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Microsporidia in exotic birds: Intermittent spore excretion of Encephalitozoon spp. in naturally infected budgerigars (Melopsittacus undulatus)
Veterinary Parasitology 168 (2010) 196–200
Contents lists available at ScienceDirect
Veterinary Parasitology journal homepage: www.elsevier.com/locate/vetpar
Microsporidia in exotic birds: Intermittent spore excretion of Encephalitozoon spp. in naturally infected budgerigars (Melopsittacus undulatus) ˇ ova´ b, Oleg Ditrich a,b Bohumil Sak a,*, Denisa Kasˇicˇkova´ b, Martin Kva´cˇ a,c, Dana Kveˇton Institute of Parasitology, Biology Centre of the Academy of Sciences of the Czech Republic, v.v.i., Branisˇovska´ 31, 37005 Cˇeske´ Budeˇjovice, Czech Republic University of South Bohemia in Cˇeske´ Budeˇjovice, Faculty of Biological Science, Branisˇovska´ 31, 37005 Cˇeske´ Budeˇjovice, Czech Republic c University of South Bohemia in Cˇeske´ Budeˇjovice, Faculty of Agriculture, Studentska´ 13, 37005 Cˇeske´ Budeˇjovice, Czech Republic a
b
A R T I C L E I N F O
A B S T R A C T
Article history: Received 14 September 2009 Received in revised form 6 November 2009 Accepted 13 November 2009
Nine naturally infected asymptomatic budgerigars (Melopsittacus undulatus) were screened daily for microsporidia spore excretion during a 30-day period and the localization of infection was evaluated using microscopy and molecular methods. While the microscopic examination revealed 2.4% positivity out of all fecal samples, using PCR the positivity was 10 higher (24.6%). All nine budgerigars excreted microsporidial spores intermittently in irregular intervals with 1–11-day long interruptions. Most of the birds were infected simultaneously with Encephalitozoon cuniculi and Encephalitozoon hellem. While histological and TEM examination failed to confirm the presence of microsporidial spores in tissues, the PCR detected microsporidial DNA mostly in the small intestine, liver and lungs of four selected budgerigars dissected. Despite the chronic infection proved using molecular methods, no clinical signs of disease were observed during monitoring and no pathological findings were found during dissection. ß 2009 Elsevier B.V. All rights reserved.
Keywords: Birds Budgerigars Melopsittacus undulatus Encephalitozoon spp. Natural infection Spore shedding PCR Sequencing
1. Introduction Microsporidia are a diverse group of eukaryotic singlecelled organisms closely related to fungi (Thomarat et al., 2004). They are intracellular ubiquitous parasites that infect a wide range of vertebrate and invertebrate hosts. The phylum Microsporidia contains approximately 150 genera and 1200 species (Keeling and Fast, 2002). Out of eight genera identified in humans to date, Enterocytozoon bieneusi and Encephalitozoon spp. are the most frequent causes of human infection (Weiss, 2003). Moreover, all these species of microsporidia have been detected in birds. Birds were considered to be the primary hosts of Encephalitozoon hellem (Snowden et al., 2001; Barton et al., 2003). In wild birds, E. hellem was found in pigeons, ducks,
* Corresponding author. Tel.: +420 387 775 419; fax: +420 385 310 388. E-mail address: [email protected] (B. Sak). 0304-4017/$ – see front matter ß 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.vetpar.2009.11.012
swans, geese, crows, cranes, puffins and hummingbirds (Tocidlowski et al., 1997; Snowden et al., 2001; Haro et al., 2005; Slodkowicz-Kowalska et al., 2006; Bart et al., 2008). In captive birds, E. hellem was identified in birds from the order Psittaciformes, which includes budgerigars, eclectus parrots, lory, lovebirds and cockatoos, and has also been detected in ostriches and gouldian finches (Black et al., 1997; Gray et al., 1998; Pulparampil et al., 1998; Suter et al., 1998; Snowden and Logan, 1999; Snowden et al., 2000; Carlisle et al., 2002; Barton et al., 2003; Phalen et al., 2006; Slodkowicz-Kowalska et al., 2006). E. hellem is primarily found in the liver, intestine and kidney, but it has also been identified in the eyes, lungs and spleens of infected birds (Poonacha et al., 1985; Black et al., 1997; Pulparampil et al., 1998; Snowden and Logan, 1999; Snowden et al., 2001; Carlisle et al., 2002; Phalen et al., 2006). Less frequently microsporidia spp. identified in birds are listed below. To date, Encephalitozoon cuniculi has been
B. Sak et al. / Veterinary Parasitology 168 (2010) 196–200
found in chickens, cockatiel and recently in pigeons (Reetz, 1993, 1999; Kasˇicˇkova´ et al., 2007; Bart et al., 2008), E. intestinalis in a few cases in pigeons and in geese (Haro et al., 2005; Slodkowicz-Kowalska et al., 2006; Bart et al., 2008) and E. bieneusi in pigeons, grey parrots, cockatiels, lovebirds, finches, chickens and falcons (Haro et al., 2006; Lobo et al., 2006; Graczyk et al., 2007; Bart et al., 2008) Despite the wide host spectrum of avian microsporidia, there is sparse information about the dynamics of microsporidial infection in birds. According to Bart et al. (2008), the prevalence of microsporidia in feces of urban feral pigeons significantly varies during highbreeding (18%) and low-breeding periods (6%). This indicates that either infection with microsporidia is transient and fluctuates in time, or that shedding occurs intermittently. In this study, excretion of microsporidial spores by nine naturally infected budgerigars (Melopsittacus undulatus), originating from a pet shop, were monitored daily using microscopy and molecular methods. 2. Materials and methods 2.1. Specimen collection Nine 2 months old naturally infected male budgerigars (M. undulatus) were selected on the basis of a wide screening of microsporidial infection in exotic birds in the Czech Republic (Kasˇicˇkova´ et al., 2009) and were obtained from a pet store during the summer of 2008. Parasitological examination (coprology, scrapings and histology) did not reveal any other pathogens. Bacteriological, virological and mycological examination of birds were not performed, however, all birds were in good health condition without any apparent signs of infection. Birds were kept in individual cages, placed in separate areas without any contact with other animals. The birds were on commercial seed mixtures and sterile water ad libitum, which were previously confirmed to be Encephalitozoon spp. free using PCR. Fecal samples were collected daily from the cages and screened for the presence of microsporidia during a 30-day long period. Two pairs of birds selected on the basis of frequency of spore shedding were examined at necropsy on day 15 or 30, respectively, and examined using histology and PCR for the presence of microsporidia development stages in the tissues. 2.2. Calcofluor M2R staining (Va´vra and Chalupsky´, 1982) Calcofluor M2R staining was used for the detection of microsporidian spores in feces. Briefly, smears made from fecal samples were fixed with methanol for 2 min and allowed to air dry. Subsequently, the slides were stained with 0.1% calcofluor M2R stain (SIGMA) in phosphate buffered saline (PBS) for 10 min, rinsed gently with PBS, stained with 0.5% Evans Blue stain (SIGMA) in distilled water for 30 s, rinsed again in PBS and allowed to air dry. Stained slides were examined by fluorescence microscopy for the presence of spores using UV light with a wavelength of 490 nm and at a magnification of 1000.
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2.3. DNA isolation Fecal or tissue samples were homogenized by 0.5 mm glass bead disruption and DNA was extracted using commercially available isolation kit QIAamp1 DNA Stool Mini Kit (QIAGEN, Hilden, Germany) or DNeasy Blood & Tissue Kit (QIAGEN, Hilden, Germany) according to the manufacturer’s instructions. Extracted DNA was stored at 20 8C. 2.4. PCR amplification The nested PCR amplification was performed according to Kasˇicˇkova´ et al. (2009) using microsporidia-specific primers described by Katzwinkel-Wladarsch et al. (1996). PCR products were visualized on a 2% agarose gel containing 0.2 mg/ml ethidium bromide. Secondary PCR products were directly sequenced using an ABI Prism Dye Terminator 3.1 cycle sequencing kit on the ABI3730XL sequence analyzer (Applied Biosystems, Foster City, CA). Nucleotide sequences were completed using Chromas Pro v 1.32 (Technelysium, Queenland, Australia), aligned using Custal X (ftp://ftpigbmc.u-strasbg.fr/pub/ClustalX/), manually edited using Bioedit (http://mbio.ncsu.edu/BioEdit/bioedit.html) and compared with sequences in GenBank. As positive controls DNA isolated from the spores of E. intestinalis (3 107 ml 1) originating from AIDS patient (Didier et al., 1996) and grown in vitro in VERO E6 cells and spores of E. bieneusi of genotype D originally isolated from a pig (Sak et al., 2008) were used. 2.5. Histology Tissues from four euthanized and dissected birds including liver, kidney, lung, small intestine, large intestine and caecum were fixed in 10% buffered formalin. The samples were processed by the usual paraffin method. Five-micron sections were routinely stained with haematoxylin and eosin (HE), Brown & Brenn Gram stain (Brown and Brenn, 1931), Weber’s modification of chromotrop (Weber et al., 1992) and Calcofluor M2R (Va´vra and Chalupsky´, 1982). 2.6. Transmission electron microscopy (TEM) For TEM, specimens were fixed in 2.5% glutaraldehyde in cacodylate buffer (0.1 M, pH 7.4) at 4 8C and post-fixed in 1% osmium tetroxide in the same buffer. Fixed samples were washed three times in cacodylate buffer, dehydrated in graded alcohols and embedded in Durcupan resin. Thin sections were stained with uranyl acetate and lead citrate and examined using JEOL 1010 microscope. 3. Results During the 30-day long observation of spore excretion, 240 fecal samples were obtained from budgerigars. Microsporidial spores were microscopically identified in only 11 fecal samples (4.6%) of six individuals (Fig. 1). In contrast, using PCR, microsporidia were detected in 58
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Fig. 1. Frequency of microsporidal spores shedding by naturally infected budgerigars (Melopsittacus undulatus).
fecal samples (24.2%) originating from all nine birds. Microscopically positive fecal samples were also identified as positive using PCR. Each of the nine budgerigars excreted microsporidial spores intermittently in irregular intervals, the frequency of positivity of samples from individual birds varied between 10 and 40% (Fig. 1). Two species of microsporidia were detected in naturally infected budgerigars, E. cuniculi and E. hellem. In six individuals mixed infection of E. hellem 1A genotype and E. cuniculi II genotype (budgerigars number 1, 2, 4, 6, 8, and 9) was observed. Additionally, mixed infection of E. hellem 1A and 3 genotype (budgerigar number 3) was identified and in budgerigars number 7 and 5 monoinfection with E. cuniculi genotype II and E. hellem 1A, respectively, was detected (Fig. 1). All ITS sequences from the study samples were a 100% match to the reference genotypes from GenBank as follows: E. hellem 1A (AF338367), E. hellem 3 (AF110328), E. cuniculi II (GQ422153). The illustrative image of PCR products of different Encephalitozoon spp. genotypes detected in examined birds presents Fig. 2.
Two budgerigars with E. hellem monoinfection were euthanised and dissected on day 15 (number 3 and 5), and the budgerigars with E. hellem an E. cuniculi mixed infection were euthanised and dissected on day 30 (number 2 and 8). Tissue samples of these birds were tested for the presence of microsporidia, but neither histology nor TEM revealed the presence of microsporidial spores. However, using PCR we were able to detect microsporidial DNA in different tissues of all dissected budgerigars (Table 1). Liver samples were the most PCR positive (three animals), lung and large intestine were positive only in two animals and kidney and caecum only in one. While E. cuniculi was identified in kidney, lung, large intestine and caecum, E. hellem was found solely in the liver and lung. No microsporidia were detected in the small intestine. Budgerigars with the most positive tissue samples (#2 and 8) were most often microscopically positive and shed the spores most frequently (9 and 12 days out of 30, respectively) (Fig. 1). Despite the proof of chronic infection using PCR, no clinical signs of disease were observed during monitoring and no pathological findings were found during dissection and histological examination. 4. Discussion
Fig. 2. Gel image of PCR products of genotypes of Encephalitozoon spp. detected in naturally infected budgerigars (Melopsittacus undulatus). Lanes 1 and 10: molecular weight marker (100 bp ladder, Fermentas); lanes 2 and 3: Encephalitozoon hellem genotype 1a; lanes 4 and 5: E. hellem genotype 3; lanes 6 and 7: E. cuniculi genotype II; lane 8: E. intestinalis (positive control); lane 9: negative control (Encephalitozoon-free sample).
Although microsporidia, as a worldwide spread, environmentally resistant opportunistic parasite, represents an important agent of complications in immunodeficient patients, especially AIDS patients, limited data concerning the course of microsporidiosis in naturally infected immunocompetent hosts has been reported. However, published results of epidemiological studies imply high prevalence of asymptomatic microsporidial infections among healthy host populations (Coyle et al., 1996; Tumwine et al., 2005; Samie et al., 2007). Although the repeated spore shedding was already described on the basis of an experimental infection of rabbits with E. cuniculi (Cox et al., 1979), wild-type mice with E. intestinalis (Achbarou et al., 1996) and pigs with E. bieneusi (Breitenmoser et al., 1999), in this study we
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Table 1 Detection of Encephallitozoon spp. in tissues of dissected budgerigars. Bird #
Day of dissection
Encephalitozoon spp. detection Spore shedding frequency in feces
Tissue
Eh 1A (2), Eh 3 (2) Eh 1A (3) Ec II (4), Eh 1A (4) Ec II (10), Eh 1A (2)
Eh 1A Eh 1A Eh 1A
Liver 3 5 2 8
15 15 30 30
Kidney
Lung
Small intestine
Eh 1A Ec II Ec II
Large intestine
Caecum
Ec II Ec II
Ec II
Ec II: Encephalitozoon cuniculi genotype II; Eh 1A: E. hellem genotype 1A; Eh 3: E. hellem genotype 3.
reported for the first time the intermittent microsporidian spore shedding in naturally infected asymptomatic birds during a 30-day period. In accordance with previously reported studies (Rinder et al., 1998; Mungthin et al., 2005), the light microscopy was less sensitive (19% compared to PCR). However, contrary to results of Mungthin et al. (2005), whose specimens from asymptomatic children, which were not positive for E. bieneusi by light microscopy, gave positive results when examined by PCR and the shedding pattern was found to be continuous using PCR, the period between two PCR positive fecal specimens from budgerigars varied from 1 to 11 days. Though none of the tested birds manifested any clinical signs of microsporidiosis, the long period of spore shedding negates the possibility of spore passage only and indicates the chronic course of the infection, which is supported by the positive PCR results obtained from tissue samples of the dissected birds. On the basis of the impossibility to detect the spores in feces microscopically and negative results of histological and TEM examination of dissected birds, the infection intensity was considered to be low, which is in agreement with the results of Miller et al. (2008), who was not able to prove the lethal microsporidiosis in falcons on the basis of several kinds of histological staining despite the evident lesions and subsequently the microsporidiosis was confirmed only by immunohistochemistry. Molecular analyses revealed that most of the examined budgerigars were simultaneously infected with E. hellem and E. cuniculi. As published elsewhere, E. hellem genotype 3 and E. cuniculi genotype II were for the first time described in birds (Kasˇicˇkova´ et al., 2009). Although disseminated infections caused by both detected Encephalitozoon spp. has been described in different hosts (Snowden and Shadduck, 1999), E. hellem in budgerigars had predominant affinity to the liver in this study. In conclusion, our finding, especially the presence of microsporidia in tissues, confirmed the hypothesis about common occurrence of latent microsporidian infections in various hosts. Nevertheless, the role of the asymptomatic hosts as the source of infection is important and might be underrated. Such epidemiological data must be compared with experiments that could solve this question definitely. It was shown, that similarly to other mycoses, risk of symptomatic microsporidiosis is much more dependent on predisposition such as immunological deficiency, than on exposition to the infectious agent because of ubiquitous occurrence of spores.
Acknowledgements This work was supported by the grant of the Academy of Sciences of the Czech Republic (project no. KJB500960701), the Grant Agency of the Czech Republic (project no. 523/ 07/P117) and research project of the Institute of Parasitology, Academy of Sciences of the Czech Republic (Z60220518). References Achbarou, A., Ombrouck, C., Gneragbe, T., Charlotte, F., Renia, L., Desportes Livage, I., Mazier, D., 1996. Experimental model for human intestinal microsporidiosis in interferon gamma receptor knockout mice infected by Encepahlitozoon intestinalis. Parasite Immunol. 18, 387– 392. Bart, A., Wentink-Bonnema, E.M., Heddema, E.R., Buijs, J., van Gool, T., 2008. Frequent occurrence of human-associated microsporidia in fecal droppings of urban pigeons in Amsterdam, the Netherlands. Appl. Environ. Microbiol. 74, 7056–7058. Barton, C.S., Phalen, D.N., Snowden, K.F., 2003. Prevalence of microsporidian spores shed by asymptomatic lovebirds: evidence for a potential emerging zoonosis. J. Avian Med. Surg. 17, 197–202. Black, S.S., Steinohrt, L.A., Bertucci, D.C., Rogers, L.B., Didier, E.S., 1997. Encephalitozoon hellem in budgerigars (Melopsittacus undulatus). Vet. Pathol. 34, 189–198. Breitenmoser, A., Mathis, A., Bu¨rgi, E., Weber, R., Deplazes, P., 1999. High prevalence of Enterocytozoon bieneusi in swine with four genotypes that differ from those identified in humans. Parasitology 118, 447– 453. Brown, J.H., Brenn, L., 1931. A method for the differential and gram negative bacteria in tissue sections. Bull. Johns Hopkins Hosp. 48, 69. ´ Donoghue, P., Prociv, P., Carlisle, M.S., Snowden, K., Gill, J., Jones, M., O 2002. Microsporidiosis in a gouldian finch (Erythrura [Chloebia] gouldiae). Aust. Vet. J. 80, 41–44. Cox, J.C., Hamilton, R.C., Attwood, H.D., 1979. An investigation of the route and progression of Encephalitozoon cuniculi infection in adult rabbits. J. Protozool. 26, 260–265. Coyle, C.M., Wittner, M., Kotler, D.P., Noyer, C., Orenstein, J.M., Tanowitz, H.B., Weiss, L.H., 1996. Prevalence of microsporidiosis due to Enterocytozoon bieneusi and Encephalitozoon (Septata) intestinalis among patients with AIDS-related diarrhea: determination by polymerase chain reaction to the microsporidian small-subunit rRNA gene. Clin. Infect. Dis. 23, 1002–1006. Didier, E.S., Rogers, L.B., Orenstein, J.M., Baker, M.D., Vossbrinck, C.R., Van Gool, T., Hartskeerl, R., Soave, R., Beaudet, L.M., 1996. Characterization of Encephalitozoon (Septata) intestinalis isolates cultured from nasal mucosa and bronchoalveolar lavage fluids of two AIDS patients. J. Eukaryot. Microbiol. 43, 34–43. Graczyk, T.K., Sunderland, D., Rule, A.M., da Silva, A.J., Moura, I.N.S., Tamang, L., Girouard, A.S., Schwab, K.J., Breysse, P.N., 2007. Urban feral pigeons (Columba livia) as a source for air- and waterborne contamination with Enterocytozoon bieneusi spores. Appl. Environ. Microbiol. 73, 4357–4358. Gray, M.L., Puette, M., Latimer, K.S., 1998. Microsporidiosis in a young ostrich (Struthio camelus). Avian Dis. 42, 832–836. Haro, M., Henriques-Gil, N., Fenoy, S., Izquierdo, F., Alonso, F., del A´guila, C., 2006. Detection and genotyping of Enterocytozoon bieneusi in pigeons. J. Eukaryot. Microbiol. 53, S58–S60. Haro, M., Izquierdo, F., Henriques-Gil, N., Andre´s, I., Alonso, F., Fenoy, S., del A´guila, C., 2005. First detection and genotyping of human-associ-
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Microsporidia in exotic birds: Intermittent spore excretion of Encephalitozoon spp. in naturally infected budgerigars (Melopsittacus undulatus) ˇ ova´ b, Oleg Ditrich a,b Bohumil Sak a,*, Denisa Kasˇicˇkova´ b, Martin Kva´cˇ a,c, Dana Kveˇton Institute of Parasitology, Biology Centre of the Academy of Sciences of the Czech Republic, v.v.i., Branisˇovska´ 31, 37005 Cˇeske´ Budeˇjovice, Czech Republic University of South Bohemia in Cˇeske´ Budeˇjovice, Faculty of Biological Science, Branisˇovska´ 31, 37005 Cˇeske´ Budeˇjovice, Czech Republic c University of South Bohemia in Cˇeske´ Budeˇjovice, Faculty of Agriculture, Studentska´ 13, 37005 Cˇeske´ Budeˇjovice, Czech Republic a
b
A R T I C L E I N F O
A B S T R A C T
Article history: Received 14 September 2009 Received in revised form 6 November 2009 Accepted 13 November 2009
Nine naturally infected asymptomatic budgerigars (Melopsittacus undulatus) were screened daily for microsporidia spore excretion during a 30-day period and the localization of infection was evaluated using microscopy and molecular methods. While the microscopic examination revealed 2.4% positivity out of all fecal samples, using PCR the positivity was 10 higher (24.6%). All nine budgerigars excreted microsporidial spores intermittently in irregular intervals with 1–11-day long interruptions. Most of the birds were infected simultaneously with Encephalitozoon cuniculi and Encephalitozoon hellem. While histological and TEM examination failed to confirm the presence of microsporidial spores in tissues, the PCR detected microsporidial DNA mostly in the small intestine, liver and lungs of four selected budgerigars dissected. Despite the chronic infection proved using molecular methods, no clinical signs of disease were observed during monitoring and no pathological findings were found during dissection. ß 2009 Elsevier B.V. All rights reserved.
Keywords: Birds Budgerigars Melopsittacus undulatus Encephalitozoon spp. Natural infection Spore shedding PCR Sequencing
1. Introduction Microsporidia are a diverse group of eukaryotic singlecelled organisms closely related to fungi (Thomarat et al., 2004). They are intracellular ubiquitous parasites that infect a wide range of vertebrate and invertebrate hosts. The phylum Microsporidia contains approximately 150 genera and 1200 species (Keeling and Fast, 2002). Out of eight genera identified in humans to date, Enterocytozoon bieneusi and Encephalitozoon spp. are the most frequent causes of human infection (Weiss, 2003). Moreover, all these species of microsporidia have been detected in birds. Birds were considered to be the primary hosts of Encephalitozoon hellem (Snowden et al., 2001; Barton et al., 2003). In wild birds, E. hellem was found in pigeons, ducks,
* Corresponding author. Tel.: +420 387 775 419; fax: +420 385 310 388. E-mail address: [email protected] (B. Sak). 0304-4017/$ – see front matter ß 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.vetpar.2009.11.012
swans, geese, crows, cranes, puffins and hummingbirds (Tocidlowski et al., 1997; Snowden et al., 2001; Haro et al., 2005; Slodkowicz-Kowalska et al., 2006; Bart et al., 2008). In captive birds, E. hellem was identified in birds from the order Psittaciformes, which includes budgerigars, eclectus parrots, lory, lovebirds and cockatoos, and has also been detected in ostriches and gouldian finches (Black et al., 1997; Gray et al., 1998; Pulparampil et al., 1998; Suter et al., 1998; Snowden and Logan, 1999; Snowden et al., 2000; Carlisle et al., 2002; Barton et al., 2003; Phalen et al., 2006; Slodkowicz-Kowalska et al., 2006). E. hellem is primarily found in the liver, intestine and kidney, but it has also been identified in the eyes, lungs and spleens of infected birds (Poonacha et al., 1985; Black et al., 1997; Pulparampil et al., 1998; Snowden and Logan, 1999; Snowden et al., 2001; Carlisle et al., 2002; Phalen et al., 2006). Less frequently microsporidia spp. identified in birds are listed below. To date, Encephalitozoon cuniculi has been
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found in chickens, cockatiel and recently in pigeons (Reetz, 1993, 1999; Kasˇicˇkova´ et al., 2007; Bart et al., 2008), E. intestinalis in a few cases in pigeons and in geese (Haro et al., 2005; Slodkowicz-Kowalska et al., 2006; Bart et al., 2008) and E. bieneusi in pigeons, grey parrots, cockatiels, lovebirds, finches, chickens and falcons (Haro et al., 2006; Lobo et al., 2006; Graczyk et al., 2007; Bart et al., 2008) Despite the wide host spectrum of avian microsporidia, there is sparse information about the dynamics of microsporidial infection in birds. According to Bart et al. (2008), the prevalence of microsporidia in feces of urban feral pigeons significantly varies during highbreeding (18%) and low-breeding periods (6%). This indicates that either infection with microsporidia is transient and fluctuates in time, or that shedding occurs intermittently. In this study, excretion of microsporidial spores by nine naturally infected budgerigars (Melopsittacus undulatus), originating from a pet shop, were monitored daily using microscopy and molecular methods. 2. Materials and methods 2.1. Specimen collection Nine 2 months old naturally infected male budgerigars (M. undulatus) were selected on the basis of a wide screening of microsporidial infection in exotic birds in the Czech Republic (Kasˇicˇkova´ et al., 2009) and were obtained from a pet store during the summer of 2008. Parasitological examination (coprology, scrapings and histology) did not reveal any other pathogens. Bacteriological, virological and mycological examination of birds were not performed, however, all birds were in good health condition without any apparent signs of infection. Birds were kept in individual cages, placed in separate areas without any contact with other animals. The birds were on commercial seed mixtures and sterile water ad libitum, which were previously confirmed to be Encephalitozoon spp. free using PCR. Fecal samples were collected daily from the cages and screened for the presence of microsporidia during a 30-day long period. Two pairs of birds selected on the basis of frequency of spore shedding were examined at necropsy on day 15 or 30, respectively, and examined using histology and PCR for the presence of microsporidia development stages in the tissues. 2.2. Calcofluor M2R staining (Va´vra and Chalupsky´, 1982) Calcofluor M2R staining was used for the detection of microsporidian spores in feces. Briefly, smears made from fecal samples were fixed with methanol for 2 min and allowed to air dry. Subsequently, the slides were stained with 0.1% calcofluor M2R stain (SIGMA) in phosphate buffered saline (PBS) for 10 min, rinsed gently with PBS, stained with 0.5% Evans Blue stain (SIGMA) in distilled water for 30 s, rinsed again in PBS and allowed to air dry. Stained slides were examined by fluorescence microscopy for the presence of spores using UV light with a wavelength of 490 nm and at a magnification of 1000.
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2.3. DNA isolation Fecal or tissue samples were homogenized by 0.5 mm glass bead disruption and DNA was extracted using commercially available isolation kit QIAamp1 DNA Stool Mini Kit (QIAGEN, Hilden, Germany) or DNeasy Blood & Tissue Kit (QIAGEN, Hilden, Germany) according to the manufacturer’s instructions. Extracted DNA was stored at 20 8C. 2.4. PCR amplification The nested PCR amplification was performed according to Kasˇicˇkova´ et al. (2009) using microsporidia-specific primers described by Katzwinkel-Wladarsch et al. (1996). PCR products were visualized on a 2% agarose gel containing 0.2 mg/ml ethidium bromide. Secondary PCR products were directly sequenced using an ABI Prism Dye Terminator 3.1 cycle sequencing kit on the ABI3730XL sequence analyzer (Applied Biosystems, Foster City, CA). Nucleotide sequences were completed using Chromas Pro v 1.32 (Technelysium, Queenland, Australia), aligned using Custal X (ftp://ftpigbmc.u-strasbg.fr/pub/ClustalX/), manually edited using Bioedit (http://mbio.ncsu.edu/BioEdit/bioedit.html) and compared with sequences in GenBank. As positive controls DNA isolated from the spores of E. intestinalis (3 107 ml 1) originating from AIDS patient (Didier et al., 1996) and grown in vitro in VERO E6 cells and spores of E. bieneusi of genotype D originally isolated from a pig (Sak et al., 2008) were used. 2.5. Histology Tissues from four euthanized and dissected birds including liver, kidney, lung, small intestine, large intestine and caecum were fixed in 10% buffered formalin. The samples were processed by the usual paraffin method. Five-micron sections were routinely stained with haematoxylin and eosin (HE), Brown & Brenn Gram stain (Brown and Brenn, 1931), Weber’s modification of chromotrop (Weber et al., 1992) and Calcofluor M2R (Va´vra and Chalupsky´, 1982). 2.6. Transmission electron microscopy (TEM) For TEM, specimens were fixed in 2.5% glutaraldehyde in cacodylate buffer (0.1 M, pH 7.4) at 4 8C and post-fixed in 1% osmium tetroxide in the same buffer. Fixed samples were washed three times in cacodylate buffer, dehydrated in graded alcohols and embedded in Durcupan resin. Thin sections were stained with uranyl acetate and lead citrate and examined using JEOL 1010 microscope. 3. Results During the 30-day long observation of spore excretion, 240 fecal samples were obtained from budgerigars. Microsporidial spores were microscopically identified in only 11 fecal samples (4.6%) of six individuals (Fig. 1). In contrast, using PCR, microsporidia were detected in 58
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Fig. 1. Frequency of microsporidal spores shedding by naturally infected budgerigars (Melopsittacus undulatus).
fecal samples (24.2%) originating from all nine birds. Microscopically positive fecal samples were also identified as positive using PCR. Each of the nine budgerigars excreted microsporidial spores intermittently in irregular intervals, the frequency of positivity of samples from individual birds varied between 10 and 40% (Fig. 1). Two species of microsporidia were detected in naturally infected budgerigars, E. cuniculi and E. hellem. In six individuals mixed infection of E. hellem 1A genotype and E. cuniculi II genotype (budgerigars number 1, 2, 4, 6, 8, and 9) was observed. Additionally, mixed infection of E. hellem 1A and 3 genotype (budgerigar number 3) was identified and in budgerigars number 7 and 5 monoinfection with E. cuniculi genotype II and E. hellem 1A, respectively, was detected (Fig. 1). All ITS sequences from the study samples were a 100% match to the reference genotypes from GenBank as follows: E. hellem 1A (AF338367), E. hellem 3 (AF110328), E. cuniculi II (GQ422153). The illustrative image of PCR products of different Encephalitozoon spp. genotypes detected in examined birds presents Fig. 2.
Two budgerigars with E. hellem monoinfection were euthanised and dissected on day 15 (number 3 and 5), and the budgerigars with E. hellem an E. cuniculi mixed infection were euthanised and dissected on day 30 (number 2 and 8). Tissue samples of these birds were tested for the presence of microsporidia, but neither histology nor TEM revealed the presence of microsporidial spores. However, using PCR we were able to detect microsporidial DNA in different tissues of all dissected budgerigars (Table 1). Liver samples were the most PCR positive (three animals), lung and large intestine were positive only in two animals and kidney and caecum only in one. While E. cuniculi was identified in kidney, lung, large intestine and caecum, E. hellem was found solely in the liver and lung. No microsporidia were detected in the small intestine. Budgerigars with the most positive tissue samples (#2 and 8) were most often microscopically positive and shed the spores most frequently (9 and 12 days out of 30, respectively) (Fig. 1). Despite the proof of chronic infection using PCR, no clinical signs of disease were observed during monitoring and no pathological findings were found during dissection and histological examination. 4. Discussion
Fig. 2. Gel image of PCR products of genotypes of Encephalitozoon spp. detected in naturally infected budgerigars (Melopsittacus undulatus). Lanes 1 and 10: molecular weight marker (100 bp ladder, Fermentas); lanes 2 and 3: Encephalitozoon hellem genotype 1a; lanes 4 and 5: E. hellem genotype 3; lanes 6 and 7: E. cuniculi genotype II; lane 8: E. intestinalis (positive control); lane 9: negative control (Encephalitozoon-free sample).
Although microsporidia, as a worldwide spread, environmentally resistant opportunistic parasite, represents an important agent of complications in immunodeficient patients, especially AIDS patients, limited data concerning the course of microsporidiosis in naturally infected immunocompetent hosts has been reported. However, published results of epidemiological studies imply high prevalence of asymptomatic microsporidial infections among healthy host populations (Coyle et al., 1996; Tumwine et al., 2005; Samie et al., 2007). Although the repeated spore shedding was already described on the basis of an experimental infection of rabbits with E. cuniculi (Cox et al., 1979), wild-type mice with E. intestinalis (Achbarou et al., 1996) and pigs with E. bieneusi (Breitenmoser et al., 1999), in this study we
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Table 1 Detection of Encephallitozoon spp. in tissues of dissected budgerigars. Bird #
Day of dissection
Encephalitozoon spp. detection Spore shedding frequency in feces
Tissue
Eh 1A (2), Eh 3 (2) Eh 1A (3) Ec II (4), Eh 1A (4) Ec II (10), Eh 1A (2)
Eh 1A Eh 1A Eh 1A
Liver 3 5 2 8
15 15 30 30
Kidney
Lung
Small intestine
Eh 1A Ec II Ec II
Large intestine
Caecum
Ec II Ec II
Ec II
Ec II: Encephalitozoon cuniculi genotype II; Eh 1A: E. hellem genotype 1A; Eh 3: E. hellem genotype 3.
reported for the first time the intermittent microsporidian spore shedding in naturally infected asymptomatic birds during a 30-day period. In accordance with previously reported studies (Rinder et al., 1998; Mungthin et al., 2005), the light microscopy was less sensitive (19% compared to PCR). However, contrary to results of Mungthin et al. (2005), whose specimens from asymptomatic children, which were not positive for E. bieneusi by light microscopy, gave positive results when examined by PCR and the shedding pattern was found to be continuous using PCR, the period between two PCR positive fecal specimens from budgerigars varied from 1 to 11 days. Though none of the tested birds manifested any clinical signs of microsporidiosis, the long period of spore shedding negates the possibility of spore passage only and indicates the chronic course of the infection, which is supported by the positive PCR results obtained from tissue samples of the dissected birds. On the basis of the impossibility to detect the spores in feces microscopically and negative results of histological and TEM examination of dissected birds, the infection intensity was considered to be low, which is in agreement with the results of Miller et al. (2008), who was not able to prove the lethal microsporidiosis in falcons on the basis of several kinds of histological staining despite the evident lesions and subsequently the microsporidiosis was confirmed only by immunohistochemistry. Molecular analyses revealed that most of the examined budgerigars were simultaneously infected with E. hellem and E. cuniculi. As published elsewhere, E. hellem genotype 3 and E. cuniculi genotype II were for the first time described in birds (Kasˇicˇkova´ et al., 2009). Although disseminated infections caused by both detected Encephalitozoon spp. has been described in different hosts (Snowden and Shadduck, 1999), E. hellem in budgerigars had predominant affinity to the liver in this study. In conclusion, our finding, especially the presence of microsporidia in tissues, confirmed the hypothesis about common occurrence of latent microsporidian infections in various hosts. Nevertheless, the role of the asymptomatic hosts as the source of infection is important and might be underrated. Such epidemiological data must be compared with experiments that could solve this question definitely. It was shown, that similarly to other mycoses, risk of symptomatic microsporidiosis is much more dependent on predisposition such as immunological deficiency, than on exposition to the infectious agent because of ubiquitous occurrence of spores.
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