2013 Chapman Et Al

Provided for non-commercial research and educational use only. Not for reproduction, distribution or commercial use.

This chapter was originally published in the book Advances in Parasitology, Vol. 83 published by Elsevier, and the attached copy is provided by Elsevier for the author's benefit and for the benefit of the author's institution, for non-commercial research and educational use including without limitation use in instruction at your institution, sending it to specific colleagues who know you, and providing a copy to your institution’s administrator.

All other uses, reproduction and distribution, including without limitation commercial reprints, selling or licensing copies or access, or posting on open internet sites, your personal or institution’s website or repository, are prohibited. For exceptions, permission may be sought for such use through Elsevier's permissions site at:

http://www.elsevier.com/locate/permissionusematerial

From H. David Chapman, John R. Barta, Damer Blake, Arthur Gruber, Mark Jenkins, Nicholas C. Smith, Xun Suo, Fiona M. Tomley, A Selective Review of Advances in Coccidiosis Research. In D. Rollinson, editors: Advances in Parasitology, Vol. 83,

Amsterdam, The Netherlands: Academic Press, 2013, pp. 93-171. ISBN: 978-0-12-407705-8

© Copyright 2013 Elsevier Ltd. Elsevier

Author's personal copy

CHAPTER TWO

A Selective Review of Advancesin Coccidiosis ResearchH. David Chapman*,1, John R. Barta†, Damer Blake{, Arthur Gruber},Mark Jenkins}, Nicholas C. Smith||, Xun Suo#, Fiona M. Tomley{*Department of Poultry Science, University of Arkansas, Fayetteville, Arkansas, United States†Department of Pathobiology, Ontario Veterinary College, University of Guelph, Guelph, Ontario, Canada{Royal Veterinary College, Hatfield, United Kingdom}Department of Parasitology, Institute of Biomedical Sciences, University of Sao Paulo, Sao Paulo, Brazil}Animal Parasitic Diseases Laboratory, Agricultural Research Service, USDA, Beltsville, Maryland, UnitedStates||Queensland Tropical Health Alliance Laboratory, Faculty ofMedicine, Health andMolecular Sciences, JamesCook University, Cairns, Queensland, Australia#National Animal Protozoa Laboratory & College of Veterinary Medicine, China Agricultural University,Beijing, China1Corresponding author: e-mail address: [email protected]

Contents

1.

AdvaISSNhttp:/

Introduction

nces in Parasitology, Volume 83 # 2013 Elsevier Ltd0065-308X All rights reserved./dx.doi.org/10.1016/B978-0-12-407705-8.00002-1

95
2. Taxonomy and Systematics 96
2.1
The genus Eimeria Schneider 1875: A melting pot of biologically diversecoccidia 97
2.2
Molecular identification and characterization of Eimeria and relatedcoccidia 99
2.3
Conclusions 104 3. Genetics 105
3.1
Markers employed in genetic studies 105 3.2 Cross-fertilization and genetic recombination 105 3.3 Genetic linkage analyses 106
4.
The ‘Omics’ Technologies 108 4.1 Genomics 108 4.2 Transcriptomics 113 4.3 Proteomics 116 4.4 The future 119
5.
Transfection 120 5.1 Transfection construct design 120 5.2 Transient transfection 121 5.3 Stable transfection 121 5.4 PiggyBac-based forward genetic system 122 5.5 Stably transfected Eimeria as a vaccine vector and beyond 123
93

http://dx.doi.org/10.1016/B978-0-12-407705-8.00002-1

mailto:[email protected]

94 H. David Chapman et al.


6.
Oocyst Biogenesis 124 6.1 Veil and WFBs 124 6.2 Oocyst wall proteins 126 6.3 Formation of the oocyst wall 126
7.
Host Cell Invasion 128 7.1 Parasite surface proteins 129 7.2 MIC proteins are adhesins and many function as multi-protein
complexes
130 7.3 Host glycan recognition by MIC proteins contributes to host
and tissue tropism
131 7.4 Regulated secretion of microneme and rhoptry organelles 132 7.5 AMAs and formation of the MJ 133
8.
Immunobiology 133 8.1 Innate responses to primary infection 134 8.2 Acquired immunity 138 8.3 Maternal immunity 138 8.4 Immunological research 139
9.
Diagnosis and Identification 140 9.1 Traditional methods 140 9.2 Early molecular methods 141 9.3 Methods based on DNA amplification by PCR 142 9.4 LAMP 146 9.5 Morphological diagnosis revisited 148 9.6 Conclusions 148
10.
Control 149 10.1 Chemotherapy 149 10.2 Vaccination 151 10.3 Strategies for the control of coccidiosis 153 10.4 Natural products 154
11.
Conclusions 154 Acknowledgement 155 References 155
Abstract

Coccidiosis is a widespread and economically significant disease of livestock caused byprotozoan parasites of the genus Eimeria. This disease is worldwide in occurrence andcosts the animal agricultural industry many millions of dollars to control. In recent years,the modern tools of molecular biology, biochemistry, cell biology and immunologyhave been used to expand greatly our knowledge of these parasites and the diseasethey cause. Such studies are essential if we are to develop new means for the controlof coccidiosis. In this chapter, selective aspects of the biology of these organisms, withemphasis on recent research in poultry, are reviewed. Topics considered include taxon-omy, systematics, genetics, genomics, transcriptomics, proteomics, transfection, oocystbiogenesis, host cell invasion, immunobiology, diagnostics and control.

95Review of Coccidiosis Research


1. INTRODUCTION

Coccidiosis is caused by protozoan parasites of the apicomplexan

genus Eimeria that occur in many vertebrate and invertebrate hosts. This dis-

ease is a major cause of mortality, poor performance and lost productivity in

domestic livestock. The parasites have an oral-faecal life cycle involving

three phases: schizogony (also known as merogony), gametogony and spo-

rogony (or sporulation). The infective transmission stage is the oocyst which

contains, when sporulated, four sporocysts each containing two sporozoites.

Following ingestion, the sporozoites are released and penetrate epithelial

cells of the intestine. This is followed by schizogony, an asexual phase of

multiplication involving several repeated generations, and gametogony,

which results in the production of a new generation of oocysts that are passed

out in the faeces. The third phase of the life cycle, sporogony, occurs in the

external environment and results in the formation of a new generation of

oocysts. Seven species are recognized in the fowl that vary according to

the number of generations of schizogony, physical characteristics such as

the size of the oocyst, and biological characteristics such as site of develop-

ment in the intestine, pathogenicity and immunogenicity. A similar number

of species have been described from the turkey. In poultry, the life cycle is

completed in about 7 days but in ruminants it may be longer. Both schizog-

ony and gametogony can cause pathology because the cells in which the par-

asites develop are functionally impaired and eventually destroyed. The

extent of such destruction is determined by the numbers of infective oocysts

ingested, which in turn depends upon the extent to which sporulation is suc-

cessful. This requires warmth, oxygen and moisture, factors often not lac-

king in commercial livestock production.

It is in the poultry industry that coccidiosis is of the greatest economic

significance because modern production methods involve the rearing of

large numbers of birds in confinement at high stocking densities, often on

built-up litter. For example, a modern broiler house may contain as many

as 20–50,000 birds under one roof at a stocking density of one bird per

0.08 m2 and as many as 10 houses may be present at one farm. Furthermore,

both the broiler industry and the turkey industry tend to be restricted geo-

graphically; for example, in the United States, one of the largest areas of pro-

duction is confined to a few counties in northwest Arkansas. Thus the

commercial conditions under which poultry are raised provide ideal condi-

tions for parasite transmission. Fortunately, control of coccidiosis can be



achieved either by the inclusion of drugs in the feed (prophylactic chemo-

therapy) or by vaccination and, because of such control measures, coccidiosis

is less a problem today than in the past (Chapman, 2003). Nevertheless, the

resilience of the oocyst ensures the continued presence of these organisms

wherever poultry are reared. The expansion of the poultry industry has pro-

vided a major source of protein to feed the growing human population and

any disease that limits this production, such as coccidiosis, will have the

potential to affect human health. There is a continued need, therefore,

for both basic and applied research into all aspects of the biology of these

organisms (Shirley and Lillehoj, 2012).

An example of basic research is the utilization of genomics and proteo-

mics to elucidate molecular aspects of invasion of host cells by sporozoites.

The apical complex of coccidia, found in motile stages of the life cycle (spo-

rozoites and merozoites), contains several sub-cellular organelles including

paired rhoptries, micronemes and dense bodies that secrete proteins

involved in the invasion process. Another example is the investigation of

biochemical events involved in the formation of the wall of the oocyst. Pro-

gress has also been made in understanding the role of CD4/CD8 lympho-

cytes and cytokines in inflammatory responses to infection; cell-mediated

immune mechanisms have been studied utilizing murine Eimeria as a model.

Molecular techniques have been utilized to develop new methods for iden-

tifying the species that infect the fowl. Much applied research has been

undertaken to demonstrate efficacy of various products for the control of

coccidiosis but in general this has not been published in peer-reviewed sci-

entific journals. In this review, some aspects of coccidiosis research in poul-

try is provided by the following contributors: taxonomy and systematics

(Barta); genetics (Blake); ‘omics’ technologies (Blake, Tomley, Gruber);

transfection (Suo, Blake); oocyst biogenesis (Smith); host cell invasion

(Tomley); immunology (Smith); diagnostics ( Jenkins, Gruber) and control

(Chapman).

2. TAXONOMY AND SYSTEMATICS

The taxonomy and systematics of eimeriid coccidia andEimeria species

infecting vertebrates is, at best, problematic. This confounding state has

arisen because of numerous poorly described species, lack of type deposition

(even of micrographs or drawings) and improper taxonomic methods.

Misidentification is a general difficulty in studies concerned with the biology

of Eimeria. Classical original descriptions of the species that infect poultry



were based upon isolates that have since been lost and, thus, there are no

surviving type specimens. A few defined laboratory strains still exist

(although not type specimens in the taxonomic sense), for example, the

‘Houghton’ and ‘Weybridge’ strains of various species, but others, such as

most ‘Beltsville’ strains, have been lost. Ideally, molecular studies should

be based upon strains derived from a single oocyst and identified using

key parasitological characters. Difficulties with the interpretation of DNA

analysis from inadequately described strains of Eimeria have been addressed

by Williams et al. (2010).

Confounding this taxonomic history is frequent application of the

‘different-host–different-parasite’ mindset. For Eimeria species in some host

groups, this principle may be justified because of relatively strict host spec-

ificities of these parasites; however, in some coccidia that infect passerine

birds (Isospora species in particular), host specificity may not be nearly as strict

and there may be many Isospora species that may ultimately be synonymized.

To further complicate matters, parasites in the genus Atoxoplasma Garnham

1950 may actually all be members of either the genus Isospora (and perhaps

synonymized with previously described Isospora species) or the genus Lank-

esterella (e.g. Barta et al., 2005; Merino et al., 2006).

2.1. The genus Eimeria Schneider 1875: A melting potof biologically diverse coccidia

The apicomplexan suborder Eimeriorina (Leger, 1911) is home to the

eimeriid coccidia (family Eimeriidae Minchin 1903) that includes the

Eimeria species and related monoxenous or facultatively heteroxenous

coccidia infecting vertebrates. A recent taxonomic definition of the family

Eimeriidae (see Upton, 2000) included apicomplexan parasites with the fol-

lowing features: ‘Homoxenous or facultatively homoxenous; merogony,

gamogony and formation of oocysts all in the same host; in vertebrates or

invertebrates’. Clearly this is a broadly applicable definition for inclusion

of parasites into this family. The definition of the genus is likewise permis-

sive: ‘oocysts with four sporocysts, each with two sporozoites’. Not surpris-

ingly, the number of Eimeria species that have been described to date exceeds

1200 and this number continues to grow. The frequently observed strict host

specificity of many Eimeria species and the infection of many hosts with mul-

tiple Eimeria species means that there remain probably tens of thousands of

undescribed Eimeria species infecting birds, herbivorous or omnivorous

mammals, reptiles, amphibia and, perhaps, even fish.



Whilst taxonomically useful (and conservative), this rather broad defini-

tion of the genus bears part of the responsibility for one of the fundamental

taxonomic difficulties with the genus Eimeria—it is not a monophyletic tax-

onomic group. There is considerable agreement that species currently

assigned to the genus Eimeria do not form a monophyletic group (e.g.

Jirku et al., 2002, 2009) whether assessed systematically using morphological

or molecular characters. Instead, the genus Eimeria is paraphyletic or poly-

phyletic. When analyzed using complete or partial nuclear 18S rDNA

sequences, the phylogenetic analysis generated a consensus tree (Fig. 2.1)

in which members of the Toxoplasmatinae (species of Cystoisospora,

Neospora, Hammondia and Toxoplasma) formed a well-supported monophy-

letic group that had a sister group relationship with the other coccidia,

including Eimeria species. The large clade of eimeriid coccidia that includes

all Eimeria species for which 18S rDNA sequence data have been obtained

includes many genera other than Eimeria, including Goussia, Caryospora,

Lankesterella, Atoxoplasma, Isospora and Cyclospora. Even the genera currently

included in the large family Eimeriidae are not found within a monophyletic

grouping; members of the heteroxenous family Lankesterellidae are found

within the clade that contains all species belonging to the Eimeriidae.

The tissue coccidia (Toxoplasma and its relatives—Toxoplasmatinae) and

all of the early branching coccidia (Hyaloklossia and Goussia spp. plus

E. tropidura) possess valvular sutures on their sporocysts and do not have

Stieda bodies (see Fig. 2.2). Eimeria arnyi from a snake and E. ranae from frogs

were found in a well-supported branch that arose near the base of the

coccidia that possess Stieda bodies in their oocysts (when sporocysts are

formed). The next branching clade consisted of: (1) Eimeria species of mar-

supials (E. trichosuri); (2) Eimeria spp. of cranes (E. gruis and E. reichenowi) and

(3) a clade containing Caryospora spp. and Lankesterella spp. This sister group

to these parasites was a large collection of Eimeria species from a wide variety

of hosts interrupted by the inclusion of a well-supported clade of

Cyclospora spp.

The genus Eimeria was observed to be polyphyletic with at least four

independent lineages of Eimeria species. Caryospora and Lankesterella species

formed a monophyletic clade as has been described previously (Barta et al.,

2001). Avian Isospora and Atoxoplasma species formed a well-supported

monophyletic clade; the mixing of Atoxoplasma and Isospora species gives

additional support for the questioning the validity of the genus Atoxoplasma.

Giving some support to the concept of host specificity, Eimeria species fre-

quently formed well-supported clades of parasites that parasitized the same



or closely related definitive hosts such as Eimeria spp. infecting swine, rabbits

or galliform birds. Among the Eimeria species infecting galliform birds, the

cecal coccidia of chickens and turkeys (i.e. E. necatrix, E. tenella and

E. adenoeides) are frequently found to form amonophyletic group of parasites

(e.g. Miska et al., 2010; Fig. 2.1) to the exclusion of the other Eimeria species

infecting the chicken. This repeated observation, and the failure to find

E. necatrix or E. tenella in wild jungle fowl (Fernando and Remmler,

1973), present the possibility that these two species of Eimeria infecting

domestic chickens may have arisen from a host transfer from some other gal-

liform host (Barta et al., 1997).

2.2. Molecular identification and characterization of Eimeriaand related coccidia

Multiple, distinct rDNA copies were described decades ago for the malar-

ial parasites, Plasmodium species (e.g. McCutchan et al., 1988; Nishimoto

et al., 2008). In these haemosporinid parasites, up to three independently

evolving paralogous rDNA copies were described that frequently demon-

strated more sequence variation between paralogous within a single para-

site species than between homologues among different species.

Interestingly, paralogous rDNA copies were expressed differentially in dif-

ferent life cycle stages. Until recently, the presence of paralogous rDNA

copies had not been observed in Eimeria species and thus the nuclear

18S rDNA locus was considered a relatively useful and stable genetic target

for species-level identification and for molecular phylogenetics. However,

Vrba et al (2011) recently showed that a single-oocyst-derived line of

E. mitis contained two paralogous types of nuclear 18S rDNA that had

considerable sequence variation (i.e. 1.3–1.7% sequence diversity between

paralogous compared with 0.3–0.6% sequence diversity among homo-

logues). Recent work with a number of single-oocyst-derived lines of sev-

eral Eimeria species infecting turkeys indicates that paralogous 18S rDNA

copies exist within the nuclear genome of at least some of these parasites as

well. For example, single-oocyst clonal lines of E. meleagrimitis, from

which polymerase chain reaction (PCR)-amplified, near-complete nuclear

18S rDNA amplicons were cloned and sequenced, demonstrated consid-

erable sequence diversity. Two paralogous groups of sequences were

obtained; each group of sequences had mean intraspecific sequence diver-

sities of about 0.5% but the mean interspecific variation between these

paralogous exceeded 2.7%. By the way of comparison, the mean interspe-

cific variation for the 18S rDNA between the widely accepted species


Figure 2.1 Maximum likelihood consensus tree resulting from the analysis of complete or near complete nuclear 18S rDNA sequences. Foreach taxon in the tree, the definitive host (DH), if known, and the GenBank Accession number for the sequence used to generate the tree areincluded. Percentage bootstrap supports for major clades are presented as numbers located at nodes on the tree. Horizontal branch lengthsare proportional to hypothesized evolutionary change with the scale indicating 10% hypothesized sequence variation.


Figure 2.2 Photomicrographs of sporulated oocysts of coccidia that each possess twosporocysts, containing four sporozoites. (A) An avian Isospora sp. that possess Stiedabodies at the apex of each sporocyst in the oocyst (arrows) and that belongs to the fam-ily Eimeriidae. (B) Cystoisospora felis, a coccidium infecting felids that has an oocyst withno Stieda bodies on its sporocysts and that belongs to the family Sarcocystidae.



E. tenella and E. necatrix is only 1.1%. Clearly, such paralogous loci can

confound both molecular phylogenetics and the taxonomic pursuits of

identification and characterization.

Recognition of intragenomic polymorphisms among 18S rDNA in

Eimeria has important ramifications for use of other regions of the rDNA

gene array for molecular diagnostics and/or characterization. For example,

the internal transcribed spacer (ITS) regions have been used to characterize

species and strains of Eimeria. However, the ITS-1 and ITS-2 regions are

likely subject to both intragenomic (e.g. Blake et al., 2006; Vrba et al.,

2011) and intraspecific (e.g. Barta et al., 1998) variations that may limit

the utility any resulting ITS sequences or molecular diagnostic methods

based on these sequences.

Ogedengbe et al. (2011a) have recently demonstrated that another

genetic locus, the mitochondrial cytochrome c oxidase subunit I gene

(cox-1, COI), is a muchmore reliable gene for molecular species delimitation

of Eimeria species and other coccidia than nuclear 18S rDNA sequences. The

recognition of intragenomic polymorphism of the 18S rDNA locus (e.g.

Vrba et al., 2011) may explain the superiority of the COI locus for species

delimitation and, potentially, identification. The COI genetic target has

additional features that argue for its use in the molecular characterization

and identification of coccidia: (1) COI is found in multiple copies within

the mitochondria of coccidia making it a good PCR target; (2) COI



demonstrates sufficient DNA sequence variability (�2–3% sequence diver-

sity between Eimeria spp.) that 500–800 bp fragments will provide sufficient

data for most identification purposes; (3) COI is sufficiently divergent from

the COI of most hosts that parasite-specific PCR primers can be readily

developed, permitting the use of this locus for parasites located within host

tissues; (4) COI is located on a genome derived from an endosymbiotic pro-

karyote meaning that the resulting COI protein coding sequences are free of

introns and can be assessed relatively easily for contiguous open reading

frames (ORFs) as an internal check for correct PCR amplification

(Ogedengbe et al., 2011a) and finally, (5) intraspecific variation among spe-

cies is relatively modest (usually <0.2%) compared with interspecific varia-

tion (usually >1.5%). However, the use of COI sequences for parasite

identification and/or species delimitation is not without its problems. Com-

pared with the 18S rDNA locus, relatively few COI sequences have been

deposited to public sequence databases, limiting the use of this locus for

identification purposes until more reference sequences become available.

In addition, PCR amplification of COI fragments from mixed parasite

DNA templates (such as litter or mixed species samples, e.g. Schwarz

et al., 2009) can generate hybrid amplicons (e.g. GenBank Accession num-

ber FJ236441). In our experience, up to 5% of COI fragments amplified by

PCR from DNA samples containing multiple Eimeria spp. may be hybrid

molecules generated during the PCR process. For Eimeria spp. of chickens

or other well-sampled taxonomic groups (such as Plasmodium spp. and

related haemosporinid parasites), this is not a major issue. In such cases,

hybrid sequences are easily detected using a simple BLAST search because

of the availability of reliable reference sequences in the public sequence data-

bases. Examining pairwise alignments of the top BLAST hits will show

sequence divergence restricted to only one portion of the new sequence

when aligned with sequences from one Eimeria sp. and with a different por-

tion of the molecule when aligned with a second Eimeria species. However,

for Eimeria or Isospora species from less well-sampled hosts, hybrid sequences

may be problematic. To avoid the influence of such hybrids, direct sequenc-

ing of PCR products is recommended—messy or ambiguous sequences

resulting from such direct sequencing likely indicates multiple species within

the sample and biological purification should be attempted (biological

enrichment or cloning of the parasite). Alternately, after cloning PCR frag-

ments from such a sample, multiple clones must be sequenced so that both

repeated COI sequences (likely valid amplicons) and potential hybrids can

be identified.



For the well-acceptedEimeria species infecting chickens it was frequently

assumed that lack of immunological cross-protection in the host was evi-

dence in support of identification of particular species. For example, chickens

immunized against Eimeria maxima through natural infection would be

expected to be protected against subsequent challenge with the same species.

However, it has been shown conclusively that immunologically distinct

strains of E. maxima can arise both spontaneously and in response to selective

immunological pressure. Despite a near complete lack of immunological

cross-protection, the Guelph and M6 strains of E. maxima have identical

COI sequences (Ogedengbe et al., 2011a) demonstrating again the utility

of this genetic locus for species identification, even when classical methods

of species identification such as immunological cross-protection can fail.

2.3. ConclusionsThe taxonomic mess that is the current genus Eimeria is likely to be resolved

slowly through the application of appropriate nuclear or mitochondrial

sequence analysis coupled with phenotypic characters. Ideally, monotypic

reference strains of Eimeria species and other coccidia of veterinary or med-

ical importance should be characterized biologically (minimally oocyst

dimensions as described by Bandoni and Duszynski, 1988), but optimally

including solid descriptions of endogenous development during experimen-

tal infections as well as molecularly (preferentially obtaining both nuclear

18S rDNA and mitochondrial COI sequences).

Such combined morphological and molecular characterizations have

been accomplished with single Isospora sp. oocysts (Dolnik et al., 2009).

However, even with both molecular and morphological data available, tax-

onomic decisions are ultimately opinions of the various authors involved.

For example, although there were significant differences for two strains of

turkey Eimeria in oocyst dimensions and 2.3% sequence divergence at the

COI locus over 767 bp, Poplstein and Vrba (2011) concluded that these par-

asites were simply variants of a single species, E. adenoeides because of some

immunological cross-protection of these parasites in turkeys. Clearly further

studies are warranted even for parasites that have been described for many

years and that infect agriculturally important animals.

Finally, the taxonomic breadth of the genus Eimeria is clearly too broad.

It is anticipated that the genus Eimeria will be divided into several genera,

each containing fewer, but biologically more homogenous, species with

each new genus.



3. GENETICS

Studies with Eimeria genomes can be divided into physical and non-

physical categories including genomic, transcriptomic and proteomic or

fundamental and applied genetics, respectively. Key among the features that

underpin such studies is the haploid state of the eimerian genome through-

out the majority of the life cycle. Thus, clonal parasites can be isolated by

passage of a single sporozoite or sporocyst and each cloned parasite line

may be considered homologous at all loci prior to fertilization and zygote

formation (Chapman andRose, 1986; Shirley andHarvey, 1996). Nonethe-

less, the brief sexual phase of the life cycle facilitates chromosomal segrega-

tion and genetic recombination ( Jeffers, 1976) and supports classical genetics

studies where a departure from the anticipated Mendelian phenotypic ratio

of inheritance informs on genotype (Sturtevant, 1913). More recently,

molecular biology has revolutionized our understanding of genomes and

their associated biology, providing new tools for genetics-led research and

creating the ‘omics’ disciplines.

3.1. Markers employed in genetic studiesVariation between and within Eimeria species has been investigated using

several phenotypic and genotypic tools as reviewed elsewhere (Beck

et al., 2009). Briefly, phenotypic tools have included differential isoenzyme

migration through starch gels or by isoelectric focusing, resistance or suscep-

tibility to defined chemotherapeutic exposure, precocious development and

the ability to escape strain-specific immune killing (Blake et al., 2005; Jeffers,

1976; Shirley et al., 1989; Smith et al., 1994a–d). Genotypic tools, including

random amplification of polymorphic DNA polymerase chain reaction

(RAPD-PCR), restriction fragment length polymorphism (RFLP), ampli-

fied fragment length polymorphism (AFLP) and gross chromosomal size

polymorphism as revealed by pulsed field gel electrophoresis (PFGE), have

also been used to define inter- and intraspecific variation (Blake et al., 2011a;

Fernandez et al., 2003a; Shirley, 2000).

3.2. Cross-fertilization and genetic recombinationEarly studies with coccidial parasites suggested the absence of genetic recom-

bination, however, molecular, genetic and microscopic studies have now

demonstrated numerous crossover events and recombination nodules,



respectively (Blake et al., 2011b; Canning and Anwar, 1968; del Cacho

et al., 2005; Shirley and Harvey, 2000). Almost 40 years ago, the first evi-

dence of genetic exchange during multi-clonal infection revealed a capacity

for cross-fertilization, independent segregation and the consequential pro-

duction of hybrid progeny ( Jeffers, 1974). In this example, concurrent infec-

tion of E. tenella strains resistant to the anticoccidial drugs amprolium or

decoquinate yielded a population resistant to both compounds. However,

the efficiency of cross-fertilization remains unclear, as the proportion of

hybrid progeny within specific crosses has been calculated to vary between

0.05% and 39.5% (Blake et al., 2004; Joyner andNorton, 1975). Proportions

of hybrid progeny, based on comparison of oocyst output in the presence or

absence of deleterious selection, depend on the underlying genetic com-

plexities of the trait (i.e. the number of contributing genetic loci). In addition,

proportions are affected by the timing of selection within the life cycle and

the magnitude of the doses of oocysts given, which can result in parasite

‘crowding’ within the intestine (Blake et al., 2004; Williams, 2001).

3.3. Genetic linkage analysesThe inheritance of polymorphic genetic markers can be used to infer a

genetic map based upon comparisons of recombination frequencies between

markers. Key assumptions include a random (independent) association

between markers on separate chromosomes and a correlation between

recombination rate and physical distance for markers linked on the same

chromosome. For the Eimeria species, early linkage studies included the cre-

ation and selection of multi-drug resistant hybrid lines by cross-fertilization,

in which it was noted that close physical linkage of contributing loci may be

an explanation for incompatible resistance combinations ( Joyner and

Norton, 1978). The development of sequence-based genetic markers per-

mitted the development of more sophisticated linkage analyses, including

the derivation of genetic linkage maps and whole genome association

(WGA) studies.

Genetic linkage maps have been produced for two Eimeria species. In the

first genome-wide linkage study, a cross was made between an attenuated

E. tenella Wisconsin line (selected for precocious development within

125 h) and aWeybridge line selected to become resistant to the anticoccidial

drug arprinocid (Shirley and Harvey, 2000). The hybrid component of the

progeny of this cross, capable of replication within 125 h in the presence of

arprinocid, were recovered and amplified by selective in vivo passage and



used to derive a panel of 22 hybrid clonal progeny lines by single sporocyst

cloning. Using RAPD-PCR, RFLP, AFLP and full karyotype PFGE, 443

parent-specific genetic markers were generated and mapped against all

22 clones. Using the free linkage software Map Manager QT (Manly and

Cudmore, 1997), 16 linkage groups were created, representing a genetic

genome size of �653 centimorgans (cM). Intriguingly, �53% of all the

polymorphic markers scored were clustered into just three linkage groups,

indicating the absence of dividing recombination events within all 22 cloned

progeny. At the time of publication, this was attributed to possible bias intro-

duced by AFLP, however, the subsequent description of a segmented

genome structure for E. tenella now suggests pronounced hot and cold spots

of genetic recombination (Blake et al., 2011b; Ling et al., 2007). More

recently, a genetic map was created for E. maxima using a similar strategy

with the selectable traits resistance to the drug robenidine and escape from

strain-specific immune killing (Blake et al., 2011b). Using a larger panel of

647 genetic markers, generated exclusively by AFLP, linkage analysis cre-

ated a map made up of 13 major linkage groups representing a genetic

genome size of 2883.9 cM.

The first WGA study on E. tenella (Shirley and Harvey, 2000) made the

key assumption that each parent-specific genetic marker is inherited in a 1:1

ratio in the absence of deleterious selection. Thus, a genetic marker distantly

linked to a locus under selection, or located on a separate chromosome,

should persist within the hybrid progeny irrespective of selection. Represen-

tation of a more closely linkedmarker should be reduced within the progeny

that survive selection, while a marker that is linkedmost intimately should be

severely under-represented or lost altogether. Clonal lines drawn from a

hybrid population under selection should conform to the same rules,

resulting in linkage disequilibrium. Following combined selection for resis-

tance to arprinocid and the ability to reproduce within 125 h, all genetic

markers incorporated into the E. tenella genetic map were found to persist

in the anticipated�1:1 ratio with the exception of linkage groups mapped to

chromosomes 1 and 2, whose inheritance correlated with drug resistance

and precocious ability, respectively (Shirley andHarvey, 2000). The number

and size of loci mapped in this study would have been influenced by the bio-

logical factors described above, as well as practical factors including the num-

ber of genetic markers and independent clones.

While the number of genetic markers available can be adjusted with rel-

ative ease, the effort required to isolate and amplify clonal lines of Eimeria

places an inherent limit on the number of clones that can be handled in



any one laboratory. Working with an uncloned hybrid progeny population

under selection, instead of a finite panel of clones, can provide a massive

increase in mapping power. Working on this hypothesis, hybrid

E. maxima populations, selected for resistance to robenidine and the ability

to escape strain-specific immune killing, were used in the first population-

based WGA strategy with Eimeria species to map loci that encode strain-

specific immunoprotective antigens (Blake et al., 2011a). Briefly,

E. maxima isolates are commonly characterized by antigenic diversity such

that immunization with one strain can induce apparently complete immune

protection against homologous challenge but incomplete protection against

challenge by an antigenically distinct strain (Smith et al., 2002). In one

extreme example, immunization of inbred Line CWhite Leghorn chickens

(maintained at the Institute for Animal Health, UK) with the Houghton (H)

or Weybridge (W) E. maxima strains yields no statistically significant cross-

protection.WGA scrutiny of this phenotype, using the uncloned progeny of

multiple crosses between the H and W E. maxima strains before and after

W strain-specific immune selection, identified six distinct genetic loci whose

inheritance correlated absolutely with the immune phenotype (Blake et al.,

2011a). Subsequent fine mapping of two of these loci identified immune

mapped protein-1 and confirmed apical membrane antigen 1 (AMA1) as

partially immunoprotective antigens. The immunoprotective capacity of

three of the remaining four loci was also demonstrated using bacterial arti-

ficial clones (BACs) that covered each mapped W strain locus. Transient

transfection with whole BAC DNA was used to genetically complement

the H strain, and transferred the ability to induce a protective immune

response against the W strain (Blake et al., 2011a).

4. THE ‘OMICS’ TECHNOLOGIES

4.1. Genomics
Three distinct DNA genomes have been defined in eimerian parasites, local-
ized to the nucleus, the mitochondrion and the apicoplast organelle. Addi-

tionally, a double stranded RNA genome associated with virus-like particles

has been commonly found in Eimeria species (Han et al., 2011; Lee and

Fernando, 2000; Shirley, 2000). Using PFGE, the nuclear genomes of all

avian Eimeria species investigated to date have been estimated to contain

between 50 and 60 Mb DNA (Blake et al., 2011b; Shirley, 2000). The

E. tenella genome is the best characterized to date, featuring a GC content



of �53% and two major ribosomal gene clusters on chromosomes 10 and

12 (�500 copies of the 5S rDNA and �140 copies of the 18S–5.8S–28S

rDNA, respectively) (Shirley, 2000). Complete mitochondrial genome

sequences have been assembled for all seven species that infect the chicken,

and comprise �6200 bp (6148–6407 bp) DNA with �65% AþT content

(Lin et al., 2011; Liu et al., 2012). The relatively high AþT content influ-

ences codon usage and, consequently, amino acid content of the three pro-

tein coding genes cox-1, cox-3 and cytB. Sequencing of the larger plastid

genome identified �35 kb DNA with a similarly high AþT content

(Cai et al., 2003). Double stranded RNA virus genomes have been iden-

tified within many isolates of chicken Eimeria species ranging in size from

�1.7 to >7.4 kb. Hybridization studies suggest the existence of multiple,

genetically distinct viral strains or species (Lee and Fernando, 2000;

Lee et al., 1996).

4.1.1 Nuclear karyotypeOocysts of the Eimeria species that infect poultry are characterized by

extremely tough walls which are resistant to chemical, detergent,

enzymatic- and temperature-based disruptions, in contrast to those that

infect mammalian hosts such as the rat (Kurth and Entzeroth, 2008). Until

recently, the oocyst wall has prevented morphological analysis of eimerian

karyotypes and chromosome replication, hindering microscopic charac-

terization (del Cacho et al., 2001). However, the recent development of

a protocol based upon oocyst incubation in a hydrochloric acid/ethanol

solution followed by multiple freeze–thaw cycling has permitted the

release and spread of intact chromosomes, confirming previous PFGE-

based estimates of a 14 chromosome karyotype for all species that have

been investigated (del Cacho et al., 2005; Shirley, 1994a). Using PFGE,

the E. tenella and E. maxima karyotypes have been shown to range from

�1 to >7 Mb and �2 to >6 Mb, respectively (Blake et al., 2011b;

Shirley, 2000). Comparison between E. tenella strains has revealed variable

sizes for chromosomes 1–4 and 11 (Sheriff et al., 2003; Shirley, 2000). Var-

iation by as much as 5% for chromosome 1 between extreme examples has

been used as a genetic marker in linkage analyses (Shirley and Harvey,

2000). The molecular basis for the observed polymorphism remains

unclear, although the highly repetitive nature of the E. tenella genome is

likely to encourage non-homologous genetic recombination, resulting

in length polymorphisms.



4.1.2 Genome sequencingThe E. tenella Houghton strain was the first eimerian to be subjected to

nuclear genome sequencing. A combination of Sanger, 454 and Illumina

sequencing technologies was used to create a whole genome assembly rep-

resenting �94% of the complete nuclear genome in 4682 contiguous

sequences (contigs). While high frequency of repetitive sequences has hin-

dered further assembly of the genome, sequencing large insert genomic

DNA libraries has permitted assembly of �90% of the genome into 1720

supercontigs. Public access to the consensus assembly is available through

the Wellcome Trust Sanger Institute website, GeneDB (www.genedb.

org) and EUPathDB (www.eupathdb.org). Additionally, chromosome 1

of E. tenella has been fully sequenced and assembled. The identification of

telomeric-like repeats at each end of the assembly suggests representation

of the majority of the chromosome, including �85% of the predicted

1.05 Mb sequence with a small number of sequence gaps (Ling et al.,

2007). Comparison with a chromosome 1 HAPPY map (mapping based

on the analysis of approximately HAPloid DNA samples using PCR) sup-

ported the validity of the assembly and provided a tool to anchor and order

the sequence contigs (Ling et al., 2007). More recently, a cloned line derived

from the E. maximaHoughton strain has been sequenced, providing the first

genomic resource for a species that parasitises the mid-intestine (Blake et al.,

2012). Sanger sequencing combined with 454 sequencing yielded�13-fold

genome coverage and a consensus assembly representing �74% of the

nuclear genome in 12,852 contigs (publically available through EmaxDB,

www.emaxdb.org). Nuclear genome sequences for the Houghton strains

of all seven species of Eimeria of the chicken, and for additional strains of

E. tenella, are now being generated using multiple next-generation sequenc-

ing technologies and during 2013 all of these sequences will be available for

public scrutiny within both the GeneDB and the EuPathDB databases.

Gross genomic comparison among apicomplexan parasites reveals a rel-

atively conserved genome size for coccidial parasites, significantly larger than

the haemosporids and piroplasms (Table 2.1). While genome size varies sig-

nificantly among the Apicomplexa, the predicted number of protein coding

genes varies less dramatically (from largest to smallest fold difference:

genome¼�eightfold, gene number¼� twofold; Table 2.1). Broader com-

parison with most other apicomplexan parasites reveals a negatively corre-

lated association between gene density and genome size, likely to be

underpinned by a core gene set essential to parasite function and survival

irrespective of genome size. Furthermore, the multi-host lifestyles of the

http://www.genedb.org


http://www.eupathdb.org

http://www.emaxdb.org

Table 2.1 Apicomplexan parasites: A current genetics and genomics summary

Organism StrainGenome size(Mb)

Chromosomeno.

Predictedproteins

Gene density(genes Mb�1)

Recombination rate(kb cM�1) Referencea

Eimeria tenella Houghton 55.0 14 8786 160 264b Blake et al.

(2011b)

Eimeria maxima Houghton 57.5 14 Not known Not known 60–120 Blake et al.

(2011b)

Toxoplasma

gondii

ME49 63.0 14 7993 127 104 Khan et al.

(2005)

Neospora

caninum

NCLiv 61.0 14 7082 116 na na

Plasmodium

falciparum

3D7 23.3 14 5538 238 17 Su et al. (1999)

Babesia bovis T2Bo 8.1 4 3706 458 na na

Theileria parva Muguga 8.3 4 4082 492 4.6 Katzer et al.

(2011)

Cryptosporidium

parvum

Iowa II 9.1 8 3805 418 10-56 Tanriverdi et al.

(2007)

aReference used to derive the rate of genetic recombination.bExpected to drop should additional markers be included as described previously for T. gondii (Khan et al., 2005; Sibley et al., 1992).na, not available.Data derived from EuPathDB (http://eupathdb.org/eupathdb/, accessed 3 July 2012).


http://eupathdb.org/eupathdb/



hemoparasites and piroplasms might indicate a requirement for a larger core

gene set. While total chromosome number remains stable within the larger

genomes, the relative rate of genetic recombination exhibits a strong neg-

ative correlation (Table 2.1), possibly indicating a minimum requirement

for recombination, irrespective of genome size.

4.1.3 Genome structureThe Eimeria species are protozoan eukaryotic organisms. Each eimerian

genome is represented by a series of chromosomes including telomeres

and centromeres (del Cacho et al., 2005). Transmission electron microscopy

studies have revealed a constant telomere length of 32 nm for E. tenella and a

consistent centromere index per chromosome among several strains,

although figures vary between chromosomes, thereby providing distinctive

identifiers (del Cacho et al., 2005). Intronic sequences are common within

most coding regions and there is proteomic evidence for alternative splicing

(Lal et al., 2009). Unusually, among apicomplexan parasites, sequences

related to eukaryotic transposable elements are readily identified. It has

been suggested that these are most similar to non-LTR LINE-like

retrotransposons (long terminal repeat and long interspersed nuclear

elements) (Ling et al. 2007). Most strikingly, the sequencing and assembly

of E. tenella chromosome 1 revealed an unusual segmented structure to

the chromosome, which contains three repeat-rich segments flanked by four

repeat-poor segments (GenBank Accession number AM269894). The

repeat-rich segments contain large numbers of simple sequence repeats

and LINE-like elements as well as highly variable AþT content, CpG to

GpC ratio and second-order Markov entropy, prompting the identifier

‘feature-rich’ (R-segment) (Ling et al., 2007). In contrast, the repeat-poor

segments exhibited less variation in all measures, prompting the identifier

‘feature-poor’ (P-segment). Whole genome HAPPYmapping now suggests

that the segmental organization described from chromosome 1 is also present

throughout the rest of the E. tenella genome (Lim et al., 2012).

Preliminary characterization of predicted gene structures within the

R- and P-segments suggests shorter coding sequences with larger exons

and less numerous introns in the latter. RFLP comparison of E. tenella geno-

mic DNA revealed polymorphism between the Houghton, Weybridge and

Wisconsin strains for all four of theR-segment probes tested, but none of the

four P-segment probes, suggesting a higher rate of genome evolution within

the feature-rich regions. Comparative studies with the E. maxima genome



suggests that R and P regions are also present throughout the genome of this

species (Blake et al., 2011b).

4.1.4 Repetitive sequencesEimerian genes are known to feature large numbers of repetitive DNA

sequences ( Jenkins, 1988) and early reports of numerous trinucleotide

GCA repeats (and alternative frame permutations CAG, AGC, TGC,

GCT and CTG) are supported by more systematic studies of published

EST data and the E. tenella chromosome 1 assembly. Almost 3000 simple

repeat units were identified on chromosome 1 and more than 60% of these

were GCA-based (Ling et al., 2007). Other common repeats included a

telomere-like heptamer AGGGTTT, representing nearly 20% of the repeats

on chromosome 1, and the palindromic octamer TGCATGCA, which has

been described previously within several other apicomplexan genomes. As

noted above, simple sequence repeats were confined largely to the

R-segments in the E. tenella chromosome 1 assembly, where triplet repeats

and AGGGTTT represent �14% of the sequence. Both EST and genomic

analyses identified frequent triplet repeats in putative coding regions, a fea-

ture confirmed by recent proteomic analysis (Lal et al., 2009; Ling et al.,

2007; Shirley, 2000). Importantly, triplet repeats do not interfere with

the coding frame in expressed sequences, unlike heptamer and octomer

repeats. To date, no functional association has been made with these trans-

lated repetitive sequences, although they have been hypothesized to play a

role in genome evolution and diversification as possible hotspots of recom-

bination (Ling et al., 2007).

Other repetitive sequences include multiple putative transposable

sequences and arrays of tandemly repeated 5S and 18S–5.8S–28S ribosomal

genes (both discussed above). The high copy number of the rDNA arrays

and the ITS regions (ITS-1 and ITS-2) has promoted their use as targets

for molecular diagnostics and phylogenetic analyses, although it is important

to note the existence of polymorphism between copies within a single

genome (Blake et al., 2006; Vrba et al., 2011).

4.2. TranscriptomicsSystematic descriptions of four Eimeria species transcriptomes have been

reported in addition to small numbers of targeted cDNA sequences from

the same and other species. By far the most thoroughly characterized has

been E. tenella, with more than 50,000 publically available expressed

sequence tag (EST) sequences (Amiruddin et al., 2012; Chen et al., 2008;



Klotz et al., 2007; Novaes et al., 2012; Wan et al., 1999). E. acervulina and

E. maxima are both well represented (e.g. Dong et al., 2011; Miska et al.,

2008; Novaes et al., 2012; Schwarz et al., 2010) and E. brunetti has been sam-

pled (Aarthi et al., 2011). Sequences have frequently been derived from the

most easily accessed oocyst and sporozoite life cycle stages, although second-

generation merozoites have commonly been prioritized given their rele-

vance to coccidiosis caused by E. tenella (Amiruddin et al., 2012; Miska

et al., 2008; Novaes et al., 2012; Schwarz et al., 2010). Key stages in the

eimerian life cycle waiting to be sampled include the gametocytes and devel-

oping intracellular schizonts.

4.2.1 Full-length cDNA sequencesAnalysis of 433 full-length cDNA sequences from E. tenellaHoughton strain

second-generation merozoites has provided the most detailed study of tran-

script structure for the Eimeria species. At 1647 bp the average transcript

length was longer than described for Toxoplasma gondii or Cryptosporidium

parvum (range 441–3083 bp), including average 50 untranslated region

(UTR), ORF and 30 UTR sizes of 342, 867 and 438 bp, respectively

(Amiruddin et al., 2012). The longer transcript length was possibly

influenced by the high frequency of simple sequence repeats. Alignment

of translation initiation sequences proximal to each predicted start codon

identified a consensus Kozak sequence of (G/C) AAAATGG.Usage analysis

identified 10 under-represented codons, UAU, UGU, GUA, CAU, AUA,

CGA, UUA, CUA, CGU and AGU, in line with previous reports for

E. tenella (Amiruddin et al., 2012; Ellis et al., 1993). Simple sequence repeats

were again found to be common throughout many of the full-length

sequences, most commonly translating as poly-glutamine tracts (CAG) or

related equivalents following frameshifts and repeat degradation.

4.2.2 Transcript identification and inter-species comparisonSignificant EST and open reading frame expressed sequence tag

(ORESTES) cDNA datasets exist for E. tenella, E. acervulina and

E. maxima in the public domain. In a recent study, 48,361 ORESTES

and EST sequences derived from a series of E. tenella zoite and oocyst stages

at multiple developmental time points were collated and assembled into

8700 contiguous and singleton sequences (Novaes et al., 2012; Rangel

et al., 2013). Comparison with the 8786 putative protein coding sequences

predicted from the E. tenella genome sequence (http://www.genedb.org;

Table 2.1) suggested a good coverage, although the absence of sequences




derived from schizont and gametocyte stages indicates distinct gaps imposed

by the difficulty in obtaining suitable parasite material. Comparison with

other coccidian parasites including Neospora caninum and T. gondii

highlighted a conserved genome-wide gene density (as discussed above;

Table 2.1). Consideration of the individual sequence read distribution across

these assemblies prompted the authors to hypothesize that each life cycle

stage is likely to be characterized by a small number of highly expressed

genes, supplemented by a larger number of genes expressed at a much lower

level (Novaes et al., 2012). The application of RNAseq technologies is now

starting to significantly improve transcriptome coverage and gene predic-

tion, as has been described for N. caninum (Reid et al., 2012), and may con-

firm this hypothesis.

Equivalent ORESTES analyses for E. acervulina and E. maxima resulted

in 3413 and 3426 assembled cDNAs, respectively (Novaes et al., 2012;

Rangel et al., 2013), overlapping with many of the 1029 and 1380 unique

contiguous and singleton EST sequences derived in other notable studies for

these species (Miska et al., 2008; Schwarz et al., 2010). E. brunetti is at present

the only other eimerian parasite whose transcriptome has been systematically

sampled, being represented by 269 unique contiguous and singleton EST

sequences (Aarthi et al., 2011). Comparison with E. tenella cDNA sequences

identified putative homologues for between 19% and 32% of these unique

expressed sequences, with more than 47% of all sequences sharing no signif-

icant similarity to currently available annotated cDNA sequences derived

from any organism. The appearance of such a large number of unknown

putative coding sequences was anticipated and has been a common feature

of many apicomplexan genomes (Reid et al., 2012; Schwarz et al., 2010).

The number of putatively genus- and species-specific expressed sequences

is consistent with the specialized life cycle and exquisitely restricted host

and tissue range of these parasites. Nonetheless, transcripts encoding homo-

logues of key apicomplexan invasion-relevant proteins including several

microneme proteins (MICs) and glideosome components have been readily

identified in at least three Eimeria species (Novaes et al., 2012; Schwarz et al.,

2010). Similarly, surface antigen (SAG) transcripts have been identified

within the E. tenella, E. acervulina and E. maxima transcriptomes.

Hierarchical clustering of transcriptomic data derived from each of the

Eimeria species sampled to date has revealed a highly conserved expression

profile between life cycle stages and a strong correlation with stage order

within the life cycle. Thus, transcripts derived from sporozoites were most

likely to be conserved within the sporulated oocyst transcriptome



(Novaes et al., 2012). Similarly, close associations have been identified

between the sampled zoite stages and between oocyst datasets collected at

different stages of sporulation.

Functional transcriptomic studies using custom designed cDNA arrays

with or without a suppression subtractive hybridization step have been

undertaken with E. tenella and E. maxima. Examples include the association

of monensin resistance with up-regulation of transcripts involved with cyto-

skeletal rearrangement and energymetabolism and a panel of 32 differentially

expressed transcripts associated with precocious parasite development (Chen

et al., 2008; Dong et al., 2011). Screening a cDNA panel derived from mul-

tiple purified zoite and unpurified chicken intestine/schizont samples against

a genome tiling array has been used to identify coding sequences within

genetic loci mapped by association with susceptibility to strain-specific

immune killing in E. maxima (Blake et al., 2011a).

4.3. ProteomicsBefore the advent of genome sequencing and annotation, the processes by

which specific proteins could be identified, characterized and analyzed

were time-consuming, laborious and low throughput, usually limited to

the study of one or two individual proteins at a time. Methods for direct

analysis of known polypeptide sequences often utilized mass spectrometry

(MS), for example, to identify specific features of the protein such as pro-

teolytic cleavage sites. However, generation of de novo protein sequence

was most commonly achieved by chromatographic analyses of peptides

following chemical treatment of purified protein to progressively remove

amino acids, usually from the N-terminus. Over the past decade, huge

advances in MS instrumentation and the availability of well-annotated

genomes has allowed the burgeoning of high-throughput proteomics

technologies. Complex mixtures of proteins can be fragmented into small

peptides by enzymatic digestion or chemical degradation, and subjected in

parallel to high-energyMS that generates large numbers of individual spec-

tra from which peptide sequences can be directly inferred. These experi-

mentally derived sequences are then mapped in silico onto databases of

predicted proteins derived from annotated genomes, allowing the

unequivocal identification of full coding sequences. This type of approach

is well suited to high-throughput experiments in which hundreds or thou-

sands of individual proteins can be identified. While it is possible to analyze

very complex mixtures, such as a whole cell lysate, it is usual to carry a



protein separation technique prior to MS to reduce sample complexity and

aid the downstream in silico analysis. For parasites, a number of approaches

have been taken including one- or two-dimensional gel electrophoresis,

and in-line liquid chromatography (LC; recently reviewed by Wastling

et al., 2012).

4.3.1 Studies using MALDI-MSHigh-throughput proteomics technologies emerged in parasitology at the

beginning of the twenty-first century and were readily adopted by the

coccidian research community. Initial studies focused on polypeptide spots

excised from polyacrylamide gels following their separation by two-

dimensional electrophoresis (2DE). Spots were subjected to in-gel digestion,

usually with trypsin, and protein identifications made by generating peptide

mass fingerprints, acquired by matrix-assisted laser desorption/ionization

(MALDI) MS. Initial proteomes were derived for tachyzoites of T. gondii

(Cohen et al., 2002) and N. caninum (Lee et al., 2003) and for sporozoites

and second-generation merozoites of E. tenella (de Venevelles et al.,

2004; Liu et al., 2009). A major disadvantage of MALDI-MS is, however,

that unambiguous protein identifications can be made only if high-quality

gene annotations are available, which was not the case for these parasites

at the time. Therefore, MALDI data were generally supplemented by

re-analysis of selected protein spots using tandem MS to generate de novo

peptide sequence data. As an adjunct to proteomics studies, de Venevelles

et al. (2004) and Liu et al. (2009) probed 2DE blots of sporozoite or mer-

ozoite proteins with hyperimmune chicken sera and identified 50 and

85 spots, respectively, that were immunogenic, numbers that are in agree-

ment with several earlier studies using Western blotting (Sutton et al., 1989;

Tomley, 1994; Xie et al., 1992).

A different proteomics approach, aimed at elucidating the proteome of

purified microneme organelles from E. tenella sporozoites (Bromley et al.,

2003), used a post-source modification of MALDI, termed chemically

assisted fragmentation, which improves fragmentation efficiency and sim-

plifies interpretation of the spectrum. Briefly, a negatively charged group

is coupled to the N-terminus of tryptic peptides so that formation of a pos-

itively charged ion requires the introduction of two protons, one of which

resides in the peptide backbone, where it can resonate and assist fragmenta-

tion. After fragmentation, only y-ions retain a positive charge, which sim-

plifies the spectrum and allows it to be used to generate de novo peptide

sequences. Using this method, 37 of 96 spots excised from 2DE gels were



successfully identified, which included proteins known to reside in the

micronemes (EtMIC1, 2, 3) as well as several novel proteins that had not

previously been linked to this organelle. In a similar type of study, de

Venevelles et al. (2006), used a combination of MALDI and tandem MS

to analyze peptides derived from partially purified refractile bodies (RB)

of E. tenella sporozoites. As well as two proteins known to reside in the

RB (the aspartyl proteinase Eimepsin and the antigen SO7), 30 additional

putative RB proteins were identified including a hydrolase, a subtilisin

and a lactate dehydrogenase.

4.3.2 Studies using high-energy MSAdvances in MS instrumentation, especially the use of high-energy,

collision-induced dissociation-based, tandem MS, has revolutionized the

field of proteomics, allowing high-throughput identification of very large

numbers of tryptic peptides obtained from gel spots, gel slices or LC frac-

tions. Using a range of complementary approaches (2DE, gel-LC linked

to tandem MS and multi-dimensional protein identification technology,

MuDPIT), whole cell proteomes with a high level of coverage have now

been obtained for four developmental stages of E. tenella (Lal et al.,

2009). In addition, there are also high-quality proteomes available for var-

ious sub-cellular fractions including the rhoptry organelles of E. tenella

(Oakes et al., 2013).

The most comprehensive proteomics study of an eimerian is that of Lal

et al. (2009) who generated proteomes for four life cycle stages of E. tenella

(unsporulated oocysts, sporulated oocysts, sporozoites and second-

generation merozoites) resulting in the unequivocal identification of 1868

proteins, which represents almost 30% of the likely total number of

E. tenella proteins. A total of 288 proteins were conserved between sporo-

zoites and merozoites, but not found in unsporulated oocysts, suggesting

these are linked to zoite-specific functions such as attachment, invasion

and egress. These included proteins known to localize to the microneme,

and rhoptry secretory organelles and proteins associated with the

‘glideosome’, which drives zoite motility and the ‘moving junction’ (MJ)

structure that is formed at the host–parasite interface during invasion.

Importantly, it was noted that stage-specific variants of key molecules

involved in the formation of the MJ are expressed, such as AMAs and

rhoptry neck proteins (RONs), suggesting strongly that sporozoites and

merozoites assemble the MJ in a stage-specific manner, something

that has since also been shown to occur in T. gondii (Fritz et al., 2012a,b)



and has been confirmed again in the rhoptry sub-cellular proteome of

E. tenella (Oakes et al., 2013). Additional protein differences between

the zoite stages included differential expression of the GPI-linked families

of SAGs, as also shown in the transcriptome analyses (Novaes et al., 2012).

The multi-stage proteome indicates that energy production throughout

most of the developmental cycle is linked strongly to gluconeogenesis

and glycolysis, with the mannitol cycle present in oocysts and sporozoites,

but not in merozoites, consistent with in-depth biochemical analysis of this

pathway throughout the E. tenella life cycle (Allocco et al., 1999). It was

also apparent that proteins linked to oxidative phosphorylation were

expressed at the highest levels in the merozoite, suggesting a metabolic

shift to use oxygen to mobilize energy production during the asexual

phases of growth and replication. There is also a higher abundance of

proteins linked to transcription, protein synthesis and nucleotide metabo-

lism in the merozoite, compared to the sporozoite, which is consistent

with observations in the transcriptome and likely to be related to the

massive replication that has taken place within the schizont, just prior to

merozoite release.

4.4. The futureProgress in the development of novel molecular technologies applicable to

nucleotide sequencing, epigenetics and proteomics support increased

understanding of eimerian parasites, their close relatives and their host inter-

actions. Improved sequencing technologies facilitating cheaper, more effec-

tive, genome sequencing and assembly are now supporting the extension of

such studies to a rapidly increasing range of coccidial parasites. Next-

generation DNA and RNA sequencing strategies also have the potential

to revolutionize genetic mapping and our biological understanding of select-

able parasite phenotypes. Importantly, many such resources have been made

publically available for the Eimeria species. Genome sequences and predicted

protein datasets are freely available through EUpathDB (http://eupathdb.

org/eupathdb/), GeneDB (http://www.genedb.org/Homepage/Etenella)

and EmaxDB (http://www.genomemalaysia.gov.my/emaxdb/). Comple-

mentary resources including annotated transcript assemblies and genetic

maps can be accessed through the Eimeria Transcript database (Rangel

et al., 2013) (http://www.coccidia.icb.usp.br/eimeriatdb/) and NCBI

Map Viewer (http://www.ncbi.nlm.nih.gov/projects/mapview/), respec-

tively. The production of novel and improved datasets will impact upon



http://www.genedb.org/Homepage/Etenella

http://www.genomemalaysia.gov.my/emaxdb/

http://www.coccidia.icb.usp.br/eimeriatdb/

http://www.ncbi.nlm.nih.gov/projects/mapview/



the development of new diagnostic and anticoccidial control strategies and

provide tools with which questions of evolutionary and population biology

may be interrogated.

5. TRANSFECTION

Transfection refers to the introduction of exogenous DNA or RNA

into cells by chemical, biological or physical means. Through transfection,

the recipient cell can gain a new genetic trait, and some of the introduced

DNA can be integrated into the genome of the recipient cell. In

apicomplexan parasites, such as Toxoplasma and Plasmodium, plasmid-

mediated transient and stable transfection systems were established in the

early 1990s, but owing to the difficulty of completing the life cycle of Eimeria

in vitro, and the lack of regulatory DNA sequences, genetic manipulation has

lagged behind that of other protozoan parasites (Hao et al., 2007; Kelleher

and Tomley, 1998; Shi et al., 2008). The first stable transfection system was

developed in Eimeria in 2008, 10 years after the first report of a transient

transfection system in this genus of parasite (Clark et al., 2008; Kelleher

and Tomley, 1998).

5.1. Transfection construct designTransfection constructs are usually based on pre-constructed, commercially

available plasmids, and contain elements including regulatory and signal

sequences. Promoters of both constitutive genes and stage-specific genes,

together with their homologous or heterologous 30 UTR sequences, have

been used successfully to drive expression of reporter genes in eukaryotic

cells, including Eimeria. Using enhanced yellow fluorescent protein (eyfp

marker gene) as a reporter, it was shown that the E. tenella microneme pro-

tein 1 (EtMIC1) promoter drives EYFP expression in sporulated but not

unsporulated oocysts, as would be expected (Yin et al., 2011). Another study

showed that three different promoter sequences originating from E. tenella

could function effectively not only in other species of Eimeria but also in

T. gondii (Kurth and Entzeroth, 2009; Zou et al., 2009). Similarly, promoters

of the ‘housekeeping’ tubulin gene and the differentially regulated surface

antigen gene (sag1) of T. gondii, were effective in driving the expression

of the EYFP marker gene in E. tenella (Zou et al., 2009). As genetic tools

are well developed for T. gondii, the mutual recognition of these promoter

sequences in Eimeria and Toxoplasma suggests that some promoter sequences

fromT. gondii could be utilized directly in Eimeria and thatT. gondii could be



used as a novel transfection system for Eimeria-rooted vectors. This has the

potential to help improve our understanding of Eimeria spp. through the

development of both forward and reverse genetic technologies.

Several signal sequences are known to exhibit conserved activity in

apicomplexan parasites. For example, parasitophorous vacuole targeting sig-

nal sequences of T. gondii GRA8 and Plasmodium falciparum repetitive inter-

spersed family proteins have been found to function effectively in transfected

E. tenella and successfully target EYFP to the parasitophorous vacuole in

E. tenella (Shi et al., 2009; Yin et al., 2011). Furthermore, the nucleus-

targeting signal of the H5N1 subtypic avian influenza virus nuclear protein

also exhibits conserved functionality in eukaryotic cells, supporting nuclear

targeting when incorporated into an E. tenella transfection construct (Yin

et al., 2011).

5.2. Transient transfectionIn transient transfection, introduced exogenous DNA does not integrate

into the genome of the cell, but is transcribed into mRNA and is subse-

quently translated to protein. Transient transfection is an efficient tool to

identify regulatory and signal sequences of genes and to screen for genes

associated with certain phenotypes. For Eimeria, transfection of sporozoites

has been achieved by electroporation with plasmids; PCR amplified DNAs

or fragmented genomic templates that encode the exogenous DNA, flanked

by Eimeria-specific regulatory sequences (Hao et al., 2007; Kelleher and

Tomley, 1998; Liu et al., 2008). The efficiency of transient expression is usu-

ally low but this has been overcome by the use of restriction enzyme-

mediated integration (REMI), which boosts transfection efficiency about

200-fold (Liu et al., 2008). Very high transfection efficiency has been

achieved in E. tenella sporozoites by using cytomix-buffered REMI and

the AMAXA nucleofection system (Clark et al., 2008).

5.3. Stable transfectionStable transfection refers either to the permanent expression of the gene of

interest through the integration of the transfected DNA into the nuclear

genome, or the maintenance of a transfected plasmid as an extra chromo-

somal replicating episome within the cell. Stable transfection has been dif-

ficult to achieve in Eimeria because of the inability to transfect oocysts or

sporocysts, the absolute requirement for in vivo amplification and selection,

and the poor survival of sporozoites in the acidic environment of the host



stomach. The latter has been overcome by gavaging birds with sodium

bicarbonate to neutralize the acidic barrier (Clark et al., 2008). Stable trans-

fection systems have also been established forE. tenella by cloacal inoculation

of sporozoites, combined with in vivo drug selection and/or fluorescence

activated cell sorting (FACS) (Clark et al., 2008; Yan et al., 2009). To date,

the mutated dihydrofolate reductase–thymidylate synthase gene is the only

drug-mediated selection marker available for the transfection of Eimeria

(Clark et al., 2008; Yan et al., 2009). The mutated gene confers resistance

to pyrimethamine, a drug used to potentiate the action of the sul-

phonamides. Drug selection, together with FACS of fluorescence reporter

proteins and the high transient transfection efficiency using REMI, contrib-

uted to the success in establishing stable transfection in Eimeria.

Integration of a transfection construct into the Eimeria genome seems to

occur at random during the production of stably transfected Eimeria as

detected by Southern blotting and plasmid rescue (Yan et al., 2009). Quan-

titative real-time PCR analysis of insertion rate post-transfection showed an

average persistence of four copies of the tandem YFP reporter cassette per

genome from the first round of replication after electroporation and REMI.

After two further cycles of in vivo amplification with both FACS and pyri-

methamine selection, an average of 10 copies per genome were detected and

remained relatively stable through five further unselected generations. In

contrast, when REMI was not used, only a single copy of the relevant

reporter gene was detected per genome in first or second-generation trans-

fected parasite populations (Clark et al., 2008). Serial selection of fluorescent

mCitrine-transfected oocysts by FACS did not notably increase copy num-

ber, although tightened gating and FACS with sporocysts in place of oocysts

increased both the expression rate and the copy number to 2–3 per genome

(Clark et al., 2008).

5.4. PiggyBac-based forward genetic systemPiggyBac is a cut-and-paste transposon that is useful for transgenesis and inser-

tional mutagenesis and has been used for stable transfection in a wide variety of

organisms. This new molecular technology has been used successfully to

achieve targeted insertional mutagenesis in Eimeria (Su et al., 2012). Using

REMI, E. tenella sporozoites were electroporated with a mix containing

the restriction enzyme AscI, an AscI-linearized helper plasmid containing

the transposase gene, and an uncut donor plasmid containing the eyfp gene.

The eyfp gene was flanked by Eimeria-specific regulatory sequences that



were further flanked by piggyBac inverted terminal repeats (ITRs). Subse-

quently, electroporated sporozoites were inoculated into chickens via the

cloacal route and transfected progeny oocysts expressing eyfp were sorted

by flow cytometry. A stable eyfp expressing population was obtained by suc-

cessive in vivo passaging and FACS selection (Su et al., 2012). Locus-specific

PCR and genome walking revealed that the ITR-restricted sequence was

successfully targeted into TTAA sites, with about seven copies per genome

(Su et al., 2012). Both reverse and forward genetic tools will hopefully allow

an in-depth analysis of Eimeria basic biology. PiggyBac-mediated efficient

TTAA targeted mutations should be an attractive tool for genetic manipula-

tion of Eimeria.

5.5. Stably transfected Eimeria as a vaccine vector and beyondThe feasibility of using genetically modified Eimeria as a vaccine vector has

been studied using model antigens such as EYFP. It was found that

E. tenella expressing EYFP stimulated both humoral and cell-mediated

immunity to the expressed protein, and that antigen compartmentalization

affects the magnitude of the immune response with microneme-targeted

EYFP stimulating a higher IgA response than cytoplasm-targeted EYFP

(Huang et al., 2011). In another study, vaccination of specific pathogen-

free chickens with a population of E. tenella expressing Campylobacter

jejuni antigen A caused a significant reduction in bacterial load following

challenge with C. jejuni compared with unvaccinated and wild-type

E. tenella vaccinated controls (Clark et al., 2012). Thus, it has been dem-

onstrated that transfected Eimeria parasites can successfully express foreign

antigens that may stimulate immunity against a target pathogen. However,

to provide complete protection, co-expression of adjuvant antigens and/or

cytokines may be necessary (Guangwen Yin and Xun Suo, unpublished

observations).

Transfection of Eimeria species is still limited by the inability to transfect

oocysts and sporocysts, the difficulty of obtaining single-sporocyst-derived

recombinant clones, and the obligate requirement of in vivo amplification

and selection of stably transfected parasites (Clark et al., 2008; Shi et al.,

2008; Yan et al., 2009). It is difficult to maintain a large number of mutated

clones as a mutated Eimeria library, which needs manpower, facilities and

financial support. Nevertheless, high transfection efficiencies (Clark et al.,

2008; Hanig et al., 2012) will boost the advance of both reverse and for-

ward genetic systems in this important group of parasites. The interchange



and development of Eimeria transfection constructs between laboratories in

countries including China, France, Germany, Japan and the UK promises

rapid development over the coming years. More advanced genetic tools

established in other protozoa, such as Toxoplasma, Plasmodium, Leishmania

and Trypanosoma may eventually be applied to research with Eimeria.

6. OOCYST BIOGENESIS

One of the defining features of the coccidia is the oocyst. There are

three crucial milestones in oocyst production: first, merozoites undergo rapid,

asexual division within the intestine, amplifying dramatically the total number

of parasites poised to develop into microgametes or macrogametes; second,

microgametes fertilize the macrogametes and third, the macrogametes mobi-

lize specialized organelle – wall forming bodies (WFBs) – to generate the

oocyst wall, one of the most remarkable biological structures known.

The oocyst wall encapsulates and protects coccidian parasites as they exit

their definitive host in faeces and, subsequently, in the harsh, external world,

while they undergo meiosis to produce infectious sporozoites. Thus, the

oocyst is the endpoint of sexual reproduction. It is also notoriously resilient,

resisting both mechanical and chemical damage and tolerating changes in

humidity and temperature for months, if not years (reviewed by Belli

et al., 2006; Fritz et al., 2012a). This resilience is critical for transmission

of coccidian parasites from host to host, via ingestion of contaminated food

or water.

6.1. Veil and WFBsThe formation of the oocyst wall proceeds via an orderly release of the con-

tents of: first, the veil forming bodies; second, wall forming bodies type 1

(WFB1) and third, wall forming bodies type 2 (WFB2) (Ferguson et al.,

2003). The contents of the veil forming bodies are undescribed but form

a loose outer veil that appears to provide a temporary scaffold or frame

around the developing oocyst wall (Ferguson et al., 2000, 2003). It is lost

before the oocyst is excreted in the faeces and, therefore, plays no role in

protecting the parasite in transit from host to host (Ferguson et al., 1975;

Pittilo and Ball, 1980).

The release of the contents of the WFBs appears to be controlled by the

rough endoplasmic reticulum/Golgi apparatus (Ferguson et al., 2003). Once

a zygote has formed, WFB1 migrate to the periphery of the parasite, align

and disaggregate rapidly, before appearing to merge together to form the

Figure 2.3 Immunofluorescent images of macrogametocytes and early oocysts ofE. maxima within the intestine of a chicken 144 h post-infection. (A) An early-stagemacrogametocyte—type 1 and type 2, wall forming bodies (WFBs) are indistinguish-able. (B) A mid-stage macrogametocyte—type 1 and type 2, WFBs are distinguishableby size (WFB1s are larger). (C) A cluster of mature gametocytes showing peripheralalignment of WFBs, with disaggregation of some type 1 WFBs evident. (D) Earlyoocysts—the outer wall has formed and disaggregation of type 2 WFBs is evident. Hostnuclei are stained with 40,6-diamidino-2-phenylindole (blue); WFBs and oocyst walls arestained with antibodies to affinity-purified gametocyte antigens (green). Image suppliedby Professor D.J.P. Ferguson (University of Oxford, UK).



outer layer of the bi-layered oocyst wall (Fig. 2.3). This outer layer may ini-

tially be as thick as 600 nm but quickly compacts to 200 nm or less (Ferguson

et al., 2003). Not long after the outer layer forms, WFB2 are also transferred

to the parasites surface, by the endoplasmic reticulum, and also align, disag-

gregate and also appear to fuse together to form the inner layer of the oocyst

wall (Ferguson et al., 2003). This layer is less electron-dense than the outer

layer and more consistent in size, being around 40 nm in most species exam-

ined (reviewed by Belli et al., 2006). The inner and outer layers are, at first,

separated by a 40 nm zone, which shrinks as the wall compacts. However,

the two layers never fuse together and are readily separated in the laboratory

(Monne and Honig, 1954).



6.2. Oocyst wall proteinsGas chromatography and MS analyses of the oocyst walls of E. maxima and

E. tenella indicate that both layers are dominated by protein (>90%) with

surprisingly low levels of carbohydrate and lipids (Mai et al., 2009). Thus,

an understanding of the structure and characteristics of proteins that com-

prise the oocysts wall is essential for genuine understanding of how the wall

forms and why it is so robust. Only a small number of oocyst wall proteins

have been identified and the origin of all of these can be traced back to the

WFBs in macrogametes (reviewed in detail by Mai et al., 2009). These pro-

teins can be grouped into, essentially, three groups.

First, there is a 22 kDa antigen in the macrogametocytes of E. tenella,

which is found in WFB2 and the inner layer of the oocyst wall (Krucken

et al., 2008). This 22 kDa protein is dominated by histidine and proline res-

idues. As yet, no information is available about if or how this protein is

incorporated into the oocyst wall, though its involvement in stabilizing

the oocyst wall via cross-links between histidine and catechols, as seen in

insect cuticles, is a distinct possibility (Krucken et al., 2008).

Second, there is a family of nine large (174–190 kDa), cysteine-rich pro-

teins that localize to WFB1 of macrogametocytes and the outer wall of the

Cryptosporidium oocyst (Spano et al., 1997; Templeton et al., 2004). It is

thought that these ‘OWPs’ form disulphide bridges and matrices within

the oocyst wall (Spano et al., 1997). The recent discovery of seven OWPs

in Toxoplasma, with at least some of these localized to the outer oocyst wall

(Possenti et al., 2010), supports the idea that OWPs are involved in wall for-

mation. Recently, two OWP homologues have been found in Eimeria, at

least one of which appears to localize to WFB1 in macrogametes, further

evidence that OWPs are destined for the outer wall (Walker, 2009).

Third, there are numerous tyrosine-rich proteins, ranging in size from

8 to 31 kDa, in the inner wall of theEimeria oocyst; all of these are derived from

precursor proteins of 56 and 82 kDa (GAM56 and GAM82) from WFB2 in

macrogametes of several species of Eimeria (Belli et al., 2003a, 2009). It has also

been discovered very recently that, although the Toxoplasma genome contains

no direct homologues of either GAM56 or GAM82, the oocyst wall of Toxo-

plasma contains up to six tyrosine-rich proteins (Fritz et al., 2012a).

6.3. Formation of the oocyst wallThe role of tyrosine-rich proteins in the formation of the Eimeria oocyst wall

has been studied in some depth, the result being the proposal of a two-step



model: in step 1, precursor proteins found in WFBs are processed by

gametocyte-specific proteases into smaller, tyrosine-rich proteins and in step

2, peroxidases and/or oxidoreductases catalyze cross-linking of these pro-

teins via their tyrosine residues, resulting in extensive dityrosine matrices

within the oocyst wall. There is some evidence for both of these proposed

reaction steps.

It has been known for more than two decades that two proteins –

GAM56 and GAM82 – dominate the protein profile of gametocytes

(Wallach et al., 1989). Both GAM56 and GAM82 are processed into small,

tyrosine-rich proteins, as demonstrated in two ways: (i) antibodies to

GAM56 and GAM82 react with proteins of these sizes in gametocytes

but react with proteins of 8–31 kDa in oocysts (Belli et al., 2003a,b,

2009); and (ii) N-terminal sequencing of these wall proteins shows that they

are ‘cleaved’ from GAM56 or GAM82 at specific points (Belli et al., 2003a).

It has been discovered recently, using an in vitro assay, that the degradation of

GAM56 into smaller proteins is largely dependent on subtilase-like serine

protease activity (Katrib et al., 2012). There are at least six subtilase-like

enzymes in the genome of E. tenella, and at least three of these are expressed

specifically in gametocytes. Thus, assembly of the oocyst wall may follow a

mechanism that is similar to that involved in the assembly of the cuticle of

nematodes (Page and Winter, 2003; Thacker et al., 2006).

After formation of the numerous, small, tyrosine-rich derivatives of

GAM56 and GAM82, peroxidases or oxidoreductases are predicted to cat-

alyze their cross-linking via dityrosine bond formation. There is substantial

circumstantial evidence indicating that oocyst walls are rich in dityrosine

bonds. First, the oocysts exhibit a vivid blue autofluorescence between

the ultraviolet excitation wavelengths of 330 and 385 nm (Fig. 2.4), which

is characteristic of dityrosine cross-linking (Belli et al., 2006). Second,

dityrosine levels have been measured in the oocyst wall of E. maxima and

found to be remarkably high (Belli et al., 2003a), begging the conclusion

that their generation within the oocyst wall is a deliberate, enzymatically

catalyzed process, initiated by the parasite (Belli et al., 2006). It has been

shown that the WFBs of E. maxima embody a highly focused region of

peroxidase activity (Belli et al., 2003a, 2006) and, while neither an endog-

enous peroxidase or oxidoreductase has yet been isolated from Eimeria, it

has been established that exogenous peroxidases can induce dityrosine

cross-linking of a truncated version of GAM56 in vitro (Mai et al., 2011).

And, an oxidoreductase has been found in the oocyst wall of T. gondii

(Fritz et al., 2012a).

Figure 2.4 Sporulated and unsporulated oocysts of E. maxima showing characteristicUV autofluorescence (blue).



The concept that dityrosine cross-linking constitutes a critical feature of

the structure of the oocyst wall helps to explain the resilience of oocysts –

dityrosine cross-linking, leading to the formation of structural matrices,

sclerotization and quinone tanning, is widespread in nature, almost always

in association with the construction of protective coatings such as inverte-

brate egg shells, cuticles, cell walls, glues and cements (reviewed by Belli

et al., 2006). Moreover, it might be predicted that interfering with this pro-

cess is a way to limit the transmission of coccidian parasites. The subunit vac-

cine, CoxAbic®, may be an example of this. This vaccine contains GAM56

and GAM82 from E. maxima and laboratory experiments have shown that

immunization of broiler breeder hens with this vaccine stimulates the pro-

duction of protective IgY (¼IgG) antibodies that are transferred to offspring

chicks via the egg yolk (Wallach et al., 2008). There are two potential expla-

nations for this: (1) the antibodies ‘protect’ GAM56 and GAM82 from pro-

teolysis and, thereby, deprive the parasite of the tyrosine-rich building

blocks it needs to form the oocyst wall; and/or (2) the antibodies interfere

with dityrosine bond formation (Sharman et al., 2010).

7. HOST CELL INVASION

Apicomplexans, including all species of Eimeria, are highly successful

obligate intracellular parasites. Unlike many microorganisms that rely on

host-cellular pathways such as phagocytosis or pinocytosis for invasion,



apicomplexans invade host cells rapidly and forcefully in a highly regulated,

parasite-driven process (reviewed by Santos and Soldati-Favre, 2011). Initial

interaction with host cells can occur with the parasite in any orientation but

commitment to invasion requires that the apical pole of the parasite makes

irreversible contact with the host cell surface. This initiates formation of a

MJ, a tight focus of constriction between the parasite and host cell mem-

brane, which migrates towards the posterior end of the parasite as invasion

proceeds (reviewed by Besteiro et al., 2011). The parasite synthesizes spe-

cialized molecular complexes at the parasite–host interface, which are essen-

tial for gliding movement (the actinomyosin-dependent glideosome) and

formation of the MJ. These complexes are assembled by regulated secretion

of microneme (MIC) and rhoptry (RON/ROP) proteins, and interact with

the parasite motor, localized in the pellicle, and with specific receptors on

the host cell surface. As the parasite propels itself forwards, surface-bound

adhesion complexes are released by proteolysis. Other proteins, derived

largely from the rhoptries, are secreted into the host cell where they contrib-

ute to the formation of a parasitophorous vacuole and its associated mem-

brane, and modify the host intracellular environment (reviewed by

Boothroyd and Dubremetz, 2008).

7.1. Parasite surface proteinsIn common with many groups of protozoa, the surfaces of Eimeria sporozo-

ites and merozoites are coated with glycosylphosphatidylinositol (GPI)-

anchored proteins that are collectively referred to as surface antigens or SAGs

(Gurnett et al., 1990; Tabares et al., 2004). In T. gondii and species of Plas-

modium, GPI-anchored proteins are implicated in the early stages of parasite

attachment, prior to apical re-orientation. This requires interaction with

sulphated glycosaminoglycans on the surface of host cells. Preliminary data

indicate that several E. tenella SAGs are able to bind a variety of cultured cells

(F. Tomley and C. Subramaniam, unpublished observations) suggesting that

they too are involved in this initial non-specific binding step.

Examination of EST sequences from E. tenella identified 37 potential

GPI-linked variant SAGs encoded by multi-gene families and differentially

expressed between sporozoites and second-generation merozoites (Tabares

et al., 2004). GPI-anchored proteins in higher eukaryotes are often found

within membrane structures called lipid rafts, which are detergent-resistant

microdomains involved in signal transduction, membrane trafficking and

molecular sorting. Potential lipid rafts were identified on the surface of



Eimeria invasive stages by staining for the lipid-raft marker flotillin-1 (del

Cacho et al., 2007). However flotillin-1 was most prominent at the apical

end of sporozoites, whereas E. tenella SAGs are expressed over the entire

sporozoite surface (Tabare et al., 2004).

Recent transcriptome, proteome and genome data indicate that E. tenella

expresses up to 80 different SAG proteins, and confirms that these are dif-

ferentially regulated during the life cycle such that second-generation mer-

ozoites are coated with more complex mixtures of SAGs than either

sporozoites or first-generation merozoites (Lal et al., 2009; Novaes et al.,

2012; A. Reid, Wellcome Trust Sanger Institute, personal communication;

F. Tomley, unpublished observations). Their surface location suggests that

SAGs may induce potent immune responses that can, for example, block

sporozoite invasion of cultured cells (Brothers et al., 1988). The

co-expression by merozoites of highly polymorphic SAGs could render

anti-SAG immune responses ineffective against these stages. A study of

10 E. tenella merozoite-expressed SAGs showed that three of these induced

an increase in nitric oxide production, IL-1b and IL-10 transcription, and

induced a decrease in IL-12 and interferon-g (IFN-g) transcription in

chicken macrophages (Chow et al., 2011). This indicates that at least a subset

of SAGs has the ability to modulate chicken innate and adaptive immune

responses, which may suppress cell-mediated immunity and also contribute

to the marked pro-inflammatory responses and associated pathology seen

during E. tenella infection.

7.2. MIC proteins are adhesins and many functionas multi-protein complexes

The repertoire and broad functions of coccidian MIC proteins, including

those from Eimeria species, has been reviewed extensively, and the reader

is referred to recent articles for more details (Carruthers and Tomley,

2008; Cowper et al., 2012). Most MICs comprise modular arrangements

of protein domains that share homology with adhesins from higher eukary-

otes and it is the specific binding of these domains to host cell glycans that

establish irreversible apical attachment. Across the Apicomplexa there are

many orthologous MIC proteins although the precise arrangement of

domains is not always conserved and there are some, such as the sialic

acid-binding MAR domains and galactose-binding Apple/PAN domains,

discussed below, that are restricted to the coccidia.

MIC proteins often associate to form multivalent heteromeric com-

plexes, which assemble within the endoplasmic reticulum before being



trafficked to the micronemes. EachMIC complex contains an ‘escorter’ pro-

tein that possesses a transmembrane domain and a short cytoplasmic tail. This

facilitates targeting to the micronemes and allows the microneme complex

to interact with the underlying glideosome ( Jewett and Sibley, 2003). Some

MIC complexes are conserved across different genera, whereas others are

not. The T. gondii complex of TgMIC2/MIC2AP, which is essential for

gliding motility, host cell attachment and invasion (Huynh and

Carruthers, 2006), is orthologous to the E. tenella complex of EtMIC1/

MIC2. The introduction of the E. tenella complex into tachyzoites of

T. gondii can partially complement for loss of endogenous TgMIC2/

M2AP, indicating conservation of function (Huynh et al., 2004). However,

the T. gondii TgMIC1/4/6 complex, in which TgMIC6 is the escorter for

the MAR-domain containing TgMIC1 and Apple-domain containing

TgMIC4, is not replicated in E. tenella. The MAR-domain containing

EtMIC3 (Labbe et al., 2005) has not been found in a complex, but instead

is secreted directly from the micronemes onto the host cell surface (Lai et al.,

2011). The Apple-domain containing EtMIC5 forms a complex with

EtMIC4, which is presumed to act as an escorter, but which also bears

adhesive thrombospondin-like domains that have the potential to bind host

receptors (Periz et al., 2005, 2007).

7.3. Host glycan recognition by MIC proteins contributesto host and tissue tropism

One of the most intriguing biological questions is why there are such huge

differences in host and tissue tropisms across members of the coccidia.

T. gondii, for example, invades virtually any nucleated cell and infects almost

all warm-blooded vertebrates, whereas each species of Eimeria infects only a

single host, replicates only in epithelial cells, and is often restricted to very

specific regions of the intestine. Recent studies on the binding of coccidian

MIC proteins to host glycans, particularly sialic acid and galactose, are now

shedding light on this issue (Cowper et al., 2012; Lai et al., 2011; Marchant

et al., 2012). Generally, there is a direct correlation between host range and

the possession of a wide repertoire of MIC proteins expressing variant

domains that are capable of binding a broad range of oligosaccharide epi-

topes. MAR domains, which are found in MICs across the coccidia, bind

a range of sialyl groups but evidence from carbohydrate microarrays, atomic

structure, and cell binding studies, reveals that those from T. gondii and

E. tenella are differentially equipped for binding. Thus E. tenella MAR

domains bind a limited range of structures, with a strong in vivo preference



for a2,3-linked sialic acid and an absence of binding to any N-glycolated

sialyl structures (which are not found in the chicken). In contrast, MAR

domains from T. gondiiMICs are more divergent, and bind a variety of oli-

gosaccharides including a2,9-linked sialic acid and N-glycolylated deriva-

tives (Lai et al., 2011). Very recent studies on carbohydrate recognition

by Apple domains, also found inMICs across the coccidia, indicates an addi-

tional contribution to host range and tissue tropism conferred by differential

recognition of galactose, another sugar that is widely distributed in animal

tissues, commonly forming b1,3 or b1,4 linkages to a preceding glucose

or galactose. While both T. gondii and E. tenella Apple domains bind gal-

actosylated structures, there is a marked preference by T. gondii for

Galb1,3GalNAc, commonly found on gangliosides which are prevalent

on many host cell surfaces (Marchant et al., 2012). The precise binding pref-

erences of E. tenella Apple domains have not yet been elucidated, but pre-

liminary data indicate that these bind predominantly to b1,4-linkedgalactose and do not recognize the more common b1,3 linkages (Cowper

et al., 2012).

7.4. Regulated secretion of microneme and rhoptry organellesMIC proteins are discharged onto the parasite surface at an early stage in

invasion. The physiological trigger that induces microneme secretion is

not known, but treating parasites with agents that cause a rise in intracellular

free calcium induces rapid secretion and this can be inhibited by treatment

with intracellular calcium chelating agents (Bumstead and Tomley, 2000;

Carruthers and Sibley, 1999;Wiersma et al., 2004). Blocking calcium release

channels by treating parasites with IP3 inhibitors or ryanodine, or blocking

activity of cyclic GMP-dependent kinase or calcium-dependent protein

kinase, all interrupt the regulated exocytosis of MICs and prevent parasite

attachment and invasion of host cells (Dunn et al., 1996; Lourido et al.,

2010; Schubert et al., 2005; Wiersma et al., 2004).

ROP proteins are also secreted in a regulated manner and, while the

exact mechanisms are unknown, it is hypothesized that the initial signal

comes via the cytoplasmic tails of membrane-bound MIC complexes, fol-

lowing their interaction with a host cell receptor. The process is complicated

because rhoptry secretion occurs in two separate ‘waves’. Proteins that reside

within the anterior neck portion of the rhoptry (RONs) are secreted early in

invasion and are critical for the formation and maintenance of the MJ

(Alexander et al., 2005; Lebrun et al., 2005). However, proteins from the



posterior bulb of the rhoptry (ROPs) are secreted slightly later, once the par-

asitophorous vacuole is formed. In T. gondii, several ROP proteins are

known to be potent virulence factors that modify and subvert host cell sig-

nalling pathways (Bradley and Sibley, 2007). The current model for differ-

ential secretion of RONs and ROPs in T. gondii is that a membrane-bound

MIC complex containing the escorter protein TgMIC8 triggers release of

the RONs (Kessler et al., 2008). Interaction of RONs with AMA1 then

triggers the release of ROPs (Tyler and Boothroyd, 2011). Regulation of

secretion of ROP/RON proteins in Eimeria, where there is no defined

orthologue of TgMIC8, has not been determined.

7.5. AMAs and formation of the MJThe MJ was visualized many years ago (Aikawa, 1978) and is a key structure

that provides an anchor, against which the parasite can generate a force that

allows forward movement into the parasitophorous vacuole. A key compo-

nent is AMA1, a transmembrane protein secreted by the micronemes onto

the parasite surface, where it interacts with secreted RONs to initiate for-

mation of theMJ (Alexander et al., 2005; Lebrun et al., 2005). Collaboration

between proteins that are secreted from different sub-cellular organelles

requires a remarkable degree of orchestration. It has emerged recently that

while AMA1 remains anchored on the parasite surface, a RON complex is

secreted into the host cell and then one of them, RON2, becomes inserted

into the host cell membrane and interacts directly with AMA1 (Tyler and

Boothroyd, 2011). Both AMA1 and the RON protein repertoire are con-

served in Eimeria indicating that the mechanism for forming the MJ is likely

to be conserved in different coccidia (Blake et al., 2011a,b; Jiang et al., 2012;

Lal et al., 2009; Oakes et al., 2013). It is worthy of note, however, that

E. tenella expresses stage-specific variants of AMA1 and several of the

RON proteins, which suggests that each zoite stage assembles a different

set of gene products with which to build the MJ (Lal et al., 2009; Oakes

et al., 2013).

8. IMMUNOBIOLOGY

It is 50 years since Elaine Rose and Peter Long published, ‘Immunity

to four species of Eimeria in fowls’, sparking a seminal year in research into

the immunology of poultry coccidiosis. By the end of 1962, it was known

that: (a) even a single infection with various species of Eimeria confers solid



resistance to reinfection and this is increased further after a second infection

(Rose and Long, 1962); (b) immunity is expressed against the very early

asexual stages of infection such that, although penetration of epithelial cells

by sporozoites may occur, subsequent development is blocked (Pierce et al.,

1962); (c) immunity is not confined to early stages of parasite development

but also affects later stage merozoites and sexual stages (Rose, 1963); (d) in

general, immunity to one species confers no protection against other species

(Rose and Long, 1962), though it was shown subsequently that some degree

of cross-protection can occur between closely related pairs ofEimeria, such as

E. maxima and E. brunetti (Rose, 1967a), and E. tenella and E. necatrix (Rose,

1967b) and (e) B cells and the antibodies they produce play little, if any, role

in resistance to reinfection, implicating cell-mediated responses in acquired

immunity (Long and Pierce, 1963). However, in reviewing this remarkable

year, Rose (1963) noted that, ‘The way in which an animal which has expe-

rienced an infection with a species of Eimeria subsequently prevents the

development of that species within its body is not yet understood’. That

statement is equally valid today.

Immunity to Eimeria is complex, multifactorial and influenced by host

and parasite, with different elements playing greater or lesser roles in three

different types or stages of immunity: innate resistance to primary infection;

acquired immunity to reinfection and maternal immunity. Many host/

parasite combinations have been used to dissect the immunobiology of coc-

cidiosis, with significant insights being gained through the use of murine

models due to advantages connected with the availability of murine immu-

nological reagents, in-depth fundamental understanding of the murine

immune system and technologies to disrupt immune response genes in mice.

In this review, key similarities rather than differences in the immunobiology

of coccidial infections will be emphasized.

8.1. Innate responses to primary infectionPrimary infections with the majority of Eimeria species, in poultry and

rodents, are self-limiting; asexual reproduction proceeds via a pre-set num-

ber of cycles of schizogony prior to differentiation into gametocytes, subse-

quent sexual reproduction and production of oocysts. This can make it

challenging to demonstrate a role for the immune system in resistance to pri-

mary infection. Nevertheless, it has been shown that increased oocyst excre-

tion, by different Eimeria species, is a consistent feature of primary infection

in immunodeficient hosts (Klesius and Hinds, 1979; Long and Rose, 1970;



Mesfin and Bellamy, 1979; Rose, 1970; Rose and Hesketh, 1979, 1986;

Rose and Long, 1970; Schito and Barta, 1997; Schito et al., 1996;

Stockdale et al., 1985). Moreover, there is one parasite and host pairing –

E. vermiformis in the mouse – where the patent period of primary infection

is clearly increased in susceptible mice, almost certainly due to a relaxation of

immune pressure on the parasite that allows additional generations of schiz-

onts to develop (Rose and Millard, 1985; Rose et al., 1984, 1985; Schito

et al., 1996). Thus, the E. vermiformismurine model of coccidiosis has proved

particularly significant for our understanding of immunological resistance to

primary infection.

Studies with E. vermiformis have demonstrated that resistance to primary

infection is associated with more rapid inflammatory responses including

increased granulocyte numbers (Ovington et al., 1990), enhanced generation

of free oxygen radicals (Ovington et al., 1990), increased Natural Killer (NK)

cell activity (Smith et al., 1994a), earlier production of pro-inflammatory

cytokines such as IFN-g, tumour necrosis factor (TNF) and others

(Ovington et al., 1995; Wakelin et al., 1993), and faster T cell responses

(Rose et al., 1990; Wakelin et al., 1993). Corollaries of these observations

also exist for other rodent (Dkhil et al., 2011; Rausch et al., 2010; Rose

and Hesketh, 1982; Rose et al., 1979a; Schito and Barta, 1997), chicken

(Hong et al., 2006a,b; Kim et al., 2008, 2010; Rose et al., 1979a;

Rothwell et al., 1995, 2000, 2004; Yun et al., 2000) and turkey (Gadde

et al., 2011) coccidioses. However, of all these immunological parameters,

only two – lymphocytes and IFN-g – appear to be indispensible for resistance.Severe combined immunodeficient mice, which are deficient in both

T and B cells, are highly susceptible to primary infection with

E. vermiformis, producing many more oocysts and harbouring parasites for

far longer than immunocompetent mice (Schito et al., 1996). Studies in

B cell-deficient mice (Smith andHayday, 1998) and bursectomized chickens

(Long and Pierce, 1963; Rose and Hesketh, 1979), suggest that B cells play a

minor, though consistent, role in this resistance. Since deficiencies in antigen

presentation also increase susceptibility (Smith and Hayday, 1998, 2000),

this relatively minor role may be via the ability of B cells to act as antigen

presenting cells rather than anything to do with antibodies. Experiments

with congenitally athymic (nude) mice, however, show that T cells play a

critical role in resistance to primary infections with E. vermiformis; infected

nude mice excrete many more oocysts over a much extended patent period

(Rose et al., 1984, 1985). Other murine Eimeria (Klesius and Hinds, 1979;

Mesfin and Bellamy, 1979; Rose and Hesketh, 1986; Stockdale et al., 1985),



as well as E. nieschulzi in rats (Rose et al., 1979b), also produce many more

oocysts in athymic animals but without any affect on patency. Experiments

with thymectomized chickens have generated inconsistent data, probably

because of the difficulty in completely removing all T cells (Rose and

Long, 1970).

Depletion of specific T cell subsets, either via antibodies with appropri-

ate, rigorous, confirmatory adoptive transfer experiments (Rose et al., 1988,

1992), or via the deletion of specific genes from mice (Roberts et al., 1996;

Smith and Hayday, 1998, 2000), show that CD4þ, and not CD8þ, T cells

are the critical T cell subset mediating resistance to primary infection with

E. vermiformis. Moreover, protective effects appear to be ab T cell-specific

(Roberts et al., 1996; Smith and Hayday, 1998), though gd T cells may play

an important role in preventing immunopathology (Roberts et al., 1996)

rather than in contributing to the control of the parasite (Roberts et al.,

1996; Rose et al., 1996). Immune CD8þmesenteric lymph node cells have

also been shown to be capable of suppressing immunopathology in

E. falciformis infections (Pogonka et al., 2010). However, similar effects

are not seen in infections with Eimeria papillata (Schito et al., 1998) and

depletion studies in chickens infected with E. acervulina or E. tenella are

somewhat equivocal, possibly confounded by relatively small numbers of

experimental chickens in each treatment group and by inefficient depletion

of CD4þ T cells (Trout and Lillehoj, 1996).

The most important role for CD4þ T cells in mediating resistance to

primary infection with E. vermiformis is most likely as the source of IFN-

g. Mice treated with an antibody to IFN-g (Rose et al., 1991a) or IFN-ggene-knockout mice (Smith and Hayday, 2000) are highly susceptible to

E. vermiformis, suffering prolonged patency, high levels of excretion of

oocysts and increased mortality. This is not so evident in infections of

IFN-g knockout mice with E. papillata where patency is not affected; in

this case, NK cells are the likely source of IFN-g (Schito and Barta,

1997). How IFN-g is controlling the parasite is not known; generation of

free oxygen radicals (Ovington et al., 1995), reactive nitrogen intermediates

(Ovington et al., 1995; Smith and Hayday, 2000) and interference with

tryptophan metabolism (Schmid et al., 2012) can all be ruled out. However,

it is known that the effects of IFN-g are mediated via the host cell rather than

a direct effect on parasites (Rose et al., 1991b).

Infections with E. pragensis or E. falciformis indicate an additional role for

IFN-g in the immunobiology of coccidiosis. Depletion of IFN-g using



monoclonal antibodies has little apparent effect on parasite load but has a

significant affect on weight loss during primary and secondary infection with

E. pragensis (Rose et al., 1991a). Similarly, IFN-g receptor knockout mice

infected with E. falciformis suffer severe intestinal immunopathology and

weight loss mediated via Th17 pathways, involving the cytokines, IL17

and IL-22 (Stange et al., 2012). Thus, IFN-gmay have an important immu-

noregulatory role in response to infection with Eimeria, helping to keep

intestinal inflammation in check.

Mice deficient in granulocyte and NK cell function are more susceptible

to primary infectionwithE. vermiformis (Rose et al., 1984). However, T cell-

mediated control of infection with E. vermiformis does not require

co-operation with granulocytes (Rose et al., 1989) and experiments with

E. papillata indicate that increased susceptibility to primary infection may

be due more to participation of NK cells than granulocytes in resistance

(Schito and Barta, 1997). However, the effects on oocyst excretion and pat-

ent period are relatively modest compared to those of lymphocyte deficiency

(Schito et al., 1996) and a role for NK cells in innate resistance is not

supported by results obtained with E. vermiformis (Rose et al., 1995;

Smith et al., 1994a).

Free oxygen radicals appear to play no role in resistance to E. vermiformis

since quenching of their activity in vivo actually leads to reduced, not

enhanced, oocyst activity (Ovington et al., 1995). Moreover, treatment

of mice with agents designed to enhance macrophage activity, including free

oxygen radical generation, leads to increased oocyst excretion (Smith and

Ovington, 1996), as does treatment with TNF (Ovington et al., 1995). Sim-

ilarly, reactive nitrogen intermediates, despite their temporal association

with resistance to poultry coccidia (Allen and Fetterer, 2002), also enhance

oocyst production in E. vermiformis (Ovington et al., 1995). These are, at first

glance, puzzling results in light of the well-established anti-protozoal effects

of TNF, free oxygen radicals and reactive nitrogen intermediates (see

Ovington and Smith, 1992). However, with our current knowledge about

involvement of an oxidative reaction in oocyst wall assembly (Belli et al.,

2006; Mai et al., 2009, 2011) it makes some sense, indicating that perhaps

Eimeria actually subjugates the host’s oxidative burst to assist it in construc-

tion of its oocysts. Intriguingly, a related proposal has been put forward

recently – it appears that E. falciformis also subverts IFN-g-inducedindoleamine 2,3-dioxygenase activity to help drive microgamete develop-

ment (Schmid et al., 2012).



8.2. Acquired immunityAcquired immunity to Eimeria is even more enigmatic than innate resistance

to primary infections. All that can be said with any certainty is that immunity

to reinfection with Eimeria is remarkably effective and is T cell dependent

(this has been realized for more than 30 years; Rose et al., 1979b), and

that B cells (and, therefore, antibodies) are not involved in acquired immu-

nity since bursectomized birds (Rose and Hesketh, 1979) and mice lacking

B cells (Rose et al., 1984; Smith and Hayday, 1998) are perfectly capable of

developing immunity to reinfection. It has proven almost impossible to

correlate any immune parameter with immunity to reinfection because

the expression of that immunity in experimental settings, at least, is so rapid

and efficient. However, studies using gene-knockout mice have proved

extremely useful in determining which factors may play a role. Thus,

as for primary infection, CD4þ ab T cells are crucial for immunity to

reinfection with E. vermiformis (Roberts et al., 1996; Smith and Hayday,

1998, 2000). Similar, though less definitive data were also obtained for

secondary infections with E. papillata (Schito et al., 1998). However, in con-

trast to primary infection, IFN-g plays no role in this acquired immunity

(Rose et al., 1991a; Schito and Barta, 1997; Smith and Hayday, 2000).

Contrarily, some studies demonstrate that CD8þ T cells can be used

to transfer immunity (e.g. to E. falciformis; Pogonka et al., 2010) or that

depletion of CD8þ T cells can increase, very slightly, susceptibility to

E. vermiformis (Rose et al., 1992). Evidence from poultry experiments

(Trout and Lillehoj, 1996) is more difficult to interpret because experiments

showing an increase in oocyst excretion in secondary infection of birds

depleted of CD8þ T cells did not include a concomitant primary infection

control, making it hard to assess how significant the increased oocyst pro-

duction really was. More, and more sophisticated, analyses of acquired

immunity to Eimeria are required to resolve the mechanism(s) that are

operating.

8.3. Maternal immunityThe immune system of young animals is ‘uneducated’ rendering them more

susceptible to infectious disease. Protection against infection during this vul-

nerable period is provided via transfer of antibodies from mother to young.

In chickens, this occurs via the egg yolk; indeed, the ability of hens to trans-

fer remarkable quantities of IgY (¼IgG) antibodies to their hatchlings has

long been appreciated, including in regard to the transfer of antibodies that



protect chicks from infection with E. tenella (Rose and Long, 1971) or

E. maxima (Rose, 1972). In many of the progeny from hens deliberately

infected with high doses of E. maxima, this maternal immunity can be abso-

lute (i.e. result in the complete absence of oocysts in the faeces of chicks), at

least during the first week post-hatching (Smith et al., 1994b). Maternal anti-

body levels (in egg yolk or chicks) are correlated with protection (Smith

et al., 1994b). Moreover, maternal immunity induced by E. maxima confers

partial protection against E. tenella, possibly via cross-recognition of con-

served proteins (or, at least, epitopes) in different Eimeria species (Smith

et al., 1994c), an idea lent further credibility by the ability of maternal immu-

nization with conserved macrogametocyte proteins to protect hatchlings

against multiple species of Eimeria (Wallach et al., 1995, 2008).

The effectiveness of maternal, antibody-mediated immunity to Eimeria

appears contradictory to the body of evidence, reviewed above, indicating

that antibodies play only a minor role in resistance to Eimeria. However,

there is actually no shortage of (often overlooked) evidence, dating back

over 40 years, showing that antibodies can protect against infection with

Eimeria. Thus, for example, sera taken 2 weeks after infection with

E. maxima can be transferred to naıve birds and, as for maternal immunity,

protect some of them almost completely against infection (Rose, 1972,

1974). The protection conferred by these convalescent sera was later dem-

onstrated to be correlated tightly with levels of parasite-specific IgG

(Wallach et al., 1994). Additionally, it was demonstrated 40 years ago

(Rose, 1972) and, again, more recently (Lee et al., 2009a,b) that Eimeria-

specific antibodies purified from egg yolks of immunized hens can be used

to transfer passive immunity against several species of poultry coccidia; the

protection can be achieved via injection or oral delivery of the antibodies.

Immune sera can even partially protect highly susceptible T cell-deficient

animals (Rose and Hesketh, 1979). Thus, antibodies certainly can protect

against Eimeria but the effect must be described as variable – from absolute

to negligible even if similar immunization regimens are used (Wallach et al.,

1994). Why this is so is anything but clear. Maternal immunization,

however, does appear to be a phenomenon that can be harnessed to control

poultry coccidiosis (Smith et al., 1994b; Wallach et al., 2008).

8.4. Immunological researchLamentably, in the decade since the retirement of Elaine Rose, research

into the immunobiology of coccidiosis has declined significantly



(notwithstanding the efforts of researchers at the United States Department

of Agriculture to understand the role of various feed additives and Eimeria

profilin in boosting immune responses to Eimeria; reviewed by Lillehoj

and Lee, 2012). This has been exacerbated by the declines, even disappear-

ance, of the coccidiosis research programmes at the UK’s Institute for Ani-

mal Health and at the University of Technology, Sydney in Australia, as well

as a distinct lack of new commercial research into vaccines. It is doubly

lamentable because immunity to Eimeria, whether innate, acquired or

maternal, is remarkable among all parasites in its effectiveness. Moreover,

coccidiosis appears to be an excellent model to study the molecular basis

of gut immunopathology. The gut epithelium is the first point of contact

with the host for many pathogens but studies of this interaction have proven

challenging experimentally, with many models employing delivery of path-

ogens via subcutaneous, intravenous or intraperitoneal injection. Infection

with poultry or murine Eimeria, being largely confined to the intestine is,

therefore, an exceptional model for genuine study of gut immunology.

Hopefully, recent promising and innovative insights into unravelling the

complexity of the host/Eimeria inter-relationship (Blake et al., 2011a,b;

Stange et al., 2012) will be followed up and exploited fully.

9. DIAGNOSIS AND IDENTIFICATION

9.1. Traditional methods
Diagnosis of coccidiosis in poultry flocks continues to rely on necropsy and
the examination of birds for intestinal lesions in different areas of the gut.

Because of the site-specificity of invasion, the presence of lesions can provide

insight into which species of coccidia is/are responsible for clinical symp-

toms. Diagnosis may be corroborated by microscopic analysis of shape

and size of Eimeria oocysts shed in faeces from infected birds. Additional

criteria classically used to characterize Eimeria species include pre-patent

period, minimum sporulation time, tissue location of parasitic forms and

immunological specificity. However, definitive identification of a particular

Eimeria species based on morphological and pathological criteria can be

tedious, requires highly qualified personnel and may be confounded by

the overlapping features observed in different Eimeria species (Long and

Joyner, 1984).

While lesion site and aspect, and oocysts size and shape, are features often

sufficient to corroborate clinical signs of coccidiosis, there are instances

when knowing precisely which Eimeria species is/are present would be



helpful in managing the disease. For example, a preponderance of

E. acervulina in a litter sample might indicate increased drug resistance in this

species. This information would be useful in choosing alternative strategies,

such as switching to another anticoccidial compound known to be effective

against E. acervulina, or to a live Eimeria oocyst vaccine. If, on the other hand,

E. mitis or E. brunetti were present, then using a vaccine that contains only

E. acervulina, E. maxima and E. tenellawould not be a particularly useful con-

trol strategy.

9.2. Early molecular methods9.2.1 Starch gel electrophoresisIn the 1970s, a biochemical approach to identification of Eimeria spp. was

developed that involved starch gel enzyme electrophoresis of enzymes, such

as lactate dehydrogenase and glucose phosphate isomerase obtained from

oocyst homogenates (Rollinson, 1975; Shirley, 1975; Shirley and

Rollinson, 1979). The technique was employed to examine field isolates

of E. tenella obtained from around the world and was used to distinguish

mixtures of at least three species when present in one sample (Chapman,

1982; Shirley et al., 1989). The former study showed for the first time that

E. praecox, a species difficult to identify because of the absence of diagnostic

lesions, had a high incidence (74% of samples) in broiler flocks. While an

interesting laboratory tool for investigation of phenomena such as intraspe-

cific variation, starch gel electrophoresis is a time-consuming technique that

requires large numbers of oocysts. In addition, protein variability is limited

by evolutionary constraints, thus limiting the observed phenotypic poly-

morphism. Due to these limitations, this technique has been superseded

by DNA-based methods for identification of Eimeria spp. in the field.

9.2.2 DNA hybridizationDNA hybridization was the first DNA-based technique proposed for the

molecular discrimination of Eimeria parasites (Shirley, 1994b). A typical pro-

tocol consisted of genomic DNA digestion with different restriction

enzymes, separation through agarose gel electrophoresis, blotting and

hybridization with DNA probes composed of repetitive regions. The final

result was a DNA fingerprinting comprisingmultiple band profiles. Similarly

to enzyme variation detection, this approach also required large numbers of

parasites and was highly time demanding. Also, the method was inherently

unable to deal with mixed samples, since overlapping band profiles are not

informative.



9.3. Methods based on DNA amplification by PCRMolecular techniques, primarily the PCR, have been developed to over-

come the limitations of morphological examination and the aforementioned

molecular techniques. Since primers can be designed to specifically amplify

DNA of any single Eimeria species, samples containing multiple species can

be properly diagnosed. Also, the high level of amplification permits the use

of low numbers of oocysts. An excellent review of early efforts to develop

PCR assays for Eimeria is available (Morris and Gasser, 2006); here we pro-

vide an update of work in this area.

9.3.1 RAPD fingerprintingThe first PCR-based diagnostic assays developed for Eimeria relied on the

use of RAPD. This technique is based on DNA amplification using arbitrary

primers under low stringency. Under this condition, primers can anneal in

multiple sites of the target genomic DNA, thus producing DNA fingerprints

that permit differentiation of polymorphic populations. Because no previous

knowledge of the nucleotide sequence is required, RAPD was employed

largely for parasite discrimination at a time when very few genomic

sequences were available. Several groups succeeded in developing RAPD

assays for species and strain differentiation of poultry Eimeria (MacPherson

and Gajadhar, 1993; Procunier et al., 1993; Shirley and Bumstead, 1994).

However, given the low stringency of the reaction, RAPD typically suffered

from a low reproducibility, especially among different laboratories, and was

superseded by more reliable and specific PCR assays. Two main approaches

have been employed by different groups to develop such specific assays:

(1) the use of ITS1 and ITS2 and (2) the conversion of anonymous RAPD

markers into Sequence-Characterized Amplified Region (SCAR) markers.

9.3.2 PCR assays directed to specific targetsA specific PCR assay directed to E. tenella 5S rDNA was reported over

20 years ago (Stucki et al., 1993). This pioneer work was followed by assays

capable of detecting and differentiating the seven Eimeria species of domestic

fowl using ITS1 (Schnitzler et al., 1998, 1999) and ITS2 rDNA (Gasser et al.,

2005) as targets. ITS1 and ITS2 are intervening sequences that are post-

transcriptionally excised from the rRNA precursor. Unlike 18S, 5.8S and

28S rRNAs, ITS1 and ITS2 are not subjected to an appreciable selective

pressure, and have undergone sufficient divergence among Eimeria species

to allow design of species-specific primers (Lew et al., 2003). A number



of assays based on amplification of ITS1 and ITS2 rDNA have been devel-

oped, and used to determine the species of Eimeria present in poultry litter

(Hamidinejat et al., 2010; Haug et al., 2007, 2008; Jenkins et al., 2006a; Lew

et al., 2003). As a word of caution in using ITS sequences, Lew et al. (2003)

found sufficient variation on ITS1 of different E. maxima isolates to require

the design and use of two distinct sets of specific primers for this species.

Combined with rapid techniques for extracting high-quality DNA from

oocysts, ITS1-specific PCR was found to provide a more accurate picture

of Eimeria distribution at poultry farms than traditional morphometric anal-

ysis (Hamidinejat et al., 2010; Haug et al., 2008).

Primers directed to conserved ribosomal DNA sequences (18S, 5.8S,

28S), flanking either the ITS1 or ITS2 regions, have also been used in com-

bination with denaturing polyacrylamide gel electrophoresis or capillary

electrophoresis (CE) for species discrimination (Cantacessi et al., 2008;

Gasser et al., 2005; Morris et al., 2007a,b). The latter method relies upon

identifying species-specific peaks in CE chromatograms that have been

established using pure cultures (Gasser et al., 2005). Using this approach,

one group has identified a genetic variant of E. maxima, and new operational

taxonomic units in oocysts isolated from poultry operations (Cantacessi

et al., 2008; Morris et al., 2007b). By employing primers directed to con-

served regions (28S, 5.8S) flanking the ITS1 sequence, these authors iden-

tified genetic variants that would have gone unnoticed using ITS1-specific

primers. Although assays based on ITS1 and ITS2 are highly sensitive due to

the large number of rDNA repeats (Vrba et al., 2010), variation in the

sequence may prevent primer binding. Nevertheless, ITS1 is still being used

as a target for the development of diagnostic assays for Eimeria parasites of

other hosts, including 4 species pathogenic for turkeys (Cook et al.,

2010) and 11 species that infect the domestic rabbit (Oliveira et al., 2011).

As an alternative to ITS1 and ITS2, Fernandez et al. (2003a) used RAPD

to develop SCAR markers for each Eimeria species of domestic fowl. In

developing this assay, the DNA sequences of individual RAPD markers

were determined and used to design longer primers, which were then tested

under highly stringent conditions for species-specific amplification of

Eimeria DNA. By combining a set of seven SCAR markers, Fernandez

et al. (2003b) developed a multiplex PCR assay that permits the simulta-

neous discrimination of all Eimeria species infecting chickens in a single-tube

reaction (Fig. 2.5). An Eimeria SCAR database containing 151 SCARs is

publicly available on the web (Fernandez et al., 2004; http://www.

coccidia.icb.usp.br/eimeriaScardb), and SCAR markers have been used

http://www.coccidia.icb.usp.br/eimeriaScardb

http://www.coccidia.icb.usp.br/eimeriaScardb

Figure 2.5 Agarose gel electrophoresis of multiplex PCR products using DNA samplesof E. acervulina (lane 1), E. brunetti (lane 2), E. tenella (lane 3), E. mitis (lane 4), E. praecox(lane 5), E. maxima (lane 6), E. necatrix (lane 7), a mixture of the seven Eimeria species(lane 8) and a control with no starting DNA (lane 9). Molecular size markers (lane M) inbase pairs are indicated on the left Reproduced from Fernandez et al. (2003b) with per-mission from Cambridge University Press.



by other groups to determine the Eimeria species composition on poultry

farms in different regions of the world (Carvalho et al., 2011a,b;

Ogedengbe et al., 2011b), and for the development of quantitative PCR

assays (Blake et al., 2008; Vrba et al., 2010). The advantage of SCARmarker

technology, unlike assays based on ITS1- and ITS2-PCR, is that highly con-

served SCAR marker sequences are available as targets of amplification

(Blake et al., 2008; Fernandez et al., 2004; Vrba et al., 2010). This avoids

the problem of false negative reactions due to poor annealing of primer

to target DNA because of variation in the target sequence. The drawback

to SCAR technology is that it may be less sensitive than assays based on

ITS1 and ITS2, which are found in multiple copies in the Eimeria genome

(Vrba et al., 2010).

Finally, strain differentiation still lacks the variety of molecular markers

already available for other apicomplexan parasites. In this direction, it is

worth mentioning that species-specific sets of microsatellite markers for

E. acervulina, E. maxima and E. tenella have been developed in Brazil

(A. Gruber and S. Fernandez, unpublished observations). This work has

been deposited as a patent (Espacenet, patent BRPI0702051). These



microsatellite markers allowed for the differentiation of both field samples

and commercial vaccines lines.

Despite the enormous impact of PCR-based methods to detect and dis-

criminate Eimeria species, some drawbacks still persist. Oocysts are the most

accessible stage of the life cycle, and the obvious choice as a source of Eimeria

DNA. Since the oocyst wall is remarkably resistant to chemical agents,

mechanical disruption with glass beads is the most common method to

extract DNA. However, DNA yield does not correlate linearly with the

number of oocysts (Fernandez et al., 2003b), due to decreased efficacy of

mechanical disruption in low-concentration suspensions. Despite some

authors having proposed alternative chemical treatments to disrupt the

oocyst wall (Haug et al., 2007; Zhao et al., 2001), this step remains the most

important limiting factor for good sensitivity. Beside faeces, litter is another

important source of Eimeria samples; however, a drawback of utilizing poul-

try litter as a source of Eimeria DNA is the presence of PCR inhibitors.

Although different extraction techniques have been developed to recover

DNA from oocysts (Carvalho et al., 2011a; Haug et al., 2007; Zhao

et al., 2001), the adequate removal of inhibitory substances from litter is

often difficult. With the goal of controlling for false negative reactions

due to PCR inhibition, a technique has been developed based on

co-amplification of ITS1 sequences and an Eimeria species-specific internal

standard ( Jenkins et al., 2006b). Using gel electrophoresis, the target and

internal standard PCR products can easily be distinguished from each other

by acrylamide gel electrophoresis (Fig. 2.6). In the event of inhibition, a sec-

ond extraction of DNA is undertaken with the goal of removing inhibitors.

9.3.3 Quantitative PCR assaysEarly assays using ITS1, ITS2 or SCAR markers relied on qualitative assays

in which identification of amplification products has been obtained by aga-

rose or polyacrylamide gel electrophoresis. Over the past 5 years, a number

of assays using quantitative real-time PCR (qPCR) amplification of ITS1,

ITS2 or SCAR markers have been developed. Blake et al. (2008) utilized

primers that were specific for SCAR markers of E. acervulina, E. necatrix

and E. tenella, and for a mic gene of E. maxima. The authors reported a sen-

sitivity varying from 1 to 10 genomes. Another group utilized ITS1 targets

to amplify E. acervulina, E. brunetti, E. maxima, E. necatrix and E. tenellaDNA

isolated from pure cultures and field samples, and achieved an assay sensitiv-

ity that was between 10 and 100 oocysts (Kawahara et al., 2008). A qPCR

assay using primers conserved among various protozoa and melting curve

Figure 2.6 Detection of Eimeria species oocysts using ITS1-PCR and internal standard (IS).Ea, E. acervulina; Eb, E. brunetti; Ema, E.maxima; Emi, E.mitis; En, E. necatrix; Et, E. tenella. kbp,fX174 HaeIII DNA standards. *Target band for each species of Eimeria. Reproduced fromJenkins et al. (2006a,b) with permission from the American Association of Avian Pathologists.



analysis could detect E. acervulina in a mixture of oocysts (Lalonde and

Gajadhar, 2011).

Multiplexing real-time PCR, using one of four different upstream

primers conserved between two species, and a conserved downstream

primer, in combination with species-specific TaqMan probes, was used to

analyze all Eimeria species in poultry litter (Morgan et al., 2009).

A greater number of species were identified than those revealed by micros-

copy. Another group reported the development of FAM-labelled TaqMan

species-specific probes, targeted to microneme 1 gene of E. maxima and

SCAR markers of the remaining species (Vrba et al., 2010). An assay sensi-

tivity of a single sporulated oocyst has been claimed. In a study using only

E. acervulina, qPCR directed to SCAR markers was applied to DNA

extracted from oocysts obtained from cloacal swabs, and compared with

oocyst counts from individual faecal droppings (Velkers et al., 2010). The

authors concluded that qPCR of cloacal swabs might be useful for determin-

ing the prevalence and identity of Eimeria oocysts in litter. A similar

approach using high-resolution melting curve analysis and qPCR directed

to ITS1 sequences was found capable of identifying all seven Eimeria species

in pure oocysts cultures (Kirkpatrick et al., 2009).

9.4. LAMPWhile qPCR is superior to conventional PCR in that it eliminates the need

for gel electrophoresis and provides quantitative results, samples must be run



on a fairly expensive real-time apparatus. In fact, the complexity of DNA

extraction from the oocyst, associated with the need for expensive

thermocycling and electrophoresis equipment, severely limit the use of

molecular assays in poultry farms. To overcome these limitations, an

Eimeria-specific PCR-based technique that utilizes loop-mediated isother-

mal amplification (LAMP) technology has been developed (Barkway et al.,

2011). Since the enzyme is isothermal, the reaction can be performed in a

simple heat block or water bath, without the need for thermocyclers. Also,

detection can be made with intercalating dyes using the naked eye for obser-

vation (Fig. 2.7), thus eliminating the requirement for electrophoresis.

Finally, instead of using oocysts, the proposed protocol employs mucosal tis-

sues collected post-mortem as samples, and DNA extraction by a simple

boiling method. Altogether, the method addresses the several limitations

Figure 2.7 Loop-mediated isothermal amplification (LAMP) specific for E. tenella.Application to a purified genomic DNA dilution series revealed a limit of detectionof between 1 and 10 E. tenella genomes using agarose gel electrophoresis (A) orhydroxynaphthol blue as a visual indicator (B, blue: positive, pink: negative). �ve,no template negative control.



of conventional molecular assays and may become a mainstream cost-

effective tool for the diagnosis of Eimeria infection in poultry flocks.

9.5. Morphological diagnosis revisitedOne of the key features of morphological diagnosis based on oocyst shape

and size is the inherent subjectivity of the method and the requirement of

skilled personnel. An early attempt to differentiate species of Eimeria in

the fowl utilized a computerized image-analysis system (Kucera and

Reznicky, 1991). The method used two measurements, length and width

of oocysts, which restricted the ability to differentiate all seven species

due to the overlap of these characters. To address these limitations,

Castanon et al. (2007) have reported the development of COCCIMORPH,

a system that implements a framework for feature extraction, shape charac-

terization and automated classification of chicken Eimeria oocyst images.

The system employs a classifier trained with thousands of oocyst images

of all species of chicken Eimeria. COCCIMORPH provides a public web

frontend (www.coccidia.icb.usp.br/coccimorph) that permits users to

upload oocyst images and obtain a reliable diagnosis in real-time. The

Bayesian classifier showed an overall correct species assignment of 85.7%,

with individual rates varying from 74.9% for E. necatrix to 99.2% for

E. maxima. While this system still has many limitations to be widely used

as a mainstream diagnostic system, it represents a proof of principle that mor-

phology may have gained a revival in the era of digital image processing and

pattern recognition methods. COCCIMORPH breaks a classical paradigm,

as it does not require sample transportation to a reference laboratory, and

photomicrographs sent through the Internet are sufficient to obtain species

diagnosis. While not competing with the accuracy, sensitivity and quantita-

tive nature of modern PCR-based methods, morphological diagnosis based

on digital image processing might represent a near-zero cost alternative to be

used by technicians in poultry farms that do not harbour expensive molec-

ular biology facilities.

9.6. ConclusionsField diagnosis of Eimeria infection in poultry will continue to rely on the

identification of intestinal lesions and microscopic examination of faecal

droppings and litter for Eimeria oocysts. However, several molecular assays

that can detect and differentiate all seven Eimeria species of the chicken are

now available, and are being used either in a research setting to study the

http://www.coccidia.icb.usp.br/coccimorph



epidemiology of avian coccidiosis, or by private companies to monitor the

purity of vaccine lines. Information gleaned from molecular assays can assist

in managing disease by allowing informed decisions on which anticoccidial

compounds or live oocyst vaccines should be used in particular poultry

farms. The development of real-time qPCR represents a step forward

towards quantitative diagnosis of Eimeria. Also, isothermal amplification

assays with colorimetric detection (e.g. LAMP), that avoids a requirement

of expensive equipment, and the development of more robust image

processing software, may provide low-cost alternatives for species diagnosis

in poultry farms.

10. CONTROL

10.1. Chemotherapy
A truly landmark contribution to poultry science was the demonstration, in
1948, that it was possible to control coccidiosis by the continuous inclusion

of an anticoccidial drug (sulphaquinoxaline) in the feed of chickens

(Grumbles et al., 1948; reviewed by Chapman, 2009). The principle

involved (prevention or prophylaxis) has had a profound impact on our abil-

ity to grow chickens and turkeys under intensive conditions. Indeed it is

possible that the modern poultry industry could never have developed to

its present extent without the advent of drugs to control coccidiosis. Today,

anticoccidial drugs are incorporated routinely into the feed of broiler

chickens and turkeys for this purpose (Chapman, 2001, 2008). For example,

data available for the United States indicates that the use of anticoccidial

drugs in broiler flocks varied from 70% to 98% depending upon the season

(AgriStats, Inc., Fort Wayne, IN, USA). In Western Europe, 91% of com-

plexes use an anticoccidial drug (C. Bostvironnois, Elanco Animal Health,

personal communication). Drug usage is similarly extensive in other major

poultry producing regions around the world. Although there may be sea-

sonal variation in the use of drugs, it is clear that chemotherapy as a means

of control is widespread. The long-term outlook for such a heavy reliance

upon chemotherapy is often stated to be uncertain because of the widespread

development of drug resistance, a problem first recognized in the 1950s, and

a concomitant lack of new drug discovery. Furthermore, some anticoccidials

have been banned (in the EU) and others are said to be under threat

(McDonald and Shirley, 2009). So far, however, such considerations do

not seem to have led to a decline in drug use.



Anticoccidial drugs fall into two categories, the synthetic compounds

(produced by chemical synthesis and popularly known as ‘chemicals’) and

the ionophore antibiotics, which are by-products of bacterial fermenta-

tion. Synthetic drugs were the first to be discovered and comprise a diverse

array of molecules that are absorbed into the blood stream of the host and

kill developing parasites in epithelial cells of villi in the intestine. One of

the oldest synthetic drugs, nicarbazin, is also one of the most successful and

is still used widely today (Chapman, 1994a). Ionophores have a different

mode of action from synthetic drugs since they are able to destroy motile

stages of the Eimeria life cycle (sporozoites and merozoites) in the gut

lumen or following cell penetration (Smith and Strout, 1979). Since the

introduction of monensin in 1972, ionophores have been the most widely

used anticoccidial drugs for the control of coccidiosis (Chapman, 2001).

The mode of action and discovery of monensin, together with matters

of importance to the poultry industry such as toxicity, pharmacology,

residues and resistance to this drug, has been reviewed recently

(Chapman et al., 2010).

Despite the availability of a dozen or so anticoccidial drugs it may be sur-

prising to know that for the majority the mode of action against coccidia is

not known (Chapman, 1997). In one case, the discovery of a biochemical

pathway unique to Eimeria (the mannitol cycle) enabled the mode of action

of nitrophenide, a drug briefly used in the 1950s, to be elucidated (Schmatz

et al., 1989). Unfortunately, resistance to this drug developed quickly, a fate

shared by most other synthetic compounds. Diclazuril and decoquinate are

synthetic drugs to which resistance can also develop. Diclazuril has recently

been shown to induce ultrastructural changes in merozoites and cause

disruption of transmembrane potential in the mitochondrion (Zhou et al.,

2010). It is not clear if this reflects a true mode of action or is just a

consequence of cell death. Decoquinate, like other members of the quino-

lone family, is known to act against the electron transport chain of coccidia

(reviewed by Chapman, 1997). This drug has recently been shown to cause

chromosomal rearrangements during meiosis in oocysts of E. tenella

(Del Cacho, et al., 2006). The mode of action of ionophores involves dis-

ruption of ion transport across the parasite cell membrane (reviewed by

Chapman, et al., 2010) and resistance has been much slower to develop.

Evidence has been obtained that resistance may be due to changes in the flu-

idity of the cell membrane of sporozoites (Wang, et al., 2006). A recent study

suggests that monensin is also able to interrupt invasion of host cells by spo-

rozoites (del Cacho et al., 2007).



Most drugs are no longer as effective as when they were first introduced

due to the development of drug resistance. For example, one recent report

indicated that 68% and 53% of field isolates of E. acervulina from chicken

flocks in the EU were resistant to the synthetic drug diclazuril, and the ion-

ophore monensin, respectively (Peek and Landman, 2006). Similar reports

of resistance have been reported worldwide. In the turkey, drug resistance

has also been shown to be widespread (Rathinam and Chapman, 2009).

Details of the emergence of resistance in the 1970s to decoquinate have been

provided retrospectively (Williams, 2006). Although many surveys have

been published indicating the extent of resistance, little research has been

conducted on the mechanisms involved. Biochemical, genetic and applied

aspects of resistance have been reviewed (Chapman, 1997).

An early insight was that use of low concentrations of certain drugs in the

feed did not necessarily prevent the acquisition of immunity (Grumbles

et al., 1948). It is now known that most drugs are effective in the field

because they only partially suppress parasite development, allowing birds

to acquire natural immunity as a consequence of exposure to parasites that

escape drug action (Chapman, 1999). An advantage of immunity develop-

ment is that it allows the safe withdrawal of drugs several weeks before the

birds are sold with considerable savings in the cost of medication and reduc-

tion of the risk of potential drug residues in poultry meat.

10.2. VaccinationVaccination as a means to control coccidiosis has a long history (see

Chapman, 2003; Williams, 2002a) but it is only in the past 20 years or so

that this has proved a practical method for the control of coccidiosis in com-

mercial broiler flocks, principally because it has proved feasible to vaccinate

chicks in the hatchery by spraying birds with controlled numbers of oocysts

within enclosed cabinets. This involves considerable cost savings compared

with traditional methods of vaccination which were carried out on the farm

by trained personnel. Most commercially available vaccines comprise live

oocysts and vary according to the number of species of Eimeria included,

the numbers of oocysts present, and whether or not they are attenuated.

Vaccines containing all species that infect the chicken are used mainly to

immunize egg laying stock whereas vaccines containing fewer species (usu-

ally E. acervulina, E. maxima and E. tenella) are used in broilers. The first vac-

cines comprised populations of wild-type oocysts that were potentially

pathogenic, but more recently, vaccines containing attenuated parasites



which have reduced pathogenicity but retain immunogenicity, have been

introduced.

The purpose of vaccination with live oocysts is to provide an early initial

stimulus of the immune response. After placement of birds on litter, new

vaccinal oocysts are shed in the faeces and, following sporulation, these

are capable of re-infecting the flock. Secondary exposure to vaccinal oocysts

and wild-type oocysts present in the litter is thought to induce protective

immunity. Development of immunity takes several weeks and some cases

of vaccination failure occur because birds are overwhelmed with exposure

to wild-type virulent oocysts before they have had time to develop an

immune response. It is obviously important that vaccination is undertaken

carefully because any chicks that are not exposed to vaccinal oocysts may be

vulnerable to potentially high numbers of virulent oocysts when placed on

litter. The objective of vaccination is to induce sufficient immunity to pre-

vent chronic infestation while still allowing sufficient Eimeria to accumulate

such that a full immune response to the local Eimeria species will develop.

Several reviews have been published that are concerned with various aspects

of vaccination (Chapman, 2000; Chapman et al., 2002; Shirley et al., 2005;

Vermeulen et al., 2001; Williams, 2002b). Guidelines have been developed

to facilitate the worldwide adoption of consistent standard procedures

for determining the efficacy and safety of live anticoccidial vaccines

(Chapman et al., 2005).

Vaccination can also be achieved by in ovo injection of sporulated oocysts

into the embyronating egg (Weber et al., 2004). This is carried out in the

hatchery using complex machines that are also able to deliver other poultry

vaccines. Surprisingly, little research has been published that explains how

the oocysts, when injected into the egg, are able to establish a patent infec-

tion in the gut of the developing embryo. It is possible that infection results

from exposure to oocysts present in the eggshells post-hatch. As with other

methods of vaccination, secondary exposure to infection via the litter fol-

lowing placement of birds on the litter is necessary for a protective immune

response.

Another approach to vaccination involves immunizing hens with

affinity-purified antigens from the WFBs of macrogametocytes of

E. maxima (Sharman et al., 2010; Wallach et al., 2008). The nature of these

antigens has already been described in the section on oocyst biogenesis

above. Maternal antibodies pass via the egg to the newly hatched chick

and provide passive protection of limited duration. This is the only subunit

vaccine currently employed against any protozoan parasite. As in the case of



live oocyst vaccines, however, full protective immunity requires exposure to

potentially pathogenic coccidia in the litter.

Considerable research has been undertaken utilizing molecular technolo-

gies to identify antigens capable of inducing an immune response but in most

cases only partial protection has been achieved and none of the vaccine candi-

dates have been proven in commercial applications (Shirley and Lillehoj, 2012).

10.3. Strategies for the control of coccidiosisMeetings aimed at poultry producers, poultry veterinarians and other pro-

fessional groups with an interest in the poultry industry, often include pre-

sentations concerned with ‘strategies’ for the control of coccidiosis.

Unfortunately, data that can be tested rigorously are rarely presented, partly

due to the difficulty in designing reproducible trials under field conditions.

Thus the almost universal presence of Eimeria in poultry flocks negates the

inclusion of an ‘uninfected’ group in any controlled study. Strategies may

involve drugs, vaccines or both. In the case of drugs, different products,

often with different modes of action, may be used in the different feeds that

are provided during the life of a flock of birds. Often referred to as ‘shuttle’

programmes, one such employed in the United States involves inclusion of

nicarbazin or a nicarbazin/narasin mixture in the first feed and monensin or

salinomycin in the second; however, there are numerous variations

(Chapman, 2001). In subsequent flocks, different drugs may be used, for

which the term ‘rotation programme’ has been coined. Use of such

programmes is widespread and may be considered an appropriate strategy

for the control of coccidiosis although rarely is evidence of long-term effi-

cacy (compared with other approaches) available.

Vaccination, and its integration with chemotherapy in control

programmes, is an alternative strategy. Evidence has been obtained that par-

tial restoration of sensitivity to drugs may occur following the use of vaccines

comprising drug-sensitive strains of Eimeria. This phenomenon has been

demonstrated for the ionophores, monensin and salinomycin, and the syn-

thetic drug, diclazuril (Chapman, 1994b; Jenkins et al., 2010; Peek and

Landman, 2006). Based upon these observations, a yearly rotation pro-

gramme has been proposed in which use of ionophores is alternated in suc-

cessive flocks with vaccination (Chapman et al., 2010); such programmes are

commonly used in the United States. There is a need for more evidence to

support the notion that rotation programmes involving vaccines and drugs

prolong the life of the latter in the field.



10.4. Natural productsThere is considerable current interest in the use of so-called ‘natural prod-

ucts’ which may include plant extracts, probiotics and so on, to reduce prob-

lems caused by coccidiosis (e.g. Allen, 2003, 2007; Faber et al., 2012;

Giannenas et al., 2012; Lee et al., 2011). For example, the antimalarial

artemisinin, a product extracted from the herb Artemisia annua, was shown

to have a deleterious effect upon macrogametocytes of E. tenella by affecting

the expression of an enzyme sarcoplasmic–endoplasmic reticulum calcium

ATPase (del Cacho et al., 2010). Improved resistance to E. acervulina was

observed when the diet of chickens was supplemented with garlic metabo-

lites (Kim et al., 2012). There are many other examples in the literature.

Most natural products, however, are said not to specifically target the parasite

but ‘improve gut health’ or act as ‘enhancers’ of some aspect of immune sys-

tem function. All natural products contain undefined chemicals that, neces-

sarily, will have to be evaluated for safety and toxicity before being

acceptable for registration authorities. There is often very little scientific

literature that supports the claims made for these products and, as far as is

known, none are used commercially at present probably because of unreal-

istic dietary inclusion rates and failure to demonstrate efficacy under con-

trolled conditions.

11. CONCLUSIONS

In this review, we have considered selective aspects of research con-

cerned with the apicomplexan parasites of the genus Eimeriawhich cause the

disease, coccidiosis, of domestic livestock. The emphasis has been on poul-

try, where coccidiosis has been shown to have an enormous economic

impact. Fortunately, control of coccidiosis in poultry has been achieved,

by a combination of improved management, the prophylactic use of drugs,

and vaccination. Nevertheless, we should not be complacent because the

parasite has not been eradicated from commercial facilities where animals

are reared and is still capable of causing production losses.

In recent years, many research projects, and publications that result, have

used the modern tools of molecular biology, biochemistry, cell biology and

immunology to expand greatly our knowledge of these parasites and the dis-

ease they cause. Such studies are essential if we are to develop newmeans for

the control of coccidiosis. Past success was achieved by research funded and

conducted by universities, government agencies and private industry.



In recent years, however, a number of government funded organizations

have terminated their coccidiosis programmes and private industry has been

reluctant to allocate funds to the animal health sector due to the enormous

costs involved in drug discovery and vaccine development. Few university

departments have the facilities to undertake costly coccidiosis research. It is

evident, therefore, that both from a practical and research perspective con-

trol of coccidiosis is at a crossroads. In the future, the ever expanding human

population will become increasingly dependent upon a source of cheap pro-

tein of which poultry will necessarily be an important component. Hope-

fully, this will not be compromised by the ubiquitous parasites of the

genus Eimeria.

ACKNOWLEDGEMENTWe would like to thank Thilakar Rathinam for help in preparing the figures.

REFERENCESAarthi, S., Raj, G.D., Raman, M., Blake, D., Subramaniam, C., Tomley, F.M., 2011.

Expressed sequence tags from Eimeria brunetti—preliminary analysis and functional anno-tation. Parasitol. Res. 108, 1059–1062.

Aikawa, M., Miller, L.H., Johnson, J., Rabbege, J., 1978. Erythrocyte entry by malarial par-asites. A moving junction between erythrocyte and parasite. J. Cell Biol. 77, 72–82.

Alexander, D.L., Mital, J., Ward, G.E., Bradley, P., Boothroyd, J.C., 2005. Identification ofthe moving junction complex of Toxoplasma gondii: a collaboration between distinctsecretory organelles. PLoS Pathog. 1 (2), e17.

Allen, P.C., 2003. Dietary supplementation with Echinacea and development of immunity tochallenge infection with coccidia. Parasitol. Res. 91, 74–78.

Allen, P.C., 2007. Anticoccidial effects of xanthohumol. Avian Dis. 51, 21–26.Allen, P.C., Fetterer, R.H., 2002. Recent advances in biology and immunobiology of

Eimeria species and in diagnosis and control of infection with these coccidian parasitesof poultry. Clin. Microbiol. Rev. 15, 58–65.

Allocco, J.J., Profous-Juchelka, H., Myers, R.W., Nare, B., Schmatz, D.M., 1999. Biosyn-thesis and catabolism of mannitol is developmentally regulated in the protozoan parasiteEimeria tenella. J. Parasitol. 85, 167–173.

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 cDNAsequences provides insights into the Eimeria tenella transcriptome. BMC Genomics 13, 21.

Bandoni, S.M., Duszynski, D.W., 1988. A plea for improved presentation of type material forcoccidia. J. Parasitol. 74, 519–523.

Barkway, C.P., Pocock, R.L., Vrba, V., Blake, D.P., 2011. Loop-mediated isothermalamplification (LAMP) assays for the species-specific detection of Eimeria that infectchickens. BMC Vet. Res. 7, 67.

Barta, J.R., Martin, D.S., Liberator, P.A., Dashkevicz, M., Anderson, J.W., Feighner, S.D.,Elbrecht, A., Perkins-Barrow, A., Jenkins, M.C., Danforth, H.D., Ruff, M.D., Profous-Juchelka, H., 1997. Phylogenetic relationships among eight Eimeria species infectingdomestic fowl inferred using complete small subunit ribosomal DNA sequences.J. Parasitol. 83, 262–271.

http://refhub.elsevier.com/B978-0-12-407705-8.00002-1/rf0005
































Barta, J.R., Coles, B.A., Schito, M.L., Fernando, M.A., Martin, A., Danforth, H.D., 1998.Analysis of infraspecific variation among five strains of Eimeria maxima from NorthAmerica. Int. J. Parasitol. 28, 485–492.

Barta, J.R., Martin, D.S., Carreno, R.A., Siddall, M.E., Profous-Juchelka, H., Hozza, M.,Powles, M.A., Sundermann, C., 2001. Molecular phylogeny of the other tissue coccidia:Lankesterella and Caryospora. J. Parasitol. 87, 121–127.

Barta, J.R., Schrenzel, M.D., Carreno, R., Rideout, B.A., 2005. The genus Atoxoplasma(Garnham 1950) as a junior objective synonym of the genus Isospora (Schneider 1881)species infecting birds and resurrection of Cystoisospora (Frenkel 1977) as the correctgenus for Isospora species infecting mammals. J. Parasitol. 91, 726–727.

Beck, H.P., Blake, D.P., Darde, M.L., Felger, I., Pedraza-Dıaz, S., Regidor-Cerrillo, J.,Gomez-Bautista, M., Ortega-Mora, L.M., Putignani, L., Shiels, B., Tait, A.,Weir, W., 2009. Molecular approaches to diversity of populations of apicomplexan par-asites. Int. J. Parasitol. 39, 175–189.

Belli, S.I., Wallach, M.G., Luxford, C., Davies, M.J., Smith, N.C., 2003a. Roles of tyrosine-rich precursor glycoproteins and dityrosine- and 3,4-dihydroxyphenylalanine-mediatedprotein cross-linking in development of the oocyst wall in the coccidian parasite Eimeriamaxima. Eukaryot. Cell 2, 456–464.

Belli, S.I., Wallach, M.G., Smith, N.C., 2003b. Cloning and characterization of the 82 kDatyrosine-rich sexual stage glycoprotein, GAM82, and its role in oocyst wall formation inthe apicomplexan parasite, Eimeria maxima. Gene 307, 201–212.

Belli, S.I., Smith, N.C., Ferguson, D.J.P., 2006. The coccidian oocyst: a tough nut to crack!Trends Parasitol. 22, 416–423.

Belli, S.I., Ferguson, D.J., Katrib, M., Slapetova, I., Mai, K., Slapeta, J., Flowers, S.A.,Miska, K.B., Tomley, F.M., Shirley, M.W., Wallach, M.G., Smith, N.C., 2009. Con-servation of proteins involved in oocyst wall formation in Eimeria maxima, Eimeria tenellaand Eimeria acervulina. Int. J. Parasitol. 39, 1063–1070.

Besteiro, S., Dubremetz, J.F., Lebrun, M., 2011. The moving junction of apicomplexan par-asites: a key structure for invasion. Cell. Microbiol. 13, 797–805.

Blake, D.P., Hesketh, P., Archer, A., Carroll, F., Smith, A.L., Shirley, M.W., 2004. Parasitegenetics and the immune host: recombination between antigenic types of Eimeria maximaas an entree to the identification of protective antigens. Mol. Biochem. Parasitol. 138,143–152.

Blake, D.P., Hesketh, P., Archer, A., Carroll, F., Shirley, M.W., Smith, A.L., 2005. Theinfluence of immunizing dose size and schedule on immunity to subsequent challengewith antigenically distinct strains of Eimeria maxima. Avian Pathol. 34, 489–494.

Blake, D.P., Hesketh, P., Archer, A., Shirley, M.W., Smith, A.L., 2006. Eimeria maxima: theinfluence of host genotype on parasite reproduction as revealed by quantitative real-timePCR. Int. J. Parasitol. 36, 97–105.

Blake, D.P., Qin, Z., Cai, J., Smith, A.L., 2008. Development and validation of real-timepolymerase chain reaction assays specific to four species of Eimeria. Avian Pathol. 37,89–94.

Blake, D.P., Billington, K.J., Copestake, S.L., Oakes, R.D., Quail, M.A., Wan, K.L.,Shirley, M.W., Smith, A.L., 2011a. Genetic mapping identifies novel highly protectiveantigens for an apicomplexan parasite. PLoS Pathog. 7 (2), e1001279.

Blake, D.P., Oakes, R., Smith, A.L., 2011b. A genetic linkage map for the apicomplexanprotozoan parasite Eimeria maxima and comparison with Eimeria tenella. Int. J. Parasitol.41, 263–270.

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.
























































Boothroyd, J.C., Dubremetz, J.F., 2008. Kiss and spit: the dual roles of Toxoplasma rhoptries.Nat. Rev. Microbiol. 6, 79–88.

Bradley, P.J., Sibley, L.D., 2007. Rhoptries: an arsenal of secreted virulence factors. Curr.Opin. Microbiol. 10, 582–587.

Bromley, E., Leeds, N., Clark, J., McGregor, E., Ward, M., Dunn, M.J., Tomley, F.M.,2003. Defining the protein repertoire of microneme secretory organelles in theapicomplexan parasite Eimeria tenella. Proteomics 3, 1553–1561.

Brothers, V.M., Kuhn, I., Paul, L.S., Gabe, J.D., Andrews, W.H., Sias, S.R.,McCaman, M.T., Dragon, E.A., Files, J.G., 1988. Characterization of a surface antigenof Eimeria tenella sporozoites and synthesis from a cloned cDNA in Escherichia coli. Mol.Biochem. Parasitol. 28, 235–247.

Bumstead, J., Tomley, F., 2000. Induction of secretion and surface capping of micronemeproteins in Eimeria tenella. Mol. Biochem. Parasitol. 110, 311–321.

Cai, X., Fuller, A.L., McDougald, L.R., Zhu, G., 2003. Apicoplast genome of the coccidianEimeria tenella. Gene 321, 39–46.

Canning, E.U., Anwar, M., 1968. Studies on meiotic division in coccidial and malarial par-asites. J. Protozool. 15, 290–298.

Cantacessi, C., Riddell, S., Morris, G.M., Doran, T., Woods, W.G., Otranto, D.,Gasser, R.B., 2008. Genetic characterization of three unique operational taxonomicunits of Eimeria from chickens in Australia based on nuclear spacer ribosomal DNA.Vet. Parasitol. 152, 226–234.

Carruthers, V.B., Sibley, L.D., 1999. Mobilization of intracellular calcium stimulates micro-neme discharge in Toxoplasma gondii. Mol. Microbiol. 31, 421–428.

Carruthers, V.B., Tomley, F.M., 2008. Microneme proteins in apicomplexans. Subcell.Biochem. 47, 33–45.

Carvalho, F.S., Wenceslau, A.A., Teixeira, M., Albuquerque, G.R., 2011a. Molecular diag-nosis of Eimeria species affecting naturally infected Gallus gallus. Genet. Mol. Res. 10,996–1005.

Carvalho, F.S., Wenceslau, A.A., Teixeira, M., Matos Carneiro, J.A., Melo, A.D.,Albuquerque, G.R., 2011b. Diagnosis of Eimeria species using traditional and molecularmethods in field studies. Vet. Parasitol. 176, 95–100.

Castanon, C.A.B., Fraga, J.S., Fernandez, S., Gruber, A., Costa, L.F., 2007. Biological shapecharacterization for automatic image recognition and diagnosis of protozoan parasites ofthe genus Eimeria. Pattern Recognit. 40, 1899–1910.

Chapman, H.D., 1982. The use of enzyme electrophoresis for the identification of the speciesof Eimeria present in field isolates of coccidia. Parasitology 85, 437–442.

Chapman, H.D., 1994a. A review of the biological activity of the anticoccidial drugnicarbazin and its application for the control of coccidiosis in poultry. Poult. Sci.Rev. 5, 231–243.

Chapman, H.D., 1994b. Sensitivity of field isolates of Eimeria to monensin following the useof a coccidiosis vaccine in broiler chickens. Poult. Sci. 73, 476–478.

Chapman, H.D., 1997. Biochemical, genetic and applied aspects of drug resistance in Eimeriaparasites of the fowl. Avian Pathol. 26, 221–244.

Chapman, H.D., 1999. Anticoccidial drugs and their effects upon the development of immu-nity to Eimeria infections in poultry. Avian Pathol. 28, 521–535.

Chapman, H.D., 2000. Practical use of vaccines for the control of coccidiosis in the chicken.World’s Poult. Sci. J. 56, 7–20.

Chapman, H.D., 2001. Use of anticoccidial drugs in broiler chickens in the USA: analysis forthe years 1995 to 1999. Poult. Sci. 80, 572–580.

Chapman, H.D., 2003. Origins of coccidiosis research in the fowl—the first fifty years. AvianDis. 47, 1–20.

Chapman, H.D., 2008. Coccidiosis in the turkey. Avian Pathol. 37, 205–223.























































Chapman, H.D., 2009. A landmark contribution to poultry science—prophylactic control ofcoccidiosis in poultry. Poult. Sci. 88, 813–815.

Chapman, H.D., Rose, M.E., 1986. Cloning of Eimeria tenella in the chicken. J. Parasitol. 72,605–606.

Chapman, H.D., Cherry, T.E., Danforth, H.D., Richards, G., Shirley, M.W.,Williams, R.B., 2002. Sustainable coccidiosis control in poultry production: the roleof live vaccines. Int. J. Parasitol. 32, 617–629.

Chapman, H.D., Roberts, B., Shirley, M.W., Williams, R.B., 2005. Guidelines for evalu-ating the efficacy and safety of live anticoccidial vaccines, and obtaining approval for theiruse in chickens and turkeys. Avian Pathol. 34, 279–290.

Chapman, H.D., Jeffers, T.K., Williams, R.B., 2010. Forty years of monensin for the controlof coccidiosis in poultry. Poult. Sci. 89, 1788–1801.

Chen, T., Zhang, W., Wang, J., Dong, H., Wang, M., 2008. Eimeria tenella: analysis of dif-ferentially expressed genes in the monensin- and maduramicin-resistant lines usingcDNA array. Exp. Parasitol. 119, 264–271.

Chow, Y.P., Wan, K.L., Blake, D.P., Tomley, F., Nathan, S., 2011. Immunogenic Eimeriatenella glycosylphosphatidylinositol-anchored surface antigens (SAGs) induce inflamma-tory responses in avian macrophages. PLoS One 6 (9), e25233.

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 Eimeriaspecies. Mol. Biochem. Parasitol. 162, 77–86.

Clark, J.D., Oakes, R.D., Redhead, K., Crouch, C.F., Francis, M.J., Tomley, F.M.,Blake, D.P., 2012. Eimeria species parasites as novel vaccine delivery vectors: anti-Campylobacter jejuni protective immunity induced by Eimeria tenella-delivered CjaA.Vaccine 30, 2683–2688.

Cohen, A.M., Rumpel, K., Coombs, G.H., Wastling, J.M., 2002. Characterisation of globalprotein expression by two-dimensional electrophoresis and mass spectrometry: proteo-mics of Toxoplasma gondii. Int. J. Parasitol. 32, 39–51.

Cook, S.M., Higuchi, D.S., McGowan, A.L., Schrader, J.S., Withanage, G.S., Francis, M.J.,2010. Polymerase chain reaction-based identity assay for pathogenic turkey Eimeria.Avian Dis. 54, 1152–1156.

Cowper, B., Matthews, S., Tomley, F., 2012. The molecular basis for the distinct host andtissue tropisms of coccidian parasites. Mol. Biochem. Parasitol. 186, 1–10. http://dx.doi.org/10.1016/j.molbiopara.2012.08.007.

del Cacho, E., Gallego, M., Monteagudo, L., Lopez-Bernad, F., Quilez, J., Sanchez-Acedo, C., 2001. A method for the sequential study of Eimerian chromosomes by lightand electron microscopy. Vet. Parasitol. 94, 221–226.

del Cacho, E., Pages, M., Gallego, M., Monteagudo, L., Sanchez-Acedo, C., 2005. Syn-aptonemal complex karyotype of Eimeria tenella. Int. J. Parasitol. 35, 1445–1451.

del Cacho, E., Gallego, M., Pages, M., Monteagudo, L., Sanchez-Acedo, C., 2006. Effect ofthe quinolone coccidiostat decoquinate on the rearrangement of chromosomes ofEimeria tenella. Int. J. Parasitol. 36, 1515–1520.

del Cacho, E., Gallego, M., Sanchez-Acedo, C., Lillehoj, H.S., 2007. Expression offlotillin-1 on Eimeria tenella sporozoites and its role in host cell invasion. J. Parasitol.93, 328–332.

del Cacho, E., Gallego, M., Francesch, M., Quılez, J., Sanchez-Acedo, C., 2010. Effect ofartemisinin on oocyst wall formation and sporulation during Eimeria tenella infection.Parasitol. Int. 59, 506–511.

de Venevelles, P., Chich, J.F., Faigle, W., Loew, D., Labbe, M., Girard-Misguich, F.,Pery, P., 2004. Towards a reference map of Eimeria tenella sporozoite proteinsby two-dimensional electrophoresis and mass spectrometry. Int. J. Parasitol. 34,1321–1331.
































http://dx.doi.org/10.1016/j.molbiopara.2012.08.007

http://dx.doi.org/10.1016/j.molbiopara.2012.08.007





















de Venevelles, P., Chich, J., Faigle, W., Lombard, B., Loew, D., Pery, P., Labbe, M., 2006.Study of proteins associated with the Eimeria tenella refractile body by a proteomicapproach. Int. J. Parasitol. 36, 1399–1407.

Dkhil, M., Abdel-Baki, A.A., Delic, D., Wunderlich, F., Sies, H., Al-Quraishy, S., 2011.Eimeria papillata: upregulation of specific miRNA-species in the mouse jejunum. Exp.Parasitol. 127, 581–586.

Dolnik, O.V., Palinauskas, V., Bensch, S., 2009. Individual oocysts of Isospora(Apicomplexa: Coccidia) parasites from avian feces: from photo to sequence.J. Parasitol. 95, 169–174.

Dong, H., Lin, J., Han, H., Jiang, L., Zhao, Q., Zhu, S., Huang, B., 2011. Analysis of dif-ferentially expressed genes in the precocious line of Eimeria maxima and its parent strainusing 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 ofcalmodulin-domain protein kinases in Eimeria tenella and Eimeria maxima. Parasitology113, 439–448.

Ellis, J., Griffin, H., Morrison, D., Johnson, A.M., 1993. Analysis of dinucleotide frequencyand codon usage in the phylum Apicomplexa. Gene 126, 163–170.

Faber, T.A., Dilger, R.N., Hopkins, A.C., Price, N.P., Fahey Jr., G.C., 2012. The effects of agalactoglucomannan oligosaccharide-arabinoxylan (GGMO-AX) complex in broilerchicks challenged with Eimeria acervulina. Poult. Sci. 91, 1089–1096.

Ferguson, D.J., Hutchison, W.M., Siim, J.C., 1975. The ultrastructural development of themacrogamete and formation of the oocyst wall of Toxoplasma gondii. Acta Pathol.Microbiol. Scand. B 83, 491–505.

Ferguson, D.J., Brecht, S., Soldati, D., 2000. The microneme protein MIC4, or an MIC4-like protein, is expressed within the macrogamete and associated with oocyst wall for-mation in Toxoplasma gondii. Int. J. Parasitol. 30, 1203–1209.

Ferguson, D.J., Belli, S.I., Smith, N.C., Wallach, M.G., 2003. The development of the mac-rogamete and oocyst wall in Eimeria maxima: immuno-light and electron microscopy.Int. J. Parasitol. 33, 1329–1340.

Fernandez, S., Costa, A.C., Katsuyama, A.M., Madeira, A.M., Gruber, A., 2003a. A surveyof the inter- and intraspecific RAPDmarkers of Eimeria spp. of the domestic fowl and thedevelopment of reliable diagnostic tools. Parasitol. Res. 89, 437–445.

Fernandez, S., Pagotto, A.H., Furtado,M.M., Katsuyama, A.M.,Madeira, A.M., Gruber, A.,2003b. A multiplex PCR assay for the simultaneous detection and discrimination of theseven Eimeria species that infect domestic fowl. Parasitology 127, 317–325.

Fernandez, S., Katsuyama, A.M., Kashiwabara, A.Y., Madeira, A.M., Durham, A.M.,Gruber, A., 2004. Characterization of SCAR markers of Eimeria spp. of domestic fowland construction of a public relational database (The Eimeria SCARdb). FEMSMicrobiol. Lett. 238, 183–188.

Fernando, M.A., Remmler, O., 1973. Four new species of Eimeria and one of Tyzzeria fromthe Ceylon Jungle fowl Gallus lafayettei. J. Protozool. 20, 43–45.

Fritz, H.M., Bowyer, P.W., Bogyo, M., Conrad, P.A., Boothroyd, J.C., 2012a. Proteomicanalysis of fractionatedToxoplasma oocysts reveals clues to their environmental resistance.PLoS One 7 (1), e29955.

Fritz, H.M., Buchholz, K.R., Chen, X., Durbin-Johnson, B., Rocke, D.M., Conrad, P.A.,Boothroyd, J.C., 2012b. Transcriptomic analysis of Toxoplasma developmentreveals many novel functions and structures specific to sporozoites and oocysts. PLoSOne 7 (1), e29998.

Gadde, U., Chapman, H.D., Rathinam, T., Erf, G.F., 2011. Cellular immune responses,chemokine, and cytokine profiles in turkey poults following infection with the intestinalparasite Eimeria adenoeides. Poult. Sci. 90, 2243–2250.























































Gasser, R.B., Skinner, R., Fadavi, R., Richards, G., Morris, G., 2005. High-throughputcapillary electrophoresis for the identification and differentiation of seven species ofEimeria from chickens. Electrophoresis 26, 3479–3485.

Giannenas, I., Papadopoulos, E., Tsalie, E., Triantafillou, E., Henikl, S., Teichmann, K.,Tontis, D., 2012. Assessment of dietary supplementation with probiotics on perfor-mance, intestinal morphology and microflora of chickens infected with Eimeria tenella.Vet. Parasitol. 188, 31–40.

Grumbles, L.C., Delaplane, J.P., Higgins, T.C., 1948. Continuous feeding of low concen-trations of sulfaquinoxaline for the control of coccidiosis in poultry. Poult. Sci. 27,605–608.

Gurnett, A., Dulski, P., Hsu, J., Turner, M.J., 1990. A family of glycolipid linked proteins inEimeria tenella. Mol. Biochem. Parasitol. 41, 177–185.

Hamidinejat, H., Shapouri, M.S., Mayahi, M., Borujeni, M.P., 2010. Characterization ofEimeria species in commercial broilers by PCR based on ITS1 regions of rDNA. IranJ. Parasitol. 5, 48–54.

Han, Q., Li, J., Gong, P., Gai, J., Li, S., Zhang, X., 2011. Virus-like particles in Eimeria tenellaare associated with multiple RNA segments. Exp. Parasitol. 127, 646–650.

Hanig, S., Entzeroth, R., Kurth, M., 2012. Chimeric fluorescent reporter as a tool for gen-eration of transgenic Eimeria (Apicomplexa, Coccidia) strains with stage specific reportergene expression. Parasitol. Int. 61, 391–398.

Hao, L., Liu, X., Zhou, X., Li, J., Suo, X., 2007. Transient transfection ofEimeria tenella usingyellow or red fluorescent protein as a marker. Mol. Biochem. Parasitol. 153, 213–215.

Haug, A., Thebo, P., Mattsson, J.G., 2007. A simplified protocol for molecular identificationof Eimeria species in field samples. Vet. Parasitol. 146, 35–45.

Haug, A., Gjevre, A.G., Thebo, P., Mattsson, J.G., Kaldhusdal, M., 2008. Coccidial infec-tions in commercial broilers: epidemiological aspects and comparison of Eimeria speciesidentification bymorphometric and polymerase chain reaction techniques. Avian Pathol.37, 161–170.

Hong, Y.H., Lillehoj, H.S., Lee, S.H., Dalloul, R.A., Lillehoj, E.P., 2006a. Analysis ofchicken cytokine and chemokine gene expression following Eimeria acervulina andEimeria tenella infections. Vet. Immunol. Immunopathol. 114, 209–223.

Hong, Y.H., Lillehoj, H.S., Lillehoj, E.P., Lee, S.H., 2006b. Changes in immune-relatedgene expression and intestinal lymphocyte subpopulations following Eimeria maximainfections of chickens. Vet. Immunol. Immunopathol. 114, 259–272.

Huang, X., Zou, J., Xu, H., Ding, Y., Yin, G., Liu, X., Suo, X., 2011. Transgenic Eimeriatenella expressing enhanced yellow fluorescent protein targeted to different cellular com-partments stimulated dichotomic immune responses in chickens. J. Immunol. 187,3595–3602.

Huynh, M.H., Carruthers, V.B., 2006. ToxoplasmaMIC2 is a major determinant of invasionand virulence. PLoS Pathog. 2 (8), e84.

Huynh, M.H., Opitz, C., Kwok, L.Y., Tomley, F.M., Carruthers, V.B., Soldati, D., 2004.Trans-genera reconstitution and complementation of an adhesion complex in Toxo-plasma gondii. Cell. Microbiol. 6, 771–782.

Jeffers, T.K., 1974. Genetic transfer of anticoccidial drug resistance in Eimeria tenella.J. Parasitol. 60, 900–904.

Jeffers, T.K., 1976. Genetic recombination of precociousness and anticoccidial drug resis-tance in Eimeria tenella. Z. Parasitenkd. 50, 251–255.

Jenkins, M.C., 1988. A cDNA encoding a merozoite surface protein of the protozoan Eimeriaacervulina contains tandem-repeated sequences. Nucleic Acids Res. 16, 9863.

Jenkins, M.C., Miska, K., Klopp, S., 2006a. Application of polymerase chain reaction basedon ITS1 rDNA to speciate Eimeria. Avian Dis. 50, 110–114.






















































Jenkins, M.C., Miska, K., Klopp, S., 2006b. Improved polymerase chain reaction techniquefor determining the species composition of Eimeria in poultry litter. Avian Dis. 50,632–635.

Jenkins, M., Klopp, S., Ritter, D., Miska, K., Fetterer, R., 2010. Comparison of Eimeria spe-cies distribution and salinomycin resistance in commercial broiler operations utilizingdifferent coccidiosis control strategies. Avian Dis. 54, 1002–1006.

Jewett, T.J., Sibley, L.D., 2003. Aldolase forms a bridge between cell surface adhesins and theactin cytoskeleton in apicomplexan parasites. Mol. Cell 11, 885–894.

Jiang, L., Lin, J., Han, H., Dong, H., Zhao, Q., Zhu, S., Huang, B., 2012. Identification andcharacterization of Eimeria tenella apical membrane antigen-1 (AMA1). PLoS One 7 (7),e41115.

Jirku, M., Modry, D., Slapeta, J.R., Koudela, B., Lukes, J., 2002. The phylogeny of GoussiaandCholeoeimeria (Apicomplexa; Eimeriorina) and the evolution of excystation structuresin coccidia. Protist 153, 379–390.

Jirku, M., Jirku, M., Obornık, M., Lukes, J., Modry, D., 2009. Goussia Labbe, 1896(Apicomplexa, Eimeriorina) in Amphibia: diversity, biology, molecular phylogenyand comments on the status of the genus. Protist 160, 123–136.

Joyner, L.P., Norton, C.C., 1975. Transferred drug-resistance in Eimeria maxima. Parasitol-ogy 71, 385–392.

Joyner, L.P., Norton, C.C., 1978. The activity of methyl benzoquate and clopidol againstEimeria maxima: synergy and drug resistance. Parasitology 76, 369–377.

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 expressionof protease genes in the apicomplexan parasite, Eimeria tenella. BMC Genomics 13, 685.http://dx.doi.org/10.1186/1471-2164-13-685.

Katzer, F., Lizundia, R., Ngugi, D., Blake, D., McKeever, D., 2011. Construction of agenetic map for Theileria parva: identification of hotspots of recombination. Int. J. Para-sitol. 41, 669–675.

Kawahara, F., Taira, K., Nagai, S., Onaga, H., Onuma, M., Nunoya, T., 2008. Detection offive avian Eimeria species by species-specific real-time polymerase chain reaction assay.Avian Dis. 52, 652–656.

Kelleher, M., Tomley, F.M., 1998. Transient expression of beta-galactosidase in differenti-ating sporozoites of Eimeria tenella. Mol. Biochem. Parasitol. 97, 21–31.

Kessler, H., Herm-Gotz, A., Hegge, S., Rauch, M., Soldati-Favre, D., Frischknecht, F.,Meissner, M., 2008. Microneme protein 8—a new essential invasion factor in Toxo-plasma gondii. J. Cell Sci. 121, 947–956.

Khan, A., Taylor, S., Su, C., Mackey, A.J., Boyle, J., Cole, R., Glover, D., Tang, K.,Paulsen, I.T., Berriman, M., Boothroyd, J.C., Pfefferkorn, E.R., Dubey, J.P.,Ajioka, J.W., Roos, D.S., Wootton, J.C., Sibley, L.D., 2005. Composite genomemap and recombination parameters derived from three archetypal lineages of Toxoplasmagondii. Nucleic Acids Res. 33, 2980–2992.

Kim,C.H., Lillehoj, H.S., Bliss, T.W., Keeler Jr., C.L., Hong,Y.H., Park, D.W., Yamage,M.,Min, W., Lillehoj, E.P., 2008. Construction and application of an avian intestinal intra-epithelial lymphocyte cDNA microarray (AVIELA) for gene expression profiling duringEimeria maxima infection. Vet. Immunol. Immunopathol. 124, 341–354.

Kim, C.H., Lillehoj, H.S., Hong, Y.H., Keeler Jr., C.L., Lillehoj, E.P., 2010. Comparison ofglobal transcriptional responses to primary and secondary Eimeria acervulina infections inchickens. Dev. Comp. Immunol. 34, 344–351.

Kim, D.K., Lillehoj, H.S., Lee, S.H., Lillehoj, E.P., Bravo, D., 2012. Improved resistance toEimeria acervulina infection in chickens due to dietary supplementation with garlic metab-olites. Br. J. Nutr. 13, 1–13.






















http://dx.doi.org/10.1186/1471-2164-13-685





























Kirkpatrick, N.C., Blacker, H.P., Woods, W.G., Gasser, R.B., Noormohammadi, A.H.,2009. A polymerase chain reaction-coupled high-resolution melting curve analyticalapproach for the monitoring of monospecificity of avian Eimeria species. Avian Pathol.38, 13–19.

Klesius, P.H., Hinds, S.E., 1979. Strain-dependent differences in murine susceptibility tococcidia. Infect. Immun. 26, 1111–1115.

Klotz, C., Gehre, F., Lucius, R., Pogonka, T., 2007. Identification of Eimeria tenella genesencoding for secretory proteins and evaluation of candidates by DNA immunisationstudies in chickens. Vaccine 25, 6625–6634.

Krucken, J., Hosse, R.J., Mouafo, A.N., Entzeroth, R., Bierbaum, S., Marinovski, P.,Hain, K., Greif, G., Wunderlich, F., 2008. Excystation of Eimeria tenella sporozoitesimpaired by antibody recognizing gametocyte/oocyst antigens GAM22 and GAM56.Eukaryot. Cell 7, 202–211.

Kucera, J., Reznicky, M., 1991. Differentiation of species of Eimeria from the fowl using acomputerized image-analysis system. Folia Parasitol. (Praha) 38, 107–113.

Kurth, M., Entzeroth, R., 2008. Improved excystation protocol for Eimeria nieschulzi(Apikomplexa, Coccidia). Parasitol. Res. 102, 819–822.

Kurth, M., Entzeroth, R., 2009. Reporter gene expression in cell culture stages and oocystsof Eimeria nieschulzi (Coccidia, Apicomplexa). Parasitol. Res. 104, 303–310.

Labbe, M., de Venevelles, P., Girard-Misguich, F., Bourdieu, C., Guillaume, A., Pery, P.,2005. Eimeria tenella microneme protein EtMIC3: identification, localisation and role inhost cell infection. Mol. Biochem. Parasitol. 140, 43–53.

Lai, L., Bumstead, J., Liu, Y., Garnett, J., Campanero-Rhodes, M.A., Blake, D.P.,Palma, A.S., Chai, W., Ferguson, D.J., Simpson, P., Feizi, T., Tomley, F.M.,Matthews, S., 2011. The role of sialyl glycan recognition in host tissue tropism of theavian parasite Eimeria tenella. PLoS Pathog. 7 (10), e1002296.

Lal, K., Bromley, E., Oakes, R., Prieto, J.H., Sanderson, S.J., Kurian, D., Hunt, L.,Yates, J.R., Wastling, J.M., Sinden, R.E., Tomley, F.M., 2009. Proteomic comparisonof four Eimeria tenella life-cycle stages: unsporulated oocyst, sporulated oocyst, sporozoiteand second-generation merozoite. Proteomics 9, 4566–4576.

Lalonde, L.F., Gajadhar, A.A., 2011. Detection and differentiation of coccidian oocystsby real-time PCR and melting curve analysis. J. Parasitol. 97, 725–730.

Lebrun, M., Michelin, A., El Hajj, H., Poncet, J., Bradley, P.J., Vial, H., Dubremetz, J.F.,2005. The rhoptry neck protein RON4 re-localizes at the moving junction duringToxo-plasma gondii invasion. Cell. Microbiol. 7, 1823–1833.

Lee, S., Fernando, M.A., 2000. Viral double-stranded RNAs of Eimeria spp. of the domesticfowl: analysis of genetic relatedness and divergence among various strains. Parasitol. Res.86, 733–737.

Lee, S., Fernando, M.A., Nagy, E., 1996. dsRNA associated with virus-like particles inEimeria spp. of the domestic fowl. Parasitol. Res. 82, 518–523.

Lee, E.G., Kim, J.H., Shin, Y.S., Shin, G.W., Suh, M.D., Kim, D.Y., Kim, Y.H., Kim, G.S.,Jung, T.S., 2003. Establishment of a two-dimensional electrophoresis map for Neosporacaninum tachyzoites by proteomics. Proteomics 3, 2339–2350.

Lee, S.H., Lillehoj, H.S., Park, D.W., Jang, S.I., Morales, A., Garcia, D., Lucio, E.,Larios, R., Victoria, G., Marrufo, D., Lillehoj, E.P., 2009a. Protective effect of hyper-immune egg yolk IgY antibodies against Eimeria tenella and Eimeria maxima infections.Vet. Parasitol. 163, 123–126.

Lee, S.H., Lillehoj, H.S., Park, D.W., Jang, S.I., Morales, A., Garcia, D., Lucio, E.,Larios, R., Victoria, G., Marrufo, D., Lillehoj, E.P., 2009b. Induction of passive immu-nity in broiler chickens against Eimeria acervulina by hyperimmune egg yolk immuno-globulin Y. Poult. Sci. 88, 562–566.






















































Lee, K.W., Li, G., Lillehoj, H.S., Lee, S.H., Jang, S.I., Babu, U.S., Lillehoj, E.P.,Neumann, A.P., Siragusa, G.R., 2011. Bacillus subtilis-based direct-fed microbials aug-ment macrophage function in broiler chickens. Res. Vet. Sci. 91, 87–91.

Lew, A.E., Anderson, G.R., Minchin, C.M., Jeston, P.J., Jorgensen, W.K., 2003. Inter- andintra-strain variation and PCR detection of the internal transcribed spacer 1 (ITS-1)sequences of Australian isolates of Eimeria species from chickens. Vet. Parasitol. 112,33–50.

Lillehoj, H.S., Lee, K.W., 2012. Immune modulation of innate immunity as alternatives-to-antibiotics strategies to mitigate the use of drugs in poultry production. Poult. Sci. 91,1286–1291.

Lim, L.S., Tay, Y.L., Alias, H., Wan, K.L., Dear, P.H., 2012. Insights into the genome struc-ture and copy-number variation of Eimeria tenella. BMC Genomics 13, 389.

Lin, R.Q., Qiu, L.L., Liu, G.H., Wu, X.Y., Weng, Y.B., Xie, W.Q., Hou, J., Pan, H.,Yuan, Z.G., Zou, F.C., Hu,M., Zhu,X.Q., 2011. Characterization of the completemito-chondrial genomes of five Eimeria species from domestic chickens. Gene 480, 28–33.

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.P., Mohamed, R., Shirley, M.W., Gruber, A.,Berriman, M., Tomley, F.M., Dear, P.H., Wan, K.L., 2007. Sequencing and analysisof chromosome 1 of Eimeria tenella reveals a unique segmental organization. GenomeRes. 17, 311–319.

Liu, X., Shi, T., Ren, H., Su, H., Yan, W., Suo, X., 2008. Restriction enzyme-mediatedtransfection improved transfection efficiency in vitro in apicomplexan parasite Eimeriatenella. Mol. Biochem. Parasitol. 161, 72–75.

Liu, L., Xu, L., Yan, F., Yan, R., Song, X., Li, X., 2009. Immunoproteomic analysis of thesecond-generation merozoite proteins of Eimeria tenella. Vet. Parasitol. 164, 173–182.

Liu, G.H., Hou, J., Weng, Y.B., Song, H.Q., Li, S., Yuan, Z.G., Lin, R.Q., Zhu, X.Q.,2012. The complete mitochondrial genome sequence of Eimeria mitis (Apicomplexa:Coccidia). Mitochondrial DNA 23 (5), 341–343.

Long, P.L., Joyner, L.P., 1984. Problems in the identification of species of Eimeria.J. Protozool. 31, 535–541.

Long, P.L., Pierce, A.E., 1963. Role of cellular factors in the mediation of immunity to aviancoccidiosis (Eimeria tenella). Nature 200, 426–427.

Long, P.L., Rose, M.E., 1970. Extended schizogony of Eimeria mivati in betamethasone-treated chickens. Parasitology 60, 147–155.

Lourido, S., Shuman, J., Zhang, C., Shokat, K.M., Hui, R., Sibley, L.D., 2010. Calcium-dependent protein kinase 1 is an essential regulator of exocytosis in Toxoplasma. Nature465, 359–362.

MacPherson, J.M., Gajadhar, A.A., 1993. Differentiation of seven Eimeria species by randomamplified polymorphic DNA. Vet. Parasitol. 45, 257–266.

Mai, K., Sharman, P.A., Walker, R.A., Katrib, M., DeSouza, D., McConville, M.J.,Wallach, M.G., Belli, S.I., Ferguson, D.J., Smith, N.C., 2009. Oocyst wall formationand composition in coccidian parasites. Mem. Inst. Oswaldo Cruz 104, 281–289.

Mai, K., Smith, N.C., Feng, Z.P., Katrib, M., Slapeta, J., Slapetova, I., Wallach, M.G.,Luxford,C.,Davies,M.J.,Zhang,X.,Norton,R.S.,Belli, S.I., 2011.Peroxidase catalyzedcross-linkingof an intrinsically unstructuredprotein via dityrosine bonds in theoocystwallof the apicomplexan parasite, Eimeria maxima. Int. J. Parasitol. 41, 1157–1164.

Manly, K., Cudmore,R., 1997.MapManagerQT, software formapping quantitative trait loci.In: Abstracts of the 11th International Mouse Genome Conference, St. Petersburg, FL.






















































Marchant, J., Cowper, B., Liu, Y., Lai, L., Pinzan, C., Marq, J.B., Friedrich, N.,Sawmynaden, K., Liew, L., Chai, W., Childs, R.A., Saouros, S., Simpson, P.,Roque Barreira, M.C., Feizi, T., Soldati-Favre, D., Matthews, S., 2012. Galactoserecognition by the apicomplexan parasite Toxoplasma gondii. J. Biol. Chem. 287,16720–16733.

McCutchan, T.F., de la Cruz, V.F., Lal, A.A., Gunderson, J.H., Elwood, H.J., Sogin, M.L.,1988. Primary sequences of two small subunit ribosomal RNA genes from Plasmodiumfalciparum. Mol. Biochem. Parasitol. 28, 63–68.

McDonald, V., Shirley, M.W., 2009. Past and future: vaccination against Eimeria. Parasitol-ogy 136, 1477–1489.

Merino, S., Martınez, J., Martınez-de la Puente, J., Criado-Fornelio, A., Tomas, G.,Morales, J., Lobato, E., Garcıa-Fraile, S., 2006. Molecular characterization of the 18srDNA gene of an avian Hepatozoon reveals that it is closely related to Lankesterella.J. Parasitol. 92, 1330–1335.

Mesfin, G.M., Bellamy, J.E.C., 1979. Thymic dependence of immunity to Eimeria falciformisvar. pragensis in mice. Infect. Immun. 23, 460–464.

Miska, K.B., Fetterer, R.H., Rosenberg, G.H., 2008. Analysis of transcripts from intracel-lular stages of Eimeria acervulina using expressed sequence tags. J. Parasitol. 94, 462–466.

Miska, K.B., Schwarz, R.S., Jenkins, M.C., Rathinam, T., Chapman, H.D., 2010.Molecular characterization and phylogenetic analysis of Eimeria from turkeys andgamebirds: implications for evolutionary relationships in Galliform birds. J. Parasitol.96, 982–986.

Monne, L., Honig, G., 1954. On the properties of the shells of the coccidian oocysts.Ark. Zool. 7, 251–256.

Morgan, J.A., Morris, G.M., Wlodek, B.M., Byrnes, R., Jenner, M., Constantinoiu, C.C.,Anderson, G.R., Lew-Tabor, A.E., Molloy, J.B., Gasser, R.B., Jorgensen, W.K., 2009.Real-time polymerase chain reaction (PCR) assays for the specific detection and quan-tification of sevenEimeria species that cause coccidiosis in chickens. Mol. Cell. Probes 23,83–89.

Morris, G.M., Gasser, R.B., 2006. Biotechnological advances in the diagnosis of aviancoccidiosis and the analysis of genetic variation in Eimeria. Biotechnol. Adv. 24,590–603.

Morris, G.M., Woods, W.G., Grant Richards, D., Gasser, R.B., 2007a. The application of apolymerase chain reaction (PCR)-based capillary electrophoretic technique providesdetailed insights into Eimeria populations in intensive poultry establishments. Mol. Cell.Probes 21, 288–294.

Morris, G.M.,Woods,W.G., Richards, D.G., Gasser, R.B., 2007b. Investigating a persistentcoccidiosis problem on a commercial broiler-breeder farm utilizing PCR-coupled cap-illary electrophoresis. Parasitol. Res. 101, 583–589.

Nishimoto, Y., Arisue, N., Kawai, S., Escalante, A.A., Horii, T., Tanabe, K., Hashimoto, T.,2008. Evolution and phylogeny of the heterogeneous cytosolic SSU rRNA genes in thegenus Plasmodium. Mol. Phylogenet. Evol. 47, 45–53.

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 analysisreveals expression profiles conserved across three Eimeria spp. of domestic fowl and asso-ciated with multiple developmental stages. Int. J. Parasitol. 42, 39–48.

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 ofEimeria tenella sporozoites. Int. J. Parasitol. 43, 181–188.

Ogedengbe, J.D., Hanner, R.H., Barta, J.R., 2011a. DNA barcoding identifies Eimeria spe-cies and contributes to the phylogenetics of coccidian parasites (Eimeriorina,Apicomplexa, Alveolata). Int. J. Parasitol. 41, 843–850.























































Ogedengbe, J.D., Hunter, D.B., Barta, J.R., 2011b. Molecular identification of Eimeria spe-cies infecting market-age meat chickens in commercial flocks in Ontario. Vet. Parasitol.178, 350–354.

Oliveira, U.C., Fraga, J.S., Licois, D., Pakandl, M., Gruber, A., 2011. Development ofmolecular assays for the identification of the 11 Eimeria species of the domestic rabbit(Oryctolagus cuniculus). Vet. Parasitol. 176, 275–280.

Ovington, K.S., Smith, N.C., 1992. Cytokines, free radicals and resistance to Eimeria. Para-sitol. Today 8, 422–426.

Ovington, K.S., Smith, N.C., Joysey, H.S., 1990. Oxygen derived free radicals and thecourse of Eimeria vermiformis infection in inbred strains of mice. Parasite Immunol. 12,623–631.

Ovington, K.S., Alleva, L.M., Kerr, E.A., 1995. Cytokines and immunological control ofEimeria spp. Int. J. Parasitol. 25, 1331–1351.

Page, A.P.,Winter, A.D., 2003. Enzymes involved in the biogenesis of the nematode cuticle.Adv. Parasitol. 53, 85–148.

Peek, H.W., Landman,W.J., 2006. Higher incidence ofEimeria spp. field isolates sensitive fordiclazuril and monensin associated with the use of live coccidiosis vaccination withparacox-5 in broiler farms. Avian Dis. 50, 434–439.

Periz, J., Gill, A.C., Knott, V., Handford, P.A., Tomley, F.M., 2005. Calcium binding activ-ity of the epidermal growth factor-like domains of the apicomplexan microneme proteinEtMIC4. Mol. Biochem. Parasitol. 143, 192–199.

Periz, J., Gill, A.C., Hunt, L., Brown, P., Tomley, F.M., 2007. The microneme proteinsEtMIC4 and EtMIC5 of Eimeria tenella form a novel, ultra-high molecular mass proteincomplex that binds target host cells. J. Biol. Chem. 282, 16891–16898.

Pierce, A.E., Long, P.L., Horton-Smith, C., 1962. Immunity to Eimeria tenella in youngfowls (Gallus domesticus). Immunology 5, 129–152.

Pittilo, R.M., Ball, S.J., 1980. The ultrastructural development of the oocyst wall of Eimeriamaxima. Parasitology 81, 115–122.

Pogonka, T., Schelzke, K., Stange, J., Papadakis, K., Steinfelder, S., Liesenfeld, O.,Lucius, R., 2010. CD8þ cells protect mice against reinfection with the intestinal parasiteEimeria falciformis. Microbes Infect. 12, 218–226.

Poplstein, M., Vrba, V., 2011. Description of the two strains of turkey coccidia Eimeriaadenoeides with remarkable morphological variability. Parasitology 138, 1211–1216.

Possenti, A., Cherchi, S., Bertuccini, L., Pozio, E., Dubey, J.P., Spano, F., 2010. Molecularcharacterization of a novel family of cysteine-rich proteins of Toxoplasma gondii and ultra-structural evidence of oocyst wall localisation. Int. J. Parasitol. 40, 1639–1649.

Procunier, J.D., Fernando, M.A., Barta, J.R., 1993. Species and strain differentiation ofEimeria spp. of the domestic fowl using DNA polymorphisms amplified by arbitraryprimers. Parasitol. Res. 79, 98–102.

Rangel, L.T., Novaes, J., Durham, A.M., Madeira, A.M.B.N., Gruber, A., 2013. TheEimeria Transcript DB: an integrated resource for annotated transcripts of protozoan par-asites of the genus Eimeria. Database 2013, http://dx.doi.org/10.1093/database/bat006,Article ID bat006.

Rathinam, T., Chapman, H.D., 2009. Sensitivity of isolates of Eimeria from turkey flocks tothe anticoccidial drugs amprolium, clopidol, diclazuril, and monensin. Avian Dis. 53,405–408.

Rausch, S., Held, J., Stange, J., Lendner, M., Hepworth, M.R., Klotz, C., Lucius, R.,Pogonka, T., Hartmann, S., 2010. A matter of timing: early, not chronic phase intestinalnematode infection restrains control of a concurrent enteric protozoan infection. Eur. J.Immunol. 40, 2804–2815.

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.,










































http://dx.doi.org/10.1093/database/bat006












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 andNeospora can-inum: Coccidia differing in host range and transmission strategy. PLoS Pathog. 8 (3),e1002567.

Roberts, S.J., Smith, A.L., West, A.B., Wen, L., Findly, R.C., Owen, M.J., Hayday, A.C.,1996. T-cell alpha betaþ and gamma deltaþ deficient mice display abnormal but distinctphenotypes toward a natural, widespread infection of the intestinal epithelium. Proc.Natl. Acad. Sci. USA 93, 11774–11779.

Rollinson, D., 1975. Electrophoretic variation of enzymes in chicken coccidiosis. Trans. R.Soc. Trop. Med. Hyg. 72, 436–437.

Rose, M.E., 1963. Some aspects of immunity to Eimeria infections. Ann. NY Acad. Sci. 113,383–399.

Rose, M.E., 1967a. Immunity to Eimeria brunetti and Eimeria maxima infections in the fowl.Parasitology 57, 363–370.

Rose, M.E., 1967b. Immunity to Eimeria tenella and Eimeria necatrix in the fowl. I. Influenceof the site of infection and the stage of parasite. II. Cross-protection. Parasitology 57,567–583.

Rose, M.E., 1970. Immunity to coccidiosis: effect of betamethasone treatment of fowls onEimeria mivati infection. Parasitology 60, 137–146.

Rose, M.E., 1972. Immunity to coccidiosis: maternal transfer in Eimeria maxima infections.Parasitology 65, 273–282.

Rose, M.E., 1974. Protective antibodies in infections with Eimeria maxima: the reduction ofpathogenic effects in vivo and a comparison between oral and subcutaneous administra-tion of antiserum. Parasitology 68, 285–292.

Rose, M.E., Hesketh, P., 1979. Immunity to coccidiosis: T-lymphocyte- or B-lymphocyte-deficient animals. Infect. Immun. 26, 630–637.

Rose, M.E., Hesketh, P., 1982. Coccidiosis: T-lymphocyte-dependent effects of infectionwith Eimeria nieschulzi in rats. Vet. Immunol. Immunopathol. 3, 499–508.

Rose,M.E., Hesketh, P., 1986. Eimerian life cycles: the patency of Eimeria vermiformis but notEimeria pragensis is subject to host (Mus musculus) influence. J. Parasitol. 72, 949–954.

Rose, M.E., Long, P.L., 1962. Immunity to four species of Eimeria in fowls. Immunology 5,79–92.

Rose, M.E., Long, P.L., 1970. Resistance to Eimeria infections in the chicken: the effects ofthymectomy, bursectomy, whole body irradiation and cortisone treatment. Parasitology60, 291–299.

Rose, M.E., Long, P.L., 1971. Immunity to coccidiosis: protective effects of transferredserum and cells investigated in chick embryos infected with Eimeria tenella. Parasitology63, 299–313.

Rose, M.E., Millard, B.J., 1985. Eimeria vermiformis: host strains and the developmental cycle.Exp. Parasitol. 60, 285–293.

Rose, M.E., Hesketh, P., Ogilvie, B.M., 1979a. Peripheral blood leucocyte responses tococcidial infection: a comparison of the response in rats and chickens and its correlationwith resistance to reinfection. Immunology 36, 71–79.

Rose, M.E., Ogilvie, B.M., Hesketh, P., Festing, M.F., 1979b. Failure of nude (athymic) ratsto become resistant to reinfection with the intestinal coccidian parasite Eimeria nieschulzior the nematode Nippostronglus brasiliensis. Parasite Immunol. 1, 125–132.

Rose, M.E., Owen, D.G., Hesketh, P., 1984. Susceptibility to coccidiosis: effect of strain ofmouse on reproduction of Eimeria vermiformis. Parasitology 88, 45–54.

Rose, M.E., Wakelin, D., Hesketh, P., 1985. Susceptibility to coccidiosis: contrasting courseof primary infections with Eimeria vermiformis in BALB/c and C57/BL/6mice is based onimmune responses. Parasite Immunol. 7, 557–566.



























































Rose,M.E., Joysey,H.S., Hesketh, P., Grencis, R.K.,Wakelin, D., 1988.Mediation of immu-nity to Eimeria vermiformis in mice by L3T4þ T cells. Infect. Immun. 56, 1760–1765.

Rose, M.E., Wakelin, D., Joysey, H.S., Hesketh, P., 1989. Immunity to coccidiosis: T-cellcontrol of infection with Eimeria vermiformis in mice does not require co-operation withinflammatory cells. Parasite Immunol. 11, 231–239.

Rose, M.E., Wakelin, D., Hesketh, P., 1990. Eimeria vermiformis: differences in the course ofprimary infection can be correlated with lymphocyte responsiveness in the BALB/c andC57BL/6 mouse, Mus musculus. Exp. Parasitol. 71, 276–283.

Rose, M.E., Smith, A.L., Wakelin, D., 1991a. Gamma interferon-mediated inhibition ofEimeria vermiformis growth in cultured fibroblasts and epithelial cells. Infect. Immun.59, 580–586.

Rose, M.E., Wakelin, D., Hesketh, P., 1991b. Interferon-gamma-mediated effects uponimmunity to coccidial infections in the mouse. Parasite Immunol. 13, 63–74.

Rose, M.E., Hesketh, P., Wakelin, D., 1992. Immune control of murine coccidiosis: CD4þand CD8þ T lymphocytes contribute differentially in resistance to primary and second-ary infections. Parasitology 105, 349–354.

Rose, M.E., Hesketh, P., Wakelin, D., 1995. Cytotoxic effects of natural killer cells have nosignificant role in controlling infection with the intracellular protozoon Eimeria ver-miformis. Infect. Immun. 63, 3711–3714.

Rose, M.E., Hesketh, P., Rothwell, L., Gramzinski, R.A., 1996. T-cell receptor gamma-delta lymphocytes and Eimeria vermiformis infection. Infect. Immun. 64, 4854–4858.

Rothwell, L., Gramzinski, R.A., Rose, M.E., Kaiser, P., 1995. Avian coccidiosis: changes inintestinal lymphocyte populations associated with the development of immunity toEimeria maxima. Parasite Immunol. 17, 525–533.

Rothwell, L., Muir, W., Kaiser, P., 2000. Interferon-gamma is expressed in both gut andspleen during Eimeria tenella infection. Avian Pathol. 29, 333–342.

Rothwell, L., Young, J.R., Zoorob, R., Whittaker, C.A., Hesketh, P., Archer, A.,Smith, A.L., Kaiser, P., 2004. Cloning and characterization of chicken IL-10 and its rolein the immune response to Eimeria maxima. J. Immunol. 173, 2675–2682.

Santos, J.M., Soldati-Favre, D., 2011. Invasion factors are coupled to key signalling eventsleading to the establishment of infection in apicomplexan parasites. Cell. Microbiol.13, 787–796.

Schito, M.L., Barta, J.R., 1997. Nonspecific immune responses and mechanisms of resistanceto Eimeria papillata infections in mice. Infect. Immun. 65, 3165–3170.

Schito, M.L., Barta, J.R., Chobotar, B., 1996. Comparison of four murine Eimeria species inimmunocompetent and immunodeficient mice. J. Parasitol. 82, 255–262.

Schito, M.L., Chobotar, B., Barta, J.R., 1998. Major histocompatibility complex class I- andII-deficient knock-out mice are resistant to primary but susceptible to secondary Eimeriapapillata infections. Parasitol. Res. 84, 394–398.

Schmatz, D.M., Baginsky, W.F., Turner, M.J., 1989. Evidence for and characterization of amannitol cycle in Eimeria tenella. Mol. Biochem. Parasitol. 32, 263–270.

Schmid, M., Lehmann, M.J., Lucius, R., Gupta, N., 2012. Apicomplexan parasite, Eimeriafalciformis, co-opts host tryptophan catabolism for life cycle progression in mouse. J. Biol.Chem. 287, 20197–20207.

Schnitzler, B.E., Thebo, P.L., Mattsson, J.G., Tomley, F.M., Shirley, M.W., 1998. Devel-opment of a diagnostic PCR assay for the detection and discrimination of four patho-genic Eimeria species of the chicken. Avian Pathol. 27, 490–497.

Schnitzler, B.E., Thebo, P.L., Tomley, F.M., Uggla, A., Shirley, M.W., 1999. PCR iden-tification of chicken Eimeria: a simplified read-out. Avian Pathol. 28, 89–93.

Schubert, U., Fuchs, J., Zimmermann, J., Jahn, D., Zoufal, K., 2005. Extracellular calciumdeficiency and ryanodine inhibit Eimeria tenella sporozoite invasion in vitro. Parasitol.Res. 97, 59–62.




























































Schwarz, R.S., Jenkins, M.C., Klopp, S., Miska, K.B., 2009. Genomic analysis of Eimeria spp.populations in relation to performance levels of broiler chicken farms in Arkansas andNorth Carolina. J. Parasitol. 95, 871–880.

Schwarz, R.S., Fetterer, R.H., Rosenberg, G.H., Miska, K.B., 2010. Coccidian merozoitetranscriptome analysis from Eimeria maxima in comparison to Eimeria tenella and Eimeriaacervulina. J. Parasitol. 96, 49–57.

Sharman, P.A., Smith, N.C.,Wallach, M.G., Katrib, M., 2010. Chasing the golden egg: vac-cination against poultry coccidiosis. Parasite Immunol. 32, 590–598.

Sheriff, R., Carroll, F., Shirley, M.W., 2003. Molecular karyotypes of Eimeria tenella resolvedby PFGE: an evaluation of different agaroses. Parasitol. Res. 89, 317–319.

Shi, T.Y., Liu, X.Y., Hao, L.L., Li, J.D., Gh, A.N., Abdille, M.H., Suo, X., 2008. Trans-fected Eimeria tenella could complete its endogenous development in vitro. J. Parasitol.94, 978–980.

Shi, T., Yan, W., Ren, H., Liu, X., Suo, X., 2009. Dynamic development of parasit-ophorous vacuole of Eimeria tenella transfected with the yellow fluorescent protein genefused to different signal sequences from apicomplexan parasites. Parasitol. Res. 104,315–320.

Shirley, M.W., 1975. Enzyme variation in Eimeria species of the chicken. Parasitology 71,369–376.

Shirley, M.W., 1994a. The genome of Eimeria tenella: further studies on its molecular orga-nisation. Parasitol. Res. 80, 366–373.

Shirley, M.W., 1994b. Coccidial parasites from the chicken: discrimination of differentpopulations of Eimeria tenella by DNA hybridisation. Res. Vet. Sci. 57, 10–14.

Shirley, M.W., 2000. The genome of Eimeria spp., with special reference to Eimeria tenella—acoccidium from the chicken. Int. J. Parasitol. 30, 485–493.

Shirley,M.W., Bumstead, N., 1994. Intra-specific variation withinEimeria tenella detected bythe random amplification of polymorphic DNA. Parasitol. Res. 80, 346–351.

Shirley, M.W., Harvey, D.A., 1996. Eimeria tenella: infection with a single sporocyst gives aclonal population. Parasitology 112, 523–528.

Shirley, M.W., Harvey, D., 2000. A genetic linkage map of the apicomplexan protozoanparasite Eimeria tenella. Genome Res. 10, 1587–1593.

Shirley, M.W., Lillehoj, H.S., 2012. The long view: a selective review of 40 years of coc-cidiosis research. Avian Pathol. 41, 111–121.

Shirley, M.W., Rollinson, D., 1979. Coccidia: the recognition and characterization ofpopulations of Eimeria. In: Problems in the Identification of Parasites and their Vectors,Symposium of the British Society for Parasitology, UK. vol. 17. pp. 7–30.

Shirley, M.W., Chapman, H.D., Kucera, J., Jeffers, T.K., Bedrnik, P., 1989. Enzyme var-iation and pathogenicity of recent field isolates ofEimeria tenella. Res. Vet. Sci. 46, 79–83.

Shirley, M.W., Smith, A.L., Tomley, F.M., 2005. The biology of avian Eimeria with anemphasis on their control by vaccination. Adv. Parasitol. 60, 285–330.

Sibley, L.D., LeBlanc, A.J., Pfefferkorn, E.R., Boothroyd, J.C., 1992. Generation of arestriction fragment length polymorphism linkage map for Toxoplasma gondii. Genetics132, 1003–1015.

Smith, A.L., Hayday, A.C., 1998. Genetic analysis of the essential components of theimmunoprotective response to infection with Eimeria vermiformis. Int. J. Parasitol. 28,1061–1069.

Smith, A.L., Hayday, A.C., 2000. Genetic dissection of primary and secondary responses to awidespread natural pathogen of the gut, Eimeria vermiformis. Infect. Immun. 68,6273–6280.

Smith, N.C., Ovington, K.S., 1996. The effect of BCG, zymosan andCoxiella burnetti extracton Eimeria infections. Immunol. Cell Biol. 74, 346–348.






















































Smith, C.K.I.I., Strout, R.G., 1979. Eimeria tenella: accumulation and retention of anti-coccidial ionophores by extracellular sporozoites. Exp. Parasitol. 48, 325–330.

Smith, A.L., Rose, M.E., Wakelin, D., 1994a. The role of natural killer cells in resistance tococcidiosis: investigations in a murine model. Clin. Exp. Immunol. 97, 273–279.

Smith, N.C., Hunt, M., Ellenreider, C., Eckert, J., Shirley, M.W., 1994b. Detection of met-abolic enzymes of Eimeria by ampholine-polyacrylamide gel isoelectric focusing. Para-sitol. Res. 80, 165–169.

Smith, N.C., Wallach, M., Miller, C.M.D., Morgenstern, R., Braun, R., Eckert, J., 1994c.Maternal transfer of immunity to Eimeria maxima: enzyme-linked immunosorbent assayanalysis of protective antibodies induced by infection. Infect. Immun. 62, 1348–1357.

Smith, N.C., Wallach, M., Petracca, M., Braun, R., Eckert, J., 1994d. Maternal transfer ofantibodies induced by infection with Eimeria maxima partially protects chickens againstchallenge with Eimeria tenella. Parasitology 109, 551–557.

Smith, A.L., Hesketh, P., Archer, A., Shirley, M.W., 2002. Antigenic diversity in Eimeriamaxima and the influence of host genetics and immunization schedule on cross-protective immunity. Infect. Immun. 70, 2472–2479.

Spano, F., Puri, C., Ranucci, L., Putignani, L., Crisanti, A., 1997. Cloning of the entireCOWP gene of Cryptosporidium parvum and ultrastructural localization of the proteinduring sexual parasite development. Parasitology 114, 427–437.

Stange, J., Hepworth, M.R., Rausch, S., Zajic, L., Kuhl, A.A., Uyttenhove, C.,Renauld, J.C., Hartmann, S., Lucius, R., 2012. IL-22 mediates host defense againstan intestinal intracellular parasite in the absence of IFN-g at the cost of Th17-drivenimmunopathology. J. Immunol. 188, 2410–2418.

Stockdale, P.G.H., Stockdale, M.H., Rickard, M.D.,Mitchell, G.F., 1985.Mouse strain var-iation and effects of oocyst dose in infection of mice with Eimeria falciformis, a coccidianparasite of the large intestine. Int. J. Parasitol. 15, 447–452.

Stucki, U., Braun, R., Roditi, I., 1993. Eimeria tenella: characterization of a 5S ribosomalRNA repeat unit and its use as a species-specific probe. Exp. Parasitol. 76, 68–75.

Sturtevant, A., 1913. The linear arrangement of six sex-linked factors in Drosophila, as shownby their mode of association. J. Exp. Zool. 14, 43–59.

Su, X., Ferdig, M.T., Huang, Y., Huynh, C.Q., Liu, A., You, J., Wooton, J.C.,Wellems, T.E., 1999. A genetic map and recombination parameters of the humanmalaria parasite Plasmodium falciparum. Science 286, 1351–1353.

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 (6), e40075.

Sutton, C.A., Shirley, M.W., Wisher, M.H., 1989. Characterization of coccidial proteins bytwo-dimensional sodium dodecyl sulphate-polyacrylamide gel electrophoresis. Parasitol-ogy 99, 175–187.

Tabares, E., Ferguson, D., Clark, J., Soon, P.E., Wan, K.L., Tomley, F., 2004. Eimeria tenellasporozoites and merozoites differentially express glycosylphosphatidylinositol-anchoredvariant surface proteins. Mol. Biochem. Parasitol. 135, 123–132.

Tanriverdi, S., Blain, J.C., Deng, B., Ferdig, M.T.,Widmer, G., 2007. Genetic crosses in theapicomplexan parasite Cryptosporidium parvum define recombination parameters. Mol.Microbiol. 63, 1432–1439.

Templeton, T.J., Lancto, C.A., Vigdorovich, V., Liu, C., London, N.R., Hadsall, K.Z.,Abrahamsen, M.S., 2004. TheCryptosporidium oocyst wall protein is a member of a mul-tigene family and has a homolog in Toxoplasma. Infect. Immun. 72, 980–987.

Thacker, C., Sheps, J.A., Rose, A.M., 2006. Caenorhabditis elegans dpy-5 is a cuticleprocollagen processed by a proprotein convertase. Cell. Mol. Life Sci. 63, 1193–1204.

Tomley, F.M., 1994. Antigenic diversity of the asexual developmental stages of Eimeriatenella. Parasite Immunol. 16, 407–413.
























































Trout, J.M., Lillehoj, H.S., 1996. T lymphocyte roles during Eimeria acervulina and Eimeriatenella infections. Vet. Immunol. Immunopathol. 53, 163–172.

Tyler, J.S., Boothroyd, J.C., 2011. The C-terminus of Toxoplasma RON2 provides the cru-cial link between AMA1 and the host-associated invasion complex. PLoS Pathog. 7 (2),e1001282.

Upton, S.J., 2000. Suborder Eimeriorina Leger, 1911. In: Lee, J.J., Leedale, G.F.,Bradbury, P. (Eds.), An Illustrated Guide to the Protozoa, second ed. Allen Press,Lawrence, KS, pp. 318–339.

Velkers, F.C., Blake, D.P., Graat, E.A., Vernooij, J.C., Bouma, A., de Jong, M.C.,Stegeman, J.A., 2010. Quantification of Eimeria acervulina in faeces of broilers: compar-ison of McMaster oocyst counts from 24 h faecal collections and single droppings to real-time PCR from cloacal swabs. Vet. Parasitol. 169, 1–7.

Vermeulen, A.N., Schapp, D.C., Schetters, T.P., 2001. Control of coccidiosis in chickens byvaccination. Vet. Parasitol. 100, 13–20.

Vrba, V., Blake, D.P., Poplstein, M., 2010. Quantitative real-time PCR assays for detectionand quantification of all seven Eimeria species that infect the chicken. Vet. Parasitol. 174,183–190.

Vrba, V., Poplstein, M., Pakandl, M., 2011. The discovery of the two types of small subunitribosomal RNA gene in Eimeria mitis contests the existence ofE. mivati as an independentspecies. Vet. Parasitol. 183, 47–53.

Wakelin, D., Rose, M.E., Hesketh, P., Else, K.J., Grencis, R.K., 1993. Immunity to coc-cidiosis: genetic influences on lymphocyte and cytokine responses to infection withEimeria vermiformis in inbred mice. Parasite Immunol. 15, 11–19.

Walker, R.A., 2009. The characterization of selected molecules expressed exclusively in thesexual stages of Eimeria tenella and Eimeria maxima. Ph.D. Dissertation. University ofTechnology, Sydney, P.O. Box 123, Broadway, NSW 2007, Australia.

Wallach, M.G., Mencher, D., Yarus, S., Pillemer, G., Halabi, A.Y., Pugatsch, T., 1989.Eimeria maxima: identification of gametocyte protein antigens. Exp. Parasitol. 68, 49–56.

Wallach, M., Smith, N.C., Miller, C.M.D., Eckert, J., Rose, M.E., 1994. Eimeria maxima:ELISA and Western blot analyses of protective sera. Parasite Immunol. 16, 377–383.

Wallach,M., Smith, N.C., Petracca,M.,Miller, C.M.D., Eckert, J., Braun, R., 1995. Eimeriamaxima gametocyte antigens: potential use in a subunit maternal vaccine against coccid-iosis in chickens. Vaccine 13, 347–354.

Wallach, M.G., Ashash, U., Michael, A., Smith, N.C., 2008. Field application of a subunitvaccine against an enteric protozoan disease. PLoS One 3 (12), e3948.

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.

Wang, Z., Shen, J., Suo, X., Zhao, S., Cao, X., 2006. Experimentally induced monensin-resistant Eimeria tenella and membrane fluidity of sporozoites. Vet. Parasitol. 138,186–193.

Wastling, J.M., Armstrong, S.D., Krishna, R., Xia, D., 2012. Parasites, proteomes and sys-tems: has Descartes’ clock run out of time? Parasitology 139, 1103–1118.

Weber, F.H., Genteman, K.C., LeMay, M.A., Lewis Sr., D.O., Evans, N.A., 2004. Immu-nization of broiler chicks by in ovo injection of infective stages of Eimeria. Poult. Sci. 83,392–399.

Wiersma, H.I., Galuska, S.E., Tomley, F.M., Sibley, L.D., Liberator, P.A., Donald, R.G.,2004. A role for coccidian cGMP-dependent protein kinase in motility and invasion. Int.J. Parasitol. 34, 369–380.

Williams, R.B., 2001. Quantification of the crowding effect during infections with the sevenEimeria species of the domesticated fowl; its importance for experimental designs and theproduction of oocyst stocks. Int. J. Parasitol. 31, 1056–1069.





















































Williams, R.B., 2002a. Fifty years of anticoccidial vaccines for poultry (1952–2002). AvianDis. 46, 775–802.

Williams, R.B., 2002b. Anticoccidial vaccines for broiler chickens: pathways to success.Avian Pathol. 31, 317–353.

Williams, R.B., 2006. Tracing the emergence of drug-resistance in coccidia (Eimeria spp.) ofcommercial broiler flocks medicated with decoquinate for the first time in the UnitedKingdom. Vet. Parasitol. 135, 1–14.

Williams, R.B., Thebo, P., Marshall, R.N., Marshall, J.A., 2010. Coccidian oocysts as type-specimens: long-term storage in aqueous potassium dichromate solution preserves DNA.Syst. Parasitol. 76, 69–76.

Xie, M., Gilbert, J.M., McDougald, L.R., 1992. Electrophoretic and immunologic charac-terization of proteins of merozoites of Eimeria acervulina, E. maxima, E. necatrix, andE. tenella. J. Parasitol. 78, 82–86.

Yan, W., Liu, X., Shi, T., Hao, L., Tomley, F.M., Suo, X., 2009. Stable transfection ofEimeria tenella: constitutive expression of the YFP–YFP molecule throughout the lifecycle. Int. J. Parasitol. 39, 109–117.

Yin, G., Liu, X., Zou, J., Huang, X., Suo, X., 2011. Co-expression of reporter genes in thewidespread pathogen Eimeria tenella using a double-cassette expression vector strategy.Int. J. Parasitol. 41, 813–816.

Yun, C.H., Lillehoj, H.S., Choi, K.D., 2000. Eimeria tenella infection induces local gammainterferon production and intestinal lymphocyte subpopulation changes. Infect. Immun.68, 1282–1288.

Zhao, X., Duszynski, D.W., Loker, E.S., 2001. A simple method of DNA extraction forEimeria species. J. Microbiol. Methods 44, 131–137.

Zhou, B., Wang, H., Xue, F., Wang, X., Fei, C., Wang, M., Zhang, T., Yao, X., He, P.,2010. Effects of diclazuril on apoptosis and mitochondrial transmembrane potential insecond-generation merozoites of Eimeria tenella. Vet. Parasitol. 168, 217–222.

Zou, J., Liu, X., Shi, T., Huang, X.,Wang, H., Hao, L., Yin, G., Suo, X., 2009. Transfectionof Eimeria and Toxoplasma using heterologous regulatory sequences. Int. J. Parasitol. 39,1189–1193.































2013 Chapman Et Al

Documents

limitation use

educational use

commercial use

university of guelph

university of arkansas

noncommercial research

elsevier ltdissn

china agricultural university