Toxocara canis (Ascaridida: Nematoda), which parasitizes (at the adult stage) the small intestine of canids, can be transmitted to a range of other mammals, including humans, and can cause the disease toxocariasis. Despite its significance as a pathogen, the genetics, epidemiology and biology of this parasite remain poorly understood. In addition, the zoonotic potential of related species of Toxocara, such as T. cati and T. malaysiensis, is not well known. Mitochondrial DNA is known to provide genetic markers for investigations in these areas, but complete mitochondrial genomic data have been lacking for T. canis and its congeners. In the present study, the mitochondrial genome of T. canis was amplified by long-range polymerase chain reaction (long PCR) and sequenced using a primer-walking strategy. This circular mitochondrial genome was 14162 bp and contained 12 protein-coding, 22 transfer RNA, and 2 ribosomal RNA genes consistent for secernentean nematodes, including Ascaris suum and Anisakis simplex (Ascaridida). The mitochondrial genome of T. canis provides genetic markers for studies into the systematics, population genetics and epidemiology of this zoonotic parasite and its congeners. Such markers can now be used in prospecting for cryptic species and for exploring host specificity and zoonotic potential, thus underpinning the prevention and control of toxocariasis in humans and other hosts.
Toxocara canis (Nematoda: Ascaridida) is the common roundworm of canids. This parasite is transmissible to humans as well as a range of other accidental or paratenic vertebrate hosts, in which (after the oral ingestion of infective eggs) the larvae of Toxocara canis invade the tissues and can cause different forms of clinical disease (toxocariasis). Although some aspects of the biology of T. canis are well understood, there are still significant gaps in our knowledge of areas including the molecular genetics, systematics, ecology and epidemiology of this and related parasitic nematodes. The present study elucidates the sequence, structure and organization of the mitochondrial genome of T. canis and provides mitochondrial gene markers for studies in these areas using molecular tools. A greater understanding of the epidemiology of species of Toxocara would improve the prevention and control of toxocariasis in humans and other animals.
Citation: Jex AR, Waeschenbach A, Littlewood DTJ, Hu M, Gasser RB (2008) The Mitochondrial Genome of Toxocara canis. PLoS Negl Trop Dis 2(8): e273. doi:10.1371/journal.pntd.0000273
Editor: Thomas Raymond Unnasch, University of South Florida, United States of America
Received: March 12, 2008; Accepted: July 10, 2008; Published: August 6, 2008
Copyright: © 2008 Jex et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This project was supported by the Australian Research Council. The funding body had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Roundworms (nematodes) belong to a large phylum (Nematoda) in the animal kingdom. This phylum contains a wide range of species with exceptionally varied life histories . Many nematodes are parasites of plants or animals , causing significant diseases and major socio-economic losses globally –. Central to the control of such parasites is knowledge of their population genetics, which has important implications for understanding many areas, including systematics, epidemiology and ecology –. The basis for investigating population structures is the accurate analysis of genetic variation, which is known to be widespread in many parasitic nematodes –,, utilizing molecular markers with sufficient levels of intraspecific sequence variability.
Mitochondrial DNA markers are particularly suited to population genetic and systematic investigations due to their high mutation rates and proposed maternal inheritance , , –. In spite of the availability of advanced DNA technologies, there is still a paucity of knowledge of mitochondrial genomes for many parasitic nematodes of socio-economic importance , such as members of the Ascaridida, which is a key group of nematodes that parasitizes humans and a range of other vertebrates. Although complete mitochondrial genome sequences are available for Anisakis simplex  and Ascaris suum , this is not the case for other parasites within this order, such as Toxocara canis, the common roundworm of dogs. This latter nematode parasitizes (at the adult stage) the small intestine of canids (definitive host) and is also transmissible to a range of other mammals, including rodents and humans, in which (after the oral ingestion of infective eggs) the larvae of Toxocara canis invade the tissues and cause covert toxocariasis, ocular larva migrans (OLM), visceral larva migrans (VLM) or neurotoxocariasis –. Although there has been a significant acquisition of knowledge about the biology of T. canis, there are still major gaps in our knowledge of the genetics, ecology and epidemiology of this enigmatic parasite . In addition, the detection of a cryptic species of “T. canis” from cats in Malaysia ,, its subsequent genetic characterization  and its description as a new species - Toxocara malayensis , emphasize the need for detailed molecular genetic studies of T. canis populations using suitable genetic markers . Exploring the mitochondrial genome of T. canis would provide such markers, as a foundation for molecular epidemiological and ecological studies, detecting cryptic species and assessing relationships of related species of Toxocara . Furthermore, the sequencing of the mitochondrial genome of T. canis provides a useful, comparative dataset to those of Anisakis simplex  and Ascaris suum  within the order Ascaridida. Building on recent progress in long PCR-coupled, automated sequencing , the present study determined the sequence and structure of the mitochondrial genome for a representative individual of T. canis from Australia and compared it with those available for Anisakis simplex and Ascaris suum as well as the sequences from related nematode groups (Spirurida and Strongylida), as a foundation for systematic, population genetic and epidemiological studies of T. canis.
Materials and Methods
An adult, male specimen of Toxocara canis (sample code Tcn2; ref. ) was collected (during a routine autopsy) from the small intestine of a fox from Victoria, Australia under the Scientific Procedures Premises License for the Faculty of Veterinary Science, University of Melbourne (SPPL045). Initially, the morphological identification of the worm was based on the presence of a post-oesophageal bulbus, the length and shape of the alae and the lengths of the spicules . Total genomic DNA was extracted from a small portion (0.5 cm) of the specimen by sodium dodecyl-sulphate/proteinase K treatment, phenol/chloroform extraction and ethanol precipitation , and purified over a spin column (Wizard Clean-Up; Promega) . In order to independently verify the identity of the specimen, the second internal transcribed spacer (ITS-2) of nuclear ribosomal DNA was amplified by the polymerase chain reaction (PCR) and sequenced according to an established method . The ITS-2 sequence obtained was a perfect match with that of T. canis (accession number Y09489; ref. ).
Using each of the primer pairs MH39F-MH42R and MH5F-MH40R ,,, two regions of the entire mitochondrial genome (of ~5 and 10 kb, respectively) were amplified by the long PCR (Expand 20 kbPLUS kit, Roche) from ~20 ng of genomic DNA from sample Tcn2. The cycling conditions in a 2400 thermocycler (Perkin Elmer Cetus) were: 92°C, 2 min (initial denaturation); then 92°C, 10 s (denaturation); 50°C, 30 s (annealing); 60°C (~5 kb region) or 68°C (~10 kb region), 10 min (extension) for 10 cycles, followed by 92°C, 10 s; 50°C, 30 s; 68°C or 60°C, 10 min for 20 cycles, with an elongation of 10 s for each cycle, and a final extension at 68°C or 60°C for 7 min . Each PCR yielded a single amplicon, detected by agarose gel electrophoresis . Each amplicon was column-purified (PCR-Preps, Promega) and subjected to automated sequencing, either directly or following cloning (TOPO XL PCR cloning kit, Invitrogen, according to instructions provided), employing a “primer-walking” strategy  (see Figure 1). Sequencing was performed using BigDye terminator (v.3.1) in a 3730 DNA Analyser (Applied Biosystems). The sequences obtained were assembled manually, aligned with the mitochondrial genome sequence of Ascaris suum  using the program Clustal X , and the circular map was drawn using the program MacVector v.9.5 (http://www.macvector.com/index.html). Amino acid sequences, translation initiation and termination codons, codon usage, transfer RNA (tRNA or trn) secondary structures, rRNA secondary structures and non-coding regions were predicted using standard approaches . The structure and organization of the mitochondrial genome of T. canis was then compared with those of the nematodes Anisakis simplex (GenBank accession number AY994157; ref. ), Ascaris suum (X53453; ref. ) (order Ascaridida); Brugia malayi (AF538716; ref. ), Dirofilaria immitis (AJ537512; ref. ) and Onchocerca volvulus (AF015193; ref. ) (order Spirurida); Ancylostoma duodenale (AJ417718; ref. ) and Necator americanus (AJ417719; ref. ) (order Strongylida).
Figure 1. A map of the circular mitochondrial genome (mtDNA) of Toxocara canis.
All 12 protein-coding genes and the large and small ribosomal subunits of the rRNA genes are indicated in italics. Each tRNA gene is identified by its anticodon (in brackets). The direction of transcription is indicated by an arrow. The positions of oligonucleotide primers (see table) used for PCR-amplification or sequencing are indicated in the map (drawn to scale).doi:10.1371/journal.pntd.0000273.g001
Features and Organization of the Mitochondrial Genome of Toxocara canis
The circular mitochondrial genome of T. canis (Figure 1) was 14162 bp in length (GenBank accession number EU730761) and contained 36 genes: 12 protein-coding genes (adenosine triphosphatase subunit 6 [atp6], the cytochrome c oxidase subunits 1, 2 and 3 [cox1–cox3], cytochrome b (cytb) and the nicotinamide dehydrogenase subunits 1–6 [nad1–nad6 and nad4l]), 22 tRNA genes (two coding for leucine and two coding for serine) and the small [rrnS] and large [rrnL] subunits of rRNA. Each protein-coding gene had an open reading frame (ORF), and all genes were located on the same strand and transcribed in the same direction (5′ to 3′) (Figure 1), consistent with the mitochondrial genomes of other secernentean nematodes characterized to date . The gene arrangement for the mitochondrial genome of T. canis was consistent with that of GA2 . This gene arrangement has been reported previously for members of the Ascaridida, including Anisakis simplex  and Ascaris suum  as well as members of the order Strongylida, such as the hookworms Ancylostoma duodenale and Necator americanus  as well as the barber's pole worm, Haemonchus contortus . However, consistent with the mitochondrial genomes characterized to date for other Ascaridida (but not the Strongylida), the AT-rich region for T. canis was located between rrnS and nad1, flanked (5′) by the genes trnS (UCN) and (3′) by trnN and trnY.
Nucleotide Contents and Codon Usage
The coding strand of the mitochondrial genome sequence of T. canis consisted of 21.6% A, 9.4% C, 22.1% G and 46.7% T (Table 1). Though AT-rich (68.4% AT), the sequence had a slightly lower AT content than has been reported for other nematode species (~70–80%; refs. ,,,,,). In the protein-coding genes, the AT-contents varied from 63.1% (cox1) to 73.4% (nad6), with the overall ranking (increasing richness) of cox1, cox3, nad1, cox2, cytb, nad4, atp6, nad5, nad2, nad4L, nad3 followed by nad6. To date, studies of secernentean nematodes have shown that the cytochrome c oxidase genes tend to have the lowest AT-contents ,,,. Although the overall AT-content of the mitochondrial genome sequence of T. canis was ~2.8% and ~3.6% less than those of Anisakis simplex (71.2%; ref. ) and Ascaris suum (72.0%; ref. ), respectively, there was no appreciable impact on the relative amino acid codon usage in the protein-coding genes. As has been reported for other secernentean nematodes (e.g., refs. ,,,,,), the usage in the protein-coding genes favoured codons with many A or T residues (e.g., 13.7% were TTT [phenylalanine]) over those with many C or G residues (e.g., none were CGA [arginine]) (data not shown).
Table 1. Lengths and A+T contents (%) of the sequences of the 12 protein-coding genes, the large and small ribosomal RNA genes, the AT-rich region and of the entire mitochondrial genome of Toxocara canis.doi:10.1371/journal.pntd.0000273.t001
All but the two serine tRNAs (AGN and UCN) had a predicted secondary structure containing a DHU arm and loop and a TV-replacement loop instead of the TψC arm and loop (Figure 2). As reported previously for secernentean nematodes ,,,, the two serine tRNAs each contained the TψC arm and loop but lacked the DHU arm and loop. The rrnL and rrnS genes were 923 and 693 bp in length, respectively; the predicted secondary structure of each of these two genes are displayed in Figure 3 (rrnL) and Figure 4 (rrnS). The AT-content of the sequences of rrnL, rrnS and the AT-rich (“control”) region were 77.9%, 66.5% and 78.1%, respectively. The relatively low AT-richness exhibited in the mitochondrial genome of T. canis was pronounced for the rRNA genes: The AT-content of the rrnL sequence was 4.2% and 4.9% less compared with Anisakis simplex (76.1%)  and Ascaris suum (76.8%) , respectively. The AT-content of the rrnS sequence of T. canis was 5.5% and 5.4% less than that reported for Anisakis simplex (72.0%)  and Ascaris suum (71.9%) , respectively. Interestingly, the AT-content of the T. canis mitochondrial rRNA genes does not alter their predicted secondary structures with respect to those of other secernentean nematodes studied to date ,,,.
Figure 3. The secondary structure predicted for the large subunit (rrnL) of the rRNA gene in the mitochondrial genome of Toxocara canis.
Bonds between C:G and U:A are indicated by a straight line and those between U:G by a closed circle (cf. ). Binding sites for the amino-acyl trn (A), peptidyl-transferase (P) or both (AP)  are indicated by lines.doi:10.1371/journal.pntd.0000273.g003
Figure 4. The secondary structure predicted for the small subunit (rrnS) of the rRNA gene in the mitochondrial genome of Toxocara canis.doi:10.1371/journal.pntd.0000273.g004
The AT-rich region (Figure 1) was 828 bp in length and was predicted to exhibit a complex secondary structure (Figure 5). In addition, the AT-rich region contained 13 regions consisting of a varying numbers of the dinucleotide (AT) repeat (n = 3 to 21) between nucleotide positions 307 and 806 (see Figure 5). The presence of multiple AT repeats was similar to that described in the AT-rich region of several parasitic nematodes, including Ascaris suum , Anisakis simplex , Ancylostoma duodenale and Necator americanus , but distinct from the repetitive elements (CR1–CR6) within the AT-rich region of the free-living nematode Caenorhabditis elegans .
Figure 5. Secondary structure predicted for the AT-rich region in the mitochondrial genome of Toxocara canis.doi:10.1371/journal.pntd.0000273.g005
Comparative Analysis with Other Nematodes
Pairwise comparisons were made among the amino acid sequences inferred from individual protein-coding genes and the nucleotide sequences of the rRNA genes in the T. canis mitochondrial genome with those representing seven other nematodes (of the orders Ascaridida, Spirurida and Strongylida) (Table 2). The amino acid sequence similarities in individual inferred proteins ranged from 70.3% (NAD2) to 94.4% (COX1) between T. canis and Ascaris suum and from 74.3% (CYTB) to 93.1% (COX1) between T. canis and Anisakis simplex. The amino acid sequence similarities between T. canis and each species of Spirurida (Brugia malayi, Dirofilaria immitis and Onchocerca volvulus) or Strongylida (Ancylostoma duodenale and Necator americanus) included ranged from 21.5% (ATP6) to 51.6% (COX1) and from 49.3% (NAD6) to 90.6% (COX1), respectively. The nucleotide sequence similarities (Table 2) in rrnS were 80.5–81.1%between T. canis and the two other species of Ascaridida, 59.0–60.0% between T. canis and the three members of the order Spirurida, and 71.8–72.4% between T. canis and the two species of Strongylida. In addition, the nucleotide sequence similarities in rrnL were 74.8–78.0%, 59.2–61.0% or 65.2–67.0% between T. canis and individual species representing the order Ascaridida, Spirurida or Strongylida, respectively (Table 2).
Table 2. Percentage of similarity in the amino acid sequences inferred from the 12 protein-coding genes and in the nucleotide sequence of each of the two ribosomal genes (rrnL and rrnS) upon pairwise comparison between Toxocara canis and seven other parasitic nematodes (representing the orders Ascaridida, Spirurida and Strongylida).doi:10.1371/journal.pntd.0000273.t002
Implications for Testing the Phylogeny of Parasitic Nematodes
Pairwise comparisons of the amino acid sequences conceptually translated from the protein-coding genes as well as the nucleotide sequences of the ribosomal RNA genes indicated that the mitochondrial genome of T. canis most closely resembles those of selected members of the order Ascaridida. However, based on pairwise comparisons of sequence data, the next most similar nematode group is the Strongylida, but not the Spirurida. This finding is consistent with previous phylogenetic analyses of mitochondrial datasets, such as concatenated amino acid sequences for all 12 protein-coding genes  or gene arrangements . These studies placed the Spirurida in a strongly supported clade separate from all other secernentean nematodes for which data were available, being in accordance with the evolutionary relationships based on traditional, taxonomic data , –. This placement of the Spirurida relative to the Ascaridida and Strongylida contrasts the classification of the Nematoda based on phylogenetic analysis of sequence data for the small subunit (SSU) of nuclear rRNA , inferring that members of the order Strongylida belong to “clade V”, whereas those of the orders Ascaridida and Spirurida are within “clade III”. The distinct taxonomic placement of the Spirurida relative to the Ascaridida and Strongylida is further evidenced by the variation in anti-codon usage in some of the mitochondrial tRNA genes of members of the Spirurida (i.e. Brugia malayi, Dirofilaria immitis and Onchocerca volvulus) as compared with the Strongylida and Ascaridida studied to date ,. The incongruence in the inferred relationships of these three nematode orders (i.e. among the Ascaridida, Strongylida and Spirurida) challenges the proposed molecular phylogeny for the Nematoda based on SSU sequence data  and stimulates further investigation of a broader range of nematodes.
Implications for Systematic, Population Genetic, Epidemiological and Ecological Studies
There is major significance in the use of mitochondrial DNA markers for investigating the genetic make-up of species of the Toxocara, particularly given that there are no morphological features which allow the specific identification of some stages (e.g., larvae)  and given that cryptic species have been detected within the Ascaridoidea , –. In nematodes, mitochondrial DNA is proposed to be maternally inherited (cf. ) and is usually more variable in sequence within a species than nuclear ribosomal DNA . Various different mitochondrial gene regions are suited to studying the population genetics of parasitic nematodes , , , –. However, surprisingly, there has been a paucity of information on the mitochondrial genomes of ascaridoid nematodes , which appeared to have related mainly to technical limitations and the cost associated with mitochondrial genome sequencing. To overcome this constraint, Hu et al. , developed the long PCR approach applied herein to T. canis, which has broad applicability to a range of ascaridoids, including other species of Toxocara, Toxascaris, Baylisascaris, Lagochilascaris and members of the Anisakis complex ,.
The characterization of the first complete mitochondrial genome sequence for T. canis, in the present study, provides a foundation for addressing ecological and epidemiological questions regarding this and related species. Conserved primers can be rationally and selectively designed to relatively conserved regions flanking “variable tracts” in the mitochondrial genome considered to be most informative (based on sequencing from a small number of individuals from particular populations, or genetic variants detected using nuclear ribosomal markers; refs. ,). Using such primers, single-strand conformation polymorphism (SSCP) analysis  can be applied to pre-screen large numbers of individuals representing different populations and, based on the ‘pre-screen’, samples representing the entire spectrum of haplotypic variability can be selected for subsequent sequencing and analyses. Such an approach has been applied effectively, for example, to explore the genetic make-up of the Ascaris populations in humans and pigs in six provinces in China . This study indicated restricted gene flow between human Ascaris and porcine Ascaris, and supported the conclusions from other previous epidemiological and experimental investigations , that pigs are not a significant source of Ascaris infection to humans in endemic regions.
Utilizing a range of mitochondrial gene markers (with differing degrees of intraspecific variability), such a mutation scanning-targeted approach is also readily and directly applicable to species of Toxocara. This is particularly relevant now, given that population variation and cryptic species have been detected within Toxocara ,, and that almost nothing is known about the transmissibility of these or other, as yet undetected, variants and/or cryptic species to humans and other hosts. For instance, early reports from Malaysia , described the occurrence of a parasite in cats, which was identified as T. canis, based on the presence of an oesophageal ventriculus and spear-shaped cervical alae in the adult . This parasite differed from the common species known to parasitize cats –, such as Toxocara cati, which has arrow-shaped cervical alae, and Toxascaris leonina, which lacks a ventriculus. Because T. canis has been found only rarely in cats elsewhere in the world –, the question arose as to the specific identity of this parasite in Malaysian cats. A molecular study, using markers in the first and second internal transcribed spacers (ITS-1 and ITS-2, respectively) of nuclear ribosomal DNA markers, was undertaken to genetically characterize specimens of this parasite, then called Toxocara sp. cf. canis . The molecular investigation indicated clearly that Toxocara sp. cf. canis from Malaysian cats was genetically distinct from T. canis and T. cati, a conclusion which was supported by a subsequent morphological study of a number of ascaridoids from Malaysia . Three morphological features (for lips, alae and spicules) were identified which consistently differentiated Toxocara sp. cf. canis from T. canis, T. cati and other congeners, such as T. tanuki (from canids), T. apodemi and T. mackerrasae (from rodents), T. paradoxura and T. sprenti (from viverrids), T. vajrasthirae (from mustelids) and T. pteropodis (from bats). Hence, the findings from the molecular and classical systematic studies supported the conclusion that Toxocara sp. cf. canis represented a distinct species, subsequently named T. malaysiensis .
Although T. canis is well recognized as the causative agent of toxocariasis in humans, including ocular larva migrans (OLM) and/or visceral larva migrans (VLM), other congeners, such as T. malaysiensis, T. cati and T. vitulorum, may have greater zoonotic importance than assumed ,. T. malaysiensis is of particular interest as a potential zoonotic pathogen, given its high prevalence (11%) in cats . The transmissibility of this species to other host species (e.g., mouse, rat, rabbit and pig) warrants assessment, together with epidemiological surveys utilizing molecular tools employing genetic markers from the mitochondrial genome of T. canis as well as specific nuclear markers in the ITS-1 and/or ITS-2. The discovery of T. malaysiensis in cats in Malaysia  also raises important questions as to the identity and zoonotic potential of ascaridoids considered to represent T. canis in cats in other geographical regions, including South Africa, Panama, the USA and Czech Republic –,, which provides a stimulus for the genetic characterization of additional Toxocara isolates from a broad range of hosts and geographical origins and to subsequently evaluate their potential to infect humans and/or other hosts.
From epidemiological and ecological perspectives, it would be interesting, utilizing mitochondrial genomic data, to confirm or refute the involvement of Toxocara in human VLM cases in Japan, currently considered to be caused by Ascaris suum based on serological evidence ,, as there has been considerable controversy as to the specific identity of the causative agent of the disease in these instances . It would also be particularly relevant to explore whether specific genotypes/haplotypes of Toxocara canis have a particular affinity to the human host and/or predilection sites in tissues to cause different types of toxocariasis and whether there are specific subpopulations of T. canis that undergo arrested development in tissues. Using molecular tools, in combination with traditional parasitological and serological methods, it should also be possible to characterize in detail experimental infections in “model host systems” (e.g., mouse, rabbit or pig) , –. Furthermore, mitochondrial markers would be useful for exploring the zoonotic risk of paratenic hosts, particularly those commonly encountered in an agricultural setting (e.g. chickens, ducks or pigs –), and determining the specific identity of eggs in the environment .
In conclusion, the present study emphasizes the relevance of the mitochondrial genome of T. canis defined herein, should provide a foundation for a range of systematic, population genetic, epidemiological, ecological and biological studies. Although the PCR-based sequencing-cloning approach used herein was effective, the PCR-coupled 454 technology platform , constructed recently for the direct sequencing of mitochondrial genomes from single nematodes , provides perhaps the most exciting development for large-scale, high throughput population genetic and mitochondrial genomic studies of nematodes and other organisms.
Conceived and designed the experiments: ARJ DTJL RBG. Performed the experiments: AW. Analyzed the data: ARJ MH RBG. Contributed reagents/materials/analysis tools: AW DTJL RBG. Wrote the paper: ARJ DTJL RBG.
- 1. Hugot JP, Baujard P, Morand S (2001) Biodiversity in helminths and nematodes as a field of study: an overview. Nematology 3: 199–208.
- 2. Anderson RC (2000) Nematode Parasites of Vertebrates. Their Development and Transmission. Wallingford: CAB International.
- 3. Albonico M, Crompton DW, Savioli L (1999) Control strategies for human intestinal nematode infections. Adv Parasitol 42: 277–341.
- 4. Eisenback JD, Triantaphyllou HH (1991) Root-knot nematodes: Meloidogyne species and races. In: Nickel WR, editor. Manual of Agricultural Nematology. New York: Marcell Dekker. pp. 191–274.
- 5. McCarthy J, Moore TA (2000) Emerging helminth zoonoses. Int J Parasitol 30: 1351–1360.
- 6. Anderson TC, Blouin MS, Beech RN (1998) Population biology of parasitic nematodes: applications of genetic markers. Adv Parasitol 41: 219–283.
- 7. Blouin MS (1998) Mitochondrial DNA diversity in nematodes. J Helminthol 72: 285–289.
- 8. Viney ME (1998) Nematode population genetics. J Helminthol 72: 281–283.
- 9. Gasser RB (2006) Molecular tools - advances, opportunities and prospects. Vet Parasitol 136: 69–89.
- 10. Blouin MS (2002) Molecular prospecting for cryptic species of nematodes: mitochondrial DNA versus internal transcribed spacer. Int J Parasitol 32: 527–531.
- 11. Avise JC (1991) Ten unothrodox perspectives on evolution prompted by comparative population genetic findings on mitochondrial DNA. Ann Rev Genet 25: 45–69.
- 12. Avise JC (1994) Molecular Markers, Natural History and Evolution. New York and London: Chapman and Hall.
- 13. Avise JC (1998) The history and purview of phylogeography: a personal reflection. Mol Ecol 7: 371–379.
- 14. Avise JC, Arnold J, Ball RM, Berminham E, Lamb T, et al. (1987) Intraspecific phylogeography: the mitochondrial DNA bridge between population genetics and systematics. Ann Rev Ecol Syst 18: 489–522.
- 15. Hu M, Chilton NB, Gasser RB (2004) The mitochondrial genomics of parasitic nematodes of socio–economic importance: recent progress, and implications for population genetics and systematics. Adv Parasitol 56: 133–212.
- 16. Hu M, Gasser RB (2006) Mitochondrial genomes of parasitic nematodes - progress and perspectives. Trends Parasitol 22: 78–84.
- 17. Kim KH, Eom KS, Park JK (2006) The complete mitochondrial genome of Anisakis simplex (Ascaridida: Nematoda) and phylogenetic implications. Int J Parasitol 36: 319–328.
- 18. Okimoto R, Macfarlane JL, Clary DO, Wolstenholme DR (1992) The mitochondrial genomes of two nematodes, Caenorhabditis elegans and Ascaris suum. Genetics 130: 471–498.
- 19. Hotez PJ (1993) Visceral and ocular larva migrans. Semin Neurol 13: 175–179.
- 20. Holland CV, Hamilton C (2005) The significance of Cerebral Toxocariasis. In: Holland CV, Smith HV, editors. Toxocara: the enigmatic parasite. Wallingford, UK: CABI International. pp. 58–73.
- 21. Taylor MRH (2005) Ocular Toxocariasis. In: Holland C, Smith HV, editors. Toxocara: the enigmatic parasite. Wallingford, UK: CABI International. pp. 127–144.
- 22. Despommier D (2003) Toxocariasis: clinical aspects, epidemiology, medical ecology, and molecular aspects. Clin Microbiol Rev 16: 265–272.
- 23. Overgaauw PA (1997) Aspects of Toxocara epidemiology: human toxocarosis. Crit Rev Microbiol 23: 215–231.
- 24. Holland CV, Smith HV (2005) Toxocara: the enigmatic parasite. Wallingford: CABI Press.
- 25. Rohde K (1962) Helminths of Cats and Dogs in Malaya. University of Singapore.
- 26. Lee CC, Cheng NABY, Bohari Y (1993) Toxocara canis from domestic cats in Kuala Lumpur. Tropical Biomed 10: 79–80.
- 27. Zhu XQ, Jacobs DE, Chilton NB, Sani RA, Cheng NABY, et al. (1998) Molecular characterization of a Toxocara variant from cats in Kuala Lumpur, Malaysia. Parasitology 117: 155–164.
- 28. Gibbons LM, Jacobs DE, Sani RA (2001) Toxocara malaysiensis n. sp. (Nematoda: Ascaridoidea) from the domestic cat (Felis catus L.). J Parasitol 87: 660–665.
- 29. Gasser RB, Zhu XQ, Hu M, Jacobs DE, Chilton NB (2005) Molecular genetic characterisation of members of the genus Toxocara (Nematoda: Ascaridoidea) - systematic, population genetic and epidemiological considerations. In: Holland CV, Smith HV, editors. Toxocara: The Enigmatic Parasite. Wallingford: CABI Press. pp. 18–31.
- 30. Hu M, Jex AR, Campbell BE, Gasser RB (2007) Long PCR amplification of the entire mitochondrial genome from individual helminths for direct sequencing. Nature Protoc 2: 2339–2344.
- 31. Zhu XQ, Gasser RB (1998) Single-strand conformation polymorphism (SSCP)-based mutation scanning approaches to fingerprint sequence variation in ribosomal DNA of ascaridoid nematodes. Electrophoresis 19: 1366–1373.
- 32. Skrjabin KI, Shikhobalova NP, Mozgovoi AA (1991) Key to Parasitic Nematoda: Volume 2 Oxyurata and Ascaridata;. In: Skrjabin KI, editor. Leiden: E. J. Brill.
- 33. Gasser RB, Chilton NB, Hoste H, Beveridge I (1993) Rapid sequencing of rDNA from single worms and eggs of parasitic helminths. Nucleic Acids Res 21: 2525–2526.
- 34. Jacobs DE, Zhu XQ, Gasser RB, Chilton NB (1997) PCR-based methods for identification of potentially zoonotic ascaridoid parasites of the dog, fox and cat. Acta Trop 68: 191–200.
- 35. Hu M, Chilton NB, Gasser RB (2002) The mitochondrial genomes of the human hookworms, Ancylostoma duodenale and Necator americanus (Nematoda: Secernentea). Int J Parasitol 32: 145–158.
- 36. Hu M, Chilton NB, Gasser RB (2002) Long PCR-based amplification of the entire mitochondrial genome from single parasitic nematodes. Mol Cell Probes 16: 261–267.
- 37. Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG (1997) The Clustal X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 24: 4876–4882.
- 38. Ghedin E, Wang S, Spiro D, Caler E, Zhao Q, et al. (2007) Draft genome of the filarial nematode parasite Brugia malayi. Science 317: 1756–1760.
- 39. Hu M, Gasser RB, Abs EL-Osta YG, Chilton NB (2003) Structure and organization of the mitochondrial genome of the canine heartworm, Dirofilaria immitis. Parasitology 127: 37–51.
- 40. Keddie EM, Higazi T, Unnasch TR (1998) The mitochondrial genome of Onchocerca volvulus: sequence, structure and phylogenetic analysis. Mol Biochem Parasitol 95: 111–127.
- 41. Hu M, Chilton NB, Gasser RB (2003) The mitochondrial genome of Strongyloides stercoralis (Nematoda) - idiosyncratic gene order and evolutionary implications. Int J Parasitol 33: 1393–1408.
- 42. Jex AR, Hu M, Littlewood TJ, Waeschenbach A, Gasser RB (2008) Using 454 technology for long-PCR based sequencing of the complete mitochondrial genome from single Haemonchus contortus (Nematoda). BMC Genomics [Epub ahead of print].
- 43. Wolstenholme DR, Macfarlane JL, Okimoto R, Clary DO, Wahleithner JA (1987) Bizarre tRNAs inferred from DNA sequences of mitochondrial genomes of nematode worms. Proc Natl Acad Sci U S A 84: 1324–1328.
- 44. Okimoto R, Wolstenholme DR (1990) A set of tRNAs that lack either the T psi C arm or the dihydrouridine arm: towards a minimal tRNA adaptor. EMBO J 9: 3405–3411.
- 45. Skrjabin KI (1992) Key to Parasitic Nematoda: Volume 4 Camallanata, Rhabditata, Tylenchata, Trichocephalata, Dioctophymata and distribution of parasitic nematodes in different hosts;. In: Skrjabin KI, editor. Leiden: E. J. Brill.
- 46. Skrjabin KI (1991) Key to Parasitic Nematodes: Volume 1 Spirurata and Filariata;. In: Raveh M, editor. Leiden: E. J. Brill.
- 47. Skrjabin KI, Shikhobalova NP, Schulz RS, Popova TI, Boev SN, et al. (1992) Key to Parasitic Nematoda: Volume 3 Strongylata;. In: Skrjabin KI, editor. Leiden: E. J. Brill.
- 48. Blaxter ML, De Ley P, Garey JR, Liu LX, Scheldeman P, et al. (1998) A molecular evolutionary framework for the phylum Nematoda. Nature 392: 71–75.
- 49. Biocca E, Nascetti G, Iori A, Costantini R, Bullini L (1978) Descrizione di Parascaris univalens, parassita degli equini, e suo differenziamento da Parascaris equorum. Accad Naz Lincei Rend Lincei Sci Fis, Mat Nat, Serie VIII 65: 133–141.
- 50. Bullini L, Nascetti G, Ciafre S, Rumore F, Biocca E (1978) Ricerche cariologiche ed elettroforetiche su Parascaris univalens e Parascaris equorum. Accad Naz Lincei Rend Lincei Sci Fis, Mat Nat, Serie VIII 65: 151–156.
- 51. Paggi L, Nascetti G, Orecchia P, Mattiucci S, Bullini L (1985) Biochemical taxonomy of ascaridoid nematodes. Parassitologia 27: 105–112.
- 52. Nascetti G, Paggi L, Orecchia P, Smith JM, Mattiucci S, et al. (1986) Electorphoretic studies on the Anisakis simplex complex (Ascaridida: Anisakidae) from the Mediterranean and north-east Atlantic. Int J Parasitol 16: 633–640.
- 53. Orecchia P, Paggi L, Mattiucci S, Smith JW, Nascetti G, et al. (1986) Electrophoretic identification of larvae and adults of Anisakis (Ascaridida: Anisakidae). J Helminthol 60: 331–339.
- 54. Paggi L, Nascetti G, Cianchi R, Orecchia P, Mattiucci S, et al. (1991) Genetic evidence for three species within Pseudoterranova decipiens (Nematoda, Ascaridida, Ascaridoidea) in the North Atlantic and Norwegian and Barents Seas. Int J Parasitol 21: 195–212.
- 55. Nascetti G, Cianchi R, Mattiucci S, D'Amelio S, Orecchia P, et al. (1993) Three sibling species with Contracaecum osculatum (Nematoda, Ascaridida, Ascaridoidea) from Atlantic Arctic-Boreal region: reproductive isolation and host preferences. Int J Parasitol 23: 105–120.
- 56. Orecchia P, Mattiucci S, D'Amelio S, Paggi L, Plötz J, et al. (1994) Two new members in the Contracaecum osculatum complex (Nematoda, Ascaridoidea) from Antarctic. Int J Parasitol 24: 367–377.
- 57. Mattiucci S, Nascetti G, Cianchi R, Paggi L, Arduino P, et al. (1997) Genetic and ecological data on the Anisakis simplex complex, with evidence for a new species (Nematoda, Ascaridoidea, Anisakidae). J Parasitol 83: 401–416.
- 58. Zhu XQ, Podolska M, Chilton NB, Gasser RB (1998) Characterisation of anisakid nematodes with zoonotic potential by nuclear ribosomal DNA sequences. Int J Parasitol 28: 1911–1921.
- 59. Zhu XQ, Chilton NB, Jacobs DE, Boes J, Gasser RB (1999) Characterisation of Ascaris from human and pig hosts by nuclear ribosomal DNA sequences. Int J Parasitol 29: 469–478.
- 60. Zhu XQ, Spratt DM, Beveridge I, Haycock P, Gasser RB (2000) Mitochondrial DNA polymorphism within and among species of Capillaria sensu lato from Australian marsupials and rodents. Int J Parasitol 30: 933–938.
- 61. Zhu XQ, Gasser RB, Chilton NB, Jacobs DE (2001) Molecular approaches for studying ascaridoid nematodes with zoonotic potential, with an emphasis on Toxocara species. J Helminthol 75: 101–108.
- 62. Zhu XQ, D'Amelio S, Palm HW, Paggi L, George-Nascimento M, et al. (2002) SSCP-based identification of members within the Pseudoterranova decipiens complex (Nematoda: Ascaridoidea: Anisakidae) using genetic markers in the internal transcribed spacers of ribosomal DNA. Parasitology 124: 615–623.
- 63. Anderson TJ, Romero-Abal ME, Jaenike J (1993) Genetic structure and epidemiology of Ascaris populations: patterns of host affiliation in Guatemala. Parasitology 107 (Pt 3): 319–334.
- 64. Anderson TJC, Blouin MS, Beech RN (1998) Populations biology of parasitic nematodes: applications of genetic markers. Adv Parasitol 41: 219–283.
- 65. Blouin MS, Yowell CA, Courtney CH, Dame JB (1995) Host movement and the genetic structure of populations of parasitic nematodes. Genetics 141: 1007–1014.
- 66. Blouin MS, Yowell CA, Courtney CH, Dame JB (1997) Haemonchus placei and Haemonchus contortus are distinct species based on mtDNA evidence. Int J Parasitol 27: 1383–1387.
- 67. Hu M, Chilton NB, Zhu XQ, Gasser RB (2002) Single-strand conformation polymorphism-based analysis of mitochondrial cytochrome c oxidase subunit 1 reveals significant substructuring in hookworm populations. Electrophoresis 23: 27–34.
- 68. Bowman DD (1987) Diagnostic morphology of four larval ascaridoid nematodes that may cause visceral larva migrans: Toxascaris leonina, Baylisascaris procyonis, Lagochilascaris sprenti, and Hexametra leidyi. J Parasitol 73: 1198–1215.
- 69. Gasser RB, Hu M, Chilton NB, Campbell BE, Jex AR, et al. (2006) Single-strand conformation polymorphism (SSCP) for the analysis of genetic variation. Nature Protoc 1: 3121–3128.
- 70. Peng W, Yuan K, Hu M, Zhou X, Gasser RB (2005) Mutation scanning-coupled analysis of haplotypic variability in mitochondrial DNA regions reveals low gene flow between human and porcine Ascaris in endemic regions of China. Electrophoresis 26: 4317–4326.
- 71. Anderson TJ (2001) The dangers of using single locus markers in parasite epidemiology: Ascaris as a case study. Trends Parasitol 17: 183–188.
- 72. Peng W, Yuan K, Hu M, Gasser RB (2007) Recent insights into the epidemiology and genetics of Ascaris in China using molecular tools. Parasitology 134: 325–330.
- 73. Sprent JF (1958) Observations on the development of Toxocara canis (Werner, 1782) in the dog. Parasitology 48: 184–209.
- 74. Sprent JF (1956) The life history and development of Toxocara cati (Schrank 1788) in the domestic cat. Parasitology 46: 54–78.
- 75. Sprent JF (1959) The life history and development of Toxascaris leonina (von Linstow, 1902) in the dog and cat. Parasitology 49: 330–371.
- 76. Sprent JF, Barrett MG (1964) Large roundworms of dogs and cats: differentiation of Toxocara canis and Toxascaris leonina. Austral Vet J 40: 166–171.
- 77. Calero MC, Ortiz OP, De Souza L (1951) Helminths in cats from Panama city and Balboa, C. Z. J Parasitol 37: 326.
- 78. Hitchcock DJ (1953) Incidence of gastro-intestinal parasites in some Michigan kittens. North Am Vet 34: 428–429.
- 79. Ash LR (1962) Helminth parasites of dogs and cats in Hawaii. J Parasitol 48: 63–65.
- 80. Parsons JC (1987) Ascarid infections of cats and dogs. Vet Clin North Am Small Anim Pract 17: 1307–1339.
- 81. Baker MK, Lange L, Verster A, van der Plaat S (1989) A survey of helminths in domestic cats in the Pretoria area of Transvaal, Republic of South Africa. Part 1: The prevalence and comparison of burdens of helminths in adult and juvenile cats. J S Afr Vet Assoc 60: 139–142.
- 82. Smith HV (1993) Antibody reactivity in human toxocariasis. In: Lewis JW, Maizels RM, editors. Toxocara and Toxocariasis: Clinical, Epidemiological and Molecular Perspectives. London: Institute of Biology. pp. 91–110.
- 83. Miyazaki I (1994) An Illustrated Book of Helminthic Zoonoses. Tokyo: International Medical Foundation of Japan.
- 84. Scholz T, Uhlirova M, Ditrich O (2003) Helminth parasites of cats from the Vientiane province, Laos, as indicators of the occurrence of causative agents of human parasitoses. Parasite 10: 343–350.
- 85. Maruyama H, Nawa Y, Noda S, Mimori T, Choi WY (1996) An outbreak of visceral larva migrans due to Ascaris suum in Kyushu, Japan. Lancet 347: 1766–1767.
- 86. Sakakibara A, Baba K, Niwa S, Yagi T, Wakayama H, et al. (2002) Visceral larva migrans due to Ascaris suum which presented with eosinophilic pneumonia and multiple intra-hepatic lesions with severe eosinophil infiltration - outbreak in a Japanese area other than Kyushu. Intern Med 41: 574–579.
- 87. Petithory JC (1996) Can Ascaris suum cause visceral larva migrans? Lancet 348: 689.
- 88. Holland CV, Cox DM (2001) Toxocara in the mouse: a model for parasite-altered host behaviour? J Helminthol 75: 125–135.
- 89. Garcia HH, Cancrini G, Bartalesi F, Rodriguez S, Jimenez JA, et al. (2007) Evaluation of immunodiagnostics for toxocarosis in experimental porcine cysticercosis. Trop Med Int Health 12: 107–110.
- 90. Sommerfelt IE, Santillan G, Mira G, Ribicich M, Betti A, et al. (2006) Toxocara canis infections in a pig model: immunological, haematological and blood biochemistry responses. J Helminthol 80: 73–77.
- 91. Fenoy S, Ollero MD, Guillen JL, del Aguila C (2001) Animal models in ocular toxocariasis. J Helminthol 75: 119–124.
- 92. Boes J, Helwigh AB (2000) Animal models of intestinal nematode infections of humans. Parasitology 121: S97–S111.
- 93. Helwigh AB, Lind P, Nansen P (1999) Visceral larva migrans: migratory pattern of Toxocara canis in pigs. Int J Parasitol 29: 559–565.
- 94. Kayes SG (1997) Human toxocariasis and the visceral larva migrans syndrome: correlative immunopathology. Chem Immunol 66: 99–124.
- 95. Taira K, Saeed I, Permin A, Kapel CM (2004) Zoonotic risk of Toxocara canis infection through consumption of pig or poultry viscera. Vet Parasitol 121: 115–124.
- 96. Morimatsu Y, Akao N, Akiyoshi H, Kawazu T, Okabe Y, et al. (2006) A familial case of visceral larva migrans after ingestion of raw chicken livers: appearance of specific antibody in bronchoalveolar lavage fluid of the patients. Am J Trop Med Hyg 75: 303–306.
- 97. Hoffmeister B, Glaeser S, Flick H, Pornschlegel S, Suttorp N, et al. (2007) Cerebral toxocariasis after consumption of raw duck liver. Am J Trop Med Hyg 76: 600–602.
- 98. Lee KT, Min HK, Chung PR, Chang JK (1976) [Studies On The Inducing Possibility Of Human Visceral Larva Migrans Associated With Eating Habit Of Raw Liver Of Domestic Animals] in Korean. Kisaengchunghak Chapchi 14: 51–60.
- 99. Mizgajska H (2001) Eggs of Toxocara spp. in the environment and their public health implications. J Helminthol 75: 147–151.
- 100. Margulies M, Egholm M, Altman WE, Attiya S, Bader JS, et al. (2005) Genome sequencing in microfabricated high-density picolitre reactors. Nature 437: 376–380.
- 101. Noller HF, Asire M, Barta A, Douthwaite S, Goldstein T, et al. (1986) Studies on the structure and function of ribosomal RNA. In: Hardesty B, Kramer G, editors. Structure, Function and Genetics of Ribosomes. New York: Springer-Verlag. pp. 143–163.
- 102. Dams E, Hendriks L, Van de Peer Y, Neefs J-M, Smits G, et al. (1988) Compilation of small ribosomal subunit RNA sequences. Nucleic Acids Res 16s: r87–r173.