Encephalitozoon hellem Infection in a Captive Juvenile Freshwater Crocodile (Crocodylus johnstoni)

J. Comp. Path. 2015, Vol. 153, 352e356

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DISEASE IN WILDLIFE OR EXOTIC SPECIES

Encephalitozoon hellem Infection in a Captive Juvenile Freshwater Crocodile (Crocodylus johnstoni) T. F. Scheelings*, R. F. Slocombe†, S. Crameri‡ and S. Hairx * Australian Wildlife Health Centre, Healesville Sanctuary, Healesville, † Veterinary Science, University of Melbourne, Werribee, ‡ Australian Animal Health Laboratory, CSIRO, Geelong, Victoria and x Animal Health Laboratories, Department of Agriculture and Food, South Perth, Western Australia, Australia

Summary Microsporidiosis is reported rarely in reptiles and has never been reported in any species of crocodilian. Microsporidiosis was diagnosed histologically in a juvenile captive freshwater crocodile (Crocodylus johnstoni) that was found suddenly dead in its enclosure. Ultrastructural and molecular testing revealed infection to be due to Encephalitozoon hellem. This is the first report of E. hellem infection in any species of reptile. Ó 2015 Elsevier Ltd. All rights reserved. Keywords: Crocodylus johnstoni; Encephalitozoon hellem; freshwater crocodile; microsporidia

The freshwater crocodile (Crocodylus johnstoni) is a medium-sized member of the family Crocodylidae and is one of two extant species of crocodilians found in Australia. They are distributed along the coast and hinterland of northern Australia, with their range extending from Queensland to Western Australia (Cogger, 2014). They are typically confined to permanent freshwater rivers, lagoons and billabongs and are opportunistic predators that forage on a range of fish, frogs, crustaceans, reptiles, birds and mammals (Cogger, 2014). Freshwater crocodiles are common in captivity in Australia and reports of pathology in this species are most often related to conditions seen in intensively farmed animals (Buenviaje et al., 1994, 1998; Shilton et al., 2011). Accounts of microsporidia infections are rare in Australian reptiles (Ladds, 2009) and to the authors’ knowledge there are no published reports of microsporidiosis in any species of crocodilian worldwide. The microsporidia are a diverse group of eukaryotic, obligate intracellular parasites belonging to the Correspondence to: T. F. Scheelings (e-mail: [email protected]). 0021-9975/$ - see front matter http://dx.doi.org/10.1016/j.jcpa.2015.08.001

phylum Microspora (Weiss, 2001; Keeling and Fast, 2002). Initially classified as Protozoa, recent DNA sequencing has determined that microsporidia originated from an ancient lineage of sexual fungi and as such, they are now considered to belong to the kingdom Fungi (Lee et al., 2008). Despite their sexual ancestry, microsporidia have evolved to reproduce asexually, with infection initiated by injection of sporoplasm into the host cell followed by a proliferating merogonic phase (Jacobson, 2007). There are 150 genera and over 1,200 species of microsporidians. They have been identified as important parasites of invertebrates and are used as biological control agents in these animals (Weiss, 2001; Keeling and Fast, 2002; Lee et al., 2008). In addition, microsporidia have been isolated from all vertebrate orders and have been implicated as an economically significant disease in aquaculture (Weiss, 2001; Keeling and Fast, 2002; Lee et al., 2008). Most notably, microsporidia have emerged as serious zoonotic pathogens, particularly in immunocompromised individuals (Mathis et al., 2005). Ó 2015 Elsevier Ltd. All rights reserved.

Encephalitozoonosis in a Crocodile

The present report describes the first case of microsporidiosis in a crocodilian. A 2.5-year-old female captive-born freshwater crocodile from the zoological collection at Healesville Sanctuary was presented to the Australian Wildlife Health Centre (AWHC) for necropsy examination following sudden death. The animal was found dead in its enclosure with no recent signs of illness. The crocodile was housed in a large mixed reptile species exhibit under conditions considered appropriate for the species. All conspecifics housed with the deceased crocodile were considered healthy, with no obvious evidence of disease within the population based on keeper observations. A necropsy examination was performed immediately on the crocodile, with no gross abnormality noted. A representative selection of a wide range of tissues was collected and fixed in 10% neutral buffered formalin for histopathology. Tissues were processed routinely and embedded in paraffin wax. Sections (5 mm) were stained with haematoxylin and eosin (HE), ZiehleNeelsen (ZN) acid-fast stain, Grocott’s methenamine silver (GMS) and by the periodic acideSchiff (PAS) reaction. For electron microscopy formalin-fixed and paraffin wax-embedded (FFPE) sections adhered to glass slides were de-waxed in xylene, rehydrated and fixed at room temperature (20 min) in 2.5% glutaraldehyde in 0.1 M Sorenson’s phosphate buffer (300 mosmol/kg, pH 7.2). The slides were then washed in buffer, post-fixed in 1% osmium tetroxide for 1 h, dehydrated in graded ethanol, embedded and polymerized inverted on blank resin stubs in Procure 812 resin (ProSciTech, Thuringowa, Queensland, Australia). After immersion in boiling water for 7 min, the glass slide was physically removed from the resin block now containing the section tissue. Ultrathin sections cut from the resultant resin block were stained with uranyl acetate and lead citrate and examined with a Philips CM120 electron microscope at 120 kV. Images were acquired using a TemCam-F416 TVIPs camera. Total genomic DNA was extracted from eight 10 mm FFPE liver sections using the QIAamp DNA FFPE Tissue kit (Qiagen, Chadstone, Victoria, Australia) and performed on the automated QIAcube extraction platform (Qiagen). Briefly, FFPE sections were collected in a 1.5 ml microfuge tube (Eppendorf, North Ryde, New South Wales, Australia) with the xylene dewaxing and tissue lysis steps performed manually as per the QIAamp protocol. Following incubation at 90 C for 1 h, to aid in the reversal of formaldehyde modification to nucleic acids, the tissue lysate was transferred to a fresh 1.5 ml Sample Tube and run on the automated QIAcube platform following the ‘isolation of genomic

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DNA from FFPE tissue sections’ protocol available from the Qiacube Web Portal (www.qiagen.com) and eluted in a volume of 40 ml. Polymerase chain reaction (PCR) was carried out using pan-microsporidian primers (Muller et al., 1999): V1 (50 -CAC CAG GTT GAT TCT GCC TGA C-30 ) and PMP2 (50 -CCT CTC CGG AAC CAA ACC CTG-30 ) targeting a 250e270 base pair DNA fragment of the small subunit (SSU) rRNA gene of the microsporidia. PCR was performed in a reaction volume of 25 ml containing 0.4 mM of each primer set, 12.5 ml of Qiagen Multiplex PCR master mix (containing HotStarTaq DNA polymerase, multiplex PCR buffer with 6 mM MgCl2 and dNTP mix) and 5 ml of extracted DNA. DNA from a previously confirmed avian case of Encephalitozoon hellem microsporidiosis was used as a positive control (unpublished). The PCR was performed in a DNA Engine (BioRad, Gladesville, New South Wales, Australia) under the following conditions: initial denaturation at 95 C for 5 min, 36 cycles of 94 C for 30 sec, 55 C for 90 sec and 72 C for 90 sec followed by a final elongation step at 72 C for 5 min. Amplified PCR products were visualized by electrophoresis on a 2% agarose gel stained with SYBR Safe (Life Technologies, Scoresby, Victoria, Australia) and viewed under ultraviolet transillumination. The positive PCR product was purified using the QIAquick PCR Purification kit (Qiagen) and run on the QIAcube extraction platform with a final elution volume of 50 ml. The purified PCR product was quantified by electrophoresis (as previously described) using the HyperLadder 100bp (Bioline, Alexandria, New South Wales, Australia) molecular weight marker as a reference. Forward and reverse sequencing reactions were prepared in 12 ml volumes containing approximately 12e18 ng of PCR product and 9.6 pmol of primers V1 or PMP2 and sent to the Perth Node of the Australian Genome Research Facility (AGRF) for Sanger sequencing. A contig was assembled from the forward and reverse quality trimmed sequences using CAP3 (Huang and Madan, 1999) and compared with nucleotide sequences in GenBank (http://www.ncbi.nlm.nih.gov) using BLAST. The ‘align two or more sequences’ function of BLAST was used for further comparison with sequences generated from three previous Australian cases of E. hellem microsporidiosis (unpublished). The assembled sequence together with 20 microsporidia SSU rRNA partial sequences (as amplified by primers V1 and PMP2) retrieved from Genbank and sequences from the previous Australian cases of E. hellem microsporidiosis were aligned using Clustal W in MEGA 6.06. A neighbour-joining tree was constructed in MEGA using the number of differences

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model with a bootstrap value of 1,000. Vittaforma corneae and Enterocytozoon bieneusi were used as outgroup species. Maximum likelihood and minimum evolution methods produced trees with highly similar topology (data not presented). Microscopically, the liver appeared congested, with hepatocytes containing little fat. There was almost complete involvement of Kupffer cells with each containing large numbers of phagocytosed, slightly refractile, oval to round organisms, often up to 20 organisms per cell (Fig. 1). The organisms had a single small nucleus. There were very large numbers of the same organisms in the mesentery, which was oedematous and contained moderate numbers of heterophils and lymphocytes. The organisms appeared to have been sequestered within phagocytic cells in the mesentery, although some appeared to be free within blood vessels. All other organs were considered microscopically normal. The organisms noted within Kupffer cells had PAS-positive cell walls, stained with GMS and were strongly positive with acid-fast stain. Acid-fast stain demonstrated low numbers of organisms in the spleen, but not in the kidney or heart. On electron microscopy, organisms within parasitophorous vacuoles with microsporidial stages and intracellular organelles were detected. The polar filament, a component of the extrusion apparatus, was seen in thin sections together with spores with finer polaroplast membranes (Fig. 2). The ultrastructure of the spores was typical for the genus Encephalitozoon (V avra and Larson, 1999). Partial sequencing of the microsporidia SSU rRNA gene identified the organism as Encephalitozoon hellem with a sequence identity of 97% across 264 nucleotides on BLAST analysis (100% query coverage). Compar-

Fig. 1. Section of liver from a freshwater crocodile (Crocodylus johnstoni). E. hellem organisms can be seen within Kupffer cells (arrows). HE. Bar, 10 mm.

Fig. 2. Parasitophorous vacuole containing spores. The polar filament (PF), a component of the extrusion apparatus, is seen in transverse section in the spore (black arrow). In addition, a spore with finer polaroplast membranes (P) is seen (white arrow). RBC, red blood cell; N, nucleus. Transmission electron microscopy. Bar, 1 mm.

ison with three previous cases of avian microsporidiosis in Western Australia and New South Wales in captive bird species showed sequence similarities of >99%. Phylogenetic analysis of the SSU rRNA region amplified by primers V1 and PMP2 showed the Australian E. hellem isolates formed a discrete cluster when compared with those in Genbank (Fig. 3). Further molecular characterization is warranted to determine if the Australian isolates represent a distinct genotype of E. hellem and whether any variation in virulence and/or host specificity exists. Microsporidian infections have been described in a range of reptiles, with the majority of these reports originating from squamate species (Jacobson, 2007). Microsporidiosis has been identified most frequently in the inland bearded dragon (Pogona vitticeps) (Jacobson et al., 1998; Mitchell and Garner, 2011; Strunk and Reavill, 2011; Richter et al., 2013), with infection caused by an organism with ultrastructural morphology and genetic similarity to mammalian strains of Encephalitozoon cuniculi (Richter et al., 2013). Other microsporidians that have been implicated in disease in reptiles include Pleistophora spp. in the tuatara (Sphenodon punctatus) (Liu and King, 1971), the European grass snake (Natrix natrix) (Guyenot and Naville, 1922, 1924) and the split keel-back snake (Atretium schistosum) (Narasimhamurti et al., 1982). Encephalitozoon lacerate has been reported in the common wall lizard (Podarcis muralis) (Canning, 1981) and the African skink (Mabuya perrotetii) (Koudela et al., 1998).

Encephalitozoonosis in a Crocodile

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Fig. 3. Neighbour-joining tree of Encephalitozoon species based on partial SSU rRNA gene sequences comparing Australian veterinary isolates of E. hellem with sequences retrieved from GenBank (accession number and host shown). Vittaforma corneae and Enterocytozoon bieneusi represent outgroup species.

More recently, infection with Heterosporis anguillarum was diagnosed in a garter snake (Thamnophis sirtalis) (Richter et al., 2014). This is a species of microsporidia that is usually associated with myositis in Japanese eels (Anguilla japonica) (Richter et al., 2014). To the authors’ knowledge there have been no reports of microsporidiosis occurring in crocodilians and this is the first report of E. hellem infection occurring in any species of reptile. E. hellem infection has been detected in birds, mice and bats and is also a serious zoonotic concern in human patients infected with human immunodeficiency virus (Mo and Drancourt, 2002). Clinical signs of microsporidiosis in reptiles are attributed typically to the predilection of the parasite for gastrointestinal tissue in these animals and may include listlessness, anorexia, diarrhoea, weight loss and sudden death (Jacobson et al., 1998; Koudela et al., 1998; Richter et al., 2013). Less commonly, microsporidian infections in reptiles may appear as non-specific granulomas in muscle or other tissues (Reece and Hartley, 1994; Strunk and Reavill, 2011). Multifocal to diffuse hepatic necrosis and intestinal villus atrophy are common histological findings in affected reptiles (Jacobson et al., 1998;

Koudela et al., 1998; Richter et al., 2013). In this case report, lesions were limited to the liver with all other tissues unaffected. Although organisms were detected in the spleen, there was no inflammation or necrosis associated with their presence in this organ. Microsporidiosis in reptiles has traditionally been thought to be an opportunistic infection (Graczyk and Cranfield, 2000); however, no evidence of concurrent disease was identified in this case. Primary microsporidiosis has now been reported in a range of reptile species (Richter et al., 2013, 2014), which might indicate that it is a true pathogen of reptiles. Transmission of microsporidia is poorly understood, but it is thought to be predominately through the faecaleoral route (Graczyk and Cranfield, 2000), with some evidence to suggest that vertical transmission may play a role in certain species such as bearded dragons (Mitchell and Garner, 2011). It remains unclear how the crocodile in this case report became infected as there have been no other instances of microsporidiosis in the reptile collection at Healesville Sanctuary and attempts to identify microsporidial oocysts in the faeces of other animals in the reptile house with the use of acid-fast stains have been unrewarding.

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Acknowledgments The authors acknowledge the technical assistance of Ms C. Holmes who prepared samples for electron microscopy and the use of the Australian Microscopy and Microanalysis Research Facilities at AAHL CSIRO. We also thank Gribbles Veterinary Pathology for histopathology services including the provision of special stains in order to make a preliminary diagnosis.

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July 17th, 2015 ½ Received, Accepted, August 10th, 2015