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Eimeria genomics: Where are we now and where are we going? Damer P. Blake ∗ Pathology and Pathogen Biology, Royal Veterinary College, Hawkshead Lane, North Mymms, Hertfordshire, AL9 7TA, UK

a r t i c l e

i n f o

Article history: Received 27 March 2015 Received in revised form 1 May 2015 Accepted 9 May 2015 Keywords: Eimeria Chickens Genomics Next-generation sequencing

a b s t r a c t The evolution of sequencing technologies, from Sanger to next generation (NGS) and now the emerging third generation, has prompted a radical frameshift moving genomics from the specialist to the mainstream. For parasitology, genomics has moved fastest for the protozoa with sequence assemblies becoming available for multiple genera including Babesia, Cryptosporidium, Eimeria, Giardia, Leishmania, Neospora, Plasmodium, Theileria, Toxoplasma and Trypanosoma. Progress has commonly been slower for parasites of animals which lack zoonotic potential, but the deficit is now being redressed with impact likely in the areas of drug and vaccine development, molecular diagnostics and population biology. Genomics studies with the apicomplexan Eimeria species clearly illustrate the approaches and opportunities available. Specifically, more than ten years after initiation of a genome sequencing project a sequence assembly was published for Eimeria tenella in 2014, complemented by assemblies for all other Eimeria species which infect the chicken and Eimeria falciformis, a parasite of the mouse. Public access to these and other coccidian genome assemblies through resources such as GeneDB and ToxoDB now promotes comparative analysis, encouraging better use of shared resources and enhancing opportunities for development of novel diagnostic and control strategies. In the short term genomics resources support development of targeted and genome-wide genetic markers such as single nucleotide polymorphisms (SNPs), with whole genome re-sequencing becoming viable in the near future. Experimental power will develop rapidly as additional species, strains and isolates are sampled with particular emphasis on population structure and allelic diversity. © 2015 Elsevier B.V. All rights reserved.

1. Introduction The word ‘genomics’ is widely considered to have appeared in 1986 (Kuska, 1998), with the first appearance of the term in the National Center for Biotechnology Information literature database appearing in 1987. By the year 2000 more than 1500 publications had appeared including the word ‘genomics’ as a searchable term, with 15,991 publications last year (2014). Many definitions of the term have been proposed, with a common theme being studies of the genome of an organism and genomewide analysis of the genes it contains using DNA sequencing and bioinformatics methods. Progress in genomics led to publication of the first eukaryotic chromosome sequence in 1992, followed by the first full eukaryotic genome in 1996 (Oliver et al., 1992; Goffeau et al., 1996), but the roll out of genomics technologies has largely been constrained by the cost and availability of Sanger (or first generation) sequencing and the expertise required for analysis. As a consequence, genomics approaches were initially only

∗ Corresponding author. Tel.: +44 1707 666041. E-mail address: [email protected]

accessible for the human genome and other model or medically relevant organisms. The advent of second or next generation sequencing technologies (NGS), providing faster, cheaper and higher throughput approaches, has unlocked genomics for use with veterinary and diagnostic purposes. While the rate of exploitation is not yet comparable with human and medical pathogen genomics, illustrated by greater fragmentation for most livestock and veterinary pathogen genomes (Table 1), sequence resources and associated opportunities are developing fast. Progress in genome sequencing and genome-wide analyses for the Eimeria species illustrates the impact of these advances and the opportunities which are only now becoming readily available. 2. Eimeria and the disease coccidiosis Eimeria are obligate protozoan parasites which have evolved to exhibit immense diversity in host range including mammals, birds, reptiles, fish and amphibians, with each parasite species commonly defined by absolute host specificity (Jirku et al., 2009; GibsonKueh et al., 2011; Yang et al., 2012; Chapman et al., 2013; Jirku et al., 2013; Kvicerova and Hypsa, 2013). As a consequence there are likely to be many thousands of Eimeria species. Eimeria that

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Table 1 Comparison of selected human, veterinary and pathogen genome sequence assemblies. Class

Species

∼Genome size (Mb)a

Genome assembly

Scaffolds (accession number)b

Av scaffolds/chromosome

Host

Homo sapiens Mus musculus Bos taurus Ovis aries Sus scrofa Gallus gallus

3,097 2,731 2,670 2,619 2,808 1,047

24 chromosomes 21 chromosomes 30 chromosomes 27 chromosomes 20 chromosomes 31 chromosomes

766 (GCA 000001405.17) 276 (GCA 000001635.4) 6,337 (GCA 000003055.5) 5,698 (GCA 000298735.1) 9,906 (GCA 000003025.4) 16,847 (GCA 000002315.2)

31.9 13.1 211.2 211.0 495.3 543.5

Parasite Apicomplexa

Neospora caninum Plasmodium falciparum Theileria parva Toxoplasma gondii Eimeria maxima Eimeria tenella Eimeria mitis

60 23 8 66 46 52 72

14 chromosomes 14 chromosomes 4 chromosomes 14 chromosomes 14 chromosomes 14 chromosomes 14 chromosomes

14 (GCA 000208865.2) 15 (GCA 000002765.1) 9 (GCA 000165365.1) 381 (GCA 000006565.1) 3,564 (GCA 000499605.1) 4,664 (GCA 000499545.1) 10,336 (GCA 000499745.1)

1.0 1.1 2.3 27.2 254.6 333.1 738.3

Parasite Other

Echinococcus granulosus Haemonchus contortus

9 chromosomes ? chromosomes

957 (GCA 000524195.1) 12,915 (GCA 000469685.1)

106.3 ?

a b

152 320

Reference sequence assembly size, Ensembl or EuPathDB, accessed 4th March, 2015. NCBI Genome Assembly report, accessed 4th March, 2015.

infect wild vertebrates can become significant for their influence on host/pathogen ecology or captive breeding of endangered species (Jeanes et al., 2013; Knowles et al., 2013), but those that infect livestock are of greatest relevance. For ruminants the economic impact of Eimeria has long been recognized with recent reports indicating a 6–9% reduction in gross margin associated with infection (Radostits and Stockdale, 1980; Lassen and Ostergaard, 2012). For poultry all 60 billion birds produced in the world each year are at risk of exposure (Blake and Tomley, 2014). The annual cost of losses to the global poultry industry attributed to clinical and sub-clinical disease, combined with the cost of control to maintain good productivity and welfare, has been predicted to exceed UK£2 billion (Dalloul and Lillehoj, 2006). Coccidiosis caused by Eimeria infection is primarily associated with enteric disease with few exceptions (Long et al., 1976; Revets et al., 1989; Novilla and Carpenter, 2004). For chickens the disease can be subdivided into haemorrhagic and malabsorptive pathologies loosely related to Eimeria brunetti, Eimeria necatrix and Eimeria tenella, or Eimeria acervulina, Eimeria maxima, Eimeria mitis and Eimeria praecox, respectively (Reid et al., 2014). The severity of disease can be influenced by multiple factors including parasite and host genotype, the size and age of oocyst dose, poultry management system and consequential level of oocyst sporulation, and previous exposure history (Blake et al., 2005; Kim et al., 2008; Williams et al., 2009; Bacciu et al., 2014). Pathognomonic signatures vary between species but poor food conversion ratio, weight loss and diarrhoea are characteristic with high morbidity common and mortality in the absence of intervention. Anticoccidial control relies primarily on routine chemoprophylaxis using ionophore and/or chemical drugs, but resistance develops rapidly and is now widespread (Chapman and Jeffers, 2014). Political and public concerns related to drug use within food production is placing further pressure on the use of chemoprophylaxis (e.g. regulation [EC] No. 1831/2003, article 11). Live anticoccidial vaccines have also proven immensely successful, although a requirement for in vivo amplification using chickens places an inherent limit on production capacity and imposes a minimum cost greater than that of drugs, discouraging use by the majority broiler industry (Blake and Tomley, 2014). As a consequence new, cost-effective anticoccidial strategies are urgently required but progress has been slow, hampered in part by the paucity of genomics resources (Blake and Tomley, 2014). Developments in sequencing technologies over the last ten years have supported a response to this deficit, culminating in the landmark publication of genome sequences for all seven Eimeria species which infect the chicken and one which infects the mouse in 2014 (Heitlinger et al., 2014; Reid et al., 2014).

3. Sequencing technologies DNA sequencing by fragmentation or chain termination first appeared in the late 1970s with the latter, commonly referred to as Sanger sequencing, becoming dominant due to methodological advantage (Maxam and Gilbert, 1977; Sanger et al., 1977). The following 25 years saw considerable innovation and improvement to the Sanger sequencing technique and associated data analysis but limitations including requirements for minimum template quality and quantity, restricted throughput capacity and high cost hindered uptake for many applications. The second wave of sequencing technologies, termed next generation sequencing (NGS), began with the appearance of pyrosequencing (454) and sequencing by synthesis (Solexa, now Illumina), followed by sequencing by oligo ligation detection (SOLiD) and ion semiconductor sequencing (Ion Torrent) (Bennett et al., 2005; Margulies et al., 2005; Valouev et al., 2008). The processes and benefits of the NGS technologies have been reviewed extensively elsewhere (e.g. Metzker, 2010; van Dijk et al., 2014). In summary all are capable of producing massive numbers of sequence reads per run, ranging from one million to several billion, at the cost of reduced read length and in some examples individual base accuracy. Four hundred fifty four pyrosequencing relies on attachment of single DNA template molecules to a primercoated bead. As a complementary DNA strand is synthesised by exposure to a flow of a single unlabelled nucleotides pyrophosphate release is used to generate light for sequence detection. Four hundred fifty four sequencing offers read lengths longer than most NGS technologies, although the cost is also greater. The Illumina approach attaches single template DNA molecules to a solid surface prior to polymerase-based amplification and the inclusion of basespecific fluorescently labelled reversible terminator bases, images of which translate into sequence detection. SOLiD sequencing operates by competing octamer ligation to a primer annealed to the DNA template, where the optimal octamer match is indicated by a fluorescent label prior to cleavage and ligation of the next octamer. Ion Torrent sequencing follows a similar process to 454 sequencing but detects hydrogen ions released during nucleotide addition in place of fluorescence. More recently third generation sequencing from single molecules has become available using Pacific Biosciences (PacBio) technology. PacBio offers extremely long reads of 20,000 bases or more, albeit with an error rate greater than the NGS technologies. Several other technologies have been described in this fast developing field and are at various stages of development (Metzker, 2010). Most recently, introduction of new NGS sequencing platforms including the Ion Torrent Personal Genome Machine (PGM), Illumina MiSeq and Pacific Biosciences RS provide opportunities for

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Fig. 1. Published genomics resources available for Eimeria species parasites. A phylogeny representing 18S rDNA sequences from nine Eimeria species, constructed by maximum likelihood with the model TN93 + G (selected based upon Bayesian Information Criterion) and 1000 bootstrap replication using MEGA v6.06 (Tamura et al., 2013). Accession numbers for the sequences used (top to bottom) U67120, EF122251, U67116, EF210324, U67118, U67119, EF210325, AF080614, U77084. Species names underlined cause haemorrhagic disease in chickens with the number of strains sequenced per species shown as n. Tier 1 = high quality genome assembly, benefitting from manual improvements post-assembly; Tier 2 = draft/improved, benefitting from automated improvements post-assembly; Tier 3 = draft only, no post-assembly improvement. Genome and transcriptome sequence resource references a: Reid et al., 2014; b: Blake et al., 2012; Schwarz et al., 2010; d: Dong et al., 2011; e: Novaes et al., 2012; f: Aarthi et al., 2011; g: Kim et al., 2010; h: Miska et al., 2008; i: Wan et al., 1999; j: Amiruddin et al., 2012; k: Tuda et al., 2011; l: Walker et al., 2015; m: Heitlinger et al., 2014; Abrahamsen et al., 1993.

longer read lengths and (for the first two) improved availability for smaller labs (Quail et al., 2012). 4. Application to eimerian genomes Genome sequencing started for Eimeria in 2002, beginning with the reference Eimeria tenella Houghton strain (Chapman and Shirley, 2003). Prior to 2002 genomic resources were scarce, largely limited to Sanger sequencing reads covering the ribosomal DNA clusters, sporozoite and second generation merozoite expressed sequence tag (EST) cDNA reads, sequenced random amplified polymorphic DNA (RAPD) markers, and a panel of specific proteincoding genes including several kinases and microneme proteins (e.g. Jenkins, 1988; Dunn et al., 1996; Wan et al., 1999; Tomley et al., 2001; Fernandez et al., 2003). For the genome, Sanger sequencing was used initially at the Wellcome Trust Sanger Institute, producing approximately eight-fold genome coverage (Shirley et al., 2004; Reid et al., 2014). Complementary projects sequencing the smallest chromosome within the E. tenella genome and open reading frame expressed sequence tag (ORESTES) cDNA reads derived from five E. acervulina, E. maxima and E. tenella lifecycle stages used the same technology (Ling et al., 2007; Novaes et al., 2012). Subsequently NGS technologies promoted an expansion of genome sequencing for Eimeria, using 454 pyrosequencing to generate more than 12-fold coverage of the E. maxima Houghton strain genome (Blake et al., 2012) and Illumina to improve the E. tenella genome assembly as well as generate the first assemblies for seven other Eimeria species/strains including one species which infects the mouse (Eimeria falciformis; Fig. 1) (Heitlinger et al., 2014; Reid et al., 2014). Prior to the NGS revolution generating such resources would have been impractical, if not impossible, for Eimeria. At the time of writing genome sequence assemblies are available for eight Eimeria species ranging from high quality (tier 1, benefitting from manual improvements post-assembly) to draft/improved (tier 2, automated improvements post-assembly) and draft only (tier 3, no

post-assembly improvement; Fig. 1) (Reid et al., 2014). Comparison with the Toxoplasma gondii genome assembly suggests the Tier 1 and 2 eimerian genomes all exceed 93% completeness, although they remain highly fragmented with most including several thousand scaffolds as a consequence of the high frequency of repetitive sequence (Heitlinger et al., 2014; Reid et al., 2014). A common feature of all eimerian genomes sequenced and analysed to date has been the frequent occurrence of trimer CAG (and other derivative) and heptamer AAACCCT/AGGGTTT repeats, with the former common as homopolymeric amino acid repeats (HAARs) within protein coding but not functional domain sequences. A segmental genome-wide occurrence of repetitive sequences permitted the description of distinct repeat-rich (R; defined as presenting a repeat density in excess of 5%) and opposing repeat-poor (P) regions for all sequenced eimerian genomes, although R region occurrence and boundaries varied between species (Reid et al., 2014). Genome-wide gene identification has predicted more than 8000 protein coding genes in each of the haemorrhagic eimerian genomes and E. mitis, with far fewer detected for the other species (Heitlinger et al., 2014; Reid et al., 2014). While absolute gene numbers may vary as the assemblies improve this balance is likely to persist. One other striking feature of the genes predicted has been the occurrence of sequences encoding families of single domain glycosylphosphatidylinositol (GPI) anchored surface antigens (SAGs). More than ten-fold variation has been detected in sag gene number between the genomes of the chicken-infecting Eimeria, displaying an incomplete association with pathogenicity and (inversely) immunogenicity (Reid et al., 2014). Comparison with the E. falciformis genome revealed the conserved occurrence of clusters of sag-like genes, albeit with distinct clades for the murine and avian Eimeria (Heitlinger et al., 2014). Metabolic pathway reconstruction confirmed close conservation between Eimeria and other sequenced coccidian parasites such as T. gondii, with established differences including the enzymatic basis of a mannitol cycle in Eimeria verified (Reid et al., 2014). The previous description

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of retrotransposon-like elements within the first sequenced E. tenella chromosome was confirmed in the avian Eimeria genomes and found to be related to the chromovirus long-terminal repeat (LTR) retrotransposons, although most were highly fragmented and unlikely to have been functional for many generations (Ling et al., 2007; Reid et al., 2014). Numerous genes containing ApiAP2 family DNA-binding domains were identified with a tentative linkage between ApiAP2 gene number and genome size rather than life cycle complexity (Reid et al., 2014).

to access these stages. Reduction in the quantity of RNA (e.g. using the transposome-based Nextera system) or the number of cells required for RNAseq (Gertz et al., 2012; Vollmers et al., 2015), as well as improved host genomic resources to screen out contaminating material, will offer further refinements. Improvements in proteomic technologies now allow validation of transcriptomic data and definition of post-transcriptional variation (Lal et al., 2009; Oakes et al., 2013; Shen et al., 2014b).

6. What next for Eimeria genomics? 5. Transcriptomic analyses Eimerian parasites follow a strict faecal-oral life cycle which features three distinct phases and no intermediate host. The first phase, termed sporogony or sporulation, occurs outside of the host and includes asexual replication as the parasite develops from a single non-infectious unsporulated oocyst into an infective sporulated oocyst, containing eight sporozoites accommodated in pairs within four sporocysts. The second phase, known as schizogony or merogony, occurs within the host and commonly includes two to four rounds of asexual replication, with the number depending on the species. For Eimeria that infect chickens this occurs in the intestine, with the exact location depending on the species (Long et al., 1976). The final phase includes sexual differentiation (gametogony) and fertilisation, leading to formation and shedding of progeny oocysts. Exogenous and extracellular endogenous life cycle forms including the oocysts (unsporulated, sporulating and sporulated), sporozoites and merozoites have been most commonly sampled for transcriptomic analyses as a consequence of their availability, first as ESTs/ORESTES and more recently by Illumina-based RNAseq (Fig. 1) (Abrahamsen et al., 1993; Wan et al., 1999; Miska et al., 2008; Kim et al., 2010; Schwarz et al., 2010; Aarthi et al., 2011; Dong et al., 2011; Tuda et al., 2011; Amiruddin et al., 2012; Novaes et al., 2012; Heitlinger et al., 2014; Reid et al., 2014; Walker et al., 2015). As described for genome sequencing, progress from Sanger to NGS technologies has provided opportunities to expand transcriptome sequencing to additional E. tenella life cycle stages and species such as E. falciformis. The data generated by RNAseq transcriptomic sequencing has been invaluable during annotation of the eimerian genomes, supporting identification of transcription start sites, intron/exon boundaries and alternatively spliced transcripts. Comparison between life cycle stages supports detection of stage-specific protein profiles while the depth of sequence coverage available allows relative assessment of transcription levels using reads per kilobase of exon per million reads mapped (RPKM; also referred to as fragments per kilobase of exon per million reads mapped – FPKM). Among the most notable findings has been the increasing stage-specific complexity of E. tenella sag gene transcription as the endogenous life cycle progresses. Until recently parasite, and thus mRNA template availability has been a major limiting factor for eimerian transcriptomics. The requirement for microgram quantities of total RNA for library preparation restricted transcriptomic sequencing to only the most accessible lifecycle stages, hindering understanding of much of the eimerian life cycle’s complexity. These problems have been overcome for some species and life cycle stages, such as E. tenella gametocytes, where specific culture and recovery protocols have allowed complementation of previous genomic analyses through development of the first RNAseq data (Katrib et al., 2012; Walker et al., 2015). Life cycle stages including trophozoites and developing schizonts are yet to be sampled, but complementary techniques including laser dissection from in vitro or in vivo cultured samples, ex vivo explant culture and fluorescence-activated cell sorting (FACS) of parasites transfected to express fluorescent reporter genes (Clark et al., 2008; Yan et al., 2009) offer good opportunities

The publication of genome sequences for all seven Eimeria species which infect chickens represented the culmination of more than a decade of work from a consortium of 20 institutions across ten countries (Reid et al., 2014). The provision of reference genome assemblies, combined with the reduced cost and greater power of NGS technologies, now provides enormous opportunity for genomics-led studies and diagnostics with these parasites. Access to a first mammalian-infecting Eimeria genome assembly can support comparative analyses and work in a convenient laboratory model (Heitlinger et al., 2014). Sequencing additional reference or field isolates of these Eimeria species has begun with two, the US Wisconsin and the Japanese Nippon-2 E. tenella strains, already released (McDougald and Jeffers, 1976; Reid et al., 2014). Collections of strains with defined phenotypes including drug resistance, precocious development, adaptation to development within the chick embryo, differential virulence and strain-specific antigenicity are available, providing resources for sequencing (e.g. Shirley and Harvey, 2000; Bedrnik, 2003; Williams et al., 2009; Blake et al., 2011). Genomic data from these and other parasites can provide invaluable insights into eimerian biology and offer realistic routes to improving anticoccidial control using existing and novel drugs or vaccines. One of the biggest challenges of working with parasite and other genome sequence assemblies remains the annotation of coding sequences. For E. tenella more than 70% of the gene models are currently annotated as hypothetical. RNAseq and proteomics have proven invaluable in the construction and validation of gene models (Reid et al., 2012), but similarity-based annotation is inherently limited to that which is already known. Prediction of features including signal peptides, specific domains and motifs can be informative, but high-throughput coding sequence annotation awaits technological advance. For Trypanosoma brucei systematic analysis of gene function using RNA interference has proven feasible, and bar coded genetic modification vectors now provide a valuable reverse genetic tool for Plasmodium berghei (Subramaniam et al., 2006; Gomes et al., 2015). For Eimeria, advances in transfection technologies including random integration of selectable markers and piggyback transposon-mediated transgenesis offer opportunities, as do older technologies including signature-tagged mutagenesis described with T. gondii (Knoll et al., 2001; Clark et al., 2008; Yan et al., 2009; Su et al., 2012). Adaptation of the prokaryotic CRISPR/Cas9 genome editing system, recently developed for use with T. gondii, may offer the greatest potential for work with Eimeria (Shen et al., 2014a; Sidik et al., 2014). Challenges remaining for the Eimeria genomes include improving the sequence assemblies by closing the many thousands of gaps (Lim et al., 2012; Reid et al., 2014). Optical mapping has been used to order many contigs into a much smaller number of super scaffolds for some species, but the highly repetitive nature of the eimerian genomes consistently hinders closure. Increased coverage using more cost-effective NGS technology and varied library size pairedend/mate-paired sequencing offers some prospects, although read length remains limiting. Thus, access to longer reads from the third generation sequencing technologies can play an important role in

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assembly improvement. Similarly, construction of de novo genome assemblies is becoming much more feasible for previously unsampled Eimeria species and other coccidians. Using single-cell and low input amplification-based genomics rare or environmental isolates can be considered for sequencing with little or no need for in vivo culture (Nair et al., 2014). Even mixed species samples can be sequenced, with reads extracted and assembled by comparison to the reference assemblies with host, microbial, dietary or environmental contamination screened out (Gandhi et al., 2012; Manske et al., 2012). Taking a broader view, NGS-based genomics and transcriptomics will also support improved understanding of host responses to eimerian infection as well as the influence these parasites may exert on their host genomes and the enteric microbiome. 7. Applications for Eimeria genomics? Addition of Eimeria to the ever-expanding club of sequenced genomes has dramatically expanded the scope for studies of biology, prophylaxis, therapy and vaccination. Comparison between genome assemblies representing related coccidians is straightforward and accessible using GeneDB, or ToxoDB within the EuPathDB resource (http://toxodb.org/toxo/http://toxodb.org/toxo/); (Gajria et al., 2008; Logan-Klumpler et al., 2012; Aurrecoechea et al., 2013). ToxoDB currently facilitates access to eight Eimeria species genomes in addition to genome assemblies for Hammondia hammondi, Neospora caninum, Sarcocystis neurona and 18 T. gondii genomes (Reid et al., 2012; Blazejewski et al., 2015). Comparison can be used to explore the conservation of relevant protein coding genes including drug targets and vaccine candidates and extrapolate experimental findings to other parasite species and genera, benefiting from the presence or absence of gene synteny where available. More specifically for Eimeria, comparison of genomic data generated from phenotypically distinct parasite lines can be used to identify the genetic basis of the observed variation (e.g. coding, promoter and controlling sequences), first by genomewide association study (GWAS) and eventually to the causative polymorphism (Kinga Modrzynska et al., 2012). Comparison of data from spatially or temporally distinct isolates can be used to explore parasite population biology and structure, informing on the likely efficacy and longevity of novel gene/protein targeted intervention strategies including sub-unit vaccination and chemoprophylaxis/therapy. In the short term genomic resources can be used to develop single nucleotide polymorphism (SNP), microsatellite, multi-locus sequence type (MLST), copy-number variation (CNV) or other genetic markers in pursuit of these objectives and as molecular diagnostics, but as the assemblies improve whole genome analyses will become routine. Ultimately, whole or targeted genome sequencing will be applicable for parasite detection and diagnosis, providing details of the likely anticoccidial susceptibility of a field population in place of costly in vivo determination by anticoccidial sensitivity test (AST; Holdsworth et al., 2004) and indicating the optimal drug and/or vaccine intervention strategy. 8. Conclusions The landmark publication of eight eimerian genome sequences in 2014 has provided a massive boost for studies with Eimeria, related apicomplexans and many protozoa. Given greater access to sequence data and the rapidly developing availability of NGS technologies with reduced cost, experimental limitations have now moved on to our capacity for data analysis and interpretation (Padmanabhan et al., 2013). Looking to the future improved bioinformatics pipelines will be required to improve accessibility to these data. Many parasites of veterinary and environmental

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significance are now experiencing similar revolutionary advances, providing immense opportunities whilst challenging our ability to handle and utilise the data generated. References Aarthi, S., Raj, G.D., Raman, M., Blake, D., Subramaniam, C., Tomley, F., 2011. Expressed sequence tags from Eimeria brunetti-preliminary analysis and functional annotation. Parasitol. Res. 108, 1059–1062. Abrahamsen, M., Clark, T., Mascolo, P., Speer, C., White, M., 1993. Developmental gene expression in Eimeria bovis. Mol. Biochem. Parasitol. 57, 1–14. Amiruddin, N., Lee, X.W., Blake, D.P., Suzuki, Y., Tay, Y.L., Lim, L.S., Tomley, F.M., Watanabe, J., Sugimoto, C., Wan, K.L., 2012. Characterisation of full-length cDNA sequences provides insights into the Eimeria tenella transcriptome. BMC Genomics 13, 21. Aurrecoechea, C., Barreto, A., Brestelli, J., Brunk, B.P., Cade, S., Doherty, R., Fischer, S., Gajria, B., Gao, X., Gingle, A., Grant, G., Harb, O.S., Heiges, M., Hu, S., Iodice, J., Kissinger, J.C., Kraemer, E.T., Li, W., Pinney, D.F., Pitts, B., Roos, D.S., Srinivasamoorthy, G., Stoeckert Jr., C.J., Wang, H., Warrenfeltz, S., 2013. EuPathDB: the eukaryotic pathogen database. Nucleic Acids Res. 41, D684–691. Bacciu, N., Bed’Hom, B., Filangi, O., Rome, H., Gourichon, D., Reperant, J.M., Le Roy, P., Pinard-van der Laan, M.H., Demeure, O., 2014. QTL detection for coccidiosis (Eimeria tenella) resistance in a Fayoumi × Leghorn F(2) cross, using a medium-density SNP panel. Genet. Sel. Evol. 46, 14. Bedrnik, P., 2003. Comment on the review anticoccidial vaccines for broiler chickens: pathways to success by R.B. Williams (2002). Avian Pathology, 31, 317–353. Avian Pathol. 32, 219. Bennett, S.T., Barnes, C., Cox, A., Davies, L., Brown, C., 2005. Toward the 1000 dollars human genome. Pharmacogenomics 6, 373–382. Blake, D.P., Alias, H., Billington, K.J., Clark, E.L., Mat-Isa, M.N., Mohamad, A.F., Mohd-Amin, M.R., Tay, Y.L., Smith, A.L., Tomley, F.M., Wan, K.L., 2012. EmaxDB: availability of a first draft genome sequence for the apicomplexan Eimeria maxima. Mol. Biochem. Parasitol. 184, 48–51. Blake, D.P., Billington, K.J., Copestake, S.L., Oakes, R.D., Quail, M.A., Wan, K.L., Shirley, M.W., Smith, A.L., 2011. Genetic mapping identifies novel highly protective antigens for an apicomplexan parasite. PLoS Pathog. 7, e1001279. Blake, D.P., Hesketh, P., Archer, A., Carroll, F., Shirley, M.W., Smith, A.L., 2005. The influence of immunizing dose size and schedule on immunity to subsequent challenge with antigenically distinct strains of Eimeria maxima. Avian Pathol. 34, 489–494. Blake, D.P., Tomley, F.M., 2014. Securing poultry production from the ever-present Eimeria challenge. Trends Parasitol. 30, 12–19. Blazejewski, T., Nursimulu, N., Pszenny, V., Dangoudoubiyam, S., Namasivayam, S., Chiasson, M.A., Chessman, K., Tonkin, M., Swapna, L.S., Hung, S.S., Bridgers, J., Ricklefs, S.M., Boulanger, M.J., Dubey, J.P., Porcella, S.F., Kissinger, J.C., Howe, D.K., Grigg, M.E., Parkinson, J., 2015. Systems-based analysis of the Sarcocystis neurona genome identifies pathways that contribute to a heteroxenous life cycle. MBio 6. Chapman, H.D., Barta, J.R., Blake, D., Gruber, A., Jenkins, M., Smith, N.C., Suo, X., Tomley, F.M., 2013. A selective review of advances in coccidiosis research. Adv. Parasitol. 83, 93–171. Chapman, H.D., Jeffers, T.K., 2014. Vaccination of chickens against coccidiosis ameliorates drug resistance in commercial poultry production. Int. J. Parasitol. Drugs Drug Resist. 4, 214–217. Chapman, H.D., Shirley, M.W., 2003. The Houghton strain of Eimeria tenella: a review of the type strain selected for genome sequencing. Avian Pathol. 32, 115–127. Clark, J.D., Billington, K., Bumstead, J.M., Oakes, R.D., Soon, P.E., Sopp, P., Tomley, F.M., Blake, D.P., 2008. A toolbox facilitating stable transfection of Eimeria species. Mol. Biochem. Parasitol. 162, 77–86. Dalloul, R.A., Lillehoj, H.S., 2006. Poultry coccidiosis: recent advancements in control measures and vaccine development. Expert Rev. Vaccines 5, 143–163. Dong, H., Lin, J., Han, H., Jiang, L., Zhao, Q., Zhu, S., Huang, B., 2011. Analysis of differentially expressed genes in the precocious line of Eimeria maxima and its parent strain using suppression subtractive hybridization and cDNA microarrays. Parasitol. Res. 108, 1033–1040. Dunn, P.P., Bumstead, J.M., Tomley, F.M., 1996. Sequence, expression and localization of calmodulin-domain protein kinases in Eimeria tenella and Eimeria maxima. Parasitology 113 (Pt 5), 439–448. Fernandez, S., Costa, A.C., Katsuyama, A.M., Madeira, A.M., Gruber, A., 2003. A survey of the inter- and intraspecific RAPD markers of Eimeria spp. of the domestic fowl and the development of reliable diagnostic tools. Parasitol. Res. 89, 437–445. Gajria, B., Bahl, A., Brestelli, J., Dommer, J., Fischer, S., Gao, X., Heiges, M., Iodice, J., Kissinger, J.C., Mackey, A.J., Pinney, D.F., Roos, D.S., Stoeckert Jr., C.J., Wang, H., Brunk, B.P., 2008. ToxoDB: an integrated Toxoplasma gondii database resource. Nucleic Acids Res. 36, D553–556. Gandhi, K., Thera, M.A., Coulibaly, D., Traore, K., Guindo, A.B., Doumbo, O.K., Takala-Harrison, S., Plowe, C.V., 2012. Next generation sequencing to detect variation in the Plasmodium falciparum circumsporozoite protein. Am. J. Trop. Med. Hyg. 86, 775–781. Gertz, J., Varley, K.E., Davis, N.S., Baas, B.J., Goryshin, I.Y., Vaidyanathan, R., Kuersten, S., Myers, R.M., 2012. Transposase mediated construction of RNA-seq libraries. Genome Res. 22, 134–141.

Please cite this article in press as: Blake, D.P., Eimeria genomics: Where are we now and where are we going? Vet. Parasitol. (2015), http://dx.doi.org/10.1016/j.vetpar.2015.05.007

G Model VETPAR-7630; No. of Pages 7 6

ARTICLE IN PRESS D.P. Blake / Veterinary Parasitology xxx (2015) xxx–xxx

Gibson-Kueh, S., Yang, R., Thuy, N.T., Jones, J.B., Nicholls, P.K., Ryan, U., 2011. The molecular characterization of an Eimeria and Cryptosporidium detected in Asian seabass (Lates calcarifer) cultured in Vietnam. Vet. Parasitol. 181, 91–96. Goffeau, A., Barrell, B.G., Bussey, H., Davis, R.W., Dujon, B., Feldmann, H., Galibert, F., Hoheisel, J.D., Jacq, C., Johnston, M., Louis, E.J., Mewes, H.W., Murakami, Y., Philippsen, P., Tettelin, H., Oliver, S.G., 1996. Life with 6000 genes. Science 274 (546), 547–563. Gomes, A.R., Bushell, E., Schwach, F., Girling, G., Anar, B., Quail, M.A., Herd, C., Pfander, C., Modrzynska, K., Rayner, J.C., Billker, O., 2015. A genome-scale vector resource enables high-throughput reverse genetic screening in a malaria parasite. Cell Host Microbe 17, 404–413. Heitlinger, E., Spork, S., Lucius, R., Dieterich, C., 2014. The genome of Eimeria falciformis – reduction and specialization in a single host apicomplexan parasite. BMC Genomics 15, 696. Holdsworth, P.A., Conway, D.P., McKenzie, M.E., Dayton, A.D., Chapman, H.D., Mathis, G.F., Skinner, J.T., Mundt, H.C., Williams, R.B., 2004. World Association for the Advancement of Veterinary Parasitology (WAAVP) guidelines for evaluating the efficacy of anticoccidial drugs in chickens and turkeys. Vet. Parasitol. 121, 189–212. Jeanes, C., Vaughan-Higgins, R., Green, R.E., Sainsbury, A.W., Marshall, R.N., Blake, D.P., 2013. Two new Eimeria species parasitic in corncrakes (Crex crex) (Gruiformes: Rallidae) in the United Kingdom. J. Parasitol. 99, 634–638. Jenkins, M.C., 1988. A cDNA encoding a merozoite surface protein of the protozoan Eimeria acervulina contains tandem-repeated sequences. Nucleic Acids Res. 16, 9863. Jirku, M., Kvicerova, J., Modry, D., Hypsa, V., 2013. Evolutionary plasticity in coccidia – striking morphological similarity of unrelated coccidia (apicomplexa) from related hosts: Eimeria spp. from African and Asian Pangolins (Mammalia: Pholidota). Protist 164, 470–481. Jirku, M., Obornik, M., Lukes, J., Modry, D., 2009. A model for taxonomic work on homoxenous coccidia: redescription, host specificity, and molecular phylogeny of Eimeria ranae Dobell, 1909, with a review of anuran-host Eimeria (Apicomplexa: Eimeriorina). J. Eukaryot. Microbiol. 56, 39–51. Katrib, M., Ikin, R.J., Brossier, F., Robinson, M., Slapetova, I., Sharman, P.A., Walker, R.A., Belli, S.I., Tomley, F.M., Smith, N.C., 2012. Stage-specific expression of protease genes in the apicomplexan parasite, Eimeria tenella. BMC Genomics 13, 685. Kim, C.H., Lillehoj, H.S., Hong, Y.H., Keeler Jr., C.L., Lillehoj, E.P., 2010. Comparison of global transcriptional responses to primary and secondary Eimeria acervulina infections in chickens. Dev. Comp. Immunol. 34, 344–351. Kim, D.K., Hong, Y.H., Park, D.W., Lamont, S.J., Lillehoj, H.S., 2008. Differential immune-related gene expression in two genetically disparate chicken lines during infection by Eimeria maxima. Dev. Biol. (Basel) 132, 131–140. Kinga Modrzynska, K., Creasey, A., Loewe, L., Cezard, T., Trindade Borges, S., Martinelli, A., Rodrigues, L., Cravo, P., Blaxter, M., Carter, R., Hunt, P., 2012. Quantitative genome re-sequencing defines multiple mutations conferring chloroquine resistance in rodent malaria. BMC Genomics 13, 106. Knoll, L.J., Furie, G.L., Boothroyd, J.C., 2001. Adaptation of signature-tagged mutagenesis for Toxoplasma gondii: a negative screening strategy to isolate genes that are essential in restrictive growth conditions. Mol. Biochem. Parasitol. 116, 11–16. Knowles, S., Fenton, A., Petchey, O., Jones, T., Barber, R., Pedersen, A., 2013. Stability of within-host-parasite communities in a wild mammal system. Proc. R. Soc. London Ser. B 280, 20130598. Kuska, B., 1998. Beer, Bethesda, and biology: how genomics came into being. J. Natl. Cancer Inst. 90, 93. Kvicerova, J., Hypsa, V., 2013. Host-parasite incongruences in rodent Eimeria suggest significant role of adaptation rather than cophylogeny in maintenance of host specificity. PLoS One 8, e63601. Lal, K., Bromley, E., Oakes, R., Prieto, J.H., Sanderson, S.J., Kurian, D., Hunt, L., Yates 3rd, J.R., Wastling, J.M., Sinden, R.E., Tomley, F.M., 2009. Proteomic comparison of four Eimeria tenella life-cycle stages: unsporulated oocyst, sporulated oocyst, sporozoite and second-generation merozoite. Proteomics 9, 4566–4576. Lassen, B., Ostergaard, S., 2012. Estimation of the economical effects of Eimeria infections in Estonian dairy herds using a stochastic model. Prev. Vet. Med. 106, 258–265. Lim, L.S., Tay, Y.L., Alias, H., Wan, K.L., Dear, P.H., 2012. Insights into the genome structure and copy-number variation of Eimeria tenella. BMC Genomics 13, 389. Ling, K.H., Rajandream, M.A., Rivailler, P., Ivens, A., Yap, S.J., Madeira, A.M., Mungall, K., Billington, K., Yee, W.Y., Bankier, A.T., Carroll, F., Durham, A.M., Peters, N., Loo, S.S., Isa, M.N., Novaes, J., Quail, M., Rosli, R., Nor Shamsudin, M., Sobreira, T.J., Tivey, A.R., Wai, S.F., White, S., Wu, X., Kerhornou, A., Blake, D., Mohamed, R., Shirley, M., Gruber, A., Berriman, M., Tomley, F., Dear, P.H., Wan, K.L., 2007. Sequencing and analysis of chromosome 1 of Eimeria tenella reveals a unique segmental organization. Genome Res. 17, 311–319. Logan-Klumpler, F.J., De Silva, N., Boehme, U., Rogers, M.B., Velarde, G., McQuillan, J.A., Carver, T., Aslett, M., Olsen, C., Subramanian, S., Phan, I., Farris, C., Mitra, S., Ramasamy, G., Wang, H., Tivey, A., Jackson, A., Houston, R., Parkhill, J., Holden, M., Harb, O.S., Brunk, B.P., Myler, P.J., Roos, D., Carrington, M., Smith, D.F., Hertz-Fowler, C., Berriman, M., 2012. GeneDB – an annotation database for pathogens. Nucleic Acids Res. 40, D98–108. Long, P., Joyner, L., Millard, B., Norton, C., 1976. A guide to laboratory techniques used in the study and diagnosis of avian coccidiosis. Folia Vet. Lat. 6, 201–217. Manske, M., Miotto, O., Campino, S., Auburn, S., Almagro-Garcia, J., Maslen, G., O’Brien, J., Djimde, A., Doumbo, O., Zongo, I., Ouedraogo, J.B., Michon, P., Mueller, I., Siba, P., Nzila, A., Borrmann, S., Kiara, S.M., Marsh, K., Jiang, H., Su,

X.Z., Amaratunga, C., Fairhurst, R., Socheat, D., Nosten, F., Imwong, M., White, N.J., Sanders, M., Anastasi, E., Alcock, D., Drury, E., Oyola, S., Quail, M.A., Turner, D.J., Ruano-Rubio, V., Jyothi, D., Amenga-Etego, L., Hubbart, C., Jeffreys, A., Rowlands, K., Sutherland, C., Roper, C., Mangano, V., Modiano, D., Tan, J.C., Ferdig, M.T., Amambua-Ngwa, A., Conway, D.J., Takala-Harrison, S., Plowe, C.V., Rayner, J.C., Rockett, K.A., Clark, T.G., Newbold, C.I., Berriman, M., MacInnis, B., Kwiatkowski, D.P., 2012. Analysis of Plasmodium falciparum diversity in natural infections by deep sequencing. Nature 487, 375–379. Margulies, M., Egholm, M., Altman, W.E., Attiya, S., Bader, J.S., Bemben, L.A., Berka, J., Braverman, M.S., Chen, Y.J., Chen, Z., Dewell, S.B., Du, L., Fierro, J.M., Gomes, X.V., Godwin, B.C., He, W., Helgesen, S., Ho, C.H., Irzyk, G.P., Jando, S.C., Alenquer, M.L., Jarvie, T.P., Jirage, K.B., Kim, J.B., Knight, J.R., Lanza, J.R., Leamon, J.H., Lefkowitz, S.M., Lei, M., Li, J., Lohman, K.L., Lu, H., Makhijani, V.B., McDade, K.E., McKenna, M.P., Myers, E.W., Nickerson, E., Nobile, J.R., Plant, R., Puc, B.P., Ronan, M.T., Roth, G.T., Sarkis, G.J., Simons, J.F., Simpson, J.W., Srinivasan, M., Tartaro, K.R., Tomasz, A., Vogt, K.A., Volkmer, G.A., Wang, S.H., Wang, Y., Weiner, M.P., Yu, P., Begley, R.F., Rothberg, J.M., 2005. Genome sequencing in microfabricated high-density picolitre reactors. Nature 437, 376–380. Maxam, A.M., Gilbert, W., 1977. A new method for sequencing DNA. Proc. Natl. Acad. Sci. U. S. A. 74, 560–564. McDougald, L.R., Jeffers, T.K., 1976. Eimeria tenella (Sporozoa, Coccidia): gametogony following a single asexual generation. Science 192, 258–259. Metzker, M.L., 2010. Sequencing technologies – the next generation. Nat. Rev. Genet. 11, 31–46. Miska, K.B., Fetterer, R.H., Rosenberg, G.H., 2008. Analysis of transcripts from intracellular stages of Eimeria acervulina using expressed sequence tags. J. Parasitol. 94, 462–466. Nair, S., Nkhoma, S.C., Serre, D., Zimmerman, P.A., Gorena, K., Daniel, B.J., Nosten, F., Anderson, T.J., Cheeseman, I.H., 2014. Single-cell genomics for dissection of complex malaria infections. Genome Res. 24, 1028–1038. Novaes, J., Rangel, L.T., Ferro, M., Abe, R.Y., Manha, A.P., de Mello, J.C., Varuzza, L., Durham, A.M., Madeira, A.M., Gruber, A., 2012. A comparative transcriptome analysis reveals expression profiles conserved across three Eimeria spp. of domestic fowl and associated with multiple developmental stages. Int J. Parasitol. 42, 39–48. Novilla, M.N., Carpenter, J.W., 2004. Pathology and pathogenesis of disseminated visceral coccidiosis in cranes. Avian Pathol. 33, 275–280. Oakes, R.D., Kurian, D., Bromley, E., Ward, C., Lal, K., Blake, D.P., Reid, A.J., Pain, A., Sinden, R.E., Wastling, J.M., Tomley, F.M., 2013. The rhoptry proteome of Eimeria tenella sporozoites. Int. J. Parasitol. 43, 181–188. Oliver, S.G., van der Aart, Q.J., Agostoni-Carbone, M.L., Aigle, M., Alberghina, L., Alexandraki, D., Antoine, G., Anwar, R., Ballesta, J.P., Benit, P., 1992. The complete DNA sequence of yeast chromosome III. Nature 357, 38–46. Padmanabhan, R., Mishra, A.K., Raoult, D., Fournier, P.E., 2013. Genomics and metagenomics in medical microbiology. J. Microbiol. Methods 95, 415–424. Quail, M.A., Smith, M., Coupland, P., Otto, T.D., Harris, S.R., Connor, T.R., Bertoni, A., Swerdlow, H.P., Gu, Y., 2012. A tale of three next generation sequencing platforms: comparison of Ion Torrent, Pacific Biosciences and Illumina MiSeq sequencers. BMC Genomics 13, 341. Radostits, O.M., Stockdale, P.H., 1980. A brief review of bovine coccidiosis in Western Canada. Can. Vet. J. 21, 227–230. Reid, A.J., Blake, D.P., Ansari, H.R., Billington, K., Browne, H.P., Bryant, J., Dunn, M., Hung, S.S., Kawahara, F., Miranda-Saavedra, D., Malas, T.B., Mourier, T., Naghra, H., Nair, M., Otto, T.D., Rawlings, N.D., Rivailler, P., Sanchez-Flores, A., Sanders, M., Subramaniam, C., Tay, Y.L., Woo, Y., Wu, X., Barrell, B., Dear, P.H., Doerig, C., Gruber, A., Ivens, A.C., Parkinson, J., Rajandream, M.A., Shirley, M.W., Wan, K.L., Berriman, M., Tomley, F.M., Pain, A., 2014. Genomic analysis of the causative agents of coccidiosis in domestic chickens. Genome Res. 24, 1676–1685. Reid, A.J., Vermont, S.J., Cotton, J.A., Harris, D., Hill-Cawthorne, G.A., Konen-Waisman, S., Latham, S.M., Mourier, T., Norton, R., Quail, M.A., Sanders, M., Shanmugam, D., Sohal, A., Wasmuth, J.D., Brunk, B., Grigg, M.E., Howard, J.C., Parkinson, J., Roos, D.S., Trees, A.J., Berriman, M., Pain, A., Wastling, J.M., 2012. Comparative genomics of the apicomplexan parasites Toxoplasma gondii and Neospora caninum: Coccidia differing in host range and transmission strategy. PLoS Pathog. 8, e1002567. Revets, H., Dekegel, D., Deleersnijder, W., De Jonckheere, J., Peeters, J., Leysen, E., Hamers, R., 1989. Identification of virus-like particles in Eimeria stiedae. Mol. Biochem. Parasitol. 36, 209–215. Sanger, F., Nicklen, S., Coulson, A.R., 1977. DNA sequencing with chain-terminating inhibitors. Proc. Natl. Acad. Sci. U. S. A. 74, 5463–5467. Schwarz, R.S., Fetterer, R.H., Rosenberg, G.H., Miska, K.B., 2010. Coccidian merozoite transcriptome analysis from Eimeria maxima in comparison to Eimeria tenella and Eimeria acervulina. J. Parasitol. 96, 49–57. Shen, B., Brown, K.M., Lee, T.D., Sibley, L.D., 2014a. Efficient gene disruption in diverse strains of toxoplasma gondii using CRISPR/CAS9. MBio 5, e01114–01114. Shen, X.J., Li, T., Fu, J.J., Zhang, K.Y., Wang, X.Y., Liu, Y.C., Zhang, H.J., Fan, C., Fei, C.Z., Xue, F.Q., 2014b. Proteomic analysis of the effect of diclazuril on second-generation merozoites of Eimeria tenella. Parasitol. Res. 113, 903–909. Shirley, M.W., Harvey, D.A., 2000. A genetic linkage map of the apicomplexan protozoan parasite Eimeria tenella. Genome Res. 10, 1587–1593. Shirley, M.W., Ivens, A., Gruber, A., Madeira, A.M., Wan, K.L., Dear, P.H., Tomley, F.M., 2004. The Eimeria genome projects: a sequence of events. Trends Parasitol. 20, 199–201.

Please cite this article in press as: Blake, D.P., Eimeria genomics: Where are we now and where are we going? Vet. Parasitol. (2015), http://dx.doi.org/10.1016/j.vetpar.2015.05.007

G Model VETPAR-7630; No. of Pages 7

ARTICLE IN PRESS D.P. Blake / Veterinary Parasitology xxx (2015) xxx–xxx

Sidik, S.M., Hackett, C.G., Tran, F., Westwood, N.J., Lourido, S., 2014. Efficient genome engineering of toxoplasma gondii using CRISPR/Cas9. PLoS One 9, e100450. Su, H., Liu, X., Yan, W., Shi, T., Zhao, X., Blake, D.P., Tomley, F.M., Suo, X., 2012. PiggyBac transposon-mediated transgenesis in the apicomplexan parasite Eimeria tenella. PLoS One 7, e40075. Subramaniam, C., Veazey, P., Redmond, S., Hayes-Sinclair, J., Chambers, E., Carrington, M., Gull, K., Matthews, K., Horn, D., Field, M.C., 2006. Chromosome-wide analysis of gene function by RNA interference in the african trypanosome. Eukaryot. Cell 5, 1539–1549. Tamura, K., Stecher, G., Peterson, D., Filipski, A., Kumar, S., 2013. MEGA6: molecular evolutionary genetics analysis version 6.0. Mol. Biol. Evol. 30, 2725–2729. Tomley, F.M., Billington, K.J., Bumstead, J.M., Clark, J.D., Monaghan, P., 2001. EtMIC4: a microneme protein from Eimeria tenella that contains tandem arrays of epidermal growth factor-like repeats and thrombospondin type-I repeats. Int. J. Parasitol. 31, 1303–1310. Tuda, J., Mongan, A.E., Tolba, M.E., Imada, M., Yamagishi, J., Xuan, X., Wakaguri, H., Sugano, S., Sugimoto, C., Suzuki, Y., 2011. Full-parasites: database of full-length cDNAs of apicomplexa parasites, 2010 update. Nucleic Acids Res. 39, D625–631. Valouev, A., Ichikawa, J., Tonthat, T., Stuart, J., Ranade, S., Peckham, H., Zeng, K., Malek, J.A., Costa, G., McKernan, K., Sidow, A., Fire, A., Johnson, S.M., 2008. A high-resolution: nucleosome position map of C. elegans reveals a lack of universal sequence-dictated positioning. Genome Res. 18, 1051–1063.

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van Dijk, E.L., Auger, H., Jaszczyszyn, Y., Thermes, C., 2014. Ten years of next-generation sequencing technology. Trends Genet. 30, 418–426. Vollmers, C., Penland, L., Kanbar, J.N., Quake, S.R., 2015. Novel exons and splice variants in the human antibody heavy chain identified by single cell and single molecule sequencing. PLoS One 10, e0117050. Walker, R., Sharman, P., Miller, C., Lippuner, C., Okoniewski, M., Eichenberger, R., Ramakrishnan, C., Brossier, F., Deplazes, P., Hehl, A., Smith, N., 2015. RNA Seq analysis of the Eimeria tenella gametocyte transcriptome reveals clues about the molecular basis for sexual reproduction and oocyst biogenesis. BMC Genomics 16, 94. Wan, K.L., Chong, S.P., Ng, S.T., Shirley, M.W., Tomley, F.M., Jangi, M.S., 1999. A survey of genes in Eimeria tenella merozoites by EST sequencing. Int. J. Parasitol. 29, 1885–1892. Williams, R.B., Marshall, R.N., Pages, M., Dardi, M., del Cacho, E., 2009. Pathogenesis of Eimeria praecox in chickens: virulence of field strains compared with laboratory strains of E. praecox and Eimeria acervulina. Avian Pathol. 38, 359–366. Yan, W., Liu, X., Shi, T., Hao, L., Tomley, F.M., Suo, X., 2009. Stable transfection of Eimeria tenella: constitutive expression of the YFP-YFP molecule throughout the life cycle. Int. J. Parasitol. 39, 109–117. Yang, R., Fenwick, S., Potter, A., Elliot, A., Power, M., Beveridge, I., Ryan, U., 2012. 2012. Molecular characterization of Eimeria species in macropods. Exp. Parasitol. 132.

Please cite this article in press as: Blake, D.P., Eimeria genomics: Where are we now and where are we going? Vet. Parasitol. (2015), http://dx.doi.org/10.1016/j.vetpar.2015.05.007