INTRODUCTION
Current knowledge suggests that Sarcocystosis is caused by 200 currently identified species of single-cell coccidian parasites in the phylum Apicomplexa, and genus Sarcocystis. These parasites infect a wide range of definitive and intermediate hosts, including carnivorous animals, domestic animals and humans (Dubey and Lindsay, Reference Dubey and Lindsay2006; Kaltungo and Musa, Reference Kaltungo and Musa2013; Dubey et al. Reference Dubey, Calero-Bernal, Rosenthal, Speer and Fayer2015). All Sarcocystis parasites have an obligatory two host life cycle (some exception such as Sarcocystis neurona exist); asexual reproduction takes place in the intermediate host and sexual reproduction occurs in the intestine of the definitive host (Dubey and Lindsay, Reference Dubey and Lindsay2006). Sarcocystis neurona has a very broad intermediate host range and for example sporocysts from opossums can infect many hosts, of which some are natural intermediate hosts (in which sarcocysts are formed), while others are aberrant hosts (in which only schizonts are formed) (Dubey et al. Reference Dubey, Lindsay, Saville, Reed, Granstrom and Speer2001a ). Transmission from definitive to intermediate host occur via the ingestion of oocysts/sporocysts from feces via contaminated food or water and transmission from intermediate to definitive host occur via the ingestion of sarcocysts, which are found in muscle tissue (Dubey and Lindsay, Reference Dubey and Lindsay2006; Gjerde and Josefsen, Reference Gjerde and Josefsen2015). However, knowledge and understanding of all the life cycle stages of Sarcocystis species in wild carnivores is incomplete and needs to be researched in more detail to help us to better understand disease pathogenesis, symptomatology and impact of parasite diversity. More research is needed to determine the range of clinical manifestations of Sarcocystis infections in wild carnivores, as there is little information available about the signs and symptoms in these host species.
Only a few Sarcocystis species have been identified in wild carnivores of the family Mustelidae (Dubey et al. Reference Dubey, Reichard, Torretti, Garvon, Sundar and Grigg2010). For example S. neurona has previously been identified in (Eurasian) otters (Enhydra lutris) (Dubey et al. Reference Dubey, Rosypal, Rosenthal, Thomas, Lindsay, Stanek, Reed and Saville2001b , Reference Dubey, Lindsay, Rosenthal and Thomas2003; Miller et al. Reference Miller, Barr, Nordhausen, James, Magargal, Murray, Conrad, Toy-Choutka, Jessup and Grigg2009; Wendte et al. Reference Wendte, Miller, Nandra, Peat, Crosbie, Conrad and Grigg2010), while S. lutrae has been found in the Eurasian otter (Lutra lutra) in Norway (Gjerde and Josefsen, Reference Gjerde and Josefsen2015). Sarcocystis lutrae has not been confirmed in another host species other than the Eurasian otter and potentially Arctic foxes (Gjerde and Schulze, Reference Gjerde and Schulze2014; Gjerde and Josefsen, Reference Gjerde and Josefsen2015). Various Sarcocystis spp. including Sarcocystis hofmanni, Sarcocystis melis, Sarcocystis cf. sebeki and Sarcocystis cf. gracilis, have previously been recorded by light microscopy (LM) and transmission electron microscopy (TEM) in heart, thigh, loin, thorax and tongue samples in European badgers (Meles meles) from Berlin (Odening et al. Reference Odening, Stolte, Walter and Bockhardt1994a , Reference Odening, Stolte, Walter, Bockhardt and Jakob b ). None of these Sarcocystis species in badgers have been identified in the UK. More recently, an unnamed species of Sarcocystis was recorded in the tongue, diaphragm and masseter muscle of Japanese badgers (M eles anakuma) using haematoxylin and eosin (H&E) staining (Kubo et al. Reference Kubo, Okano, Ito, Tsubota, Sakai and Yanai2009). To date LM of fresh muscle tissue and TEM have been examined to identify the Sarcocystis spp. found in badgers, meaning no DNA sequences are available for the Sarcocystis species previously identified in badgers. Only a few Sarcocystis species found in wild carnivores have been examined using molecular methods, these include species, such as Sarcocystis arctica, Sarcocystis lutrae, Sarcocystis kalvikus, and Sarcocystis kitikmeotensis (Dubey et al. Reference Dubey, Calero-Bernal, Rosenthal, Speer and Fayer2015). Techniques, such as polymerase chain reactrion (PCR) and sequence analysis are more frequently used to identify Sarcocystis species (Gjerde and Josefsen, Reference Gjerde and Josefsen2015). Polymorphisms in the 18S rDNA and internal transcribed spacer (ITS1) region may help with the speciation and discrimination of the different species within the Sarcocystis genus. The aim of this study was to determine the prevalence and species of Sarcocystis in muscle samples from European badgers (M. meles).
MATERIALS AND METHODS
Collection of samples
In total 54 European badger (M. meles) carcasses were collected from around the Lothians and Borders regions of Scotland, following fatal collisions with vehicles (badgers were collected with the knowledge and permission of Scottish Natural Heritage) (Bartley et al. Reference Bartley, Wright, Zimmer, Roy, Kitchener, Meredith, Innes and Katzer2013). Carcasses were stored at −20 °C prior to processing, full necropsies were performed when possible where samples of neck muscle, tongue, spleen, submandibular lymph node, liver, lung, brain, heart, blood and spinal cord were collected.
DNA extraction
DNA was extracted from muscle samples of 54 badgers. From those badgers, 54 neck samples and 32 tongue samples derived from the same animals were extracted. Approximately 1 g of each thawed tissue was transferred into a separate CK22 Precellys tissue homogenizer tube (Cepheid, Stretton Derbyshire, UK), containing 1 mL Nuclei Lysis Solution (Promega, Madison, WI, USA). Samples were homogenized for 2 × 50 s at 6500 rpm using a Precellys 24 tissue homogenizer (Depheid, Stretton Derbyshire, UK). 400 µL of each homogenised tissues were added to a further 900 µL of nuclei lysis solution and incubated at 55°C overnight. Samples were then processed using the Wizard® genomic DNA (Promega, Madison WI, USA) purification protocol, which was adapted to use 0·4 g of starting material (Bartley et al. Reference Bartley, Wright, Zimmer, Roy, Kitchener, Meredith, Innes and Katzer2013).
Detection of protozoan DNA by 18S PCR and ITS1 PCR
Parasite DNA was detected using a nested PCR, targeting the multi-copy 18S rDNA of the ribosomal RNA gene family. The first round PCR used external primers that recognized various apicomplexan parasites including Neospora caninum, Toxoplasma gondii and Sarcocystis spp. (Table 1). Briefly, each 20 µL reaction contained 2 µL of 10× custom PCR mix- (45 mm Tris-HCl, 11 mm (NH4)2SO4, 4·5 mm MgCl2, 0·113 mg mL−1 BSA, 4·4 µ m EDTA and 1·0 mm each of dATP, dCTP, dGTP and dTTP) (ABgene, Epsom, Surrey, UK), 0·25 pm of each primer (Eurofins MWG Operon), 0·75 units of BioTaq (Bioline, London, UK), 13·85 µL of water and 2 µL of sample DNA (Burrells et al. Reference Burrells, Opsteegh, Pollock, Alexander, Chatterton, Evans, Walker, McKenzie, Hill, Innes and Katzer2016). The PCR conditions for the first round were 95 °C for 5 min followed by 35 cycles at 95 °C for 1 min, 56 °C for 1 min and 72 °C for 1 min, with the final extension period at 72 °C for 5 min. The primary PCR amplicons were diluted with 100 µL DNase/RNase free water and 2 µL of the diluted primary amplification product was added as template DNA for the second round amplification. Second round primers were designed to amplify only Sarcocystis spp. (Table 1). The specificity of the primers were tested using S. neurona, S. lutrae, S. gigantea, Sarcocystis tenella, Sarcocystis rileyi, Sarcocystis fayeri, N. caninum and T. gondii DNA samples (data not shown). The reaction conditions for the second round PCR were identical to the first round, with the exception that internal forward and reverse primers were used. Each batch of samples analysed, contained a positive control: S. lutrae (obtained from this study) and negative controls: N. caninum, T. gondii and water and were tested in duplicates. With each batch of badger samples extracted, a negative (water) ‘extraction control’ was tested (Bartley et al. Reference Bartley, Wright, Zimmer, Roy, Kitchener, Meredith, Innes and Katzer2013). Badgers that showed strong positive bands for both neck and tongue samples in the 18S rDNA PCR, were tested further using the ITS1 PCR. Here, Sarcocystidae were detected using the primers ‘SU1F’ and ‘5.8SR2’ that amplify the ITS1 region (~1000 bp) and targets the adjacent 18S and 5.8S rDNA genes, respectively (Gjerde, Reference Gjerde2014) (Table 1). The reaction conditions for the ITS1 PCR were identical to those of the 18S rDNA PCR. The ITS1 region was selected to differentiate members of the group Sarcocystidae as it is highly polymorphic compared with the 18S rDNA. PCR products (6 µL) were analysed by 2% agarose gel electrophoresis, stained with gel red (1 : 10 000) (Biotonium, Hayward, USA) and visualised using ultraviolet light. Each batch of samples analysed by ITS1 PCR contained a positive control: S. lutrae and negative control water.
Sequences and specificity of primers used for the detection of Sarcocystis spp DNA in badger samples
Cloning, DNA sequencing and sequence assembly
The PCR products from 12 animals (both positives for tongue (n = 12) and neck muscle (n = 12)) using the 18S external primers and six positive PCR products from four animals (positives tongue (n = 4) and neck (n = 2)) using the external ITS1 PCR, were purified using the commercially available Wizard® SV Gel and PCR Clean-up System (Promega, Madison WI, USA). The PCR products were eluted in 50 µL of DNase/RNase free water and the nucleic acid concentration was determined by spectrophotometer (Nanodrop, ND1000). For each sample, 100 ng of DNA was sent for sequencing (Eurofins MWG Operon). The 18S amplicons were sequenced with the 18S primers and the ITS1 amplicons were sequenced with the ITS1 primers (Table 1).
Three first round PCR amplicons from the 18S rDNA PCR (tongue n = 1, neck n = 2) and a further 3 PCR amplicons from the ITS1 PCR (tongue n = 2, neck n = 1) were cloned using the pGEM®-T Easy Vector System (Promega, Madison WI, USA) as previously described (Bartley et al. Reference Bartley, Hamilton, Wilson, Innes and Katzer2016) with the following alterations. Two microliter (64 ng) of the purified product were ligated into the pGEM®-T Easy Vector (1 µL at 50 ng μL−1) (Promega, Madison WI, USA) according to the manufacturer's instructions. Following ligation, 1 µL (8 ng) of ligated vector/insert was used to transform 40 µL of high-efficiency competent JM109 cells (⩾1 × 108 cfu μg−1 DNA) (Promega, Madison, WI, USA) using manufacturer's instructions. A successful transformation was confirmed using LB agar plates containing 100 µg mL−1 ampicillin, spread with 100 µL of IPTG (Isopropyl β-D-1-thiogalactopyranoside) (100 mm) and 20 µL of X-Gal (5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside) (50 mg mL−1). White colonies were screened by PCR using the 18S external primers and the SU1F and 5.8SR2 primers to confirm the presence of the Sarcocystis 18S rDNA and ITS1 region insert. Three clones from each of the three badger samples from each of the 18S (n = 9 clones) and ITS1 (n = 9 clones) PCR were sequenced (Eurofins, MWG Operon) using T7 and SP6 primers. Additional internal S-ITS1-F and S-ITS1-R primers (Table 1) were used for the ITS1 clones to ensure a double stranded consensus sequence of over 1000 bp was generated. Overall consensus sequences were generated for the 18S and ITS1 amplicons from each badger.
A Basic Local Alignment Search Tool (BLAST) search was completed to determine percentage identity of the generated sequences against previously published sequences. Multiple sequence alignments were performed using the BioEdit sequence alignment editor 7.1.3.0. to show the difference between the closely related Sarcocystis spp. Phylogenetic analyses were performed on both the 18S rDNA and ITS1 consensus sequences using MEGA6 software (Tamura et al. Reference Tamura, Stecher, Peterson, Filipski and Kumar2013). The evolutionary history was inferred by using the Maximum Likelihood method based on the Tamura–Nei model (Tamura and Nei, Reference Tamura and Nei1993). Initial tree(s) for the heuristic search were obtained by applying the Neighbour–Joining method to a matrix of pairwise distances estimated using the Maximum Composite Likelihood. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site and all positions containing gaps and missing data were eliminated. The phylogeny was tested with the bootstrap method, using 1000 bootstrap replications.
Statistical analysis
The proportion of positive samples (prevalence), with confidence intervals (95% CI) was calculated for the presence of Sarcocystis DNA in the tongue and neck muscle samples from badgers. The numbers of badgers where either tongue, or neck muscle sample were positive and those animals where both samples were positive were also calculated. All of the calculations were carried out using the Minitab 17 software (v17.1.0.0).
RESULTS
Screening of samples for the presence of protozoan DNA using the 18S rDNA PCR
Five badgers were initially screened and DNA samples from leg, neck muscle, tongue, sub-mandibular lymph node, liver, lung, brain, heart and spleen, were tested using the 18S external primers in a single round PCR. Positive PCR amplicons were observed for 1/5 leg muscle, brain and lung sample, 2/5 neck muscle samples and 3/5 tongue and spinal cord samples. Sequencing PCR amplicons from one neck muscle and two tongue samples showed identity to S. lutrae (accession KM657770). The PCR products from tongue and neck were the only samples to produce identifiable sequences, and thus these organs were selected for further testing. Spinal cord was not selected for further analysis, due to the limited numbers of samples available (n = 12).
Verification of PCR specificity and sequencing
The Sarcocystis specific 18S rDNA nested PCR was used to screen all muscle samples available, tongue (n = 32) and neck muscle samples (n = 54) from 54 badgers. The results showed that 36/54 (67%) (95% CI: 52·5–78·9%) neck samples and 24/32 (75%) (95% CI: 56·5–88·5%) tongue samples tested positive for Sarcocystis DNA. Twenty badger samples showed positive PCR results for Sarcocystis DNA in both neck and tongue samples using the 18S rDNA PCR (20/32) (95% CI: 43·6–78·9%). Forty badgers tested positive with the Sarcocystis specific 18S rDNA PCR with at least one tissue (40/54, 74%) (95% CI: 60·3–85·0%). No amplified products were observed for the negative controls: water, T. gondii and N. caninum. Badgers (n = 20) that showed positive results for both tongue and neck in the 18S rDNA PCR were tested using the ITS1 PCR (Gjerde, Reference Gjerde2014). The ITS1 PCR revealed positive results for 12/20 (60%) (95% CI: 36·0–80·8%) neck and 10/20 (50%) (95% CI: 27·1–72·8%) tongue samples for Sarcocystidae DNA. No PCR amplicons were generated for 4/20 badgers tested using the ITS1 primers.
Consensus sequences were generated for the 18S rDNA from 9 clones: tongue (n = 3) and neck (n = 6) and for the ITS1 region from 9 clones: neck (n = 3); and tongue (n = 6). These clones were used to create consensus sequences for the 18S rDNA and ITS1 amplicons for each animal. The 3 consensus sequences, each for the 18S rDNA and the ITS1 amplicons, were identical to each other and were used to create a general consensus sequence for both the 18S rDNA and ITS1 regions. The general consensus sequences for the 18S rDNA (468 bp) and the ITS1 region (1074 bp) were submitted to Genbank (KX229728 and KX431307, respectively). When the 18S rDNA (KX229728) and ITS1 (KX431307) sequences generated during this study, were compared on NCBI BLAST against published DNA sequences, it was found that the 18S rDNA fragments showed 100% identity to isolates of S. lutrae (18S rDNA: KM657770). The ITS1 sequence showed 99·2–100% identity to the 22 ITS1 sequences of S. lutrae found in the Eurasian otter (Lutra lutra) (Gjerde and Josefsen, Reference Gjerde and Josefsen2015).
Phylogenetic relationship and multiple sequence alignments of S. lutrae and related species
Phylogenetic analysis revealed that the S. lutrae rDNA found in badgers appears in the same clade as the S. lutrae found in otters, as well as the closely related species S. rileyi and Sarcocystis turdusi (Fig. 1A). The multiple sequence comparison demonstrated that the 18S rDNA fragment found in badgers (KX229728) is identical to S. lutrae found in otters (KM657775) (Fig. 2A). The sequence alignment of the 18S rDNA (Fig. 2A) shows polymorphic and conserved regions for the closely related Sarcocystis species. The alignment shows that our sequence and S. lutrae are identical to each other but are distinct from the other closely related species sequences by one additional ‘T’ base in comparison with Sarcocystis corvusi and S. arctica and Sarcocystis turdusi, and multiple base pair differences from S. rileyi, Sarcocystis lacerate, Sarcocystis mucosa and Sarcocystis neurona (Fig. 2A). The ITS1 region was also used for sequence alignments and phylogenetic analysis, since the 18S rDNA gave poor discrimination of closely related species. The ITS1 phylogenetic analysis showed a clearer differentiation from the closely related Sarcocystis spp. (S. corvusi, S. arctica, S. neurona, S. turdusi and S. rileyi), however Sarcocystis kalvikus was found in the same clade as S. lutrae from badgers and S. lutrae from otters (Fig. 1B). Yet, when using the ITS1 multiple sequence analysis, it can be seen that S. kalvikus can be distinguished from the S. lutrae found in badgers and S. lutrae found in otters. The ITS1 region is more polymorphic compared with the 18S rDNA and the ITS1 sequence comparison showed a clear differentiation of S. lutrae, S. kalvikus, S. turdusi, S. corvusi and S. arctica (Fig. 2B). From the multiple sequence analysis of both the 18S rDNA and ITS1 regions, it can be clearly seen that the Sarcocystis spp. rDNA fragments found in the sample of badgers in this study are identical to the S. lutrae found in otters.
Molecular Phylogenetic analysis by Maximum Likelihood method for selected members of the Sarcocystidae. (A) 18S rDNA with the highest log likelihood (−1280·9024) and (B) ITS1 spacer region with the highest log likelihood (−2396·9991). The percentage of trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) is shown next to the branches.
Multiple sequence alignment of the polymorphic sections of the 18S and the ITS1 region amplified in this study: (A) 18S rRNA gene region, (B) ITS1 region. Boxing shows identical sequences of S. lutrae (18S KM657769 and ITS1 KM657775) and the Sarcocystis sequence detected in the badger samples (18S KX229728 and ITS1 KX431307). Dots represent identical base pairs and dashed lines represent gaps in the alignments. Numbers given above each alignment correspond to the nucleotide position in the sequence (A) KM657769 and (B) KM657775, respectively.
DISCUSSION
In this paper we report the detection of 18S rDNA and ITS1 region in tongue and neck muscles of European badgers (M. meles) collected from around the Lothians and Borders regions of Scotland, that show 100% sequence identity to DNA from S. lutrae (KM657770 and KM657775). Sarcocystis lutrae has previously been identified in tongue, skeletal muscle and diaphragm in the Eurasian otter (Lutra lutra) (Gjerde and Josefsen, Reference Gjerde and Josefsen2015). Moreover, Gjerde and Schulze (Reference Gjerde and Schulze2014), found that the cox1 sequence of S. lutrae from otters were identical with one of the cox1 sequence from an arctic fox harbouring S. arctica. One of those cox1 sequences was initially assigned to S. arctica even though it differed slightly from the other cox1 sequences obtained and was later re-assigned to S. lutrae and thus the presence of S. lutrae in Arctic foxes can be disputed (Gjerde and Josefsen, Reference Gjerde and Josefsen2015). This study used the 18S rDNA and ITS1 region, to verify and identify the 18S rDNA fragments found in badger samples. Phylogenetic and multiple sequence alignments have shown that the ITS1 region is more polymorphic, compared with the 18S rDNA gene. Using multiple loci, such as the 18S rDNA gene and the ITS1 region will help species identification and is more reliable than using one locus alone. Few polymorphic regions in the 18S rDNA gene have previously been shown in especially closely related Sarcocystis species (i.e. S. lutrae, S. turdusi, S. arctica, Sarcocystis wobeseri) and little sequence data has been generated. It has been shown that the ITS1 region gives a clearer differentiation for these species (Gjerde and Schulze, Reference Gjerde and Schulze2014; Gjerde and Josefsen, Reference Gjerde and Josefsen2015). Since the ITS1 region is not a gene, higher mutation densities are tolerated, making this region highly variable among species and thus a useful marker for species identification for some, however, not all Sarcocystis spp.
From this study it can be confirmed that the 18S rDNA and ITS1 region identified in badgers showed 100% sequence identity to Sarcocystis lutrae (KM657770 and KM657775), indicating that the Sarcocystis species detected is likely to be S. lutrae. Active infections may have been detected if LM of fresh muscle tissue, such as tongue muscle, were analysed. S. hofmanni, S. melis, S. cf. sebeki, S. cf. gracilis and an unnamed Sarcocystis species have previously been recorded in European badgers (M. meles) and Japanese badgers (Meles anakuma) using TEM and, LM of fresh muscle tissue (Kubo et al. Reference Kubo, Okano, Ito, Tsubota, Sakai and Yanai2009; Odening et al. Reference Odening, Stolte, Walter and Bockhardt1994a , Reference Odening, Stolte, Walter, Bockhardt and Jakob b ). Since those studies were conducted before molecular techniques were used in such research, no 18S or ITS1 sequences were generated for these species. Testing both neck muscle and tongue for the detection of Sarcocystis DNA proved advisable, as both these tissues showed a high presence of Sarcocystis DNA and if only one tissue was tested the overall prevalence would have been lower. The density of sarcocysts may vary in different types of muscle tissues, such as the diaphragm, oesophagus, tongue and heart. These tissues are commonly used to demonstrate the presence of sarcocysts in hosts (Dubey et al. Reference Dubey, Calero-Bernal, Rosenthal, Speer and Fayer2015).
Identification of Sarcocystis species based on morphology employs looking at structural characteristics, such as sarcocyst wall and morphology; however, more than one Sarcocystis spp. may have the same sarcocysts morphology and the same species can occur in different hosts (Dubey et al. Reference Dubey, Speer and Fayer1989, Reference Dubey, Calero-Bernal, Rosenthal, Speer and Fayer2015). More recently molecular methods of Sarcocystis spp. have been particular useful to distinguish between morphologically indistinguishable species in closely related intermediate hosts, such as water buffaloes and cattle, and different cervids. Ideally, individual sarcocysts should be excised from fresh muscle tissue, examined in wet mounts by LM, used to extract DNA for molecular characterization and fixed to study them using TEM. Using both identification methods would allow phenotypic and genotypic data to be combined and be linked to the species description. Using morphological characteristics alone for the identification may prove difficult as size and shape is subject to change depending on the age of the Sarcocysts but this observation can be strengthened by sequencing DNA amplicons from different regions of the parasite genome (Dubey et al. Reference Dubey, Calero-Bernal, Rosenthal, Speer and Fayer2015).
The data presented in this study shows that the DNA detected in European badgers showed sequence identity at two different loci to S. lutrae found in otters. This shows that badgers from the Lothians and Borders regions of Scotland are frequently infected with S. lutrae. Badgers are omnivores, and it is likely that they become infected through the ingestion of sporocysts shed by definitive host (predator/scavenger) (Dubey and Lindsay, Reference Dubey and Lindsay2006). Birds, such as the white-tailed (sea) eagle (Haliaeetus albicilla) suggested by (Gjerde and Josefsen, Reference Gjerde and Josefsen2015) or birds of the family Corvidae, as well as other badgers and foxes (Vulpes vulpes) may act as a definitive host of S. lutrae. Further research, involving microscopic analysis, as well as multiple locus sequence typing, is needed to confirm whether S. lutrae is widely distributed across Great Britain and whether S. lutrae is only found in badgers, (Eurasian) otters and potentially arctic foxes.
FINANCIAL SUPPORT
We are grateful for the support by the Biotechnology and Biological Science Research Council (BBSRC), Zoetis and the Scottish Government's Rural and Environment Science and Analytical Services Division (RESAS) for funding this work.
