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Cyst formation and faecal–oral transmission of Dientamoeba fragilis – the missing link in the life cycle of an emerging pathogen
International Journal for Parasitology 43 (2013) 879–883
Contents lists available at SciVerse ScienceDirect
International Journal for Parasitology journal homepage: www.elsevier.com/locate/ijpara
Succinctus
Cyst formation and faecal–oral transmission of Dientamoeba fragilis – the missing link in the life cycle of an emerging pathogen Varuni S. Munasinghe a, Nicole G.F. Vella b, John T. Ellis a,⇑, Peter A. Windsor c, Damien Stark a,d a
School of Medical and Molecular Biosciences and the i3 Institute, University of Technology Sydney, Broadway, New South Wales 2007, Australia Microscopy Unit, Faculty of Science, Macquarie University, North Ryde, New South Wales 2109, Australia c Faculty of Veterinary Science, University of Sydney, Camden, New South Wales 2560, Australia d Division of Microbiology, Sydpath, St. Vincent’s Hospital, Darlinghurst, New South Wales 2010, Australia b
a r t i c l e
i n f o
Article history: Received 6 May 2013 Received in revised form 23 June 2013 Accepted 24 June 2013 Available online 19 July 2013 Keywords: Dientamoeba Trichomonad Transmission Cyst Diarrhoea Irritable bowel
a b s t r a c t Dientamoeba fragilis is a protozoan parasite emerging as a cause of diarrhoea and ‘‘irritable-bowel-like’’ gastrointestinal disease in humans with a propensity for establishing long-term, chronic infections in humans. Although Dientamoeba was discovered over a century ago its life cycle and mode of transmission is not known. No cyst stage has been described and no animal models are presently available for the study of this parasite. Here we describe the establishment of an animal model using laboratory rodents, the fulfilling of Koch’s postulates, and the discovery of a new cyst stage in the life cycle of D. fragilis. Our demonstration of long-term parasite carriage by rodents and prolonged shedding of cysts, together with elevated levels of calprotectin in the stool, confirms the capacity of this organism to cause disease and indicates dientamoebiasis should be considered in the differential diagnosis of gastrointestinal diseases such as Inflammatory Bowel Syndrome (IBS). Finally, we suggest that the cyst stage described here is the vehicle that mediates faecal–oral transmission of D. fragilis between hosts. Ó 2013 Australian Society for Parasitology Inc. Published by Elsevier Ltd. All rights reserved.
Dientamoeba fragilis is increasingly recognised as a relatively common cause of human diarrhoea and long-term chronic infections are a feature of infection by this parasite (Stark et al., 2010b). The mode of transmission and life cycle of D. fragilis has yet to be definitively described (Stark et al., 2006; Barratt et al., 2011). It was initially suggested that D. fragilis was transmitted via the ova of nematodes (Dobell, 1940). Reports of a higher than anticipated rate of co-infection between D. fragilis and Enterobius vermicularis led others to postulate pin worm may be a probable vector of transmission (Girginkardesler et al., 2008; Roser et al., 2013). In contrast others have demonstrated no co-infections with D. fragilis and helminths (Stark et al., 2010b). The absence of strong evidence in support of transmission by helminths, coupled with high rates of co-infection with other faecal orally transmitted bacteria and protozoa, would suggest faecal–oral transmission is the most likely mechanism of D. fragilis transmission (Stark et al., 2005b). Here we present a study on the mode of transmission of Dientamoeba using rodents and report the discovery of a new cyst stage in the life cycle of D. fragilis. All animal experiments were approved by the University of Technology, Sydney (UTS) Australia Animal Care and Ethics Committee. BALB/c mice and Sprague–Dawley rats were obtained from
⇑ Corresponding author. Tel.: +61 2 9514 4161; fax: +61 2 9514 8206. E-mail address: [email protected] (J.T. Ellis).
the Animal Resource Centre (Perth, Australia) housed in independently vented, filter-top cages and provided sterile rodent chow and water ad libitum. The cage bedding was changed every day to avoid and reduce the potential for faecal contamination occurring during the experiment. All animals were confirmed as specific pathogen-free by microscopy and PCR. Animal stool fixed in sodium acetate–acetic acid–formalin (SAF) was examined daily for 1 week prior to infection by D. fragilis for the presence of parasites using permanent iron haematoxylin stained smears. DNA was also extracted from frozen, unfixed stool using the Bioline stool minikit (Bioline, catalogue No. BIO-52037, USA) according to the manufacturer’s recommendations. PCR amplification of stool DNA targeting the D. fragilis ssrDNA was performed as described previously (Stark et al., 2005a). The D. fragilis isolate 4 (Barratt et al., 2010) was cultured on serum slopes with a PBS overlay (Munasinghe et al., 2012). BALB/c mice were orally infected with cultured trophozoites (doses ranging from 103–109/mouse; groups of two) and stools were monitored for the presence of parasites p.i. As a control group a similar number of mice were orally administered with a suspension containing bacteria from the D. fragilis in vitro culture. In order to prepare bacterial cultures for use as controls and to ensure the absence of D. fragilis trophozoites in the preparation, the D. fragilis culture overlay was plated onto blood agar. All of the bacteria isolated as colonies were uniformly returned to Loeffler’s media and incubated anaerobically at 37 °C overnight. All inocula were
0020-7519/$36.00 Ó 2013 Australian Society for Parasitology Inc. Published by Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.ijpara.2013.06.003
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Fig. 1. Iron haematoxylin stained Dientamoeba fragilis cysts present in the feces of infected mice from day 1 p.i. (A) Binucleated (n) D. fragilis cyst from mice infected with D. fragilis. Note the distinct cyst wall (w) and the zone of clearance (c) around the cyst. (B) Dientamoeba fragilis cyst from mice infected with feces of infected mice to establish Koch’s postulates. (C and D) Dientamoeba fragilis cysts present in the feces of rats. Scale bar = 5 lm.
centrifuged at 10,000g (bacteria) or 1,000g (D. fragilis) for 15 min and then resuspended in Earle’s Balanced Salt Solution (EBSS) for administration orally to mice. Post infection all mice were individually weighed daily. To test for significant differences in weight loss between the infected and the control groups of mice a student’s paired t-test was used to calculate P values in SPSS version 19. After day 1 p.i. all mice infected with D. fragilis were intermittently shedding binucleated cysts that measured 5 lm in diameter
as shown in the iron haematoxylin smears (Fig. 1A). Mice infected with D. fragilis showed an average weight loss of 2.38 g (12%) compared with the control group (P < 0.05) from day 1 through to day 6. At this point in time the mean group weights became similar. The infected group also showed symptoms of gastrointestinal disturbance with mice excreting unformed faeces intermittently, compared with the control group which were asymptomatic and produced normally formed stools during the experimental period. In addition white blood cells were observed in the stool smears
Fig. 2. Transmission electron micrographs of Dientamoeba fragilis cysts showing the cyst wall and the encysted parasite. (A) Transverse section across a whole cyst showing the outer fibrillar cyst wall (o), peritrophic space (ps) and the enclosed parasite (p). (B) Fibrillar dense outer cyst wall (o) contains a fine meshwork of fibrils. (C) Inner membrane of the cyst wall (i) which is double layered, enclosing the parasite. Note the peritrophic space (ps) and the Encystation Specific Vesicles (v) scattered beneath the outer cyst wall. (D) Transmission electron micrograph showing the double membrane-bound Encystation Specific Vesicles (v) scattered throughout the peritrophic space (ps). Note the inner membrane of the cyst wall (i) enclosing the parasite. Scale bar = 200 nm.
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Fig. 3. Transmission electron micrographs of Dientamoeba fragilis cysts showing different organelles in the encysted parasite. (A) Transverse section across a mononucleated cyst showing the cyst wall (w), nucleus (nu), and the nucleolus (arrowhead). Note the peritrophic space (ps) and the encysted parasite (p). (B) Transverse section across a binucleated cyst showing the peritrophic space (ps), cyst wall (w), nuclei (nu), nuclear membrane (nm) and the nucleolus (arrowheads). (C) Transverse section across hydrogenosomes (h). (D) Longitudinal section across the axostyle showing the microtubules (a). (E) Transverse section across a flagellar axoneme (ax) with the 9 + 2 microtubule arrangement. (F) longitudinal section across the flagellar axonemes (f). (G) Longitudinal section across the pelta (pe). (H) Longitudinal section across the costa (c). Scale bars = 1 lm for A and B; 100 nm for (C–H).
from D. fragilis-infected mice. Real time PCR also confirmed the presence of D. fragilis DNA in stool (Stark et al., 2010a). To study the duration of shedding of the organism, BALB/c mice (n = 16) were infected orally with doses of D. fragilis (as low as 600 trophozoites/mouse) and observed for a period of 6 months. All of the infected mice shed D. fragilis intermittently and
continued shedding for 6 months p.i. (Fig. 1B), while the control group (n = 2) remained uninfected. Subsequently two naïve, uninfected mice were infected orally with a suspension of mouse feces containing cysts. Iron haematoxylin smears (and subsequent PCR) confirmed the shedding of D. fragilis in these two mice from day 1 p.i. These cysts were morphologically similar to the cysts observed
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previously in mouse stool (Fig. 1A). The infection in these two mice became chronic and lasted 4 months, when the experiment was terminated. Mice were euthanised by carbon dioxide asphyxiation. Small intestine and large intestine from each mouse were collected, fixed in 4% paraformaldehyde and embedded in histowax. Sections (5 lm) were stained with H&E and the slides were viewed using an Olympus light microscope under 60 objective. Histopathological examination of the large intestine of mice infected with D. fragilis showed minor pathological changes with slight inflammation of the submucosa present. Levels of calprotectin, a marker of inflammatory and neoplastic disease in the lower gastrointestinal tract (Tøn et al., 2000), were determined in stool by ELISA over the period 28 days p.i. Stools (100 mg) were homogenised in extraction buffer (weight:volume 1:50) and faecal calprotectin levels were measured using an ELISA (Alpco Diagnostics, USA). The average calprotectin levels present in stool from D. fragilis infected mice (69 ng/ml ± 22.22 SD) was twice that of control mice (33 ng/ ml ± 13.24 SD) (P < 0.05), indicative of the presence of intestinal inflammation. Sprague–Dawley rats were also investigated as an alternative host. Two rats that were administered with trophozoites did not become infected during the course of the experiment. Four rats given a suspension of D. fragilis cysts from infected mice became infected and shed Dientamoeba cysts intermittently from days 5 through to 26 p.i. These cysts were morphologically similar to the cysts that were present in the mouse feces with a distinct cyst wall and a diameter varying between 5 and 6 lm (Fig. 1C and D). This infected group exhibited a statistically significant (P < 0.05) mean weight loss of 7.05 g (4%) compared with the control group over days 1 to 28 p.i. with the main period of weight loss being days 12 through to 22 p.i. Infected rats also produced unformed stools and had white blood cells in their feces. A total number of 10 stool samples were screened by transmission electron microscopy for cysts. Feces from infected mice and rats were collected and fixed in 3% glutaraldehyde in PBS (pH 7.2), embedded in 2% low melt agarose and fixed, stained and infiltrated with osmium tetroxide, uranyl acetate and LR White Resin, respectively, by standard procedures. Semi-thin (1 lm) and ultrathin (70 nm) sections were cut using an ultramicrotome and mounted on 200 mesh copper grids (ProScitech, Australia). Ultrathin sections were stained with saturated aqueous uranyl acetate (7.7%) and Reynold’s lead citrate (Reynolds, 1963). Semi-thin sections were stained with 1% methylene blue in 0.6% sodium bicarbonate/40% glycerol and viewed using an Olympus BH2 light microscope. Ultra-thin sections were examined using a Philips CM10 transmission electron microscope and images captured using a Megaview G2 digital camera (Olympus Soft Imaging Solutions, Germany). Cysts had a distinct thick cyst wall (Fig. 2A) and contained one or two nuclei (Fig. 3A and B). The cyst wall was filamentous in nature, similar to that of Giardia (Fig. 2B) (Chavez-Munguia et al., 2004). The inner cyst wall was more membranous, irregular and located directly adjacent to the encysted parasite (Fig. 2C). A peritrophic space (Fig. 2D) existed between the outer cyst wall and the encysted parasite. Inside the peritrophic space there were numerous spherical, double membrane-bound vesicles approximately 50 nm in diameter (Fig. 2C) resembling the Encystation Specific Vesicles (ESVs) of Giardia (Benchimol, 2004; Hehl and Marti, 2004). The parasite enclosed within the cyst wall had an amoebic appearance (Fig. 2A). The nucleus was bounded by a nuclear membrane and included a centrally located nucleolus (Fig. 3A and B). Hydrogenosomes, located in the cytoplasm of the cysts, were double membrane-bound, electron dense organelles with diameter 0.15–0.35 lm (Fig. 3C). The basal body structure of D. fragilis cysts contained other
organelles such as an axostyle, flagellar axonemes, pelta and a costa. Flagella axonemes comprised of the 9 + 2 microtubule arrangement were present in the periphery of the cytoplasm of the cyst, viewed in both transverse and longitudinal section (Fig. 3E and F). External flagella were absent. The observation of flagella components only within the cyst and not in the trophozoite stage (Banik et al., 2012) provides support for the suggestion that D. fragilis has secondarily adapted to life in the gut by losing dependence on flagella in favour of adopting an amebic appearance and style of locomotion in the gut (Gerbod et al., 2001). The axostyle comprised an axial ribbon of longitudinally oriented microtubules which ran from the anterior region to the posterior region of the cyst (Fig. 3D). A pelta was observed in the anterior region of the cyst as a crescent shaped sheet of microtubules (Fig. 3G). A costa, also commonly found in trichomonads (Benchimol, 2010), was present along the periphery of the encysted parasite, forming a stack of fine filaments within an electron dense, rod shaped skeletal structure (Fig. 3H). In summary, we believe this is the first report of a cyst stage in the life cycle of D. fragilis. Rodent experiments were also able to establish three criteria of the Koch’s postulate. Firstly, all mice inoculated with D. fragilis became infected in contrast to the negative controls which remained uninfected. Secondly, mice infected by D. fragilis developed a mild inflammation in the large intestinal mucosa; a significant transient weight loss in these animals was also detected. In addition there were significantly higher levels of calprotectin present in the feces of infected mice compared with the controls. An increase in faecal calprotectin has been reported in patients suffering from intestinal disorders such as Inflammatory Bowel Disease (IBD) and Inflammatory Bowel Syndrome (IBS) (Costa et al., 2003). Dientamoeba has also been implicated as a possible etiological agent in IBS (Stark et al., 2010b). Thirdly, cysts could establish a new infection in naïve mice or rats when given orally. Hence the results presented here using mice and rats successfully demonstrated several aspects of Koch’s postulates highlighting the infectious nature of this organism and its ability to cause disease in two animal species. These new findings have major implications for further study into the diagnosis and epidemiology of D. fragilis infections, since the most probable route of transmission of D. fragilis to humans involves the faecal–oral transmission of the cyst stage described here. The role of other animal species, such as pigs (Caccio et al., 2012), in the transmission of D. fragilis to humans remains to be determined, however it is now highly likely that dientamoebiasis is a zoonotic disease (Barratt et al., 2011).
Acknowledgements This work was performed in partial fulfilment of the Ph.D. degree at the University of Technology, Sydney (UTS), Australia by VM. This work was financially supported by Sydpath, St. Vincent Hospital Sydney, Australia and UTS. We wish to thank Mrs. Debra Birch for advice and assistance in the transmission electron microscopy.
References Banik, G.R., Birch, D., Stark, D., Ellis, J.T., 2012. A microscopic description and ultrastructural characterisation of Dientamoeba fragilis: an emerging cause of human enteric disease. Int. J. Parasitol. 42, 139–153. Barratt, J.L.N., Banik, G.R., Harkness, J., Marriott, D., Ellis, J.T., Stark, D., 2010. Newly defined conditions for the in vitro cultivation and cryopreservation of Dientamoeba fragilis: new techniques set to fast track molecular studies on this organism. Parasitology 137, 1867–1878. Barratt, J.L.N., Harkness, J., Marriott, D., Ellis, J.T., Stark, D., 2011. The ambiguous life of Dientamoeba fragilis: the need to investigate current hypotheses on transmission. Parasitology 138, 557–572.
V.S. Munasinghe et al. / International Journal for Parasitology 43 (2013) 879–883 Benchimol, M., 2004. The release of secretory vesicle in encysting Giardia lamblia. FEMS Micro. Lett. 235, 81–87. Benchimol, M., 2010. The mastigont system in trichomonads. In: de Souza, W. (Ed.), Structures and Organelles in Pathogenic Protists, Microbiology Monographs, 17. Springer, Berlin, Heidelberg, pp. 1–26. Caccio, S.M., Sannella, A.R., Manuali, E., Tosini, F., Sensi, M., Crotti, D., Pozio, E., 2012. Pigs as natural hosts of Dientamoeba fragilis genotypes found in humans. Emerg. Infect. Dis. 18, 838–841. Chavez-Munguia, B., Cedillo-Rivera, R., Martinez-Palomo, A., 2004. The ultrastructure of the cyst wall of Giardia lamblia. J. Eukaryot. Microbiol. 51, 220–226. Costa, F., Mumolo, M., Bellini, M., Romano, M., Ceccarelli, L., Arpe, P., Sterpi, C., Marchi, S., Maltinti, G., 2003. Role of faecal calprotectin as non-invasive marker of intestinal inflammation. Dig. Liv. Dis. 35, 642–647. Dobell, C., 1940. Researches of the intestinal protozoa of monkeys and man. X. The life history of Dientamoeba fragilis: observations, experiments and speculations. Parasitology 32, 417–461. Gerbod, D., Edgcomb, V.P., Noel, C., Zenner, L., Wintjens, R., Delgado-Viscogliosi, P., Holder, M.E., Sogin, M.L., Viscogliosi, E., 2001. Phylogenetic position of the trichomonad parasite of turkeys, Histomonas meleagridis (Smith) Tyzzer, inferred from small subunit rRNA sequence. J. Eukaryot. Microbiol. 48, 498–504. Girginkardesler, N., Kurt, O., Kilimcioglu, A.A., Ok, U.Z., 2008. Transmission of Dientamoeba fragilis: evaluation of the role of Enterobius vermicularis. Parasitol. Int. 57, 72–75. Hehl, A.B., Marti, M., 2004. Secretory protein trafficking in Giardia intestinalis. Mol. Microbiol. 53, 19–28.
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Munasinghe, V.S., Stark, D., Ellis, J.T., 2012. New advances in the in-vitro culture of Dientamoeba fragilis. Parasitology 139, 864–869. Reynolds, E.S., 1963. The use of lead citrate at high pH as an electron-opaque stain in electron microscopy. J. Cell Biol. 17, 208–212. Roser, D., Nejsum, P., Carlsgart, A.J., Nielsen, H.V., Stensvold, C.R., 2013. DNA of Dientamoeba fragilis detected within surface-sterilized eggs of Enterobius vermicularis. Exp. Parasitol. 133, 57–61. Stark, D., Beebe, N., Marriott, D., Ellis, J., Harkness, J., 2005a. Detection of Dientamoeba fragilis in fresh stool specimens using PCR. Int. J. Parasitol. 35, 57–62. Stark, D., Beebe, N., Marriott, D., Ellis, J., Harkness, J., 2005b. Prospective study of the prevalence, genotyping, and clinical relevance of Dientamoeba fragilis infections in an Australian population. J. Clin. Microbiol. 43, 2718–2723. Stark, D.J., Beebe, N., Marriott, D., Ellis, J.T., Harkness, J., 2006. Dientamoebiasis: clinical importance and recent advances. Trends Parasitol. 22, 92–96. Stark, D., Barratt, J., Roberts, T., Marriott, D., Harkness, J., Ellis, J., 2010a. Comparison of microscopy, two xenic culture techniques, conventional and real-time PCR for the detection of Dientamoeba fragilis in clinical stool samples. Eur. J. Clin. Microbiol. Infect. Dis. 29, 411–416. Stark, D., Barratt, J., Roberts, T., Marriott, D., Harkness, J., Ellis, J., 2010b. A review of the clinical presentation of dientamoebiasis. Am. J. Trop. Med. Hyg. 82, 614– 619. Tøn, H., Brandsnes, Ø., Dale, S., Holtlund, J., Skuibina, E., Schjønsby, H., Johne, B., 2000. Improved assay for fecal calprotectin. Clin. Chim. Acta 292, 41–54.
Contents lists available at SciVerse ScienceDirect
International Journal for Parasitology journal homepage: www.elsevier.com/locate/ijpara
Succinctus
Cyst formation and faecal–oral transmission of Dientamoeba fragilis – the missing link in the life cycle of an emerging pathogen Varuni S. Munasinghe a, Nicole G.F. Vella b, John T. Ellis a,⇑, Peter A. Windsor c, Damien Stark a,d a
School of Medical and Molecular Biosciences and the i3 Institute, University of Technology Sydney, Broadway, New South Wales 2007, Australia Microscopy Unit, Faculty of Science, Macquarie University, North Ryde, New South Wales 2109, Australia c Faculty of Veterinary Science, University of Sydney, Camden, New South Wales 2560, Australia d Division of Microbiology, Sydpath, St. Vincent’s Hospital, Darlinghurst, New South Wales 2010, Australia b
a r t i c l e
i n f o
Article history: Received 6 May 2013 Received in revised form 23 June 2013 Accepted 24 June 2013 Available online 19 July 2013 Keywords: Dientamoeba Trichomonad Transmission Cyst Diarrhoea Irritable bowel
a b s t r a c t Dientamoeba fragilis is a protozoan parasite emerging as a cause of diarrhoea and ‘‘irritable-bowel-like’’ gastrointestinal disease in humans with a propensity for establishing long-term, chronic infections in humans. Although Dientamoeba was discovered over a century ago its life cycle and mode of transmission is not known. No cyst stage has been described and no animal models are presently available for the study of this parasite. Here we describe the establishment of an animal model using laboratory rodents, the fulfilling of Koch’s postulates, and the discovery of a new cyst stage in the life cycle of D. fragilis. Our demonstration of long-term parasite carriage by rodents and prolonged shedding of cysts, together with elevated levels of calprotectin in the stool, confirms the capacity of this organism to cause disease and indicates dientamoebiasis should be considered in the differential diagnosis of gastrointestinal diseases such as Inflammatory Bowel Syndrome (IBS). Finally, we suggest that the cyst stage described here is the vehicle that mediates faecal–oral transmission of D. fragilis between hosts. Ó 2013 Australian Society for Parasitology Inc. Published by Elsevier Ltd. All rights reserved.
Dientamoeba fragilis is increasingly recognised as a relatively common cause of human diarrhoea and long-term chronic infections are a feature of infection by this parasite (Stark et al., 2010b). The mode of transmission and life cycle of D. fragilis has yet to be definitively described (Stark et al., 2006; Barratt et al., 2011). It was initially suggested that D. fragilis was transmitted via the ova of nematodes (Dobell, 1940). Reports of a higher than anticipated rate of co-infection between D. fragilis and Enterobius vermicularis led others to postulate pin worm may be a probable vector of transmission (Girginkardesler et al., 2008; Roser et al., 2013). In contrast others have demonstrated no co-infections with D. fragilis and helminths (Stark et al., 2010b). The absence of strong evidence in support of transmission by helminths, coupled with high rates of co-infection with other faecal orally transmitted bacteria and protozoa, would suggest faecal–oral transmission is the most likely mechanism of D. fragilis transmission (Stark et al., 2005b). Here we present a study on the mode of transmission of Dientamoeba using rodents and report the discovery of a new cyst stage in the life cycle of D. fragilis. All animal experiments were approved by the University of Technology, Sydney (UTS) Australia Animal Care and Ethics Committee. BALB/c mice and Sprague–Dawley rats were obtained from
⇑ Corresponding author. Tel.: +61 2 9514 4161; fax: +61 2 9514 8206. E-mail address: [email protected] (J.T. Ellis).
the Animal Resource Centre (Perth, Australia) housed in independently vented, filter-top cages and provided sterile rodent chow and water ad libitum. The cage bedding was changed every day to avoid and reduce the potential for faecal contamination occurring during the experiment. All animals were confirmed as specific pathogen-free by microscopy and PCR. Animal stool fixed in sodium acetate–acetic acid–formalin (SAF) was examined daily for 1 week prior to infection by D. fragilis for the presence of parasites using permanent iron haematoxylin stained smears. DNA was also extracted from frozen, unfixed stool using the Bioline stool minikit (Bioline, catalogue No. BIO-52037, USA) according to the manufacturer’s recommendations. PCR amplification of stool DNA targeting the D. fragilis ssrDNA was performed as described previously (Stark et al., 2005a). The D. fragilis isolate 4 (Barratt et al., 2010) was cultured on serum slopes with a PBS overlay (Munasinghe et al., 2012). BALB/c mice were orally infected with cultured trophozoites (doses ranging from 103–109/mouse; groups of two) and stools were monitored for the presence of parasites p.i. As a control group a similar number of mice were orally administered with a suspension containing bacteria from the D. fragilis in vitro culture. In order to prepare bacterial cultures for use as controls and to ensure the absence of D. fragilis trophozoites in the preparation, the D. fragilis culture overlay was plated onto blood agar. All of the bacteria isolated as colonies were uniformly returned to Loeffler’s media and incubated anaerobically at 37 °C overnight. All inocula were
0020-7519/$36.00 Ó 2013 Australian Society for Parasitology Inc. Published by Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.ijpara.2013.06.003
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Fig. 1. Iron haematoxylin stained Dientamoeba fragilis cysts present in the feces of infected mice from day 1 p.i. (A) Binucleated (n) D. fragilis cyst from mice infected with D. fragilis. Note the distinct cyst wall (w) and the zone of clearance (c) around the cyst. (B) Dientamoeba fragilis cyst from mice infected with feces of infected mice to establish Koch’s postulates. (C and D) Dientamoeba fragilis cysts present in the feces of rats. Scale bar = 5 lm.
centrifuged at 10,000g (bacteria) or 1,000g (D. fragilis) for 15 min and then resuspended in Earle’s Balanced Salt Solution (EBSS) for administration orally to mice. Post infection all mice were individually weighed daily. To test for significant differences in weight loss between the infected and the control groups of mice a student’s paired t-test was used to calculate P values in SPSS version 19. After day 1 p.i. all mice infected with D. fragilis were intermittently shedding binucleated cysts that measured 5 lm in diameter
as shown in the iron haematoxylin smears (Fig. 1A). Mice infected with D. fragilis showed an average weight loss of 2.38 g (12%) compared with the control group (P < 0.05) from day 1 through to day 6. At this point in time the mean group weights became similar. The infected group also showed symptoms of gastrointestinal disturbance with mice excreting unformed faeces intermittently, compared with the control group which were asymptomatic and produced normally formed stools during the experimental period. In addition white blood cells were observed in the stool smears
Fig. 2. Transmission electron micrographs of Dientamoeba fragilis cysts showing the cyst wall and the encysted parasite. (A) Transverse section across a whole cyst showing the outer fibrillar cyst wall (o), peritrophic space (ps) and the enclosed parasite (p). (B) Fibrillar dense outer cyst wall (o) contains a fine meshwork of fibrils. (C) Inner membrane of the cyst wall (i) which is double layered, enclosing the parasite. Note the peritrophic space (ps) and the Encystation Specific Vesicles (v) scattered beneath the outer cyst wall. (D) Transmission electron micrograph showing the double membrane-bound Encystation Specific Vesicles (v) scattered throughout the peritrophic space (ps). Note the inner membrane of the cyst wall (i) enclosing the parasite. Scale bar = 200 nm.
V.S. Munasinghe et al. / International Journal for Parasitology 43 (2013) 879–883
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Fig. 3. Transmission electron micrographs of Dientamoeba fragilis cysts showing different organelles in the encysted parasite. (A) Transverse section across a mononucleated cyst showing the cyst wall (w), nucleus (nu), and the nucleolus (arrowhead). Note the peritrophic space (ps) and the encysted parasite (p). (B) Transverse section across a binucleated cyst showing the peritrophic space (ps), cyst wall (w), nuclei (nu), nuclear membrane (nm) and the nucleolus (arrowheads). (C) Transverse section across hydrogenosomes (h). (D) Longitudinal section across the axostyle showing the microtubules (a). (E) Transverse section across a flagellar axoneme (ax) with the 9 + 2 microtubule arrangement. (F) longitudinal section across the flagellar axonemes (f). (G) Longitudinal section across the pelta (pe). (H) Longitudinal section across the costa (c). Scale bars = 1 lm for A and B; 100 nm for (C–H).
from D. fragilis-infected mice. Real time PCR also confirmed the presence of D. fragilis DNA in stool (Stark et al., 2010a). To study the duration of shedding of the organism, BALB/c mice (n = 16) were infected orally with doses of D. fragilis (as low as 600 trophozoites/mouse) and observed for a period of 6 months. All of the infected mice shed D. fragilis intermittently and
continued shedding for 6 months p.i. (Fig. 1B), while the control group (n = 2) remained uninfected. Subsequently two naïve, uninfected mice were infected orally with a suspension of mouse feces containing cysts. Iron haematoxylin smears (and subsequent PCR) confirmed the shedding of D. fragilis in these two mice from day 1 p.i. These cysts were morphologically similar to the cysts observed
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previously in mouse stool (Fig. 1A). The infection in these two mice became chronic and lasted 4 months, when the experiment was terminated. Mice were euthanised by carbon dioxide asphyxiation. Small intestine and large intestine from each mouse were collected, fixed in 4% paraformaldehyde and embedded in histowax. Sections (5 lm) were stained with H&E and the slides were viewed using an Olympus light microscope under 60 objective. Histopathological examination of the large intestine of mice infected with D. fragilis showed minor pathological changes with slight inflammation of the submucosa present. Levels of calprotectin, a marker of inflammatory and neoplastic disease in the lower gastrointestinal tract (Tøn et al., 2000), were determined in stool by ELISA over the period 28 days p.i. Stools (100 mg) were homogenised in extraction buffer (weight:volume 1:50) and faecal calprotectin levels were measured using an ELISA (Alpco Diagnostics, USA). The average calprotectin levels present in stool from D. fragilis infected mice (69 ng/ml ± 22.22 SD) was twice that of control mice (33 ng/ ml ± 13.24 SD) (P < 0.05), indicative of the presence of intestinal inflammation. Sprague–Dawley rats were also investigated as an alternative host. Two rats that were administered with trophozoites did not become infected during the course of the experiment. Four rats given a suspension of D. fragilis cysts from infected mice became infected and shed Dientamoeba cysts intermittently from days 5 through to 26 p.i. These cysts were morphologically similar to the cysts that were present in the mouse feces with a distinct cyst wall and a diameter varying between 5 and 6 lm (Fig. 1C and D). This infected group exhibited a statistically significant (P < 0.05) mean weight loss of 7.05 g (4%) compared with the control group over days 1 to 28 p.i. with the main period of weight loss being days 12 through to 22 p.i. Infected rats also produced unformed stools and had white blood cells in their feces. A total number of 10 stool samples were screened by transmission electron microscopy for cysts. Feces from infected mice and rats were collected and fixed in 3% glutaraldehyde in PBS (pH 7.2), embedded in 2% low melt agarose and fixed, stained and infiltrated with osmium tetroxide, uranyl acetate and LR White Resin, respectively, by standard procedures. Semi-thin (1 lm) and ultrathin (70 nm) sections were cut using an ultramicrotome and mounted on 200 mesh copper grids (ProScitech, Australia). Ultrathin sections were stained with saturated aqueous uranyl acetate (7.7%) and Reynold’s lead citrate (Reynolds, 1963). Semi-thin sections were stained with 1% methylene blue in 0.6% sodium bicarbonate/40% glycerol and viewed using an Olympus BH2 light microscope. Ultra-thin sections were examined using a Philips CM10 transmission electron microscope and images captured using a Megaview G2 digital camera (Olympus Soft Imaging Solutions, Germany). Cysts had a distinct thick cyst wall (Fig. 2A) and contained one or two nuclei (Fig. 3A and B). The cyst wall was filamentous in nature, similar to that of Giardia (Fig. 2B) (Chavez-Munguia et al., 2004). The inner cyst wall was more membranous, irregular and located directly adjacent to the encysted parasite (Fig. 2C). A peritrophic space (Fig. 2D) existed between the outer cyst wall and the encysted parasite. Inside the peritrophic space there were numerous spherical, double membrane-bound vesicles approximately 50 nm in diameter (Fig. 2C) resembling the Encystation Specific Vesicles (ESVs) of Giardia (Benchimol, 2004; Hehl and Marti, 2004). The parasite enclosed within the cyst wall had an amoebic appearance (Fig. 2A). The nucleus was bounded by a nuclear membrane and included a centrally located nucleolus (Fig. 3A and B). Hydrogenosomes, located in the cytoplasm of the cysts, were double membrane-bound, electron dense organelles with diameter 0.15–0.35 lm (Fig. 3C). The basal body structure of D. fragilis cysts contained other
organelles such as an axostyle, flagellar axonemes, pelta and a costa. Flagella axonemes comprised of the 9 + 2 microtubule arrangement were present in the periphery of the cytoplasm of the cyst, viewed in both transverse and longitudinal section (Fig. 3E and F). External flagella were absent. The observation of flagella components only within the cyst and not in the trophozoite stage (Banik et al., 2012) provides support for the suggestion that D. fragilis has secondarily adapted to life in the gut by losing dependence on flagella in favour of adopting an amebic appearance and style of locomotion in the gut (Gerbod et al., 2001). The axostyle comprised an axial ribbon of longitudinally oriented microtubules which ran from the anterior region to the posterior region of the cyst (Fig. 3D). A pelta was observed in the anterior region of the cyst as a crescent shaped sheet of microtubules (Fig. 3G). A costa, also commonly found in trichomonads (Benchimol, 2010), was present along the periphery of the encysted parasite, forming a stack of fine filaments within an electron dense, rod shaped skeletal structure (Fig. 3H). In summary, we believe this is the first report of a cyst stage in the life cycle of D. fragilis. Rodent experiments were also able to establish three criteria of the Koch’s postulate. Firstly, all mice inoculated with D. fragilis became infected in contrast to the negative controls which remained uninfected. Secondly, mice infected by D. fragilis developed a mild inflammation in the large intestinal mucosa; a significant transient weight loss in these animals was also detected. In addition there were significantly higher levels of calprotectin present in the feces of infected mice compared with the controls. An increase in faecal calprotectin has been reported in patients suffering from intestinal disorders such as Inflammatory Bowel Disease (IBD) and Inflammatory Bowel Syndrome (IBS) (Costa et al., 2003). Dientamoeba has also been implicated as a possible etiological agent in IBS (Stark et al., 2010b). Thirdly, cysts could establish a new infection in naïve mice or rats when given orally. Hence the results presented here using mice and rats successfully demonstrated several aspects of Koch’s postulates highlighting the infectious nature of this organism and its ability to cause disease in two animal species. These new findings have major implications for further study into the diagnosis and epidemiology of D. fragilis infections, since the most probable route of transmission of D. fragilis to humans involves the faecal–oral transmission of the cyst stage described here. The role of other animal species, such as pigs (Caccio et al., 2012), in the transmission of D. fragilis to humans remains to be determined, however it is now highly likely that dientamoebiasis is a zoonotic disease (Barratt et al., 2011).
Acknowledgements This work was performed in partial fulfilment of the Ph.D. degree at the University of Technology, Sydney (UTS), Australia by VM. This work was financially supported by Sydpath, St. Vincent Hospital Sydney, Australia and UTS. We wish to thank Mrs. Debra Birch for advice and assistance in the transmission electron microscopy.
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