutch blood donors (van Gool et al. 1995). In the human immunodeficiency virus (HIV) positive Czech patients 5.3% were seropositive to E. cuniculi and 1.3% to E. hellem (Pospisilova et al. 1997). In Slovakia, 5.1% of slaughterhouse workers were seropositive to Encephalitozoon sp (Cislakova et al. 1997). In a survey of blood donors in the US, 5% of donors had antibodies to E. hellem PTP1 antigen (L.M. Weiss, unpublished observations). Overall, these studies suggest that exposure to Microsporidia is common and that asymptomatic infection may be more common than originally suspected.

Cases of microsporidiosis have been identified from all continents except Antarctica (Morakote et al. 1995; van Gool et al. 1995; Brazil et al. 1996; Aoun et al. 1997; Bryan and Schwartz 1999; Deplazes et al. 2000). Human pathogenic Microsporidia have been found in municipal water supplies, tertiary sewage effluent, and ground water consistent with the concept that waterborne transmission occurs (Avery and Undeen 1987; Dowd et al. 1998; Cotte et al. 1999; Graczyk et al. 2007a). In addition, water contact has been found to be an independent risk factor for microsporidiosis in some studies (Enriquez et al. 1998; Hutin et al. 1998). However this has not been a consistent finding in other studies (Wuhib et al. 1994; Conteas et al. 1998a). Microsporidian spores remain viable in water for prolonged periods of time. Spores may be killed, however, by exposure to 70% ethanol, 1% formaldehyde, or 2% Lysol or by autoclaving (Waller 1979; Li and Fayer 2006). Natural transmission of most of these infections occurs when spores are ingested. However, viable spores are present in other body fluids (e.g. stool, urine, respiratory secretions) during infection, suggesting that person-to-person transmission could occur and that ocular infection may be transmitted by external autoinoculation due to contaminated fingers (Schwartz et al. 1993b). It has been possible to transmit E. cuniculi via rectal infection in rabbits, suggesting the possibility of sexual transmission (Fuentealba et al. 1992). Encephalitozoon hellem has been demonstrated in the respiratory mucosa as well as in the prostate and urogenital tract of patients, raising the possibility of respiratory and sexual transmission in humans (Schwartz et al. 1992c; 1994). Person-to-person transmission is supported by concurrent infections in cohabiting homosexual men (Bryan and Schwartz 1999). Transplacental transmission of E. cuniculi has been demonstrated in rabbits delivered by Caesarian section and reared in germ-free isolators (Hunt et al. 1972) and in mice similarly delivered and fostered to germ-free animals (Innes et al. 1962). Similar congential infections have been seen in dogs, horses, alpaca, foxes, and squirrel monkeys (Zeman and Baskin 1985), however, congenital transmission has not yet been demonstrated in humans (Hunt et al. 1972).

It is highly likely that the Microsporidia are zoonotic infections in humans (see Table 49.1) (Mathis et al. 2005). Encephalitozoon are found in many mammals and birds, and the onset of human infection has been associated with exposure to livestock, fowl, and pets (Yee et al. 1991). Encephalitozoon hellem infections have been described in pet birds including budgerigars (parakeets) (Black et al. 1997), and there is a well documented human infection in a patient from pet lovebirds (Yee et al. 1991). Dogs in animal shelters have been demonstrated to excrete microsporidia (Deplazes et al. 2000). Encephalitozoon were found in the stools of many animals in an epidemiologic survey in Mexico (Enriquez et al. 1998). E. cuniculi isolates from various animal species have been identified and separated based on the number of tetranucleotide repeats (5´GTTT3´) in the intergenic spacer region of their rRNA genes and these animal isolates have been found in humans (Didier et al. 1995b). Enterocytozoon bieneusi has been reported in pigs (Deplazes et al. 1996), dogs (del Aguila et al. 1999), chickens (Reetz et al. 2002), pigeons (Graczyk et al. 2007b), falcons, and simian immunodeficiency virus (SIV)-infected rhesus monkeys (Mansfield et al. 1997). There is a documented case of transmission of Ent. bieneusi between a child and guinea pigs (Cama et al. 2007). Differences have also been found in the intergenic spacer region of rRNA genes of Ent. bieneusi and have been used to identify isolates associated with particular animals or environments (Santin and Fayer 2009). Currently all of the different isolates that have been identified are considered to be the same species. However, it is possible that there may be more than one species of Enterocytozoon in its mammalian and avian hosts. Enterocytozoonidae such as Nucleospora (previously Enterocytozoon) salmonis are pathogens found in fish (Kent et al. 1996). Nosema and Vittaforma infections are associated with traumatic inoculation of environmental spores of insect pathogens into the cornea (Shadduck et al. 1990; Silveira and Canning 1995b; Deplazes et al. 1998).

Surveys of pathogens seen in stool samples in Africa, Asia, South America, and Central America have demonstrated that Microsporidia are often found during careful stool examinations. In immune competent hosts, microsporidiosis usually presents as self-limited diarrhoea and both Ent. bieneusi and E. intestinalis are now appreciated to be etiologic agents of traveller’s diarrhoea (Sandfort et al. 1994; Weber and Bryan 1994; Raynaud et al. 1998; Cotte et al. 1999; Wichro et al. 2005). In immune deficient hosts, such as patients with acquired immunodeficiency syndrome (AIDS) or organ transplantation, presentations have been more severe and included diarrhoea with a wasting syndrome as well as disseminated infections, depending on the species of Microsporidia. Reported prevalence rates in the studies conducted on patients with AIDS before the widespread use of active antiretroviral therapy (ART) varied between 2% and 70% depending on the symptoms of the population studied and the diagnostic techniques employed (Weber et al. 1994a; Weiss 1995; Kyaw et al. 1997; Bryan and Schwartz 1999; Deplazes et al. 2000). These studies suggest that asymptomatic carriage can occur in immune compromised patients. Co-infection with different Microsporidia or other enteric pathogens can occur. When combined, these studies identified 375 Ent. bieneusi infections among 2,400 patients with chronic diarrhoea, for a prevalence of 15% in this population. It is clear that since the institution of active antiretroviral therapy and its associated immune reconstitution the prevalence and incidence of microsporidiosis in this population has decreased.

Infection with E. cuniculi in many mammals results in chronic infection with persistently high antibody titres and ongoing inflammation. In immunocompetent murine models of E. cuniculi infection, ascites develops and then clears. However, if corticosteroids are administered, the mice redevelop ascites, consistent with latent persistence of infection (Didier et al. 1994). Cell mediated immunity is important in microsporidian infection. In SCID or athymic mice, infection with E. cuniculi results in death, with visceral dissemination of the organism and persistent ascites (Koudela et al. 1993). Adoptive transfer of sensitized syngeneic T-enriched spleen cells protects athymic or SCID mice against lethal E. cuniculi infection (Schmidt and Shadduck 1984; Hermanek et al. 1993). Interferon g and interleukin-12 are important for protective immunity against a number of microsporidia and other intracellular pathogens (Khan et al. 1999, 2001; Moretto et al. 2001, 2007). Phenotypic analysis of the spleen cells from infected animals revealed an increase in the CD8 T cell population with no significant increase in CD4 T cells. Mice deficient in CD8 cells, but not CD4 succumb to the parasitic challenge. The protective effect of CD8 T cells is mediated by their ability to produce cytokines and to reduce the parasite load by killing the infected targets in the host tissue via the perforin pathway (Khan et al. 1999; Moretto et al. 2000; Khan et al. 2001). Humoral immunity is not sufficient for protection against E. cuniculi infection, as adoptive transfer of immune B lymphocytes into either athymic or SCID mice, or passive transfer of hyperimmune serum into athymic mice does not protect these animals from death after infection. Nonetheless, maternal antibodies protect newborn rabbits from infection with E. cuniculi during the first 2 weeks of life (Bywater and Kellett 1979). Overall, it is probable that antibodies play a role in limiting infection in the host, although they are clearly not sufficient to prevent mortality or to cure infection. There are scant data to confirm the immune response to Microsporidia in humans. It is clear that a strong humoral response occurs during infection and that it includes antibodies that react with the spore wall and polar tube. The immunosuppressive states associated with microsporidiosis (e.g. AIDS and transplantation) are those that inhibit cell-mediated immunity. Microsporidiosis is usually seen in HIV-infected patients when there is a profound defect in cell-mediated immunity (e.g. a CD4 cell count less than 100/mm³); spontaneous cure of microsporidiosis can be induced by immune reconstitution with active antiretroviral therapy (Goguel et al. 1997; Conteas et al. 1998b; Foudraine et al. 1998).

Infection of the small intestine and biliary epithelium is the most frequent presentation of microsporidiosis; with the majority of these infections being due to Ent. bieneusi and the remainder caused by E. intestinalis (Bryan and Schwartz 1999; Deplazes et al. 2000; Weber et al. 2000; Franzen and Muller 2001) (Fig. 49.3). Granulomatous hepatitis due to E. cuniculi is commonly seen in mammals infected with this organism, and granulomatous hepatitis due to Encephalitozoon has been reported in patients with HIV infection (Terada et al. 1987). Hepatitis due to Ent. bieneusi with infection of the biliary system including the portal triad and gallbladder epithelium has been reported in SIV-infected rhesus macques (Schwartz et al. 1998). Ent. bieneusi and E. intestinalis infections of the biliary tract can result in sclerosing cholangitis in AIDS patients (Orenstein et al. 1992b; Pol et al. 1993). Chronic asymptomatic infection of the billiary epithelium due to Ent. bienesui occurs in pigs. Overall, this suggests that biliary epithelium may be a reservoir for relapse of Ent. bieneusi and perhaps other Microsporidia.

Encephalitozoon spp. infects the genitourinary system in most mammals (Weber and Bryan 1994; Gunnarsson et al. 1995; Molina et al. 1995; Schwartz et al. 1996). Urinary shedding of spores is often seen in infections presenting due to symptoms related to damage in other organs. For example, in HIV-infected patients with keratitis there is usually asymptomatic infection of the urinary tract. The most frequent finding in the kidney is granulomatous interstitial nephritis composed of plasma cells and lymphocytes, which is often associated with tubular necrosis, with the lumen of the tubules containing amorphous granular material. Occasionally microabscesses and granulomas form around necrotic tubules. Spores are located in the necrotic tubes, sloughing tubular epithelial cells and occasionally in the interstitium. Glomerular involvement is rarely seen (Guerard et al. 1999; Gumbo et al. 1999a; Latib et al. 2001; Mohindra et al. 2002). Spores shed into the urine can infect other epithelial cells of the urogenital tract resulting in prostatis, necrotizing ureteritis or cystitis (Schwartz et al. 1996). Dissemination can occur from the urogenital epithelial cells into the adjacent mucosal macrophages, muscle, and supporting fibroblasts.

Granulomatous encephalitis is the classic presentation of microsporidiosis in rabbits and is due to Encephalitozoon cuniculi. This infection has been seen in other mammals and has been reported in humans with HIV infection where it was initially mistaken for encephalitis due to Toxoplasma gondii (Weber et al. 1997). Both E. cuniculi type III (dog strain) and type II (rabbit strain) have been reported to cause human encephalitis (Mertens et al. 1997). On autopsy multiple organs were involved, but while spores were seen in the cerebral parenchyma, perivacular spaces, and macrophages they were not present in oligodendrocytes, neurons, astrocytes, or meningeal cells. Other microsporidia can also cause encephalitis and in humans there have been two case reports of encephalitis due to Trachiplestophora anthropopthera (Yachnis et al. 1996; Vavra et al. 1998). Both of these patients had multiple ring-enhancing lesions on computed tomography (CT) scans. On pathologic

examination there was extensive necrosis with 2.0 × 2.8 μm birefringent spores located in the gray matter including astrocytes. Other organs were involved including heart, kidney, pancreas, thyroid, parathyroid, liver, bone marrow, lymph nodes, and spleen.

Punctate keratopathy and conjunctivitis (e.g. superficial epithelial keratitis) has been described with Encephalitzoonidae and Trachipliestophora anthropopthera (Pariyakanok and Jongwutiwes 2005) (Fig. 49.4). In these infections spores are present in corneal and conjunctival epithelium, but do not invade the corneal stroma and there are few associated inflammatory cells. Ocular disease may be the presenting manifestation when there is disseminated infection (Terada et al. 1987; Cali et al. 1991b; Didier et al. 1991; Schwartz et al. 1993a). In a study of patients presenting with keratitis due to E. hellem, spores of this organism were present in many of these patients’ sputum samples in the absence of respiratory symptoms (Schwartz et al. 1992c). These infections have been seen in both immune compromised and immune competent hosts. Most of these cases in immune competent patients have occurred in contact lens wearers, however, epidemic conjuctivitis in India has recently been linked to microsporidiosis (Loh et al. 2009; Reddy et al. 2009a,b). Other species of microsporidia have also been associated with ocular infection, but have involved deeper levels of the corneal stroma. These infections have often been associated with trauma, uveitis and have occurred in immune competent hosts. The species involved have been: Vittaforma corneae, Microsporidium africanum, Microsporidium celonensis, and Nosema ocularum (Joseph et al. 2006; Loh et al. 2009; Reddy et al. 2009a, b). Pathologic changes have included necrosis and acute inflammatory cells with some giant cells in several cases.

Microsporidian myositis with inflammation has been described in humans and has included cases of Pleistophora ronneafiei, Pleistophora sp., Trachipleistophora hominis, Anncaliia vesicularum, and Anncaliia algerae infection (Chupp et al. 1993; Field et al. 1996; Cali et al. 1998; Cali and Takvorian 2003; Coyle et al. 2004). The Pleistophora sp. infections demonstrated atrophic and degenerating muscle fibers infiltrated with focal clusters of large microsporidian spores that were up to 3.4 μm in length associated with a mixed inflammatory response consisting of plasma cells, lymphocytes, eosinophils, and histiocytes (Ledford et al. 1985; Grau et al. 1996; Cali and Takvorian 2003). The T. hominis infection occurred in a patient with AIDS and was associated with degeneration, atrophy, scarring, and intense inflammation (Cali and Takvorian 2003). The A. vesicularum infection occurred in an AIDS patient and was associated with cytolysis around the spores in the muscle fibers, but no cellular immune response was seen (Cali et al. 1998). The A. algerae infection occurred in a patient with rheumatoid arthritis treated with steroids and monoclonal antibody to tumor necrosis factor alpha (TNF-α) (Coyle et al. 2004). There was a minimal cellular response to the numerous spores present in the muscle fibers. Clinically these patients have had myalgias, weakness, elevated serum CPK and aldolase levels, and abnormal electromyography consistent with inflammatory myopathy (Ledford et al. 1985; Chupp et al. 1993; Field et al. 1996; Grau et al. 1996; Cali et al. 1998; Cali and Takvorian 2003).

In Encephalitozoonidae infections respiratory disease has been described with presentations including rhinitis, sinusitis, and nasal polyposis in any combination (Dunand et al. 1997; Gritz et al. 1997; Moss et al. 1997). Encephalitozoonidae spores have been described in the epithelial cells, neutrophils in the bronchiolar wall, cells lining the aveoli, and extracellularly in the alveolar spaces associated with trachetis and bronchiolitis (Schwartz et al. 1992a). All of the Encephalitozoonidae have been associated with chronic sinusitis and spores have been seen in biopsy material in the epithelium and supporting structures (Dunand et al. 1997; Gritz et al. 1997; Moss et al. 1997) associated with a variable inflammatory response including lymphocytes, neutrophils, macrophages, and occasional granuloma formation. There have been two reports of respiratory tract infection and one report of rinosinusitis due to Ent. bieneusi with spores being found in stool, brochoalveolar lavage fluid, and transbrochial biopsy specimens (Weber et al. 1992b; Hartskeerl et al. 1993; del Aguila et al. 1997). These cases may reflect contamination and colonization of the respiratory tract due to vomiting rather than dissemination of this organism from the gastrointestinal tract. A tongue ulcer containing E. cuniculi spores has been reported in a patient with disseminated microsporidiosis (Degroote et al. 1995).

In a child with leukemia, cellulitis was due to A. algerae, with spores infecting all of the cellular elements of the dermis (Visvesvara et al. 1999). In a second case an Encephalitozoon sp. was reported to be the cause of nodular skin lesions (Kester et al. 2000).

Enterocytozoon bieneusi (Desportes et al. 1985) infection results in variable degrees of villous blunting and crypt hyperplasia, but is not invasive. The organism is located on the apical surface of the enterocytes of the small intestine and epithelial cells of the biliary tract and pancreas. Spores are rarely found on the basal surface or in the lamina propria and are associated with increased intraepithelial lymphocytes and epithelial immaturity and disarray (Schwartz et al. 1995b; 1996). Clinical manifestations include watery, nonbloody diarrhoea; nausea; diffuse abdominal pain; and fever. Infection is associated with malabsorption. Diarrhoea is self-limited in immune competent patients, but is persistent in patients with immune suppression. The most common symptom of infection is diarrhoea (Desportes-Livage et al. 1998; Enriquez et al. 1998; Raynaud et al. 1998; Bryan and Schwartz 1999; Rabodonirina et al. 2003) and the presentation classically involves chronic diarrhoea (Weber et al. 1992c), anorexia, weight loss, and bloating without associated fever. It is most frequently seen in AIDS patients with CD4 counts less than 50 cells/mm (Molina et al. 1993a; Weber et al. 1994a, 2000). The mortality of patients with advanced HIV disease and chronic diarrhoea with wasting has been reported to be in excess of 50% (Molina et al. 1993b). Ent. bieneusi has been identified as a cause of self-limited diarrhoea in immune competent hosts including travellers (Bryan and Weber 1993; Sandfort et al. 1994; Weber and Bryan 1994; Raynaud et al. 1998; Bryan and Schwartz 1999; Lopez-Velez et al. 1999; Wichro et al. 2005); and in epidemiologic studies has been identified in 1 to 10% of African children with diarrhoea (Orenstein et al. 1990; Tumwine et al. 2005). Although originally thought to invade only enterocytes, it has been demonstrated that Ent. bieneusi can also invade cholangioepithelium (Pol 1993). When present in the cholangioepithelium, this organism has been associated with sclerosing cholangitis, AIDS cholangiopathy, and cholecystitis (Beaugerie et al. 1992). Presentations include abdominal pain, nausea, vomiting, and fever; jaundice is rarely seen. Infections with Ent. bieneusi have been reported in patients with liver or heart-lung transplantation; and Encephalitozoon sp. infections have been reported in patients with kidney, pancreas, liver, or bone marrow transplantation (Kelkar et al. 1997; Guerard et al. 1999; Gumbo et al. 1999b; Latib et al. 2001; Sing et al. 2001; Mohindra et al. 2002; Rabodonirina et al. 2003).

Spores of Ent. bieneusi are smaller (1.0 × 1.5 μm) than those of Encephalitozoon spp. (1.2 × 2.2 μm) and more difficult to find in tissue sections. Other intestinal pathogens may occur simultaneously or sequentially with the presence of this or other Microsporidia (Hewan-Lowe et al. 1997). A characteristic feature of Ent. bieneusi is the presence of electron-lucent inclusions with a lamellar structure. These inclusions are closely associated with the nuclear envelope, the endoplasmic reticulum, or both. The earliest intraepithelial stages of the parasite are rounded proliferative cells limited by a typical unit membrane in direct contact with the host cell cytoplasm. Nuclear division is not immediately followed by cytokinesis in these cells, resulting in the production of multinucleate proliferative plasmodia. After the production of multiple nuclei, the parasites form electron-dense disk-like structures that cluster in stacks of three to six, eventually forming the coiled portion of the polar tube. When these multinucleated sporagonial plasmodia divide by invagination of the plasmalemma, multiple spores are formed. In mature spores, the polar tubule has five to seven coils that appear in two rows when seen in cross sections by transmission electron microscopy.

Encephalitozoonidae are widely distributed among animals (Didier et al. 2000) and human infection has been caused by E. cuniculi, E. hellem, and E. intestinalis (previously known as Septata intestinalis). The Encephalitozoonidae have the capacity to disseminate widely in their hosts, and involvement in most organs has now been documented (Orenstein et al. 1997; Wittner and Weiss 1999; Weber et al. 2000; Franzen and Muller 2001). All three organisms have been demonstrated to grow in tissue culture and rodent models exist for all of these pathogens. These organisms have been associated with gastroenteritis, keratitis, sinusitis, bronchiolitis, nephritis, cystitis/ureteritis, urethritis, prostatitis, hepatitis, fulminant hepatic failure, peritonitis, cerebritis, nodular skin lesions, corneal lesions and disseminated infection (Wittner and Weiss 1999; Weber et al. 2000; Franzen and Muller 2001; Zender et al. 1989; Mertens et al. 1997; Sheth et al. 1997; Weber et al. 1997; Silverstein 1998; Kester et al. 2000).

Encephalitozoon intestinalis (Cali et al. 1993) is commonly found in the apical and basal sides of infected intestinal enterocytes as well as in cells in the lamina propria, including fibroblasts, endothelial cells, and macrophages (Orenstein et al. 1992a). E. intestinalis-infected cells have a unique parasite-secreted fibrillar network surrounding the developing organisms so the parasitophorous vacuole appears septate. E. intestinalis was found in 7.8% of the stools of patients in a survey regarding the etiology of diarrhoea in Mexico (Enriquez et al. 1998) and has been described in travellers with chronic diarrhoea (Raynaud et al. 1998). The major syndrome associated with E. intestinalis is diarrhoea (van Gool et al. 1994) although it can also cause cholangitis (Cali et al. 1993; Willson et al. 1995), keratoconjunctivitis (Lowder et al. 1996), osteomyelitis of the mandible (Belcher et al. 1997), upper respiratory infections, renal failure, keratoconjunctivitis, and disseminated infection in AIDS patients (Dore et al. 1995; Molina et al. 1995; Schwartz et al. 1996). Elimination of this parasite by treatment with albendazole correlates with the resolution of symptoms (Weber et al. 1994b; Molina et al. 1995). Dissemination can result in necrosis of areas of the bowel, with a presentation resembling an acute abdomen (Orenstein et al. 1997; Soule et al. 1997). E. intestinalis spores are easier to detect than Ent. bieneusi spores because of their larger size, strong birefringence, and bluish color on haematoxylin and eosin staining. Sporogony is tetrasporous, and tubular appendages originate from the sporont surface and terminate in an enlarged bulb-like structure. Mature spores in cross section have a single row of four to seven coils of polar tubules.

Encephalitozoon cuniculi (Levaditti, Nocolau and Schoen 1923) has been associated with hepatitis (Terada et al. 1987), peritonitis (Schwartz et al. 1996), hepatic failure (Sheth et al. 1997), disseminated disease with fever (Mertens et al. 1997), renal insufficiency, and intractable cough (De Groote et al. 1995). Cerebral infections due to E. cuniculi are commonly described in many animals, but have been reported only rarely in immunocompetent humans. Encephalitozoon infection was demonstrated in a 3 year old boy with seizures and hepatomegaly by positive immunoglobulin G (IgG) and IgM indirect immunofluorescence assays using E. cuniculi (Bergquist et al. 1984a). Infection with Encephalitozoon sp. was also reported in a 9 year old Japanese boy with headache, vomiting, and spastic convulsions (Matsubayashi et al. 1959). Several cases of encephalitis and seizures due to E. cuniculi have been reported in AIDS patients (Mertens et al. 1997; Weber et al. 1997). E. hellem and E. cuniculi have similar developmental life cycles (Desportes-Livage 2000). The genus is characterized by the presence of a phagosome-like parasitophorous vacuole, unpaired nuclei, meronts which divide repeatedly by binary fission and sporonts which divide into two sporoblasts that mature into spores. No tubular appendages or fibrillar networks are produced. In cross section the mature spore has five to seven coils in single rows.

Encephalitozoon hellem (Didier et al. 1991) is associated with renal failure, nephritis, pneumonia, bronchitis, disseminated infection, and keratoconjunctivitis (Schwartz et al. 1992a; Weber et al. 1993; Visvesvara et al. 1994). Punctate keratoconjunctivitis is the most common clinical manifestation of E. hellem infection (Lowder et al. 1990; Rastrelli et al. 1994), however, occasionally punctate ketatitis can be due to either E. cuniculi or E. intestinalis (Lowder et al. 1996; Mertens et al. 1997). Clinically, patients have coarse punctate epithelial keratopathy and conjunctival inflammation resulting in redness, foreign body sensation, photophobia, excessive tearing, blurred vision, and changes in visual acuity. This is usually a superficial process restricted to the corneal epithelium and conjunctiva which rarely progresses to corneal ulceration. Infection may be either bilaterial or unilaterial. Slit-lamp examination usually demonstrates punctate epithelial opacities, granular epithelial cells with irregular fluorescein uptake, conjunctival injection, superficial corneal infiltrates, and a non-inflamed anterior chamber. Infection is often associated with disseminated disease (Lacey et al. 1992; Schwartz et al. 1992b; Weber et al. 1993; Degroote et al. 1995; Franzen et al. 1995) and examination of patient’s urine often reveals microsporidian spores (Lacey et al. 1992; Schwartz et al. 1992b; Weber et al. 1993; Degroote et al. 1995; Franzen et al. 1995). E. hellem can also cause infection of the nasal epithelium, presenting as sinusitis (Franzen et al. 1996).

Trachipleistophora hominis (Field et al. 1996; Hollister et al. 1996) is a pansporoblastic microsporidian that has been described in several patients with disseminated disease in the setting of AIDS (Field et al. 1996; Hollister et al. 1996) and can cause myositis, sinusitis, and keratoconjunctivitis. The predominant feature of all these parasites is the presence of a thick surface coat on all stages, which finally separates from the plasma membrane to become an envelope (sporophorous vesicle) enclosing the spores in groups of two to many. It is likely that the Pleistophora sp. described by Chupp et al. (1993) is also T. hominis, but the one seen by Ledford et al. (1985) may be different (Ledford et al. 1985; Chupp et al. 1993). In T. hominis the surface coat extends into lysed host cell cytoplasm as complex networks and merogonic and sporogonic divisions are by repeated binary fissions. The sporophorous vesicle grows to accommodate the increasing number of uninucleate sporoblasts and spores. The spores measure 4.0 × 2.4 μm and have about 11 coils of the polar tube.

Trachipleistophora anthropophthera (Vavra et al. 1997) infection presents as encephalitis, myositis, and keratoconjunctivitis (Yachnis et al. 1996; Vavra et al. 1998; Pariyakanok and Jongwutiwes 2005). Morphologically it differs from Trachipleistophora hominis in that two types of spore are formed. One type, formed in sporophorous vesicles with varying numbers of spores, often 8, measures 3.7 × 2.0 μm (fixed) and has a polar tube of about 7 thick coils and 2 narrower, posterior coils (anisofilar). Spores of the second type, of which only two are formed in each sporophorous vesicle, are nearly spherical 2.2–2.5 × 1.8–2.0 μm. They are thin walled and have only 4–5 isofilar coils of the polar tube (Vavra et al. 1998). This is the first reported occurrence of a dimorphic microsporidium in mammals. Several of these patients responded clinically to albendazole.

Vittaforma corneae (Shadduck et al. 1990) can cause keratitis, urinary tract infection, disseminated infection and prostatitis (Shadduck et al. 1990; Deplazes et al. 1998). This species was isolated into culture from a corneal biopsy and was named Nosema corneum (Shadduck et al. 1990). All stages have diplokaryotic nuclei and are completely surrounded by a cisterna of host endoplasmic reticulum bearing ribosomes on the outer membrane. The cisterna divides with the parasites so that each stage is isolated in its own cisterna. Originally it was thought that sporogony was disporoblastic but Silveira and Canning (1995) demonstrated that the sporonts are multinucleate and divide to produce several linearly arranged sporoblasts (Silveira and Canning 1995b). On the basis of the multisporous sporogony and investment by host endoplasmic reticulum Silveira and Canning (1995) transferred this organism to a new genus as Vittaforma corneae (Silveira and Canning 1995b). Spores measure about 3.7 × 1.0 μm and have 5–7 coils of the polar tube.

Pleistophora ronneafiei (Cali and Takvorian 2003) and Pleistophora sp. have been identified in the skeletal muscle of an HIV-negative patient as well as HIV-positive patients with myositis associated with normal creatine phosphokinase (CPK) levels (Ledford et al. 1985; Chupp et al. 1993; Grau et al. 1996; Cali and Takvorian 2003). In Pleistophora the surface coat is thicker and more uniform, the sporonts are multinucleate plasmodia and the sporophorous vesicle size is fixed by the size of the plasmodium at the onset of sporogony.

Anncaliia algerae (Nosema algerae,  Vavra and Undeen 1970) infection of the skin has been seen in a patient with leukemia (Visvesvara et al. 1999) and in another patient with myositis who had significant elevations in CPK and muscle pain; the latter patient had rheumatoid arthritis treated with steroids and antibody to TNFα (Coyle et al. 2004; Vavra and Undeen 1970). Anncaliia algerae infection of the cornea has also been reported (Visvesvara et al. 1999). Anncalia connori (Sprague 1974) has been described from a single case in an athymic infant. Spores measuring 4.0 × 2.0 μm have nuclei in diplokaryotic arrangement and a polar tube with about 11 coils in a single rank. Anncaliia vesicularum (Brachiola vesicularum, Cali et al. 1998) was reported from a patient with AIDS and myositis. It has diplokaryotic nuclei in all stages. Pre-spore stages have thick surface coats with extensions of complex tubular secretions into lysed host tissue. Spores are 2.5 × 2.0 μm and have about 9 coils of the polar tube in 2 ranks.

Microsporidium sp. and Nosema ocularum (Cali et al. 1991a). Microsporidium is used at the generic level for Microsporidia of unknown phylogenetic placement. In 1973 and 1981, two cases of corneal microsporidiosis due to Microsporidium africanus in Botswana (Pinnolis et al. 1981) and Microsporidium ceylonesis in Sri Lanka (Ashton and Wirasinha 1973) were described. Microsporidium ceylonensis spores, measuring 3.5 × 1.5 μm in fixed tissue were found free and in macrophages in the corneal stroma. These spores had up to 12 coils of the polar tube and there is a single nucleus lying laterally to the posterior region of the polaroplast which is lamellar (Canning and Curry, unpublished observations). Spores lie in direct contact with host cell cytoplasm. Microsporidium africanum (Pinnolis et al. 1981) spores, measured about 4.5 × 3.0 μm, were present mainly in the cytoplasm of histiocytes in the cornea and in direct contact with the stroma and spores had a single nucleus with 11 to 13 coils of the polar tube.

Additional cases of microsporidian keratitis have been identified in immunocompetent hosts (Wittner and Weiss 1999; Joseph et al. 2006). One of these organisms was classified as N. ocularum (Cali et al. 1991a), and the other, which was successfully propagated in vitro, was named N. corneum (Shadduck et al. 1990) (now V. cornea,  Silveira and Canning 1995b). Nosema ocularum was observed in the corneal stroma of the patient, who had experienced visual problems and had a corneal ulcer but was otherwise healthy. Only spores were present, these being distributed in the host cell cytoplasm and in direct contact with it. Spores, measuring about 5.0 × 3.0 μm had diplokaryotic nuclei and 9–12 coils of the polar tube in a single rank. In size and number of coils of the polar tube this parasite resembles M. africanum. Among these immunologically normal patients with corneal infections the outcomes included: enucleation (Pinnolis et al. 1981), unsuccessful penetrating keratoplasty (Ashton and Wirasinha 1973), successful treatment with a corneal transplant (Cali et al. 1991a), and therapy with topical agents until keratoplasty (Davis et al. 1990).

Diagnosis of gastrointestinal microsporidiosis is obtained by light microscopic examination of stool specimens using staining methods that produce differential contrast between the spores of the Microsporidia and the cells and debris in clinical samples in which Microsporidia are found. Visualization requires adequate magnification as spores range in size from 1 to 3 µm (Fig. 49.3). Chromotrope 2R, calcofluor white (fluorescent brightener 28) (Vavra et al. 1993), and Uvitex 2B (van Gool et al. 1993) are useful selective stains for Microsporidia in stool specimens and other body fluids. There are several useful chromotrope 2R-based methods. The technique of Weber et al. (1992a) is a modification of a standard trichrome stain using a 10-fold higher chromotrope 2R concentration and a longer staining time. Ryan et al. (1993) utilizes aniline blue in place of fast green and Kokoskin et al. (1994) use a higher temperature. Staining with chromotrope 2R results in spores which appear light pink with a belt-like stripe girding them diagonally and equatorially against a green (Weber et al. 1992a) or blue (Ryan et al. 1993) background. Microsporidian spores can also be visualized by ultraviolet (UV) microscopy using chemofluorescent optical brightening agents such as Calcofluor white M2R or Uvitex 2B which stain chitin in the spore wall. These chitin stains will also stain fungi and other faecal elements. However, microsporidian spores can be distinguished from yeast as they do not bud. In patients with proven microsporidiosis, both the chromotrope 2R and chemofluorescent brightening stains identified 100% of specimens if at least 50 high power fields were examined (van Gool et al. 1993; Didier et al. 1995a). The sensitivity of chemofluorescent brightener-based stains is slightly higher than chromotrope-based stains (especially when low numbers of spores are present in a sample). However, the specificity of the chemofluorescent stains is lower (Didier et al. 1995a). The limit of detecting Microsporidia by these techniques appears to be 50, 000 organisms/ml (Didier et al. 1995a). Monoclonal antibodies to E. hellem (Croppo et al. 1998), E. intestinalis (Beckers et al. 1996), and Ent. bieneusi (Accoceberry et al. 1999; Sheoran et al. 2005; Singh et al. 2005; Zhang et al. 2005) have been described and can be used for immunofluorescence techniques. Detection kits for microsporidia in stool and environmental samples using antibodies to Encephalitozoonidae and Ent. bieneusi are commercially available (e.g. Waterborne, Inc., New Orleans, LA).

As renal involvement with shedding of spores in the urine is common in Encephalitozoonidae infections and other disseminating Microsporidia, in addition to stool specimens all patients should have urine examinations for Microsporidia. This has therapeutic implications as finding Microsporidia in the urine suggests the infection will be due to a species of Microsporidia that would respond to albendazole. Neither the chromotrope nor the chemofluorescent stain provides information on the species of Microsporidia being identified. Definitive identification of the Microsporidia causing an infection can be done using either ultrastructural examination (e.g. electron microscopy) or molecular techniques (e.g. species-specific polymerase chain reaction (PCR)). If stool examination is negative in the setting of chronic diarrhoea (more than 2 months’ duration), endoscopy should be performed.

Microsporidia in body fluids other than stool (e.g. urine, cerebrospinal fluid), bile, duodenal aspirates, bronchoalveolar lavage fluid, sputum) can be visualized using Chromotrope 2R, chemofluorescent optical brightening agents, Giemsa, Brown-Hopps Gram stain, acid-fast staining, or Warthin-Starry silver staining (Weber et al. 2000; Field 2002). As microsporidian infections usually involve mucosa or epithelium, cytologic preparations, such as corneal swabs, are especially useful for diagnosis (Weber et al. 2000). On histopathological analysis spores are discernible with a modified tissue chromotrope 2R or tissue Gram stain (Brown-Hopp or Brown-Brenn) in tissue sections (Fig. 49.3). Other stains that may be useful include periodic acid-Schiff, Giemsa, and Steiner silver stains. Some Microsporidia are acid-fast stain-positive. Fresh tissue can be examined by phase contrast microscopy as unstained spores are refractile, appearing green; and birefringent. All biopsy or autopsy material should be examined by electron microscopy when microsporidiosis is suspected, as the definitive diagnosis of species is based on ultrastructural information.

Several molecular diagnostic tests have been developed for pathogenic Microsporidia. For a review of the PCR tests for microsporidiosis, see Weiss and Vossbrinck (1998). Over a hundred Microsporidia rRNA sequences are now in the GenBank database allowing the design of PCR primers to identify Microsporidia at the species level in clinical samples without the need for ultrastructural examination. Two main approaches have been employed for constructing PCR primers for Microsporidia: the use of universal pan-Microsporidia primers and of species-specific primer pairs. These PCR techniques have been applied to urine, cultures, and stool specimens (Fedorko et al. 1995; Katzwinkel-Wladarsch et al. 1997b; Ombrouck et al. 1997; Franzen and Muller 1999; Weiss 2000; Joseph et al. 2006). Biopsy specimens can also be analysed using PCR techniques, but this is best done on either unfixed frozen tissue or tissue fixed in ethanol. Currently, these molecular tests are available in reference laboratories such as the Centers for Disease Control (CDC) (Atlanta, GA, USA).

Serologic tests are useful for epidemiologic studies, but have, for the most part, not been proven useful for diagnosing microsporidiosis as infection often occurs in the setting of immune deficiency. For example, in a study of 12 AIDS patients with Ent. bieneusi, 2 AIDS patients with E. intestinalis, and 2 immunocompetent patients with Vit. corneae, enzyme-linked immunosorbent assay (ELISA) titers for E. hellem, E. cuniculi, or Vit. corneae were not useful for diagnosis (Didier 2000). False-negative titers were present in seven of these 14 patients.

The isolation of Microsporidia from clinical specimens is not a routine procedure, but is available in specialized research laboratories. Several of the pathogenic microsporidia including Vittaforma corneae, E. cuniculi, E. hellem, T. hominis, T. anthropopthera, A. anncalia and E. intestinalis have been cultivated in vitro (reviewed by Visvesvara 2002). Small animal models exist for many of these microsporidia. This has enabled these organisms to be used for screening for therapeutic agents (Beauvais et al. 1994; Franssen et al. 1995; Silveira and Canning 1995a). The most common cause of human infection is Ent. bieneusi and this pathogen has not been cultivated continuously in vitro, although limited in vitro cultivation has been reported (Dr Saul Tzipori, personal communication; Visvesvara et al. 1995; Feng et al. 2006). Experimental infection of SIV-infected rhesus monkeys with Ent. bieneusi from human tissue has been demonstrated (Katzwinkel-Wladarsch et al. 1997a) and serial progation has been described in immunocompromised rodents (Feng et al. 2006).

For a review of drugs used against microsporidiosis in humans and animals, see Costa and Weiss (2000). Both fumagillin and albendazole have demonstrated clinical efficacy in human infections with various Microsporidia (see Table 49.2) (Gritz et al. 1997; Molina et al. 1998, 2002; Dore et al. 1995; Dieterich et al. 1994; Corcoran et al. 1996; Didier 1997; Didier et al. 2006). Medications used without success to treat microsporidiosis are azithromycin, paromomycin (Microsporidia lack the rRNA binding site for this drug), and quinacrine. Prophylaxis with trimethoprim-sulfamethoxazole is not effective for preventing microsporidiosis, and this drug has no in vitro or in vivo activity against these organisms (Albrecht et al. 1995). While a few initial case reports indicated that metronidazole was effective, subsequent studies have demonstrated that this drug is not effective against clinical microsporidiosis and there is no activity seen in vitro (Molina et al. 1997; Eeftinck Schattenkerk et al. 1991; Beauvais et al. 1994; Gunnarsson et al. 1995). Atovaquone was reported to have some limited clinical efficacy, but it has no in vitro activity (Beauvais et al. 1994; Anwar-Bruni et al. 1996; Molina et al. 1997). Transient clinical remission of microsporidiosis has been reported with furazolidone and with nitazoxanide (1,000 mg BID) (Schwartz et al. 1995a; Bicart-See et al. 2000). Sparfloxacin and chloroquine have in vitro activity against Microsporidia but have not been used clinically (Beauvais et al. 1994). Thalidomide and octreotide have both been reported to decrease diarrhoea in some patients with microsporidiosis. However, biopsy studies demonstrate that there is no effect on parasite numbers, therefore, the effect on diarrhoea probably is due to direct effects on eneterocytes (Sharpstone et al. 1997).

Albendazole binds to β-tubulin and is active against all of the Encephalitozoonidae (E. hellem, E. cuniculi, E. intestinalis), but does not have consistent activity for treatment of infections due to Enterocytozoon bieneusi (Didier 1997). This is consistent with the sequence of Encephalitozoon β-tubulin genes which have the amino acid residues associated with sensitivity to benzimidazoles (Li et al. 1996), while both Enterocytozoon (Akiyoshi et al. 2007; Akiyoshi et al. 2009) and Vittaforma (Franzen and Salzberger 2008) demonstrate amino acids associated with albendazole resistance. There are numerous case reports demonstrating the efficacy of 2 to 4 weeks of albendazole 400 mg BID for infections due to Encephalitozoonidae. Treatment with albendazole (400 mg BID for 3 weeks) in a double-blind, placebo-controlled trial of eight patients with AIDS and diarrhoea due to E. intestinalis resulted in resolution of the diarrhoea and elimination of the organism in all eight patients, which confirms the activity of albendazole demonstrated in case reports (Weber et al. 1994b; Dore et al. 1995; Gunnarsson et al. 1995; Molina et al. 1995, 1998; Sobottka et al. 1995; Gritz et al. 1997, 1998). For E. hellem infections reported as chronic sinusitis, respiratory infection, and disseminated infection treatment with 400 mg of albendazole twice daily was effective therapy (Lecuit et al. 1994; Visvesvara et al. 1994). Similar efficacy was seen in a patient with disseminated E. cuniculi infection involving the central nervous sysyem (CNS), conjunctiva, sinuses, kidneys, and lungs (Weber et al. 1997). Albendazole has also been reported to be effective in cases of urethritis (Corcoran et al. 1996), renal failure (Aarons et al. 1994), and disseminated infection (Degroote et al. 1995). In addition to its efficacy for the Encephalitozoonidae, disseminated infection accompanied by myositis due to T. hominis and infection with myositis due to Anncaliia vesicularum have both been reported to respond to albendazole (Field et al. 1996; Cali et al. 1998). In contrast, albendazole has displayed only limited efficacy against Ent. bieneusi infection. In two studies examining 66 patients with diarrhoea due to Ent. bieneusi 50% of the albendazole treated patients had some improvement in diarrhoea, but Ent. bieneusi persisted during treatment in all patients and there was no improvement in any patient’s D-xylose absorption test (Blanshard et al. 1991; Dieterich et al. 1994; Lecuit et al. 1994; Sobottka et al. 1995; Li et al. 1996; Molina et al. 1998). Diarrhoea rapidly recurred upon discontinuing albendazole therapy in patients who reported symptom alleviation. Other studies have found that albendazole had no efficacy against Ent. bieneusi infection (Leder et al. 1998).

Fumagillin, isolated from Aspergillus fumigatus, was used to treat amebiasis in the 1950s, but is no longer commercially available for humans. Fumagillin is available commercially for the treatment of microsporidiosis due to Nosema apis in honeybees and has also been used to treat microsporidiosis in aquaculture (Kano and Fukui 1982; Higgins et al. 1998). Fumagillin and its analogues bind in a selective, covalent fashion to the metalloprotease methionine aminopeptidase type 2 (MetAP2). Data from the E. cuniculi genome project (Akiyoshi et al. 2009) indicates that E. cuniculi does not have a methionine aminopeptidase type 1 gene (MetAP1), unlike mammalian cells, which have both MetAP1 and MetAP2; therefore MetAP2 is an essential enzyme in microsporidia. The crystal structure of E. cuniculi MetAP2 has recently been determined (MMDB ID: 63862, PDB ID: 3CMK) (Alvarado et al. 2009). Fumagillin and its analogues such as TNP-470 have significant in vitro and in vivo activity against human pathogenic Microsporidia (Molina et al. 1997, 2002; Didier 1997; Coyle et al. 1998; Didier et al. 2006). A dose-escalation trial of fumagillin performed on AIDS patients with diarrhoea due to Ent. bieneusi has demonstrated that a dose of 60 mg/day for 14 days resulted in over 80% of patients clearing this organism from their stool that was associated with resolution of diarrhoea (Molina et al. 2000). A subsequent randomized trial on 12 patients with either AIDS or transplantation, confirmed that 60 mg/day (given as 20 mg TID) effectively treated Ent. bieneusi intestinal infection with resolution of diarrhoea, clearance of spores and improvement in histology and bowel permeability (Molina et al. 2002). The limiting toxicity of fumagillin was thrombocytopenia, which was reversible on stopping treatment. Microsporidian infection often occurs in immunocompromised hosts, particularly in those with HIV infection and CD4 cell counts of less than 50 mm³. Studies have demonstrated that improvement in immune function can lead to the elimination of Microsporidia and normalization of intestinal architecture (Goguel et al. 1997; Maggi et al. 2000;  Miao et al. 2000). Therefore, part of the primary treatment of microsporidiosis in the setting of AIDS is the institution of effective antiretroviral therapy with restoration of immune function. There has been one report of immune reconstitution syndrome in an AIDS patient with microsporidiosis who was started on antiretroviral therapy.

There are limited data on effective preventive strategies for microsporidiosis. Microsporidian spores can survive and remain infective in the environment for prolonged periods. Experiments with E. cuniculi have demonstrated that they can survive for years in the environment with the correct humidity and temperature (Waller 1979). Spores are rendered noninfectious by a 30-minute exposure to most disinfectants, so the procedures used to clean most hospital rooms should be sufficient to limit infection. Spores are also killed by the commonly used methods employed for sterilization. Hand washing and general hygienic habits probably reduce the chance of contamination of the conjunctiva and cornea with microsporidian spores. The usual sanitary measures that prevent contamination of food and water with animal urine and faeces should decrease the chance for infection by water or foodborne routes, severely immune compromised patients may wish to consider using bottled or filtered water in some settings. Currently, no prophylactic agents have been identified for these organisms and infection has occurred in patients on trimethoprim-sulfamethoxazole prophylaxis (Albrecht et al. 1995), dapsone, pyrimethamine, itraconazole, azithromycin, and atovaquone (Beauvais et al. 1994; Conteas et al. 1998b). Several studies in AIDS patients have demonstrated that antiretroviral therapy with immune restoration can produce remission of intestinal microsporidiosis and has decreased the prevalence and incidence of infection in these patients (Goguel et al. 1997; Foudraine et al. 1998; Maggi et al. 2000; Miao et al. 2000).